Application of Consistent Massage-Like Perturbations on Mouse Calves and Monitoring the Resulting Intramuscular Pressure Changes

* These authors contributed equally
Immunology and Infection

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Summary

Here we describe the protocols for applying defined mechanical loads to mouse calves and for monitoring the concomitant intramuscular pressure changes. The experimental systems that we have developed can be useful for investigating the mechanism behind the beneficial effects of physical exercise and massage.

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Sakitani, N., Maekawa, T., Saitou, K., Suzuki, K., Murase, S., Tokunaga, M., Yoshino, D., Sawada, K., Takashima, A., Nagao, M., Ogata, T., Sawada, Y. Application of Consistent Massage-Like Perturbations on Mouse Calves and Monitoring the Resulting Intramuscular Pressure Changes. J. Vis. Exp. (151), e59475, doi:10.3791/59475 (2019).

Abstract

Massage is generally recognized to be beneficial for relieving pain and inflammation. Although previous studies have reported anti-inflammatory effects of massage on skeletal muscles, the molecular mechanisms behind are poorly understood. We have recently developed a simple device to apply local cyclical compression (LCC), which can generate intramuscular pressure waves with varying amplitudes. Using this device, we have demonstrated that LCC modulates inflammatory responses of macrophages in situ and alleviates immobilization-induced muscle atrophy. Here, we describe protocols for the optimization and application of LCC as a massage-like intervention against immobilization-induced inflammation and atrophy of skeletal muscles of mouse hindlimbs. The protocol that we have developed can be useful for investigating the mechanism underlying beneficial effects of physical exercise and massage. Our experimental system provides a prototype of the analytical approach to elucidate the mechanical regulation of muscle homeostasis, although further development needs to be made for more comprehensive studies.

Introduction

Massage is generally recognized to be beneficial for both pain relief and improvement of the physical performance among competitive athletes and non-athletes alike1,2. In fact, previous studies have shown that massage suppresses local inflammation3 and prompts recovery from the post-exercise muscle damage4,5. Molecular mechanisms underlying the beneficial effects of massage remain largely unknown.

One of the difficulties with the mechanistic investigation on massage relates to the reproducibility of experimental techniques by which massage-like interventions are tested. In previous studies, experimental procedures that mimic massage mostly involve the application of physical interventions using practitioners’ body parts, such as palms and fingers6,7,8. This makes it is difficult to precisely reproduce their magnitude, frequency, duration, and mode.

Many devices have been developed to apply defined mechanical loads to the target tissues. For example, Zeng et al. have developed a pneumatic system for the length-wise mechanical loading to rats’ hindlimbs9 and Wang et al. have developed a mechatronic device that can apply massage-like mechanical loads to hindlimbs of rats and rabbits with real-time feedback control10. Compared to them, our local cyclical compression (LCC) system is much simpler, demanding far less cost for construction. Nonetheless, we can reproduce the intramuscular pressure changes that are generated during the mild muscle contraction. Using this device, we have successfully demonstrated that the massage-like mechanical interventions modulate local interstitial fluid dynamics and alleviate immobilization-induced muscle atrophy11.

Here, we describe the details of our device and the protocol, which may help explore the molecular mechanisms behind the positive effects of exercises and massages. The schematics of the protocol is presented as Supplementary Figure 1.

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Protocol

All animal experiments were conducted under the approval by the Institutional Animal Care and Use Committee of the National Rehabilitation Center for Persons with Disabilities.

1. Immobilization of the mouse bilateral hindlimbs

NOTE: Male C57BL/6 mice were used for experiments at the age of 11 - 12 weeks after acclimation for at least 7 days.

  1. Adequately anesthetize a mouse using sodium pentobarbital (50 mg/kg i.p.). Make sure that mice do not respond to a hindlimb toe pinch.
    NOTE: Conduct the procedure of immobilization between 10 a.m. and 7 p.m. to minimize the possible effects on the feeding activity of mice.
  2. Apply surgical tapes to the bilateral hindlimbs of the mouse laid in a supine position with the knee joints extended and ankle joints plantar-flexed.
  3. Place an aluminum wire (see Table of Materials) on the trunk at L4-5 spine level and coil the wire in a spiral configuration around the hindlimbs with 5 mm gaps between each turn of the spiral layer (Figure 1A). Make sure not to coil the wire too tightly and avoid disturbing the local blood flow.
  4. To minimize the possibility of escape from wiring, immobilize the hip joints at the position of 90° abduction by manually adjusting the configuration of aluminum wire.
  5. Return the mice to their original cages. 3 h later, make sure that they recover from anesthesia and access to food and water as usual.
  6. House 3 - 6 immobilized mice per cage as before immobilization.

