A method to test contractility of the supraspinatus muscle in mouse, rat, and rabbit

Abstract

The rotator cuff (RTC) muscles not only generate movement but also provide important shoulder joint stability. RTC tears, particularly in the supraspinatus muscle, are a common clinical problem. Despite some biological healing after RTC repair, persistent problems include poor functional outcomes with high retear rates after surgical repair. Animal models allow further exploration of the sequela of RTC injury such as fibrosis, inflammation, and fatty infiltration, but there are few options regarding contractility for mouse, rat, and rabbit. Histological findings can provide a “direct measure” of damage, but the most comprehensive measure of the overall health of the muscle is contractile force. However, information regarding normal supraspinatus size and contractile function is scarce. Animal models provide the means to compare muscle histology, imaging, and contractility within individual muscles in various models of injury and disease, but to date, most testing of animal contractile force has been limited primarily to hindlimb muscles. Here, we describe an in vivo method to assess contractility of the supraspinatus muscle and describe differences in methods and representative outcomes for mouse, rat, and rabbit.

the shoulder joint (glenohumeral joint) consists of a large humeral head articulating with a relatively small, shallow socket. Thus the surrounding rotator cuff (RTC) muscles not only generate movement, but also provide important joint stability. RTC tears, particularly in the supraspinatus muscle, are a common problem encountered in orthopedics (12, 81). Large RTC tears can lead to irreversible muscle atrophy and fatty infiltration, especially in older patients (26, 38). Despite some biologic healing after RTC repair, problems include poor functional outcomes and retear rates after surgical repair, reportedly as high as 90% (24).

Muscle damage, which occurs after a RTC tear, has been defined and measured in many ways (e.g., inflammation, fatty infiltration, atrophy, changes in cell structure, etc.). Structural damage is evident in histological findings (9, 29, 46, 47, 63), but one problem with many of the biological markers used to assess muscle injury, including those used in animal studies, is that they may not correlate with the loss of force. There is a plethora of biological markers to assess muscle damage such as indicators of membrane damage (e.g., Evans blue dye) (29, 59, 77); disruption of the muscle fiber cytoskeleton (e.g., loss of desmin or titin) (42, 43, 71); changes in excitation-contraction coupling (33, 78, 84); alterations in force-generating structures or force-transmitting structures (32, 64, 80); increases in serum markers (21); alterations in composition of the extracellular matrix (7, 71, 76); changes in neuromuscular junction structure and function (17, 35, 67); altered muscle fiber morphology (13, 30); and altered signals with noninvasive imaging, such as MRI (20, 57, 73, 75). Muscle damage is often defined within the context of the assay used to examine it; however, no single biological marker can account for the changes in contractility. Since full contractile function can persist despite the presence of biological markers indicating damage, muscle function may be the most valid and comprehensive measure of muscle health (4).

Several animal models are available to measure muscle contractility, but these have been limited to testing small, thin muscles in vitro (33), or testing the ankle (48, 49, 51) or knee (65, 66) muscles in vivo. Recently published works have provided information regarding the size and/or whole muscle contractility of the supraspinatus, but the number of papers is limited and it is difficult to obtain an overview from the piecemeal information (Table 1). Fatty infiltration after a RTC tear is a common clinical problem that also occurs in rabbits (25, 60, 70, 82), but not reliably in rats (18). Despite this advantage of the rabbit as an animal model for fatty atrophy, we could only identify two publications that examine contractility in the rabbit (15, 16). There are only two or three studies providing data on whole muscle contractility for the rat, and none for the mouse (Table 1). Of the handful of studies in all three species, the lack of data on both muscle mass and contractility makes it difficult to normalize force to muscle size.

Table 1.

Published studies that include supraspinatus muscle weight and/or whole muscle contractile force

