Ex Vivo Whole Muscle Physiology for the Assessment of Skeletal Muscle Function in Mice
Article information
Abstract
PURPOSE
The purpose of this study was to describe ex vivo muscle physiology techniques and their application in assessing skeletal muscle function under controlled experimental conditions.
METHODS
Male C57BL/6 mice aged 3 months were used, and isolated skeletal muscles (EDL and soleus) were prepared under controlled experimental conditions using Krebs-Ringer buffer. After dissection, muscles were mounted in an isolated muscle system and adjusted to the optimal muscle length (Lo). Electrical stimulation was applied using supramaximal intensity with controlled frequency, duration, and intervals. Muscle function was evaluated by measuring twitch and tetanic force, force-frequency relationship, and fatigue and recovery using electrical stimulation at the optimal muscle length.
RESULTS
Functional parameters provided complementary information for evaluating intrinsic muscle function. Twitch and tetanic force reflected basic contractile capacity. The force-frequency relationship indicated responsiveness to stimulation and calcium handling characteristics. Fatigue and recovery provided insight into energy metabolism and resistance to functional decline. Standardized experimental conditions contributed to reliable and reproducible measurements. Data were analyzed using absolute force, normalized force, and percentage of maximal force.
CONCLUSIONS
Ex vivo muscle physiology techniques provide a reliable and reproducible approach for quantitatively evaluating skeletal muscle function by enabling direct assessment of intrinsic contractile properties under controlled conditions.
INTRODUCTION
Skeletal muscle constitutes about 40% of body mass. Skeletal muscle is essential for maintaining homeostasis, including movement, glucose metabolism, thermoregulation, and energy storage [1]. Multinucleated muscle fibers compose skeletal muscle and produce force for movement through processes such as neuromuscular transmission, intracellular calcium regulation, and interactions among sarcoplasmic proteins [2]. Therefore, skeletal muscle research requires quantitative assessment of muscle function together with molecular and structural analyses [3]. Such integrated evaluations improve understanding of the relationship between muscle structure, molecular alterations, and contractile function [3].
Traditional in vivo studies have limitations because it is difficult to completely exclude systemic factors, including the nervous system, hormones, and blood flow [4]. Molecular biological analyses alone are limited in directly reflecting actual muscle function [4]. Accurate methods are required to evaluate the intrinsic contractile properties and functional changes of muscle.
Mouse models are widely used in skeletal muscle research because they closely mimic human physiology and allow genetic manipulation to investigate mechanisms underlying skeletal muscle function and dysfunction [1,5,6].
Muscle function can be assessed using in vivo, ex vivo, in vitro, and in situ approaches [7]. Each approach has distinct advantages and limitations for evaluating skeletal muscle physiology [7]. In vivo approaches preserve the physiological environment [7]. However, systemic influences make it difficult to isolate the intrinsic contractile properties of skeletal muscle [8]. In vitro approaches provide excellent experimental control but do not preserve the structural and functional integrity of intact muscle [9]. In situ approaches preserve anatomical continuity with surrounding tissues at the expense of more complex surgical procedures and continued systemic influences [10]. Among these, ex vivo muscle physiology techniques are used to quantitatively analyze the contractile function of isolated whole muscles [1]. These techniques allow quantitative evaluation of isolated muscle contractile function without systemic influences, such as neural activation, hormonal regulation, and blood flow [1]. Various mechanical parameters can be quantitatively measured in isolated whole muscles (e.g., extensor digitorum longus and soleus), such as twitch force, tetanic force, force-frequency relationship, contraction and relaxation rates, and fatigue resistance [11]. Consequently, ex vivo approaches provide direct insight into muscle function that is difficult to obtain from in vivo experiments alone [1,11].
Ex vivo systems preserve muscle structure and allow controlled experimental conditions [12]. This approach provides a balance between physiological relevance and experimental control [12]. These features are particularly important for evaluating muscle function under conditions such as aging, disease, or genetic modification [13,14]. In addition, these properties minimize the influence of compensatory mechanisms that may occur in vivo and allow more precise interpretation of intrinsic functional changes in muscle [13,14].