2. Measurement of intramuscular pressure of mouse gastrocnemius muscles

NOTE: Several different weights of cylindrical units (36 g, 66 g, and 200 g) were tested in the pressure-monitoring experiments combined with LCC. This measurement was conducted separately from the experiments to analyze muscle inflammation and atrophy (see step 3 - 5 for more details) i.e., the mice subjected to pressure measurement were not used for histological analyses.

  1. Because pressure measurement involves more invasive procedures (e.g., skin incision and needle insertion) as compared to hindlimb wiring and LCC, use a mixture of three anesthetic agents (medetomidine 0.75 mg/kg, midazolam 4.0 mg/kg, and butorphanol 5.0 mg/kg, i.p.). Make sure that mice do not respond to the hindlimb toe pinch.
  2. Lay the mouse in a prone position, make a 2-mm incision with a scalpel on the posterior calf after depilating with an electric shaver and semi-sterilizing the skin surface with 70% ethanol- and Chlorhexidine-soaked absorbent cotton.
  3. Insert a 20 G indwelling needle into the gastrocnemius muscle at an obtuse angle (150° – 170°) to the skin surface.
  4. Using the plastic sheath of the needle as a guide, place a sensor of the blood pressure telemeter (see Table of Materials) in the mid-belly of gastrocnemius muscle, and then remove the sheath from the muscle.
  5. After suturing the skin with 4-0 nylon suture, apply LCC with several different weights of cylindrical units to the calf in the mice (see step 3 for more details), and monitor the intramuscular pressure using software for biological signal analysis (see Table of Materials).
  6. Return the mice to their original cages. 3 h later, make sure that they recover from anesthesia/analgesia and have access to food and water as usual.

3. Local cyclical compression (LCC) on mouse calves

  1. Except for the intramuscular pressure measurement and euthanizing (i.e., cervical dislocation), use sodium pentobarbital (50 mg/kg i.p.) for anesthesia.
  2. Disengage the mouse from hindlimb wiring and lay it in a prone position with the knee joints extended and the ankle joints plantar-flexed so that the calves faced upward. Do not fix the mouse hindlimbs on the stage.
  3. Apply LCC to the calf by vertically moving a cylindrical weight unit (Figure 1B) covered with a cushion pad (Figure 1C) at 1 Hz for 30 min per day, 7 days.
  4. After each bout of daily LCC, re-wire the mouse hindlimbs.

4. Immunohistochemical analysis of gastrocnemius

  1. Euthanize the mouse by cervical dislocation under adequate anesthesia/analgesia by intraperitoneal injection of a mixture of three anesthetic agents (medetomidine 0.75 mg/kg, midazolam 4.0 mg/kg, and butorphanol 5.0 mg/kg).
  2. After depilating the posterior calf surface, make a skin incision, and dissect gastrocnemius muscles by separating from tibio-fibular bone using a surgical scissor and quickly freeze them in an optimal cutting temperature compound solution.
  3. Using a cryostat, prepare cryo-section samples of gastrocnemius muscles on glass slides. Store the samples in a -80 °C freezer until analysis.
  4. Take out the gastrocnemius cryo-section samples to be analyzed from the freezer and dehydrate them by air drying at room temperature.
  5. Use a liquid blocker pen to draw an area that includes all the cryo-sections on the slide. The circle will prevent the solutions from flowing off the slide.
  6. Avoid drying of the samples by placing the slides in a tray in which a moist environment is created with water-soaked paper cloth.
  7. Apply 100 µL of blocking buffer (phosphate-buffered saline (PBS) containing 0.25% casein, carrier protein, and 0.015 M sodium azide) for 30 min at room temperature.
  8. Rinse the slides twice by incubating with PBS-T (PBS containing 0.1% polyoxyethylene sorbitan monolaurate (see Table of Materials) for 5 min.
  9. Apply 100 µL of primary antibody diluted with PBS on each sample, cover the tray with a lid, and incubate overnight at room temperature.
  10. Wash 3 times with PBS-T (5 min for each wash).
  11. Apply 100 µL of secondary antibody diluted with PBS on each sample and incubate for 1 h at room temperature.
    NOTE: For anti-laminin staining, use Alexa Fluor 568-conjugated secondary antibody. For anti-F4/80, anti-MCP-1, and anti-TNF-α, use Alexa Fluor 568- or 488-conjugated secondary antibody.
  12. Wash 3 times with PBS-T (5 min for each wash).
  13. Apply 100 µL of DAPI solution diluted with PBS-T on each sample and incubate for 3 min at room temperature.
  14. Wash 3 times with PBS-T (5 min for each).
  15. Mount the samples with mounting medium and cover them with coverslips.