Authors (Ref No.)	PMID	Species	Sex	Age	Body Mass	Muscle Mass	Contractile Force
Mathewson et al. (58)	24072803	Rabbit	—	—	3.16 kg	7.9 g	—
Rowshan et al. (70)	20926720	Rabbit	Male	—	3–4 kg	6.09 g	—
Uhthoff et al. (82a)	24743593	Rabbit	Female	—	3.4–4.3 kg	8 g*	—
Fabis et al. (16)	9930099	Rabbit	Male	—	3.2–4.5 kg	—	8.5–11.1 N
Fabis et al. (15)	10888165	Rabbit	Male	—	3.7–4.6 kg	—	§
Ditsios et al. (11a)	24981552	Rat	Male	13 mo	500 g	—	1.93 N†
Mannava et al. (56b)	21445691	Rat	Male	—	400–450 g	—	5.1 N
Mannava et al. (56a)	21938374	Rat	—	—	400–450 g	—	§
Plate et al. (64a)	23791493	Rat	Male	12 mo	—	—	5.4 N
Mathewson et al. (58)	24072803	Rat	—	—	570 g	770 mg	—
Sato et al. (71)	25834081	Rat	Male	—	—	500 mg*	—
Liu et al. (47)	20949443	Rat	Female	—	250 g	310–400 mg*	—
Davies et al. (9)	25974842	Rat	Female	—	—	418.3 mg	—
Gumucio et al. (27)	22696414	Rat	Male	6 mo	—	740 mg	—
Liu et al. (46)	22488625	Mouse	Female	12 wk	—	34.6 mg	—
Mathewson et al. (58)	24072803	Mouse	—	—	30 g	30 mg	—

PMID, PubMed ID.

^*Estimated specific muscle mass from bar graph, as specific values were not provided.

^†Published in units of grams, but converted to newtons for ease of comparison.

^§Contractile testing was performed, but data were expressed as a percentage of maximal contractile force, rather than absolute force.

The overall goal of this work is to share detailed methods for testing contractile function of the supraspinatus muscle in commonly used research models such as mouse, rat, and rabbit. We also report normative values of muscle mass and contractile function for a given age and body weight. To the best of our knowledge, this is the first report of whole muscle contractile measurements for the supraspinatus muscles in mice, and one of only a handful of studies for rats and rabbits. We also provide new information on tendon morphometry and contractile characteristics such as maximal twitch and tetanic tension, force-frequency comparisons, and rates of fatigue. This protocol, together with the contractile data provided for mouse, rat, and rabbit muscles, should be valuable to researchers currently studying the biology of RTC pathology and wishing to assess the functional outcomes or therapeutic interventions.

MATERIALS AND METHODS

Contractile function.

All protocols were approved by the University of Maryland Institutional Animal Care and Use Committee. We used male mice (C57BL/10ScSn, body weight 23.6 ± 0.6 g, Jackson Laboratory, Bar Harbor, ME, N = 8), rats (Sprague-Dawley, body weight 242 ± 11 g, Charles River Laboratories, Germantown, MD, N = 8), and rabbits (New Zealand white, body weight 2.3 ± 0.6 kg, Charles River Laboratories, Germantown, MD, N = 8), all ∼3 mo of age. Before each experiment, the animal was anesthetized (∼ 4-5% isoflurane in an induction chamber, then ∼ 2% isoflurane via a nosecone for maintenance) using a precision vaporizer (cat. no. 91103, Vet Equip, Pleasanton, CA). During the procedure, the animal was kept warm by use of a heat lamp.

In the anesthetized animal, the suprascapular nerve was stimulated via subcutaneous needle electrodes (J05 Needle Electrode Needles, 36BTP, Jari Electrode Supply, Gilroy, CA) placed at the suprascapular notch. Proper electrode position was determined by a series of isometric twitches. Impulses 1 ms in duration were generated by an S48 square-pulse stimulator (Grass Instruments, West Warwick, RI) and passed through a PSIU6 stimulator isolation unit (Grass Instruments, West Warwick, RI). In preliminary experiments (data not shown), the suprascapular nerve was dissected free through a small incision and clamped with a subminiature electrode (Harvard Apparatus, Holliston, MA), which was used to stimulate the supraspinatus. There were no differences between placement of needle electrodes and the subminiature clamp electrode.

Contractile function of the isolated supraspinatus muscle was measured before tissue harvesting, similar to methods described previously (50, 52). Although contractile testing is performed in vivo, the tendon was released and this was a terminal procedure. After proper anesthetic depth was confirmed by lack of a deep tendon reflex (no foot withdrawal in response to pinching the foot), we used a custom-built rig to stabilize the scapula. This entails incising through the middle trapezius and rhomboid muscles to access the vertebral border of the scapula to place a clamp along the vertebral border near the infraspinatus fossa for complete immobilization of the scapula (Fig. 2). A second rig was specifically developed for the rabbit, as the devices used for rodents were insufficient to preclude scapula movement with the large forces generated by the rabbit supraspinatus. Once the scapula was securely immobilized, the tendon of the supraspinatus muscle was released and attached to a load cell. Single twitches (rectangular pulse, 1 ms) were applied at different muscle lengths to determine the optimal length (resting length, L₀). The rat and rabbit tendons were attached to the load cell via suture (sizes 4.0 and 0 Ethicon silk suture, respectively), but the mouse tendons can be difficult to suture; thus we left the tendon attached to the humeral head and sutured through the head for attachment to the load cell. At L₀, maximally fused tetanic contraction was obtained at ∼100 Hz (300-ms train duration of 1-ms pulses at a constant current of 5 mA). We used 150% of the maximal stimulation intensity to induce maximal activation of contraction, P₀. We also generated a force-frequency plot, obtained by progressively increasing the frequency of pulses during a 200-ms pulse train. To provide an index of fatigue, maximal tetanic contractions were performed repeatedly (every 2 s) with the final tension expressed as percentage of P₀. The entire procedure, from anesthesia to the completion of contractile testing takes ∼30 min for each animal, regardless of species.