Furthermore, ex vivo muscle physiology provides a basis for integration with molecular biological approaches (e.g., proteomic and transcriptomic analyses) following functional assessment of muscle tissue [9]. This approach can be used to elucidate the relationship between functional and molecular alterations in muscle and provide an important basis for understanding the mechanisms of muscle dysfunction in various pathological conditions, including sarcopenia, cachexia, and metabolic diseases [15].
Although several methodological papers have described ex vivo muscle physiology techniques, they primarily focus on individual experimental procedures or specific contractile measurements. However, few studies have integrated experimental preparation, muscle dissection, functional assessment, data analysis, and validation into a standardized workflow. This gap indicates the need for a comprehensive methodological framework that integrates the key components of ex vivo muscle physiology.
Therefore, ex vivo muscle physiology techniques are essential for analyzing the structure-function relationship of skeletal muscle and elucidating how molecular changes lead to functional impairments. The purpose of this study is to describe ex vivo muscle physiology and its application in assessing skeletal muscle function.
METHOD DEVELOPMENT
1. Experimental preparation
1) Preparation of Krebs-Ringer buffer and organ bath
Krebs-Ringer buffer was used to maintain the viability of mouse muscle tissue. Prepare Krebs-Ringer buffer by dissolving 115 mM NaCl, 5.9 mM KCl, 1.2 mM MgCl₂, 1.2 mM NaH₂PO₄, 1.2 mM Na₂SO₄, 2.5 mM CaCl₂, 25 mM NaHCO₃, and 10 mM dextrose in distilled water [16].
Adjust the pH to 7.4 using HCl or NaOH and confirmed the pH with a calibrated pH meter.
Store the buffer at 4°C until use. Before the experiment, fill the organ bath with the Krebs-Ringer buffer and prewarm it for 30 minutes using a circulating pump. Maintain the bath temperature was set to 25°C [16].
Continuously bubble the solution with a gas mixture of 95% O₂ and 5% CO₂ after the bath temperature has stabilized.
2) Animal preparation
Use male C57BL/6 mice aged 3 months for the experiment. All animal procedures were approved by the Institutional Animal Care and Use Committee (IACUC) of Hanyang University (Approval No. 2026-0133A) and were performed in accordance with the institutional guidelines for the care and use of laboratory animals.
Induce anesthesia by placing ether-soaked cotton in a conical tube and exposing the mouse to 100% diethyl ether vapor until an adequate level of anesthesia is achieved [17].
Place the anesthetized mouse on a warming pad and maintain the body temperature at 36°C throughout the procedure.
Position the mouse in the supine postion and secure the limbs before muscle dissection.
3) Muscle dissection
The Mouse hindlimb muscle dissection is summarized in Fig. 1. Make a skin incision at the ankle and carefully remove the skin toward the knee to expose the lower hindlimb muscles. Gently remove the fascia to clearly identify each muscle.
Mouse hindlimb muscle dissection procedure. (A) Exposure of the Lower hindlimb muscles. (B) Identification of the Achilles tendon. (C) Insertion of a suture into the Achilles tendon. (D) Traction of the Achilles tendon using the suture. (E) Ligation of the Achilles tendon with the suture. (F) Elevation of the attached to the soleus tendon. (G) Dissection around the soleus tendon. (H) Ligation of the opposite end of the soleus tendon. (I) Isolation of the soleus tendon.
Prepare 5-0 silk sutures before initiating muscle dissection.
Identify the Achilles tendon and tie the distal tendon with a 5-0 silk suture. Cut the tendon using fine scissors and dissect the soleus muscle proximally while minimizing contact with the surrounding connective tissue.
Identify the tibialis anterior (TA) muscle on the anterior aspect of the hindlimb. Cut the distal tendon at the ankle and carefully separate the muscle toward the proximal end. Complete the dissection by cutting the proximal tendon near the knee.
Retract the TA muscle laterally to expose the EDL muscle. Cut the distal tendon on the dorsal side of the foot and carefully dissect the muscle toward the proximal end while removing surrounding connective tissue. Complete the dissection by cutting the proximal tendon near the knee.
Throughout the dissection, handle only the tendons to minimize damage to the muscle tissue.
Immediately transfer the isolated muscles into oxygenated Krebs-Ringer buffer to preserve physiological viability. Keep the muscles continuously moistened with Krebs-Ringer buffer to prevent drying and preserve both tendons at sufficient length for subsequent ex vivo muscle physiology analysis.