5. Histo-morphometric analysis of gastrocnemius

  1. Place the sample slides on a fluorescence microscope (see Table of Materials) and view the samples using a 20× objective with appropriate filters (DAPI-B, 360/40 nm for excitation and 460/50 nm for emission; GFP-B, 470/40 nm for excitation and 535/50 nm for emission; FRITC, 540/25 nm for excitation and 605/55 nm for emission.
  2. Using the software for image analysis (see Table of Materials), measure the cross-sectional area (CSA) of each myofiber, and count the number of F4/80-, MCP-1-, and TNF-α-positive cells.
    NOTE: Determine CSA of each myofiber by tracing the internal margin of the basement membrane visualized with anti-laminin-2 immunostaining.

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Representative Results

Consistent with our previous observations12, the CSA of gastrocnemius myofibers were significantly decreased by hindlimb immobilization (Figure 2A,B). Furthermore, our immunofluorescence staining analysis revealed that cells expressing MCP-1 and TNF-α, both of which play key roles in regulating inflammatory processes13,14, significantly increased in gastrocnemius muscle tissues of immobilized hindlimbs (MCP-1: Figure 2C,F,H; TNF-α: Figure 2D,G,I). Together with the increase in cells positively stained with F4/80, a marker for macrophages (Figure 2C-E,H,I), hindlimb immobilization appeared to instigate calf muscle atrophy involving local inflammatory responses including macrophage accumulation. We then sought to examine whether LCC, a massage-like mechanical intervention, modulated this immobilization-induced muscle inflammation and atrophy.

Among several different LCC magnitudes that we tested by changing the weight of the cylindrical unit, the one corresponding to 50 mmHg intramuscular pressure waves (LCC with 66 g, Figure 3A) appeared to most efficiently alleviate the immobilization-induced decrease in myofiber CSA and increase in macrophage accumulation in gastrocnemius muscles (Figure 3B). Based on the results of myofiber CSA and macrophage accumulation, we employed 66 g LCC for further studies. Notably, the LCC-induced intramuscular pressure waves, whose peak magnitudes were dependent on the cylindrical unit weight, were highly uniform (Figure 3A), indicating the consistency and reproducibility of LCC as a mechanical intervention on skeletal muscles.

LCC (1 Hz, 30 min per day, 7 days) significantly alleviated the immobilization-induced decreases in myofiber CSA of gastrocnemius muscles (Figure 4A,B). Furthermore, LCC partially tempered the immobilization-induced decrease in contracting force of triceps surae muscles (Figure 4C). In addition, LCC tempered the increases in F4/80-positive, TNF-α-positive, F4/80-, MCP-1-, and TNF-α-positive cells in gastrocnemius muscle tissues of immobilized hindlimbs (F4/80, Figure 4D,F; MCP-1, Figure 4D,G; TNF-α, Figure 4E,H). Collectively, LCC, which generates intramuscular pressure waves with an amplitude of 50 mmHg, alleviated immobilization-induced muscle atrophy and local inflammatory responses including macrophage accumulation.

Figure 1
Figure 1: Mouse bilateral hindlimb immobilization and local cyclical compression (LCC) application.
(A) Bilateral mouse hindlimbs were immobilized by spiral wiring with the hip joints abducted, the knee joints extended, and the ankle joints plantar-flexed. (B) LCC device. (C) Experimental set-up for LCC on the mouse calf. Please click here to view a larger version of this figure.

Figure 2
Figure 2: Mouse hindlimb immobilization, which atrophies calf muscles, induces a local inflammatory response.
(A) Cross-sectional micrographic images of anti-laminin-2 immunofluorescence staining of gastrocnemius muscles. High magnification images (right) refer to the areas indicated by rectangles in low magnification images (left). Scale bars, 100 µm. (B) Immobilization induced muscle atrophy. CSA of gastrocnemius myofibers decreased with the period of hindlimb immobilization. To quantify CSA, 100 myofibers were randomly chosen. Data are presented as means ± S.D. *, P < 0.05, one-way ANOVA with post hoc Bonferroni test (n = 3 mice for each group). (C and D) Micrographic images of anti-MCP-1 (green in C) and anti-TNF-α (green in D) and anti-F4/80 (red) immunostaining. For merged presentation (green and red), low and high magnification images are laid as in (A). Arrows point to double positive cells for F4/80 and MCP-1 (C) or TNF-α (D) Scale bars, 100 µm. (E-I) Quantification of anti-MCP-1, anti-TNF-α, and anti-F4/80 immunostaining. Effects of immobilization were analyzed with reference to the period of bilateral hindlimb immobilization. Statistical analysis was conducted with reference to the ‘Day 0’ samples (gastrocnemius muscles from mice that were not subjected to immobilization). Data are presented as means ± S.D. *, P < 0.05, one-way ANOVA with post hoc Bonferroni test (n = 3 mice for each group). This figure has been modified with permission11. Please click here to view a larger version of this figure.