Fig. 2.

Apparatus used to assess in vivo contractile function. A: drawing illustrates the general setup used to assess supraspinatus force. With care to avoid the supraspinatus muscle, the scapula is stabilized and the distal tendon of the supraspinatus is attached to the load cell. The load cell is mounted to a micromanipulator so that the muscle can be adjusted to resting length and aligned properly (i.e., adjusting the position between the origin and insertion of the muscle and the load cell so that there is a straight line of pull) in the X, Y, and Z directions. Subcutaneous electrodes are inserted at the suprascapular notch to stimulate the suprascapular nerve. Single twitches (1-ms duration) are induced at different muscle lengths (thus a length-tension, or L-T, curve) to determine optimal length (L_o). A maximal tetanic contraction is obtained to determine maximal contractile tension (P_o). [Used with permission.] B: the apparatus is shown without the animal. The custom-made devices we use to immobilize the scapula are adjustable in all 3 anatomical planes. A commercial clamp (provided it does not clasp the supraspinatus) would likely work too, but the forces generated by the rabbit are quite large, requiring a different rig (i.e., much more stable) than the one used for rodents.

Supraspinatus muscle-tendon length.

Muscle length (origin of the muscle belly at superior angle of the scapula to the muscle-tendon junction) and tendon length (muscle-tendon junction to insertion on the humerus) were measured in situ using digital calipers to the nearest hundredth of a millimeter. We have used this method before to examine even smaller structures (22). Although reliability was not formally tested here, all measurements were performed by the same investigator and confirmed by a second investigator. After the supraspinatus muscle was harvested, we followed the tendon into the muscle belly via microdissection to examine the length of the tendon, which we refer to as internal tendon length.

Statistical analysis.

Statistical analysis was not performed on obvious differences between species, such as body weight, muscle mass, and muscle force. To evaluate potential differences between groups for such variables as fatigue, normalized force, muscle length:tendon length ratios, external tendon:internal tendon ratios, and tetany:twitch ratios, a 1-way ANOVA was used (SigmaStat, San Rafael, CA). Significance was set at P < 0.05.

RESULTS

Anatomy.

Figure 1 shows the general anatomy of the area. The supraspinatus muscle is completely covered by the upper trapezius muscle and its tendon of insertion is obscured by the deltoid (Fig. 1A). For instructional purposes, Fig. 1B shows a detached scapula with overlying muscles removed, allowing a view of the supraspinatus and its distal tendon disappearing under the acromion bone. For experiments, the supraspinatus tendon is identified and then its tendon released for contractile testing [separation of the RTC tendons is straightforward in animals; the tendons still form a “cuff”, but they have a more distinct insertion than in humans (11, 72)]. Figure 2A shows the overall apparatus setup, and Fig. 2B provides further orientation and detail regarding scapula immobilization.

Fig. 1.

General anatomy of the shoulder. A: a euthanized animal is shown with the overlying skin removed. The major superficial muscles such as the trapezius, deltoid, and triceps are indicated, but the rotator cuff muscles are deeper and not visible. B: a detached scapula from an animal postmortem is shown with the superficial muscles removed. The supraspinatus can be visualized, but its tendon of insertion is difficult to see due to the overlying acromion and clavicle bones (AC joint). As discussed in the text, the individual rotator cuff (RTC) tendons in lower species are more distinct than in humans.

Muscle mass and tendon length.

As expected, there was a marked difference in muscle mass between the mouse, rat, and rabbit (32.5 ± 0.9 mg, 407 ± 9 mg, and 8,035 ± 150 mg, respectively, Fig. 3). The muscle length (length from proximal muscle belly at superior angle of scapula to the muscle-tendon junction) and total tendon length (external portion seen in situ and internal portion of the tendon observed after harvesting the muscle) were measured at the time of euthanasia. The overall muscle length:tendon length ratios were almost identical for all three species (1.57 ± 0.08), but interestingly, the mouse had the greatest ratio of external:internal tendon length (P < 0.05, Fig. 3B).