2. Experimental analysis
1) Muscle mounting and determination of optimal length
Use the Aurora Scientific 1200A Isolated Muscle System (Vertical, Aurora Scientific Inc., Newmarket, Canada, catalog number: 800C-1). Immediately transfer the isolated muscle to an organ bath containing Krebs-Ringer buffer and allow it to equilibrate for 10-15 minutes.
Attach one tendon to the fixed hook and secure the opposite tendon to the force transducer using a 5-0 silk suture (Fig. 2). Align the muscle fibers carefully to prevent twisting and avoid excessive handling during mounting to minimize mechanical damage.
Muscle mounting with optimal length setting. (A) Attachment of the muscle tendon to a force transducer. (B) Determination of optimal length.
Apply electrical stimulation to induce muscle contraction and gradually increase the muscle length until the optimal length is reached (Fig. 2). Determine optimal length by repeatedly applying single-twitch stimulation while measuring twitch force at each muscle length [18]. As muscle length increases, twitch force progressively increases to a maximum value [18]. Define the muscle length that produces the maximum twitch force as optimal length and perform all subsequent functional assessments at this length.
2) Stimulation parameters
Electrical stimulation was used to induce muscle contraction and evaluate muscle function under controlled stimulation conditions, including stimulation frequency, intensity, pulse duration and stimulation interval.
Position the electrodes on the dual-mode lever system and maintain the muscle in oxygenated Krebs-Ringer buffer at the controlled bath temperature throughout the experiment [19].
Adjust the stimulation intensity to a supramaximal level to ensure maximal muscle contraction. Set the stimulation train duration according to the muscle type, using approximately 900 ms for the soleus and 300 ms for the EDL [20]. Set the interval between successive stimuli to allow sufficient recovery and minimize muscle fatigue. Perform all functional assessments while maintaining the muscle at its optimal length.
3) Twitch and tetanic force
Single twitch contractions and tetanic contractions were measured to evaluate the fundamental characteristics of the muscles. Apply a single electrical stimulus to record the isometric twitch force [21].
Gradually increase the stimulation frequency to elicit maximal tetanic force (Fig. 3A) [21]. Stimulation frequencies were set at 10, 40, 80, 120, 150, and 180 Hz for the EDL and 10, 40, 80, 100, 120, and 150 Hz for the soleus [21].
Comprehensive muscle contractile function assessment. (A) Tetanic force (B) Force-frequency (C) Fatigue (D) Recovery.
Allow a 1-minute rest period between successive contractions to minimize muscle fatigue.
4) Force-frequency
The Force-frequency relationships were evaluated by measuring muscle force in response to progressively increasing stimulation frequencies (Fig. 3B).
Apply electrical stimulation while maintaining a constant pulse duration and supramaximal stimulation intensity [18]. Increase the stimulation frequency sequentially to 10, 40, 80, 120, 150, and 180 Hz for the EDL and 10, 40, 80, 100, 120, and 150 Hz for the soleus [21]. Record the maximal isometric force generated at each stimulation frequency.
Set the stimulation train duration according to muscle type, using 300 ms for the EDL and 900 ms for the soleus [18]. Allow the muscle to rest for 1 minute between successive stimulations to ensure sufficient recovery and minimize muscle fatigue.
5) Fatigue and recovery
Repeated electrical stimulation was used to evaluate muscle fatigue and recovery (Fig. 3C and 3D).
Apply repeated 80 Hz tetanic stimulation to both the EDL and soleus muscles until tetanic force declines to 50% of the initial peak force (Fig. 3C) [21]. Induce successive tetanic contractions at 5 seconds intervals throughout the fatigue protocol. Quantify muscle fatigue by calculating the area under the force-time curve (AUC). Following the fatigue protocol. assess recovery using 80 Hz stimulation for the EDL and 150 Hz stimulation for the soleus at 5 minutes intervals (Fig. 3D) [21].
Record tetanic force at each recovery time point.
Calculate the initial recovery rate as the slope of force increase between the end of the fatigue protocol and the first recovery contraction [21]. Determine the total recovery force during the recovery period.
