Am J Physiol Cell Physiol 306: C889–C898, 2014. First published March 5, 2014; doi:10.1152/ajpcell.00383.2013.

CALL FOR PAPERS

Cellular Mechanisms of Tissue Fibrosis

Collagen content does not alter the passive mechanical properties of fibrotic skeletal muscle in mdx mice Lucas R. Smith and Elisabeth R. Barton Department of Anatomy and Cell Biology, School of Dental Medicine, Pennsylvania Muscle Institute, University of Pennsylvania, Philadelphia, Pennsylvania Submitted 17 December 2013; accepted in final form 4 March 2014

Smith LR, Barton ER. Collagen content does not alter the passive mechanical properties of fibrotic skeletal muscle in mdx mice. Am J Physiol Cell Physiol 306: C889 –C898, 2014. First published March 5, 2014; doi:10.1152/ajpcell.00383.2013.—Many skeletal muscle diseases are associated with progressive fibrosis leading to impaired muscle function. Collagen within the extracellular matrix is the primary structural protein providing a mechanical scaffold for cells within tissues. During fibrosis collagen not only increases in amount but also undergoes posttranslational changes that alter its organization that is thought to contribute to tissue stiffness. Little, however, is known about collagen organization in fibrotic muscle and its consequences for function. To investigate the relationship between collagen content and organization with muscle mechanical properties, we studied mdx mice, a model for Duchenne muscular dystrophy (DMD) that undergoes skeletal muscle fibrosis, and age-matched control mice. We determined collagen content both histologically, with picosirius red staining, and biochemically, with hydroxyproline quantification. Collagen content increased in the mdx soleus and diaphragm muscles, which was exacerbated by age in the diaphragm. Collagen packing density, a parameter of collagen organization, was determined using circularly polarized light microscopy of picosirius redstained sections. Extensor digitorum longus (EDL) and soleus muscle had proportionally less dense collagen in mdx muscle, while the diaphragm did not change packing density. The mdx muscles had compromised strength as expected, yet only the EDL had a significantly increased elastic stiffness. The EDL and diaphragm had increased dynamic stiffness and a change in relative viscosity. Unexpectedly, passive stiffness did not correlate with collagen content and only weakly correlated with collagen organization. We conclude that muscle fibrosis does not lead to increased passive stiffness and that collagen content is not predictive of muscle stiffness. fibrosis; collagen; skeletal muscle; passive mechanics WITHIN SKELETAL MUSCLE, THE extracellular matrix (ECM) plays a critical role in transmitting force generated by muscle cells in addition to its role as a cell scaffold in other tissues (25). Thus muscle pathologies that disrupt the ECM are likely to have profound functional consequences. Fibrosis is the pathologic accumulation of ECM components within a tissue leading to disrupted architecture and loss of function. Collagen is the major structural protein of skeletal muscle ECM, with fibrillar collagen types I and III predominately responsible for providing tensile strength (13). In skeletal muscle fibrosis, collagen is upregulated and occupies a larger proportion of muscle vol-

Address for reprint requests and other correspondence: E. R. Barton, Dept. of Anatomy and Cell Biology, School of Dental Medicine, Univ. of Pennsylvania, Philadelphia, PA (e-mail: [email protected]). http://www.ajpcell.org

ume. Skeletal muscle fibrosis is observed in response to a number of muscle pathologies including the muscular dystrophies. Duchenne muscular dystrophy (DMD), caused by the loss of dystrophin, is one of the most prevalent of the muscular dystrophies, with severe progressive fibrosis of skeletal muscle. The mdx mouse is a model of DMD and the most commonly studied model of dystrophic muscle (28). Many of the hallmarks of DMD are exhibited in this model, including heightened muscle fragility, reduced strength, and cumulative fibrosis. The extent of fibrosis in skeletal muscle is typically quantified using histological methods (picosirius red or trichrome staining) to define the fractional area of a muscle cross section that is occupied by the ECM (2, 3, 46). Biochemical quantification of collagen is often performed with a hydroxyproline assay to assess the degree of fibrosis within a whole muscle (43, 45). These methods provide quantitative data on the amount of collagen in skeletal muscle tissue; however, they do not provide information on how collagen is organized. Crosslinking is a measure of collagen organization that has been demonstrated to play a critical role in determining collagen stiffness (4). As collagen is cross-linked into larger and more tightly crowded fibrils, the density of collagen packing is increased. Direct measurement of collagen cross-linking is technically challenging albeit feasible in skeletal muscle (16, 21, 34). However, measurement of collagen packing is relatively high throughput and provides morphometric data in addition to the extent of collagen packing (10). Collagen packing can be measured by viewing picosirius red-stained tissue sections under circularly polarized light, under which densely packed collagen fibers are birefringent with a red hue, while loosely packed collagen fibers have a green hue (36, 39). Polarized light microscopy of picosirius red-stained sections has been used in a variety of tissues including cardiac muscle (40, 44, 50, 51), but to our knowledge, it has not been applied to skeletal muscle. Thus we sought to investigate how collagen packing is altered in a model of skeletal muscle fibrosis. Study of skeletal muscle fibrosis has focused on the upstream events involving muscle injury leading to inflammation and transforming growth factor-␤ expression, which induces activation of fibroblasts to secrete collagen and other ECM components (25, 28, 53). However, relatively little is known regarding how increased collagen leads to a disruption of skeletal muscle function. Certainly, replacement of contractile tissue with fibrotic scar tissue leads to a reduction in active force-generating capacity of muscle. It is often stated that fibrosis also leads to an increase in skeletal muscle passive

0363-6143/14 Copyright © 2014 the American Physiological Society

C889

C890

COLLAGEN CONTENT DOES NOT CAUSE STIFFNESS IN SKELETAL MUSCLE

stiffness; however, the link between collagen content and stiffness is yet to be resolved (25). Passive stiffness in mammalian skeletal muscle is thought to be determined largely by ECM collagen, although intracellular elements are also known to contribute (13, 14, 30). It is well established that skeletal muscle has viscoelastic passive mechanical properties, which are critical components of muscle function in vivo, yet the viscous and elastic components are not commonly studied (1, 27). The extensor digitorum longus (EDL) muscle of mdx mice has been shown to have increased viscoelastic stiffness and viscosity along with increased collagen content (19), yet other muscles are known to undergo unique pathologies in the mdx mouse. For instance, the EDL is marginally affected compared with the diaphragm, which has been shown to succumb to extensive fibrosis along with an associated increase in passive stiffness (42). We hypothesized that muscle elastic and viscous passive mechanical properties would be positively correlated with the collagen content. Furthermore, we hypothesized that the degree of densely packed collagen provides further correlation with passive mechanical properties of these muscles. Here, we test these hypotheses using both collagen area fraction and hydroxyproline assays to determine collagen content in addition to circularly polarized light microscopy to determine the density of collagen packing. These collagen parameters are associated with elastic and viscous mechanical properties of EDL, soleus, and diaphragm muscles of fibrotic mdx and control C57 mice. MATERIALS AND METHODS