Fig. 3.

Comparison of supraspinatus muscles from mouse, rat, and rabbit. A: supraspinatus muscles from a mouse, rat, and rabbit are shown for size comparison. The figure shows the underside of the muscles, where the full length of the tendon diving into the muscle belly (“internal tendon”) was identified by microdissection and measured. B: the table shows weights of supraspinatus muscles from all mice, rats, and rabbits (2–3 mo old, N = 8 each species). In addition to muscle mass, muscle length and tendon length (combined length of both the visible external tendon and dissected internal tendon) are provided, as well as the ratios between the length that was external (L_ext, measured in situ from the superficial aspect) to the length that was internal (L_int, determined through microdissection). *P < 0.05.

Contractile function.

Figure 4A shows representative trace recordings from a mouse (green), rat (red), and rabbit (blue) supraspinatus muscle. The high forces generated by the rabbit supraspinatus muscle necessitate the rabbit scapula having its own separate stabilization device (see materials and methods and Fig. 2B). Maximal isometric tension in the mouse, rat, and rabbit (0.26 ± 0.03, 2.98 ± 0.60, and 15.3 ± 0.86 N, respectively, Fig. 4B) mirror the respective differences in mass and the previously reported cross-sectional area in these species (58). The tetany:twitch ratio was lower in the rabbits than in rodents (P < 0.05).

Fig. 4.

Comparison of maximal isometric contractile force from mouse, rat, and rabbit supraspinatus muscles. A: representative in vivo trace recordings from a rabbit (blue), rat (red), and mouse (green) are shown for the supraspinatus muscle. In this example, muscles were stimulated for 200 ms at optimal muscle length (L₀) to induce a maximal tetanic contraction (P₀). B: contractile data from mouse, rat, and rabbit supraspinatus muscles. Such absolute values shown can be normalized to muscle physiological cross-sectional area. For experiments, care should be taken to evaluate animals that are age-matched, sex-matched, and of the same strain and species. We did not compare obvious differences in absolute force between species, but statistical analysis for tetany:twitch ratio showed a lower ratio for rabbit compared with rodent. *P < 0.05.

Representative traces of increasing muscle force with increasing stimulation frequencies are shown in Fig. 5A. The mean force-frequency relationship for all animals is plotted on the graph in Fig. 5A and indicates a shift to the right (arrows) for the rabbits compared with rodents. We also compared the degree of supraspinatus muscle fatigue after repeated tetanic contractions (representative recording is shown in Fig. 5B). No significant differences were found in the rates of fatigue (bar graph in Fig. 5B), but our test of repeated contractions was limited to 2 min, so differences at longer time points might still exist.

Fig. 5.

A: comparison of force-frequency data from mouse, rat, and rabbit supraspinatus muscles. A, left: curves shown were produced from maximal contractions (200-ms duration) at incrementing higher frequencies (Hz) in the supraspinatus muscle in a rat. Note that maximal force is generated at ∼80–100 Hz, which diminishes with higher frequency (not shown). This optimal frequency can vary depending on the muscle group tested and the conditions (e.g., direct nerve stimulation as here vs. field stimulation when muscles are removed for in vitro experiments, not shown here). A, right: rabbit force-frequency curves showed a shift to the right (arrows) compared with mouse and rabbit. B, left: maximal tetanic tension can be performed repeatedly over time with the final tension expressed as percentage of the starting optimal tension (P_o), providing an index of fatigue at a desired point in time. In this example, the supraspinatus was isolated, adjusted to optimal length (L_o), and then stimulated with a 200-ms tetanic contraction every other second for 2 min. B, right: comparison of fatigue from mouse, rat, and rabbit supraspinatus muscles. There were no significant differences in muscle fatigue.

DISCUSSION

Methods for muscle function testing for some hindlimb muscles are well described, with a profuse amount of data generated by contractility studies (1, 3, 5, 10, 37, 39, 40, 42, 54, 61, 67, 68). The shoulder presents some unique challenges for testing RTC muscles, such as the depth of the supraspinatus tendon and the overlying acromion (Fig. 1). The shape of the scapula provides a challenge for bony fixation compared with the long lever arms in the lower extremity, but the apparatus (Fig. 2) and methods to test the supraspinatus muscle described here are relatively straightforward. In the present study, we describe an in vivo (nonsurvival) model to assess supraspinatus muscle contractility (e.g., twitch, tetany, force-frequency, and fatigue) in the mouse, rat, and rabbit. We have also provided contractile data and morphological measurements of the muscle and tendon lengths in all three animal models.