3. Data analysis
1) Twitch and tetanic force
Twitch and tetanic force were analyzed based on isometric contraction data obtained at the optimal length. Twitch force was determined from the contractile response elicited by a single electrical stimulus [22]. Tetanic force was defined as the maximal force generated under highfrequency stimulation [22]. Force reached a plateau with increasing stimulation frequency. Active force was calculated as the difference between maximal and baseline force for all contractions [1]. For twitch contractions, time to peak tension (TPT), half-relaxation time (HRT), and contraction time were analyzed as measures of contractile kinetics from the force-time curve [23]. For tetanic contractions, the maximal force and the force-time integral were calculated to evaluate force-generating capacity and maintenance characteristics [23]. All force values were expressed in absolute units (g) and normalized to Cross-Sectional Area (CSA) to calculate specific force (g/mm²) [21]. Following completion of the contractile measurements, the isolated muscle was carefully removed from the apparatus, gently blotted to remove excess Krebs-Ringer buffer and weighed. Muscle cross-sectional area (CSA) was subsequently calculated from muscle mass and optimal muscle length (Lo) using the following equation:
where 1.06 g/cm³ represents the density of mammalian skeletal muscle. Specific force was calculated by normalizing the absolute force to CSA and expressed as g/mm² [21,22].
2) Force-frequency
The force-frequency relationship was assessed by stimulating the muscle at frequencies ranging from a single twitch (1Hz) to the maximal tetanic frequency [23]. Active force was calculated as the difference between maximal and baseline force at each stimulation frequency [24]. The force-frequency curve was generated by plotting active force against stimulation frequency [23]. Data were analyzed in three ways: (1) absolute force (g), (2) specific force normalized to CSA (g/mm²), (3) percentage of the maximal tetanic force (%) for each muscle [23,24].
Collected data can be exported to Excel and curve fitting can be performed using it. The data can be expressed as absolute force (g), normalized to CSA (g/mm²), and Percentage of the maximal tetanic force (%) for each muscle [23,24].
3) Fatigue and recovery
Fatigue was assessed by applying repeated stimulation until tetanic force declined to 50% of the initial peak force [25]. During the fatigue protocol, maximal force was measured for each contraction or at specified time intervals. The initial force was set to 100% and the decline in force during the protocol was expressed as a percentage of the initial force to generate a fatigue curve [26]. Time to reach 75%, 50%, and 25% of the initial force was analyzed as an additional index of fatigue [26]. Data were compared using absolute values or values normalized to CSA.
Recovery was assessed by applying tetanic stimulation at defined time intervals following the fatigue protocol [25]. The maximal tetanic force was expressed as a percentage of the pre-fatigue initial value at each time point [25]. The recovery curve represents the restoration of force during the recovery period and was used to evaluate the muscle’s functional recovery capacity [25]. Total recovered force was quantified over the entire recovery period to provide an integrated assessment of recovery capacity [25].
VALIDATION
Ex vivo muscle physiology techniques are established methods for quantitatively evaluating the contractile function of skeletal muscle under controlled experimental conditions. [18] These techniques provide high validity for analyzing the intrinsic functional properties of muscle [27]. Various functional parameters can be measured using isolated whole muscles, including twitch force, tetanic force, and fatigue and recovery [18]. Functional indices are closely associated with molecular and structural characteristics such as muscle fiber type, sarcomeric protein structure, and calcium handling capacity [28]. In addition, these techniques are performed under conditions that minimize systemic influences such as nervous system, hormones and blood flow. Therefore, ex vivo muscle physiology techniques provide a valid assessment to directly evaluate the intrinsic function of the muscle.
The extensor digitorum longus (EDL) and soleus muscles exhibit distinct physiological and functional characteristics that should be considered when designing and interpreting ex vivo muscle physiology experiments [29]. The EDL consists predominantly of fast-twitch glycolytic fibers (type II), which generate rapid contractions and high maximal force but fatigue relatively quickly. In contrast, the soleus is composed primarily of slow-twitch oxidative fibers (type I), which produce slower contractions, lower maximal force, and greater fatigue resistance [29]. These intrinsic differences influence contractile kinetics, calcium handling properties, and metabolic characteristics, which require muscle-specific stimulation protocols [29]. Consequently, different stimulation frequencies, pulse durations, and fatigue-recovery protocols are applied to achieve optimal muscle activation and to accurately evaluate contractile function [30].