The University of Pennsylvania Animal Care Committee approved the experiments in this study. Mouse strains included the mdx mouse, a model for DMD, and the C57 mouse, a wild-type control. Only male mice were utilized for experiments. Mice were anesthetized with ketamine/xylazine. Muscles were removed and placed in a bath of modified Ringer’s solution (119 mM NaCl, 4.74 mM KCl, 3.36 mM CaCl2, 1.18 mM KH2PO4, 1.18 mM MgSO4, 25 mM HEPES, and 2.75 mM glucose) gas-equilibrated with 95% O2-5% CO2. Silk suture 6 – 0 was tied as close as possible to the myotendinous junction with a minimal distance to a rigid metal chain to minimize series compliance of nonmuscle elements. In diaphragm muscle preparations, the suture was attached to the central tendon and rib. For EDL and soleus, left limb muscles were used for mechanics and histology, and right limb muscles were used for hydroxyproline assays. For the diaphragm, separate pieces were used for mechanics, histology, and hydroxyproline assays. Active muscle mechanics. Muscles were subjected to isolated mechanical measurements using a previously described apparatus (Aurora Scientific, Ontario, Canada) (5) and bathed in modified Ringer’s solution gas-equilibrated with 95% O2-5% CO2 and bath temperature maintained at 22°C (35). Optimum muscle length (Lo) was determined with iterative manual adjustments of length to achieve maximum twitch force with supramaximal stimulation (31). Muscle physiological cross-sectional area (PCSA) was determined using the following formula: PCSA ⫽ m ⁄ [Lo ⫻ (Lf ⁄ Lo) ⫻ ␳] where m is muscle mass (m), Lo is muscle length, Lf/Lo is the ratio of fiber length to optimal muscle length, and the density of muscle is ␳ ⫽ 1.06 g/cm3 (29). Lf/Lo was 0.45 for EDL, 0.69 for soleus, and 1.0 for diaphragm. Maximum isometric twitch was measured in the muscles after 20 s by maximum isometric tetanus during a 500-ms stimulation. The maximum twitch and tetanus were measured a total of three times

with 5-min intervals between tests, and the individual maximum value was used. Passive muscle mechanics. Following the active mechanical protocol, a passive mechanical protocol was conducted. For each strain value a cyclical strain of the given peak amplitude was performed at 1 Hz for 5 s to precondition the muscle. After preconditioning, the muscle underwent a ramp strain to the given strain at 1 Lo/s and held for 2 min to allow for stress relaxation. During this strain, the peak force was taken to represent the dynamic passive force, and the force at 2 min was taken to represent the elastic passive force. This procedure was executed at strains of 2.5, 5, 7.5, 10, 12.5, and 15% with a period of at least 20 s separating given strain procedures. During stress relaxation, the maximum stress reached is referred to as the dynamic stress (see Fig. 3A, squares) and the steady-state stress after 2 min is referred to as elastic stress (see Fig. 3A, squares). The elastic stiffness, or modulus, is determined from the tangent of the quadratic fit of elastic stress to strain at 10% strain, and the dynamic stiffness modulus is determined equivalently from dynamic stress (see Fig. 3C, tangents). The elastic and dynamic stiffness are normalized values of stiffness and referred to as stiffness (mN/mm2) throughout this article, while the raw value is referred to as the absolute stiffness (N/mm). Dynamic stiffness is purely a passive mechanical property without a contractile component that is a measure of the nonsteadystate stress response to an applied strain. The passive mechanical protocols are based on previous studies (19, 49, 54) and within a physiological range of strains and strain rates (9). Picosirius red staining. Picosirius red staining was performed similarly to previous studies (2, 3, 46). Samples were rinsed in phosphate-buffered saline, blotted, weighed, covered in mounting medium before freezing in melting isopentane, and stored at ⫺80°C. Cryosections of 10 ␮m were cut in cross section from the mid belly of the muscle at ⫺20°C (Leica CM3050 S). Sections were fixed in 4% paraformaldehyde for 10 min, rinsed, air dried, and stained for 1 h in 0.1% (wt/vol) sirius red (Sigma-Aldrich) dissolved in saturated aqueous picric acid (Sigma-Aldrich). Sections were then washed in two changes of 0.5% acetic acid, dehydrated in three changes of 100% ethanol, cleared with Citra Solv, and mounted with Cytoseal. A series of micrographs from each muscle were captured using a ⫻10 objective on a Leica DMLP (Leica) scope with a Micropublisher 5.0 (Q Imaging) camera to obtain an image of the entire cross-section, which was reconstituted by merging the images using Photoshop. A custom script was written in MATLAB (Mathworks) to determine number of pixels stained red and total number of pixels stained. To evaluate the collagen packing density, picosirius red-stained muscle sections were also viewed under circularly polarized light using the Leica DMLP (Leica) scope, rotating polarizer, rotating analyzer, and dual quarter wave plates. Whole muscle images were obtained as with light microscopy. A custom script was written in MATLAB to determine the number of pixels stained above an intensity threshold in bins determined by 8-bit hue values (245–255 and 0 –34 for red, 35– 49 for yellow, and 50 –115 for green; modified from Ref. 26). Hydroxyproline content. The hydroxyproline assay was performed as described in previous studies for muscle samples (11, 22). Muscles were rapidly frozen in liquid nitrogen and stored at ⫺80°C. The tissue was pulverized on dry ice with careful attention to remove any attached tendon. The pulverized muscle was weighed, and hydrolyzed overnight in 1 ml of 6 M hydrochloric acid at 105°C. Ten microliters of hydrolysate were mixed with 150 ␮l isopropanol followed by 75 ␮l of 1.4% chloramine-T (Sigma, St. Louis, MO) in citrate buffer and oxidized at room temperature for 10 min. The samples were then mixed with 1 ml of a 3:13 solution of Ehrlich’s reagent [1.5 g of 4-(dimethylamino) benzaldehyde (Sigma); 5 ml ethanol; 337 ␮l sulfuric acid] to isopropanol and incubated for 45 min at 55°C. Quantification was determined by extinction measurement of the resulting solution at 558 nm. A standard curve (0 –1,000 ␮M trans-4-hydroxy-

AJP-Cell Physiol • doi:10.1152/ajpcell.00383.2013 • www.ajpcell.org

C891

COLLAGEN CONTENT DOES NOT CAUSE STIFFNESS IN SKELETAL MUSCLE L-proline; Sigma) was included in each assay. Results are reported as micrograms of hydroxyproline per milligrams of tissue wet weight. Statistics. A two-way ANOVA was utilized for comparisons of muscle measurements with main effects of genotype and age for each muscle tested individually. Tukey’s multiple comparison tests were utilized to determine differences between groups. Pearson correlation was used to identify significant correlations between parameters within a muscle. Statistical significance was accepted for P ⬍ 0.05.