Figure 3
Figure 3: Effects of LCC with different magnitudes on immobilization-induced muscle atrophy and inflammation response.
(A) Application of different magnitudes of LCC by changing the weight of the cylindrical unit. Scale bar, 1 s. 36-g, 66-g and 200-g cylindrical units produced 45 mmHg, 50 mmHg and 140 mmHg intramuscular pressure waves, respectively. (B) Comparison of the effects of LCC application to immobilized hindlimbs with 36-g, 66-g and 200-g cylindrical units. CSA of gastrocnemius myofibers (left) and F4/80-positive cells (right) of LCC-applied calf were quantified as relative values to those of the control hindlimb, which was not exposed to LCC, in each mouse. Data are presented as means ± S.D. *, P < 0.05, one-way ANOVA with post hoc Bonferroni test (n = 4 mice for each group). This figure has been modified with permission11. Please click here to view a larger version of this figure.

Figure 4
Figure 4: LCC attenuates immobilization-induced muscle atrophy and inflammatory response.
(A,B) Alleviation of immobilization-induced muscle atrophy by LCC application. CSA of gastrocnemius myofibers (B) was analyzed as in Figure 2B. Data are presented as means ± S.D. *, P < 0.05; **, P < 0.01, one-way ANOVA with post hoc Bonferroni test (n = 6 mice for each group). (C) The decrease in contracting force of triceps surae muscles after immobilization and its partial restoration by LCC. Data are presented as means ± S.D. *, P < 0.05, paired Student’s t test (n = 4 mice for control, n = 5 mice for immobilization group). (D,E) Micrographic images of anti-MCP-1 (green in D), anti-TNF-α (green in E) and anti-F4/80 (red) immunofluorescence staining of gastrocnemius muscles of mobilized (top) and immobilized hindlimbs without (middle) and with (bottom) LCC application are presented as in Figure 2C,D. Scale bars, 100 µm. (F-H) Quantification of anti-MCP-1, anti-TNF-α, and anti-F4/80 immunostaining. We compared calf muscles of immobilized hindlimbs with and without LCC application. Data are presented as means ± S.D. *, P < 0.05; **, P < 0.01, one-way ANOVA with post hoc Bonferroni test (n = 6 mice for each group). This figure has been modified with permission11. Please click here to view a larger version of this figure.

Supplementary Figure 1: Schematic representation of experimental protocols. Please click here to download the figure.

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Discussion

We have described a method for applying a massage-like mechanical stimulus, which has anti-inflammatory effects. Our system has following advantages even when compared with those reported previously. First, previous studies did not quantitatively define the mechanical forces applied2 or defined their magnitudes based on the measurement at the body surface, but not inside the tissues10. In contrast, we measured intramuscular pressure using a blood pressure telemeter. Second, the simple structure of our device (Figure 1B) allowed us to construct the system with high consistency and reproducibility (Figure 3A) at a relatively low cost. Third, our intervention (LCC) relates to physical activity (mild muscle contraction) with regard to intramuscular pressure changes (50 mmHg15). Our approach will provide a scientific basis for massage-like intervention as a possible therapeutic/preventative procedure that lessens the demerit of physical inactivity16.

The most critical step in our protocol is the positioning of mouse hindlimbs (Protocol step 3.3). We need to apply LCC in the direction perpendicular to calf muscles; otherwise, muscle tissues will be partly squeezed and damaged even when the 66-g cylindrical unit is used.

The limitation of the LCC method includes the requirement of anesthesia, which may have some effects on muscle metabolism. Also, we cannot entirely preclude the influences of tiny muscle contraction that may be caused as a reflex to sharp impacts during LCC application.

In conclusion, we have demonstrated that interstitial fluid movement mediates the LCC effects11. We may be able to induce interstitial flow more efficiently by modifying the mode of cyclical compression. For example, compression of sinusoidal mode may be better as compared to sharp strokes used in our current study.