There are advantages and disadvantages when studying various species. Large animals such as the rabbit can be easier to work with, including ease of testing contractility, feasibility of surgical interventions, and providing large structures for in vivo imaging. Furthermore, the high forces generated by the rabbit supraspinatus also make it an appealing model to work with, as differences after injury, tenotomy, repair, or treatment will likely be easier to discern. The small structures and correspondingly small forces generated by the rat, and especially mouse, can make them more difficult to work with. However, there are advantages to working with smaller animals including ready availability, comparability to previous studies in the literature, and having a known genome. Because of access to many genetically altered models, mouse studies can be better suited to study mechanisms underlying changes in muscle function.

As expected, the overall tendon length (internal and external combined, Fig. 3B) scales with animal size, but the ratio of external:internal tendon length was proportional in all three species, and it was greatest for the mouse. While normative data are available for supraspinatus muscle mass and architecture (11, 15, 16, 26, 58, 74), information specific to the supraspinatus tendon length has not been reported. The supposition that the tendon length scales with overall body size and muscle mass among the three species assumes that supraspinatus function is identical in all three. However, differences between species in upper extremity use patterns during gait (8, 31) could preclude identical muscle and tendon architecture in all quadrupeds.

In addition to providing methods for testing contractility, this study provides normal values of muscle weight and contractile data for investigators wishing to study whole muscle contractility of the supraspinatus in various animal models. The absolute quantity of muscle mass is generally well correlated to muscle strength (23), and specific tension (contractile force normalized to physiological cross-sectional area, or PCSA) is similar in most mammals (44). Thus the stepwise increase in muscle mass and contractile force from mouse to rat to rabbit was not surprising, but there were some unexpected findings. For instance, the tetany:twitch ratio was lower in the rabbits than in rodents. This could be due to muscle fiber type composition, as muscles with a higher number of slow muscle fibers can have a lower tetany:twitch ratio (6). Information regarding fiber type composition of the supraspinatus in animals is scarce; there are, to the best of our knowledge, only a few studies that report on the supraspinatus of the rat (2, 27, 28) and even fewer for the rabbit (14, 70), with little to no data regarding fiber type composition in the mouse supraspinatus. Despite the paucity of information, the consensus appears to be that, like humans, the fiber type in the supraspinatus of rodent and rabbit is mixed. However, specific or quantitative comparisons are difficult based on the small number of studies, which have used a variety of methods (2, 14, 70).

The relationship between a muscle's activation frequency and isometric force is evident in the sigmoidal force-frequency curve, which shows a steep rise in force with increasing stimulation frequencies (Fig. 5A). This curve can be shifted to the left or right (reaching peak force at lower or higher frequencies, respectively) for several reasons, such as changes in passive tension during tests or differences in fiber type composition within a muscle (19, 55). We observed a shift to the right of the force-frequency relationship in the rabbit compared with the rodents. This shift results in a reduced summation of force at lower stimulation frequencies in rabbit muscle, which is consistent with a lower tetany:twitch ratio, especially if there is a higher percentage of slow fibers within the rabbit supraspinatus. There are several alternative explanations for the lowered tetany:twitch ratio in rabbits compared with rodents. For example, length-dependent calcium sensitivity seems to be a major factor determining the magnitude of the shift of optimal muscle length (69), and differences have been documented in the geometry and function of the sarcoplasmic reticulum rodents and rabbits (34, 62), which could affect intracellular Ca²⁺ kinetics. It is possible that the response of force to different frequencies is also affected by potentiation of the contractile system, distribution of sarcomere length, and interactions between force exerted and aponeurosis length (69).