RESULTS

Body mass, muscle size, and active muscle force. To investigate the relationship between passive mechanical measures and measures of fibrosis, we compared young adult (20 wk) and aged adult (1 yr) fibrotic mdx and control C57 mice. Muscle mass of the EDL and soleus was significantly greater in the mdx mouse at both ages (Table 1). There was no corresponding change in Lo between mdx and C57 mice, leading to significantly greater PCSA in the mdx mice with greater muscle mass. There was also a significant increase in PCSA from 20 wk to 1 yr in mdx and C57 mice in both EDL and soleus. The maximum tetanic force generation of mdx compared with C57 was not different in EDL or soleus. However, as expected, specific force, which is maximum tetanic force normalized to PCSA, was compromised in mdx compared with C57 mice in all muscles with a significant genotype effect observed. There was also a significant decrease in specific force for 1-yr mdx mice compared with their 20-wk-old counterparts in the diaphragm muscle, with the same trend seen in EDL and soleus muscles. Collagen content. The collagen content measured as a fraction of total muscle area was determined using picosirius red

staining (Fig. 1, A and B). There was no statistical difference in collagen area fraction of EDL muscles between C57 and mdx mice at either age; however, the soleus and diaphragm had highly significant increases in collagen area (Fig. 1C). Collagen area had a significant main effect of age for both soleus and diaphragm and a significant interaction of age and genotype in the diaphragm. This led to post hoc tests with a significant increase in collagen area in the 1-yr mdx diaphragm compared with 20 wk of age. Biochemical determination of whole muscle collagen content with a hydroxyproline assay produced largely similar results to collagen area fraction. There was not a statistically significant increase of hydroxyproline in mdx EDL muscle, but the difference was significant in both soleus and diaphragm (Fig. 1D). Age did not have a significant effect on hydroxyproline content of EDL or soleus muscle, but age and interaction between age and genotype were significant factors for the diaphragm. Post hoc tests revealed the 1-yr-old mdx diaphragm had significantly greater hydroxyproline than at 20 wk. Collagen packing. The degree of collagen packing was determined by viewing the picosirius red-stained sections under circularly polarized light (Fig. 2, A and B). The results were first analyzed as the extent of collagen packing with respect to the fraction of collagen area. In both the EDL and the soleus muscles, a smaller fraction of collagen area was occupied by densely packed collagen in 1-yr-old mdx mice (Fig. 2, C and E). However, in the diaphragm, the proportions of densely packed collagen were not significantly different at either age or between C57 and mdx mice (Fig. 2G). A second set of comparisons was performed on collagen packing with respect

Table 1. Morphometric properties of experimental animals 20 wk

Age, wk Body weight, g EDL Samples Muscle mass, mg PCSA, mm2 Length, mm Tetanic force, mN Specific force, mN/mm2 Absolute elastic stiffness, N/mm Soleus Samples Muscle mass, mg PCSA, mm2 Length, mm Tetanic force, mN Specific force, mN/mm2 Absolute elastic stiffness, N/mm Diaphragm Samples Muscle mass, mg PCSA, mm2 Length, mm Tetanic force, mN Specific force, mN/mm2

1 yr

C57

mdx

C57

mdx

20 ⫾ 4 26.8 ⫾ 2.2

20 ⫾ 2 30.4 ⫾ 1.3*

53 ⫾ 3 33 ⫾ 4†

7(7) 10.6 ⫾ 0.6 1.73 ⫾ 0.11 12.8 ⫾ 0.4 351 ⫾ 35 204 ⫾ 21 3.6 ⫾ 0.57

7(7) 13.5 ⫾ 0.7* 2.13 ⫾ 0.09* 13.3 ⫾ 0.4 373 ⫾ 30 176 ⫾ 16* 10.17 ⫾ 2.1*

7(7) 11.9 ⫾ 0.7† 1.88 ⫾ 0.08† 13.3 ⫾ 0.3 365 ⫾ 42 196 ⫾ 29 4.26 ⫾ 1.59

8(7) 14.8 ⫾ 0.9*† 2.34 ⫾ 0.15*† 13.3 ⫾ 0.5 392 ⫾ 28 168 ⫾ 13* 11.61 ⫾ 3.79*

8(5) 9.4 ⫾ 0.7 1.08 ⫾ 0.08 11.8 ⫾ 0.4 184 ⫾ 23 170 ⫾ 19 4.29 ⫾ 0.98

7(4) 12.9 ⫾ 1.2* 1.51 ⫾ 0.08* 11.6 ⫾ 0.6 216 ⫾ 37 143 ⫾ 20 4.26 ⫾ 1.02

7(4) 10.7 ⫾ 1.5† 1.26 ⫾ 0.18† 11.7 ⫾ 0.6 198 ⫾ 23 158 ⫾ 15 2.75 ⫾ 18.9

8(4) 13.8 ⫾ 0.7* 1.74 ⫾ 0.19*† 10.9 ⫾ 0.8 194 ⫾ 48 114 ⫾ 38* 4.45 ⫾ 2.04

8(5) 4.1 ⫾ 0.5 0.47 ⫾ 0.05 8.3 ⫾ 1.1 66 ⫾ 10 141 ⫾ 17

7(4) 5.8 ⫾ 1.3 0.67 ⫾ 0.11 8.1 ⫾ 0.9 75 ⫾ 19 110 ⫾ 18*

7(4) 5.2 ⫾ 0.5 0.60 ⫾ 0.14 8.5 ⫾ 1.3 76 ⫾ 19 137 ⫾ 30

8(5) 6.3 ⫾ 1 0.74 ⫾ 0.15 8.1 ⫾ 0.5 50 ⫾ 17 67 ⫾ 14*†

55 ⫾ 4 31.5 ⫾ 2

Values are means ⫾ SD. Samples in parentheses underwent passive mechanical measurements. The diaphragm muscle mass, physiological cross-sectional area (PCSA), length, and tetanic force are dependent on muscle dissection, and thus statistical analysis was not performed for these parameters. EDL, extensor digitorum longus. *P ⬍ 0.05, mdx mice are significantly different from age-matched C57 mice; †P ⬍ 0.05, 1 yr mice are significantly different from 20 wk mice of the same genotype. AJP-Cell Physiol • doi:10.1152/ajpcell.00383.2013 • www.ajpcell.org