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Disclosures

The authors declare that there are no competing interests associated with the manuscript.

Acknowledgments

We thank K. Nakanishi, K. Hamamoto, N. Kume, and K. Tsurumi for their consistent support throughout the project. This work was in part supported by Intramural Research Fund from the Japanese Ministry of Health, Labour and Welfare; Grants-in-Aid for Scientific Research from the Japan Society for the Promotion of Science; MEXT-Supported Program for the Strategic Research Foundation at Private Universities, 2015-2019 from the Japanese Ministry of Education, Culture, Sports, Science and Technology (S1511017).

Materials

Name Company Catalog Number Comments
Aluminum wire DAISO JAPAN B028 An aluminum wire is used to avoid escaping restriction by the wire
Blood pressure telemeter Millar SPR-671 A blood pressure telemeter is used to mesure intramuscular pressure.
DAPI Thermo Fisher Scientific D1306 DAPI is a fluorescent probe which is commonly used to stain DNA for fluorescent microscopy.
Goat anti-rabbit Alexa Fluor 488 (Dilution ratio, 1:500) Invitrogen A11034 Antibody for immunohistochemical staining.
Goat anti-rat Alexa Fluor 568 (Dilution ratio, 1:500)) Invitrogen A11077 Antibody for immunohistochemical staining.
ImageJ NIH N/A Analysis software for image
LabChart8 ADInstrumens   Analysis software for acquiring biological signals.
Prolong gold Thermo Fisher Scientific P36930 Prolong gold is for mounting stained samples.
Protein Block Serum-Free Dako X090930-2 For blocking non-specific background staining in immunohistochemical procedures.
Rat monoclonal anti-laminin-2 antibody (Dilution ratio, 1:1000) Sigma Aldrich L0663 Antibody for immunohistochemical staining.
Rat monoclonal anti-F4/80 antibody (Dilution ratio, 1:500) Abcam ab6640 Antibody for immunohistochemical staining.
Rabbit polyclonal anti-MCP-1 antibody (Dilution ratio, 1:1000) Abcam ab25124 Antibody for immunohistochemical staining.
Rabbit polyclonal anti-TNF-α antibody (Dilution ratio, 1:1000) Abcam ab66579 Antibody for immunohistochemical staining.
Surgical tape 3M Japan 1530EP-0 Surgical tape is used to restrict joint movement.

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References

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  3. Waters-Banker, C., Butterfield, T. A., Dupont-Versteegden, E. E. Immunomodulatory effects of massage on nonperturbed skeletal muscle in rats. Journal of Applied Physiology. 116, (2), 164-175 (2014).
  4. Haas, C., et al. Massage timing affects postexercise muscle recovery and inflammation in a rabbit model. Medicine & Science in Sports & Exercise. 45, (6), 1105-1112 (2013).
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  8. Mantovani Junior, N., et al. Effects of massage as a recuperative technique on autonomic modulation of heart rate and cardiorespiratory parameters: a study protocol for a randomized clinical trial. Trials. 19, (1), 459 (2018).
  9. Zeng, H., Butterfield, S., Agarwal, F., Haq, T., Zhao, Y. An engineering approach for quantitative analysis of the lengthwise strokes in massage therapies. Journal of Medical Devices. 2, (4), (2008).
  10. Wang, Q., et al. A mechatronic system for quantitative application and assessment of massage-like actions in small animals. Annals of Biomedical Engineering. 42, (1), 36-49 (2014).
  11. Saitou, K., et al. Local cyclical compression modulates macrophage function in situ and alleviates immobilization-induced muscle atrophy. Clinical Science. 132, (19), 2147-2161 (2018).
  12. Onda, A., et al. A New mouse model of skeletal muscle atrophy using spiral wire immobilization. Muscle Nerve. 54, (4), 788-791 (2016).
  13. Luster, A. D. Chemokines--chemotactic cytokines that mediate inflammation. The New England Journal of Medicine. 338, 436-445 (1998).
  14. Reid, M. B., Li, Y. P. Tumor necrosis factor-α and muscle wasting: a cellular perspective. Respiratory Research. 2, (5), 269-272 (2001).
  15. Baumann, J. U., Sutherland, M. D., Hangg, A. Intramuscular pressure during walking: An experimental study using the wick catheter technique. Clinical Orthopaedics Related Research. 145, 292-299 (1979).
  16. Lee, I., et al. Effect of physical inactivity on major non-communicable diseases worldwide: an analysis of burden of disease and life expectancy. Lancet. 380, 219-229 (2012).

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