Furthermore, ex vivo muscle physiology techniques can detect consistent functional differences across a variety of disease and experimental models while preserving the characteristic functional properties of individual muscles, such as the EDL and soleus [23]. This characteristic supports the convergent validity of these techniques and enhances their applicability for investigating muscle-specific adaptations under physiological and pathological conditions.
In terms of reproducibility, the reliability of ex vivo muscle physiology depends on the standardization of experimental conditions [23]. Optimal muscle length (Lo) must be accurately determined, and stimulation parameters should be consistently maintained. In addition, the temperature of the experimental solution and oxygen supply must be properly controlled. Controlled conditions ensure low variability and high reproducibility in key functional measurements [18,23,27].
Moreover, normalizing force based on muscle cross-sectional area or mass is essential to improve comparability between samples and ensure consistent data interpretation across experiments [21,23].
Overall, Ex vivo muscle physiology techniques provide a reliable and reproducible approach for quantitatively evaluating skeletal muscle function under controlled experimental conditions. Standardized experimental protocols and consideration of muscle-specific physiological characteristics improve the validity and interpretation of contractile measurements.
DISCUSSION
Ex vivo muscle physiology techniques provide a reliable approach for quantitatively evaluating the contractile properties of skeletal muscle [28]. This technique provides important information for analyzing the intrinsic functional characteristics of muscle [8]. In particular, measurements are performed under conditions that minimize systemic factors such as neural input, hormones, and blood flow [8]. Therefore, this technique allows direct assessment of functional changes within the muscle itself.
Various functional parameters provide complementary information. These integrated analysis enables a more comprehensive understanding of muscle function [28]. Twitch and tetanic force reflect basic contractile capacity. The force-frequency relationship is closely related to calcium handling capacity and muscle fiber type characteristics [31,32]. Fatigue and recovery assessments indicate energy metabolism and fatigue resistance [33]. Integrated analysis of these parameters facilitates precise interpretation of the complex functional properties of muscle that cannot be captured by a single parameter.
Standardized experimental conditions improve the reproducibility and reliability of the data [28]. All measurements were performed at the optimal muscle length (Lo). Stimulation parameters, temperature, and oxygen supply were consistently maintained. These conditions reduce variability between experiments and enhance the comparability of results [28]. They also contribute to consistent data interpretation.
However, these techniques require careful interpretation due to the nature of the experimental conditions. Although ex vivo approaches preserve muscle structure, they still lack the full range of physiological regulatory mechanisms present in vivo, which may limit the ability to fully replicate physiological [8]. In addition, the results may be influenced by mechanical stress during muscle dissection and minor variations in experimental conditions [8,28]. Therefore, these factors should be considered when interpreting the results.
Future studies should interpret ex vivo findings in conjunction with in vivo models [34]. In addition, integration with molecular biological analyses improves understanding of the relationship between functional and molecular changes [28]. This approach provides an important basis for understanding the mechanisms underlying muscle dysfunction in pathological conditions such as sarcopenia, cachexia, and metabolic diseases.
CONCLUSION
Ex vivo muscle physiology techniques provide a reliable method for quantitatively evaluating the contractile function of skeletal muscle under controlled experimental conditions. These techniques enable direct assessment of intrinsic muscle function by minimizing systemic influences such as neural input, hormones, and blood flow. Multiple functional parameters allow comprehensive evaluation of muscle function. Standardized experimental conditions contribute to high reproducibility and reliability of the data. Thus, these characteristics support the use of ex vivo muscle physiology techniques as a reliable approach for evaluating skeletal muscle function.
Notes
CONFLICT OF INTEREST
The authors declare that they have no conflicts of interest and received no financial support from any organization for this work.
AUTHOR CONTRIBUTIONS
Conceptualization: D Lee, D Kwak; Data curation: D Lee, B So, D Kwak; Formal analysis: D Lee, B So, D Kwak; Funding acquisition: None; Methodology: D Lee, B So, D Kwak; Project administration: D Kwak; Visualization: D Lee, B So; Writing - original draft: D Lee, D Kwak; Writing - review & editing: D Lee, B So, D Kwak.