C892

COLLAGEN CONTENT DOES NOT CAUSE STIFFNESS IN SKELETAL MUSCLE

Fig. 1. Characterization of the collagen content of extensor digitorum longus (EDL), soleus, and diaphragm muscles in mdx and C57 mice at 20 wk and 1 yr. Representative examples of whole muscle picosirius red staining from soleus muscle of 1-yr-old (A) C57 and (B) mdx mice. Black scale bar ⫽ 100 ␮m. C: collagen area fraction as determined from picosirius red straining of whole muscle cross section after mechanical testing. D: hydroxyproline content as a measure of overall collagen content in whole skeletal muscle from contralateral limb of mechanics testing. *P ⬍ 0.05, significant differences between mdx and aged-matched C57 mice. †P ⬍ 0.05, significant differences between 1-yr and 20-wk-old mice of the same genotype.

to the fraction of muscle area. The difference in densely packed collagen of EDL muscles at 1 yr was not present when taken as a fraction of total muscle area; however, there was a significant increase in loosely packed collagen in EDL muscles from 1-yr-old mdx mice (Fig. 2D). The same was true of soleus

muscle with both ages having significantly higher proportions of loose collagen in mdx mice (Fig. 2F). As a consequence of having much greater collagen area, the mdx diaphragm had a significant effect of genotype with increases in both densely and loosely packed collagen at 20 wk and 1 yr of age of mdx

Fig. 2. Characterization of collagen packing of EDL, soleus, and diaphragm muscles in mdx and C57 mice at 20 wk and 1 yr. The same representative examples of whole muscle picosirius red staining as viewed through circularly polarized light microscopy from soleus muscle of 1-yr-old C57 (A) and mdx (B) mice from Fig. 1. White scale bar ⫽ 100 ␮m. C–H: red represents densely packed collagen, yellow represents intermediately packed collagen, and green represents loosely packed collagen. C, E, and G: collagen packing as a fraction of collagen staining for EDL, soleus, and diaphragm, respectively. D, F, and H: collagen packing as a fraction of total muscle area. *P ⬍ 0.05, significant differences between mdx and aged-matched C57 mice. Comparisons were not made on the degree of intermediate collagen packing. AJP-Cell Physiol • doi:10.1152/ajpcell.00383.2013 • www.ajpcell.org

COLLAGEN CONTENT DOES NOT CAUSE STIFFNESS IN SKELETAL MUSCLE

C893

ANOVA revealed a significant difference in dynamic stiffness of diaphragm muscle in mdx mice; however, post hoc tests were not significant for either age. The ratio of the elastic stiffness to the dynamic stiffness provides an index of viscosity, with a larger value representing the elastic stress being a greater proportion of the total stress and thus less relative viscosity. The mdx EDL muscles had a significantly higher ratio becoming proportionally less viscous, while there was no change in the soleus muscle (Fig. 4C). The mdx muscles of the diaphragm had the opposite effect of EDL muscles and were significantly more viscous than controls. Correlations. Measuring fibrotic and functional parameters on the same muscle, or contralateral muscles in the case of hydroxyproline assays, permitted correlation between each

Fig. 3. Description of passive mechanical protocol with example data from 20-wk-old C57 EDL muscle. A: after preconditioning stress relaxation is held for 2 min with () representing maximum stress used to determine dynamic stiffness () representing stress at 2 min used to determine elastic stiffness. B: strains of 2.5, 5, 7.5, 10, and 15% with a ramp stretch of 1 fiber length/second and held for 2 min. C: stiffness is determined from a quadratic fit to the stress strain curves taking the tangent modulus at 10% strain for both dynamic and elastic stiffness.

mice. There was a significant interaction of age and genotype of densely packed collagen in the diaphragm; however, post hoc tests did not reveal a significant change with age for either genotype (Fig. 2H). Thus all muscles from the mdx mice displayed increases in loosely packed collagen, yet there was no consistent trend in tightly packed collagen across the muscle groups. Passive mechanics. The dynamic and elastic stiffness as well as an index of viscosity were determined for each muscle group (Fig. 3). The elastic stiffness was much greater for mdx muscles in the EDL at both ages (Fig. 4A). Surprisingly, there was no statistical difference in elastic stiffness of mdx muscles in either the soleus or diaphragm muscles. The result for dynamic stiffness, which includes a viscous component, was similar with the EDL having a highly significant increase in mdx muscles with no change in soleus muscles. A two-way

Fig. 4. Characterization of passive mechanical properties of EDL, soleus, and diaphragm muscles in mdx and C57 mice at 20 wk and 1 yr. Dynamic stiffness (A) and elastic stiffness (B) values. C: ratio of elastic to dynamic stiffness, with higher values representing more proportionally elastic to viscous stiffness. *P ⬍ 0.05, significant differences between mdx and aged-matched C57 mice.

AJP-Cell Physiol • doi:10.1152/ajpcell.00383.2013 • www.ajpcell.org

C894

COLLAGEN CONTENT DOES NOT CAUSE STIFFNESS IN SKELETAL MUSCLE

parameter. As expected, there was a negative correlation between collagen area and specific force for each muscle, which was highly significant for soleus and diaphragm muscles (Fig. 5, A–C). Surprisingly, the correlation between collagen area fraction and elastic stiffness was minimal for each muscle tested (Fig. 5, D–F). Correspondingly there was no significant correlation between hydroxyproline content and elastic stiffness for any muscle tested (R2 ⫽ 0.124, P ⫽ 0.061 for EDL; R2 ⫽ 0.049, P ⫽ 0.393 for soleus; and R2 ⫽ 0.012, P ⫽ 0.661 for diaphragm). Further, there was no significant correlation across groups for dense collagen muscle fraction and elastic stiffness (Fig. 5, G–I). For the EDL, however, there was a significant positive correlation between dense collagen muscle fraction and elastic stiffness for 20-wk C57 and 1-yr mdx mice. DISCUSSION

Fibrosis is an occurrence common to nearly all muscle pathologies (28). Skeletal muscles in patients with DMD have extensive replacement of muscle fibers with fibrotic and fatty tissue (23), which leads to loss of muscle strength, impaired regenerative capacity, and potentially increased stiffness. In addition to muscle weakness, patients with DMD also commonly develop muscle contractures, in which muscle passive stiffness can contribute to the loss of range of motion about a joint, leading to further loss of function. It is commonly suggested that an increase in collagen of the fibrotic muscle underlies increased passive stiffness (25). We tested the hy-

pothesis that collagen content and active and passive mechanical properties were well correlated in the mdx mouse model of DMD. While there was a strong correlation between collagen content and active force generation, we found little to no correlation between collagen content, measured histologically or biochemically, and passive muscle stiffness. Further, using circularly polarized light microscopy to obtain an index of collagen packing yielded only a minor correlation between densely packed collagen and passive stiffness. Finally, we found that collagen organization and passive mechanical properties of different muscles respond to the dystrophic environment in unique ways. It is generally accepted that the mdx mouse recapitulates only a subset of the symptoms associated with DMD, with limb muscles undergoing successful regeneration and a mild disease phenotype (8). In contrast, the diaphragm has been shown to undergo severe fibrosis in the mdx model, more characteristic of the human disease (42). Our results on collagen content reflect these concepts with the limb muscles (EDL and soleus) exhibiting moderate fibrosis in terms of collagen area fraction and biochemical collagen content, and the diaphragm having extensive fibrosis (19, 42). The accumulation of fibrosis is known to be progressive in DMD and the mdx model, corresponding with our results showing increased collagen from 20 wk to 1 yr in the mdx mice, with little to no change during the same time span of C57 mice. The fact that both collagen area fraction and hydroxyproline quantification yield very similar