Fiber type composition affects the speed of a muscle contraction, but less so the specific tension (force per unit area). Specific tension of skeletal muscle is considered relatively constant (53, 56). Force depends not only on the size and number of the fibers in the supraspinatus, but also on muscle architecture, and such variables have been well described for humans and animals (58, 83). Muscle mass can vary significantly based on species, age, and health of the animal. Since the maximal force per unit of cross-sectional area (specific tension) of skeletal muscle is considered relatively constant, contractile force of a skeletal muscle can be estimated based on its physiological cross-sectional area (PCSA) (41), represented by the equation: PCSA (mm²) = M(g) × cosθ/ρ(g/mm³) × L_f (mm), where M is muscle mass, θ represents the angle of the fibers (pennation), ρ is muscle density (1.056 g/cm³ in mammalian muscle) and L_f represents fiber length (estimated from length of the measured fiber bundle). We gleaned muscle fiber pennation angle and relative fiber length from published studies that have measured these architectural variables in mouse, rat, and rabbit supraspinatus muscles (58). Dividing P₀ by PCSA, the calculated specific tension was, as expected, similar between the mouse, rat, and rabbit supraspinatus muscles (4.48 ± 0.97 kg/cm²). Although not significantly different, the rabbit generated the highest specific tension (5.42 kg/cm²). This could be due to differences in architecture between rodents and rabbits. Mathewson et al. (58) did a careful study that compared RTC architecture between mouse, rat, and rabbits (among other species). They found that the architectural difference index (ADI), a combined measure of fiber length-to-moment arm ratio, fiber length-to-muscle length ratio and the fraction of the total RTC physiological cross-sectional area contributed by each RTC muscle, was higher (less like human architecture) for the rabbit than mouse or rat. Such differences between rabbit and rodent could help explain slight changes in normalized force.

There are large differences between humans and nonprimates when comparing the percentage of the supraspinatus mass relative to total RTC mass. As expected, the supraspinatus in quadruped animals is comparatively larger than those of bipedal animals, such as primates and humans (58). Thus, even though the ADI of the rodent more closely resembles the human, the size of the supraspinatus in the rabbit is closest to the overall size of the human supraspinatus. There are also differences in the bony anatomy and, compared with rabbits, the rodent bony anatomy is more similar to the human (11, 74). Analogous to the human shoulder, the rodent acromion projects anteriorly over the humeral head to the clavicle, creating an enclosed arch over the supraspinatus tendon. The bony anatomy of rabbit diverges from human, in that the acromion, clavicle, and the coracoid process are generally minimal or nonexistent and do not cover the RTC (11).

The majority of full-thickness RTC tears present in patients over 50 years of age (12, 81). Chronic tears can lead to fatty degeneration of the muscle, which is a poor prognostic factor for healing (45, 79). Despite many advantages of using a rodent model, there is no consensus regarding which animal model best mimics the pathophysiology of human RTC tears. The rabbit shoulder is well established as an animal model for chronic cuff tear and an induced tear (surgical tenotomy) in the rabbit RTC muscles results in fatty infiltration (60, 70, 82). Fatty infiltration after a RTC tear is a common clinical problem (26, 38) and desirable in an animal model. Substantial fatty infiltration does not occur in the rodent after a supraspinatus tear (18) unless both the muscle and its nerve (suprascapular nerve) are injured (36, 46, 47), whereas the rabbit mimics the human condition, with fatty infiltrate occurring after tenotomy alone (60, 70, 82). Despite this advantage of the rabbit as a model for fatty atrophy and ease of surgical interventions, there is a paucity of information regarding average strength of the rabbit supraspinatus muscle (Table 1).

In summary, this work describes methods to immobilize the scapula bone and assess supraspinatus muscle contractility in a variety of animal models. Even though contractile force of a healthy skeletal muscle can be roughly estimated based on its mass and architecture, this assumption only stands for healthy muscle, and makes it difficult to compare between species. The methods detailed here provide the investigator with normal values for the supraspinatus mass and contractility, as well as detailed techniques that can be used to compare uninjured, healthy supraspinatus masses and forces to those obtained after various interventions or to compare healthy, injured, and dystrophic muscles.

GRANTS

This work was supported by grants from the National Institutes of Health to A. P. Valencia (T32-AG-000268-15S1), to S. R. Iyer (AR-07592-20), and to R. M. Lovering (1R-01-AR-059179).

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the author(s).

AUTHOR CONTRIBUTIONS

Author contributions: A.P.V., S.J.P., M.N.G., and R.M.L. conception and design of research; A.P.V., S.J.P., M.N.G., and R.M.L. performed experiments; A.P.V. and R.M.L. analyzed data; A.P.V., S.J.P., M.N.G., and R.M.L. interpreted results of experiments; A.P.V. and R.M.L. prepared figures; A.P.V., S.R.I., and R.M.L. drafted manuscript; A.P.V., S.R.I., S.J.P., M.N.G., and R.M.L. edited and revised manuscript; A.P.V., S.R.I., S.J.P., M.N.G., and R.M.L. approved final version of manuscript.