Fig. 5. Correlation of fibrotic and mechanical parameters of EDL, soleus, and diaphragm muscles in mdx and C57 mice at 20 wk and 1 yr. A–C: correlation between specific force and collagen area fraction for EDL, soleus, and diaphragm muscles respectively. D–F: correlation between elastic stiffness and collagen area fraction EDL, soleus, and diaphragm muscles respectively. G–I: correlation between elastic stiffness and fraction of whole muscle dense collagen area. G: significant (P ⬍ 0.05) positive correlation exists between elastic stiffness and dense collagen area within 20-wk-old C57 and 1-yr mdx mice. ***P ⬍ 0.001, significant correlation within a muscle between parameters across genotype and age of mice. AJP-Cell Physiol • doi:10.1152/ajpcell.00383.2013 • www.ajpcell.org

COLLAGEN CONTENT DOES NOT CAUSE STIFFNESS IN SKELETAL MUSCLE

results argues that either is a useful assay of collagen content. The EDL muscle had trends for more collagen using both assays, but neither reached significance. Previous studies have shown significantly more collagen in the mdx EDL at similar ages (17–19), but it is possible our study was underpowered to detect these smaller differences. As a whole results on collagen content of our study correspond well with the existing literature. Measuring collagen packing using circularly polarized light on picosirius red-stained sections is a novel method in skeletal muscle, although it has been used successfully in other tissues (33, 36, 47). Collagen organization, including cross-linking, is known to have important effects on its material properties (4); however, collagen organization has not been measured in DMD patients or mdx mice. When performed alongside typical picosirius red staining for collagen area fraction, quantification of collagen packing represents a complementary method of collecting further data on the state of the ECM and collagen organization. While collagen packing provides only one aspect of collagen organization, these alterations serve as an index of the fibrotic state. In the current study, we observed a range of the quantity and quality of fibrosis in the different muscles from wild-type and mdx mice. Limb muscles from aged mdx mice exhibited modest but significant decreases in the proportion of densely packed collagen and an increase in loosely packed collagen. Thus limb muscle fibrosis appears as a looser, more disorganized, ECM. However, the diaphragm exhibited a different trend including an increase in both the loosely and tightly packed collagen in the diaphragms from mdx mice, which was evident when collagen packing was viewed as a percentage of total muscle area. Given that collagen production precedes collagen cross-linking, we anticipate that the proportion of tightly packed collagen would increase further in diaphragms of older mdx mice. However, it is surprising that the limb muscles show a proportional decrease in tightly packed collagen with age. This may be due to variable expression and activity of cross-linking enzymes across different muscles, variable expression and activity of collagenases, or simply that we have captured only a snapshot of a common progression of fibrosis that warrants further study. Taken together, the limb muscles and the diaphragm represent a spectrum of collagen content and organization that can be investigated to evaluate how fibrosis and mechanical properties are related. While not as commonly studied as active mechanics, the passive mechanical properties of mdx mouse muscle have been debated in the literature. These studies are difficult to compare as the mechanical protocols are not standardized and the tests are performed on different muscles. For the EDL muscle early studies showed a trend for increased passive stiffness (6, 24). Later studies suggested that limb muscle was not stiffer when undergoing small strains (7, 52). Previous studies have also demonstrated that the passive mechanical properties of EDL are not altered up to 7 wk of age (52). More recently, higher powered studies with a similar stretch relaxation protocol that used here have more conclusively established an increase in passive stiffness of EDL muscle down to 10% strain (17–19), a physiological level that does not produce appreciable muscle damage. Although our analysis was dissimilar, using the tangent modulus for stress stain, the data are consistent with Hakim et. al. (17–19) for EDL muscle. Hakim et. al. also

C895

measured the viscoelastic properties of EDL muscle using stress relaxation, and reported an increase in viscosity of mdx muscle. Our data describe a decrease in the viscosity for EDL muscle, but it is important to recognize that we looked at viscosity in proportion to the viscoelastic (dynamic) stiffness while Hakim et al. looked at the absolute viscosity. In both cases, the relative change in dynamic stress was greater than the change in viscosity. In the diaphragm, Stedman et al. (42) first demonstrated a dramatic increase in passive tension of mdx diaphragm at 18 mo with only 7.5% strain. However, Kumar et. al. showed increased compliance of the diaphragm in mdx mice with a much younger 2-wk-old time point (23a). Our results fall between these studies with no effects on elastic stiffness and a small increase in dynamic stiffness at both 20 wk and 1 yr of the mdx diaphragm. These discrepancies could be due to progressive stiffening that occurs over the lifetime of the mouse, where each study reflects a different time point, or it is possible that the variations among passive mechanical protocols produce different results. The 22°C temperature used in these experiments provides a reference to examine the relationship between collagen and passive tension. However, this is below the physiological temperature of limb muscles and to a greater degree in the diaphragm, which may lead to altered passive mechanical properties in vivo. Whether or not the absence of dystrophin alters the response of passive mechanical properties to temperature is an open question. Based on previous work assessing the active mechanical properties of C57 and mdx diaphragm muscle, which are altered in a constant ratio through a range of temperatures (32), we anticipate that a similar relationship among passive mechanical properties, fibrosis, and temperature would emerge. However, this remains to be tested in future studies. Notably, we are not aware of any studies on the passive mechanical properties of the soleus muscle in mdx and thus the first to report no effect in the mdx mice. It is clear that dystrophic patients demonstrate decreased active force generation in conjunction with the replacement of contractile muscle with fibrotic tissue, as seen with a clear correlation between collagen area fraction and specific force. The common assertion is that increased fibrosis also leads to increased passive stiffness; however, the relationship is unclear. A multitude of studies show increases in fibrosis in skeletal muscle pathology and also an increase in passive stiffness (19, 41) but are rarely paired together to determine if there is a correlation. The structures responsible for passive tension in skeletal muscle are also not entirely elucidated. Recent studies have suggested that the ECM is primarily responsible for passive stiffness in mammalian skeletal muscle (13, 14, 30); however, intracellular components may also contribute (37). Our results clearly demonstrate that collagen content is not the primary determinant of passive stiffness, with no correlation between collagen content and dynamic or elastic stiffness. This is made readily apparent by the mdx EDL having little change in collagen and a highly significant increase in passive stiffness, while the mdx soleus has a highly significant increase in collagen and no change in passive stiffness. We are thus able to reject our hypothesis that collagen content is correlated with passive stiffness in fibrotic skeletal muscle. Clearly, parameters beyond collagen content are contributing to passive stiffness. We tested the hypothesis that the amount of dense collagen fibril packing would more readily

AJP-Cell Physiol • doi:10.1152/ajpcell.00383.2013 • www.ajpcell.org

C896

COLLAGEN CONTENT DOES NOT CAUSE STIFFNESS IN SKELETAL MUSCLE

account for passive stiffness. Dense collagen packing did not correlate significantly with passive stiffness across the range of muscle groups for any muscle. When investigating with individual groups, there was predominantly a positive correlation, but it was only significant in the EDL for 20-wk-old C57 mice and 1-yr-old mdx mice. With the use of z-scores to remove effects of muscle, genotype, or age, there is a small but significant correlation between dense collagen and elastic stiffness, which is not true for collagen area or hydroxyproline (Fig. 6, A–C). While the role collagen packing plays in establishing passive stiffness is yet to be conclusively determined, there are clearly other important factors. This study does not rule out the importance of collagen in passive stiffness of skeletal muscle. The extent of cross-linking is known to be an important determinant of collagen tensile strength (4) and thus may be key to understanding fibrosis in skeletal muscle. The orientation of collagen fibrils is also critical to determining their function (38). Disorganized ECM with collagen aligned orthogonal to the muscle axis is a possible explanation for the decreased stiffness of mdx soleus muscle. This study focuses on the contribution of collagen to passive tension; however, intracellular components also affect passive mechanical properties of skeletal muscle. Titin is purported to be the primary contributor to intracellular skeletal muscle passive mechanics, with mechanical stiffness dependent on alternative splicing (48). However, there is evidence that extracellular components bear most of the passive load in whole mammalian muscle,

particularly at 10% strain (13, 15). It is currently unknown how titin splice variants are affected in mdx skeletal muscle and how titin alteration might affect mdx muscle stiffness. Certainly, more work is required to elucidate the parameters that define the passive mechanical properties of skeletal muscle in healthy and diseased states. The results of this study also demonstrate that muscles respond to the lack of dystrophin in unique ways, as has been shown previously (20). This is perhaps most clearly demonstrated by the changes in the ratio of elastic to dynamic stiffness, a measure of the relative viscosity of the tissue. Since the ratio is elastic to dynamic stiffness is internally normalized and not dependent on measurements of mass and length, there is comparatively little variation within a muscle group. Our results clearly show the EDL to become more elastic, the soleus to be unchanged, and the diaphragm to be become more viscous. The reason for this to be the case is unknown. It is interesting to point out, however, that across conditions there is little correlation between viscosity and muscle fraction of dense collagen. However, when the z-scores were used to remove the effects of muscle, genotype, and age, there is a highly significant correlation between dense collagen area and ratio of elastic to dynamic stiffness (Fig. 6, D–F). This suggests that, in the absence of other factors, dense collagen packing leads to relatively more elastic muscle properties while loose disorganized collagen packing leads to relatively more viscous mechanical properties. It is important to note that the

Fig. 6. Correlation of fibrotic parameters with elastic stiffness using z-scores (A–C) and proportion of elastic stiffness (D–F). The z-score is determined within each muscle, age, and genotype. A: correlation between elastic stiffness and collagen area fraction. B: correlation between elastic stiffness and hydroxyproline. C: correlation between elastic stiffness and fraction of whole muscle dense collagen area. D: correlation between ratio of elastic to dynamic stiffness and collagen area fraction. E: correlation between ratio of elastic to dynamic stiffness and hydroxyproline. F: correlation between ratio of elastic to dynamic stiffness and fraction of whole muscle dense collagen area. *P ⬍ 0.05, significant positive correlation exists with dense collagen area and both elastic stiffness and ratio of elastic to dynamic stiffness. AJP-Cell Physiol • doi:10.1152/ajpcell.00383.2013 • www.ajpcell.org

COLLAGEN CONTENT DOES NOT CAUSE STIFFNESS IN SKELETAL MUSCLE

increased intrinsic muscle elasticity in conjunction with the increased muscle size of the mdx EDL compound to create much larger total muscle stiffness that is experienced by the joint in vivo (Table 1), which could lead to contractures observed in mdx mice or increased susceptibility to damage (12). The muscle-specific adaptations that take place in the degree of viscosity and the alterations in stiffness, collagen content, and collagen organization may be protective in the environment of the muscle. For example, the passive mechanical properties of the diaphragm are key components of respiration (42), and a decrease in the elastic to dynamic stiffness ratio in the diaphragm would allow greater tuning of stiffness to respiratory rate. From a broader perspective, this type of assessment can reveal differential responses of passive mechanical properties not only to the loss of dystrophin, but it can also illuminate mechanisms underlying the loss of other proteins causing neuromuscular diseases. For instance, a recent study targeting COL12A1 in mice causing the loss of collagen XII resulted in significantly reduced passive tension and greater protection against loss of force following eccentric contractions (54), supporting the hypothesis that alterations in ECM properties can affect muscle fragility. In the current study, we could not address a potential correlation between passive mechanical properties and susceptibility to contractile injury due to the impact of eccentric contractions on the histological measures of collagen packing. However, future studies can be designed to determine how passive force and muscle fragility are related. Summary. In comparison to C57 control mice, we determined in 20-wk and 1-yr-old mdx mice that changes in collagen content varied between muscles, with little increase in EDL, a significant increase in soleus, and a dramatic increase in diaphragm muscle collagen content. The EDL and soleus muscles had a decreased proportion of densely packed collagen, while all dystrophic muscles had an increased total amount of loosely packed collagen. Only the EDL had significantly greater elastic stiffness in the mdx muscle than control. Both EDL and diaphragm had increased dynamic stiffness; however, the EDL became relatively more elastic and the diaphragm became relatively more viscous in the mdx mice. Surprisingly there was no correlation between collagen content and passive stiffness for any muscle. The density of collagen packing was also not able to account for much of the variability in passive stiffness. These results show that the fibrotic response to dystrophy and resulting passive mechanics are distinct to individual muscles and demonstrate that the contributors to passive mechanics warrant further investigation. ACKNOWLEDGMENTS We thank Dr. Min Liu and Zuozhen Tian from the Physiological Assessment Core at the University of Pennsylvania, Mike Hast and Dr. Louis Soslowsky for establishing circularly polarized light microscopy at the Penn Center for Musculoskeletal Disorders, and Dr. Ahlke Heydemann for assistance in developing the hydroxyproline assay for our use. GRANTS This work was supported by National Institute of Arthritis and Musculoskeletal and Skin Diseases Grants AR-057363 and AR-052646 (to E. R. Barton) and Training Grant AR-053461 (to L. R. Smith). Polarized light microscopy was supported by National Institute of Arthritis and Musculoskeletal and Skin Diseases Center Grant AR-050950 to the Penn Center for Musculoskeletal Disorders.

C897

DISCLOSURES No conflicts of interest, financial or otherwise, are declared by the authors. AUTHOR CONTRIBUTIONS Author contributions: L.R.S. and E.R.B. conception and design of research; L.R.S. performed experiments; L.R.S. analyzed data; L.R.S. and E.R.B. interpreted results of experiments; L.R.S. prepared figures; L.R.S. drafted manuscript; L.R.S. and E.R.B. edited and revised manuscript; L.R.S. and E.R.B. approved final version of manuscript. REFERENCES 1. Abbott BC, Lowy J. Stress relaxation in muscle. Proc R Soc Lond B Biol Sci 146: 281–288, 1956. 2. Acuna MJ, Pessina P, Olguin H, Cabrera D, Vio CP, Bader M, Munoz-Canoves P, Santos RA, Cabello-Verrugio C, Brandan E. Restoration of muscle strength in dystrophic muscle by angiotensin-1–7 through inhibition of TGF-beta signalling. Hum Mol Genet 25: 1237– 1249, 2014. 3. Ardite E, Perdiguero E, Vidal B, Gutarra S, Serrano AL, MunozCanoves P. PAI-1-regulated miR-21 defines a novel age-associated fibrogenic pathway in muscular dystrophy. J Cell Biol 196: 163–175, 2012. 4. Avery NC, Bailey AJ. Enzymic and non-enzymic cross-linking mechanisms in relation to turnover of collagen: relevance to aging and exercise. Scand J Med Sci Sports 15: 231–240, 2005. 5. Barton ER, Morris L, Kawana M, Bish LT, Toursel T. Systemic administration of L-arginine benefits mdx skeletal muscle function. Muscle Nerve 32: 751–760, 2005. 6. Berquin A, Schmit P, Moens P, Lebacq J. Compliance of normal, dystrophic and transplanted mouse muscles. J Biomech 27: 1331–1337, 1994. 7. Bobet J, Mooney RF, Gordon T. Force and stiffness of old dystrophic (mdx) mouse skeletal muscles. Muscle Nerve 21: 536 –539, 1998. 8. Bulfield G, Siller WG, Wight PA, Moore KJ. X chromosome-linked muscular dystrophy (mdx) in the mouse. Proc Natl Acad Sci USA 81: 1189 –1192, 1984. 9. Burkholder TJ, Lieber RL. Sarcomere length operating range of vertebrate muscles during movement. J Exp Biol 204: 1529 –1536, 2001. 10. de Jong S, van Veen TA, de Bakker JM, van Rijen HV. Monitoring cardiac fibrosis: a technical challenge. Neth Heart J 20: 44 –48, 2012. 11. Flesch M, Schiffer F, Zolk O, Pinto Y, Rosenkranz S, Hirth-Dietrich C, Arnold G, Paul M, Bohm M. Contractile systolic and diastolic dysfunction in renin-induced hypertensive cardiomyopathy. Hypertension 30: 383–391, 1997. 12. Garlich MW, Baltgalvis KA, Call JA, Dorsey LL, Lowe DA. Plantarflexion contracture in the mdx mouse. Am J Phys Med Rehabil 89: 976 –985, 2010. 13. Gillies AR, Lieber RL. Structure and function of the skeletal muscle extracellular matrix. Muscle Nerve 44: 318 –331, 2011. 14. Gillies AR, Smith LR, Lieber RL, Varghese S. Method for decellularizing skeletal muscle without detergents or proteolytic enzymes. Tissue Eng Part C Methods 17: 383–389, 2011. 15. Gindre J, Takaza M, Moerman KM, Simms CK. A structural model of passive skeletal muscle shows two reinforcement processes in resisting deformation. J Mech Behav Biomed Mater 22: 84 –94, 2013. 16. Gosselin LE, Martinez DA, Vailas AC, Sieck GC. Passive length-force properties of senescent diaphragm: relationship with collagen characteristics. J Appl Physiol 76: 2680 –2685, 1994. 17. Hakim CH, Duan D. Gender differences in contractile and passive properties of mdx extensor digitorum longus muscle. Muscle Nerve 45: 250 –256, 2012. 18. Hakim CH, Duan D. Truncated dystrophins reduce muscle stiffness in the extensor digitorum longus muscle of mdx mice. J Appl Physiol 114: 482–489, 2013. 19. Hakim CH, Grange RW, Duan D. The passive mechanical properties of the extensor digitorum longus muscle are compromised in 2- to 20-mo-old mdx mice. J Appl Physiol 110: 1656 –1663, 2011. 20. Haslett JN, Kang PB, Han M, Kho AT, Sanoudou D, Volinski JM, Beggs AH, Kohane IS, Kunkel LM. The influence of muscle type and dystrophin deficiency on murine expression profiles. Mamm Genome 16: 739 –748, 2005. 21. Haus JM, Carrithers JA, Trappe SW, Trappe TA. Collagen, crosslinking, and advanced glycation end products in aging human skeletal muscle. J Appl Physiol 103: 2068 –2076, 2007.

AJP-Cell Physiol • doi:10.1152/ajpcell.00383.2013 • www.ajpcell.org

C898

COLLAGEN CONTENT DOES NOT CAUSE STIFFNESS IN SKELETAL MUSCLE

22. Heydemann A, Huber JM, Demonbreun A, Hadhazy M, McNally EM. Genetic background influences muscular dystrophy. Neuromuscul Disord 15: 601–609, 2005. 23. Klingler W, Jurkat-Rott K, Lehmann-Horn F, Schleip R. The role of fibrosis in Duchenne muscular dystrophy. Acta Myol 31: 184 –195, 2012. 23a.Kumar A, Khandelwal N, Malya R, Reid MB, Boriek AM. Loss of dystrophin causes aberrant mechanotransduction in skeletal muscle fibers. FASEB J 18: 102–113, 2004. 24. Law DJ, Caputo A, Tidball JG. Site and mechanics of failure in normal and dystrophin-deficient skeletal muscle. Muscle Nerve 18: 216 –223, 1995. 25. Lieber RL, Ward SR. Cellular mechanisms of tissue fibrosis. 4. Structural and functional consequences of skeletal muscle fibrosis. Am J Physiol Cell Physiol 305: C241–C252, 2013. 26. MacKenna DA, Omens JH, McCulloch AD, Covell JW. Contribution of collagen matrix to passive left ventricular mechanics in isolated rat hearts. Am J Physiol Heart Circ Physiol 266: H1007–H1018, 1994. 27. Magnusson SP. Passive properties of human skeletal muscle during stretch maneuvers. A review. Scand J Med Sci Sports 8: 65–77, 1998. 28. Mann CJ, Perdiguero E, Kharraz Y, Aguilar S, Pessina P, Serrano AL, Munoz-Canoves P. Aberrant repair and fibrosis development in skeletal muscle. Skelet Muscle 1: 21, 2011. 29. Mendez J, Keys A. Density and composition of mammalian muscle. Metabolism 9: 184 –188, 1960. 30. Meyer GA, Lieber RL. Elucidation of extracellular matrix mechanics from muscle fibers and fiber bundles. J Biomech 44: 771–773, 2011. 31. Moorwood C, Liu M, Tian Z, Barton ER. Isometric and eccentric force generation assessment of skeletal muscles isolated from murine models of muscular dystrophies. J Vis Exp 71: e50036, 2013. 32. Murray JD, Canan BD, Martin CD, Stangland JE, Rastogi N, RafaelFortney JA, Janssen PM. The force-temperature relationship in healthy and dystrophic mouse diaphragm; implications for translational study design. Front Physiol 3: 422, 2012. 33. Nyska A, Dayan D. Ameloblastic fibroma in a young cat. J Oral Pathol Med 24: 233–236, 1995. 34. Palokangas H, Kovanen V, Duncan A, Robins SP. Age-related changes in the concentration of hydroxypyridinium crosslinks in functionally different skeletal muscles. Matrix 12: 291–296, 1992. 35. Petrof BJ, Stedman HH, Shrager JB, Eby J, Sweeney HL, Kelly AM. Adaptations in myosin heavy chain expression and contractile function in dystrophic mouse diaphragm. Am J Physiol Cell Physiol 265: C834 – C841, 1993. 36. Pierard GE. Sirius red polarization method is useful to visualize the organization of connective tissues but not the molecular composition of their fibrous polymers. Matrix 9: 68 –71, 1989. 37. Prado LG, Makarenko I, Andresen C, Kruger M, Opitz CA, Linke WA. Isoform diversity of giant proteins in relation to passive and active contractile properties of rabbit skeletal muscles. J Gen Physiol 126: 461–480, 2005. 38. Provenzano PP, Vanderby R Jr. Collagen fibril morphology and organization: implications for force transmission in ligament and tendon. Matrix Biol 25: 71–84, 2006. 39. Rich L, Whittaker P. Collagen and picosirius red staining: a polarized light assessment of fibrillar hue and spatial distribution. Braz J Morphol Sci 22: 97–104, 2005.

40. Shirani J, Pick R, Roberts WC, Maron BJ. Morphology and significance of the left ventricular collagen network in young patients with hypertrophic cardiomyopathy and sudden cardiac death. J Am Coll Cardiol 35: 36 –44, 2000. 41. Smith LR, Lee KS, Ward SR, Chambers HG, Lieber RL. Hamstring contractures in children with spastic cerebral palsy result from a stiffer extracellular matrix and increased in vivo sarcomere length. J Physiol 589: 2625–2639, 2011. 42. Stedman HH, Sweeney HL, Shrager JB, Maguire HC, Panettieri RA, Petrof B, Narusawa M, Leferovich JM, Sladky JT, Kelly AM. The mdx mouse diaphragm reproduces the degenerative changes of Duchenne muscular dystrophy. Nature 352: 536 –539, 1991. 43. Swaggart KA, Heydemann A, Palmer AA, McNally EM. Distinct genetic regions modify specific muscle groups in muscular dystrophy. Physiol Genomics 43: 24 –31, 2011. 44. Tabel GM, Whittaker P, Vlachonassios K, Sonawala M, Chandraratna PA. Collagen fiber morphology determines echogenicity of myocardial scar: implications for image interpretation. Echocardiography 23: 103–107, 2006. 45. Tirrell TF, Cook MS, Carr JA, Lin E, Ward SR, Lieber RL. Human skeletal muscle biochemical diversity. J Exp Biol 215: 2551–2559, 2012. 46. Trensz F, Haroun S, Cloutier A, Richter MV, Grenier G. A muscle resident cell population promotes fibrosis in hindlimb skeletal muscles of mdx mice through the Wnt canonical pathway. Am J Physiol Cell Physiol 299: C939 –C947, 2010. 47. Vij R, Vij H, Rao NN. Evaluation of collagen in connective tissue walls of odontogenic cysts–a histochemical study. J Oral Pathol Med 40: 257–262, 2011. 48. Wang K, McCarter R, Wright J, Beverly J, Ramirez-Mitchell R. Regulation of skeletal muscle stiffness and elasticity by titin isoforms: a test of the segmental extension model of resting tension. Proc Natl Acad Sci USA 88: 7101–7105, 1991. 49. Ward SR, Tomiya A, Regev GJ, Thacker BE, Benzl RC, Kim CW, Lieber RL. Passive mechanical properties of the lumbar multifidus muscle support its role as a stabilizer. J Biomech 42: 1384 –1389, 2009. 50. Whittaker P, Boughner DR, Kloner RA. Analysis of healing after myocardial infarction using polarized light microscopy. Am J Pathol 134: 879 –893, 1989. 51. Whittaker P, Kloner RA, Boughner DR, Pickering JG. Quantitative assessment of myocardial collagen with picrosirius red staining and circularly polarized light. Basic Res Cardiol 89: 397–410, 1994. 52. Wolff AV, Niday AK, Voelker KA, Call JA, Evans NP, Granata KP, Grange RW. Passive mechanical properties of maturing extensor digitorum longus are not affected by lack of dystrophin. Muscle Nerve 34: 304 –312, 2006. 53. Zhou L, Lu H. Targeting fibrosis in Duchenne muscular dystrophy. J Neuropathol Exp Neurol 69: 771–776, 2010. 54. Zou Y, Zwolanek D, Izu Y, Gandhy S, Schreiber G, Brockmann K, Devoto M, Tian Z, Hu Y, Veit G, Meier M, Stetefeld J, Hicks D, Straub V, Voermans NC, Birk DE, Barton ER, Koch M, Bonnemann CG. Recessive and dominant mutations in COL12A1 cause a novel EDS/myopathy overlap syndrome in humans and mice. Hum Mol Genet 2014 Jan 20 [Epub ahead of print].

AJP-Cell Physiol • doi:10.1152/ajpcell.00383.2013 • www.ajpcell.org

Copyright of American Journal of Physiology: Cell Physiology is the property of American Physiological Society and its content may not be copied or emailed to multiple sites or posted to a listserv without the copyright holder's express written permission. However, users may print, download, or email articles for individual use.