Histological Evidence of Muscle Degeneration in Advanced Human Rotator Cuff Disease

Gibbons, Michael C. MS; Singh, Anshu MD; Anakwenze, Oke MD; Cheng, Timothy MD; Pomerantz, Maxwill BS; Schenk, Simon PhD; Engler, Adam J. PhD; Ward, Samuel R. PT, PhD

Journal of Bone & Joint Surgery - American Volume: 1 February 2017 - Volume 99 - Issue 3 - p 190–199
doi: 10.2106/JBJS.16.00335
Scientific Articles

Background: Cellular remodeling in rotator cuff muscles following a massive rotator cuff tear is poorly understood. The aim of the current study was to provide histological evidence to elucidate the mode of muscle loss in advanced human rotator cuff disease and to assess tissue-level changes in relation to findings on noninvasive imaging.

Methods: Rotator cuff muscle biopsy samples were taken from the scapular fossae from 23 consecutive patients undergoing reverse total shoulder arthroplasty in order to evaluate muscle composition in severe rotator cuff disease. Markers of vascularity; inflammation; fat distribution; and muscle atrophy, degeneration, and regeneration were quantified.

Results: The samples primarily consisted of dense, organized connective tissue (48.2% ± 19.1%) and disorganized, loose connective tissue (36.9% ± 15.9%), with substantially smaller fractions of muscle (10.4% ± 22.0%) and fat (6.5% ± 11.6%). Only 25.8% of the biopsy pool contained any muscle fibers at all. Increased inflammatory cell counts (111.3 ± 81.5 macrophages/mm2) and increased vascularization (66.6 ± 38.0 vessels/mm2) were observed across biopsies. Muscle fiber degeneration was observed in 90.0% ± 15.6% of observable muscle fascicles, and the percentage of centrally nucleated muscle fibers was pathologically elevated (11.3% ± 6.3%). Fat accumulation was noted in both perifascicular (60.7% ± 41.4%) and intrafascicular (42.2% ± 33.6%) spaces, with evidence that lipid may replace contractile elements without altering muscle organization.

Conclusions: Dramatic degeneration and inflammation of the rotator cuff muscles are characteristics of the most chronic and severe rotator cuff disease states, suggesting that muscle loss is more complicated than, and distinct from, the simple atrophy found in less severe cases.

Clinical Relevance: In order to address degenerative muscle loss, alternative therapeutic approaches directed at muscle regeneration must be considered if muscle function is to be restored in late-stage rotator cuff disease.

1Departments of Bioengineering (M.C.G. and A.J.E.), Orthopaedic Surgery (T.C., M.P., S.S., and S.R.W.), and Radiology (S.R.W.), University of California San Diego, San Diego, California

2Kaiser Permanente, San Diego, California

E-mail address for S.R. Ward: srward@ucsd.edu

Article Outline

The lifetime prevalence of rotator cuff tear is estimated to be as high as 20% of the general population, where risk increases significantly with age1. Because rotator cuff tears may remain asymptomatic for years, and the onset of symptoms is often insidious, many patients do not seek treatment until the disease has progressed to a more chronic state. When accompanied by other factors such as smoking, diabetes, and age, this late-stage intervention results in surgical failure rates as high as 50%2. While the composition, quality, and function of the muscle in rotator cuff disease plays an important role in disease progression and recovery potential, the biological features of the muscle remain poorly defined on the cellular level, particularly in late-stage disease.

Relative to muscle, the response of the rotator cuff tendons to tear has been better studied biomechanically, biochemically, and histologically3-5. Those studies suggested that, while the tendon may look healthy macroscopically, at the cellular and molecular level, the torn tendon is degenerated, with signs including increased vascularity, adipose content, inflammation, and apoptosis3-5. We hypothesized that, as rotator cuff disease progresses, muscle as well as tendon is lost to degenerative processes. Specifically, in advanced rotator cuff disease, we expected to see a degenerative muscle phenotype demonstrating disrupted muscle fiber structure, myophagocytosis, and increased central nuclei, all signs of muscle degeneration-regeneration6 not reported in earlier disease states.

While atrophy and degeneration are distinct mechanisms of muscle loss often involving different pathways and drivers7-9, the terms fatty atrophy, fatty degeneration, and the more generic fatty infiltration are consistently and interchangeably found in the literature. Muscle atrophy is a normal response to disuse, whether that disuse is caused by tenotomy-induced mechanical unloading10 or inactivity11, and under most conditions can be nearly reversed by muscle reloading12. In contrast, degeneration is an accumulation of muscle damage and cell death driven by multiple factors including inflammation, abnormal mechanical forces, and altered vascularization and is classically seen in the inflammatory myopathies and dystrophies8,9. It is indisputable that fat accumulates in diseased rotator cuff muscle. Indeed, clinical and surgical decisions are often made on the basis of computed tomography (CT) or magnetic resonance imaging (MRI) evaluation of the rotator cuff muscles, with Goutallier scores being the primary clinical measure of the rotator cuff muscle quality and overall disease severity13,14.

Regarding the terms fatty atrophy versus fatty degeneration, previous studies of the relationship between muscle loss and fat accumulation are limited and have focused primarily on early to mid-stage rotator cuff disease. Those studies demonstrated decreased muscle fiber diameter15 and specific force production16 along with disrupted subcellular architecture, including small (∼1-μm) lipid droplet accumulation in individual muscle fibers following a tendon tear16,17. No degenerative features such as myophagocytosis, disrupted muscle fiber membranes, split fibers, centralized nuclei, or other classic features of muscle degeneration-regeneration6 have been reported15-17. While previous studies clearly demonstrated atrophic changes in the muscle, no clear evidence has emerged to suggest a degenerative mechanism of muscle loss.

To address these fundamental gaps in the literature regarding advanced rotator cuff disease, the aim of the current study was to histologically characterize the tissues found in the muscular spaces of patients with chronic and severe rotator cuff disease in order to determine whether degenerative processes are present in rotator cuff muscle. These data may provide evidence to challenge clinical assumptions about the state of muscle in rotator cuff disease and broaden our understanding of the relationships between muscle and fat in advanced rotator cuff disease.

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Materials and Methods

Twenty-three patients undergoing reverse total shoulder arthroplasty (Table I) provided consent for muscle biopsy to be performed. Approval from the appropriate institutional review boards was obtained. A fellowship-trained shoulder surgeon assigned Goutallier scores13 for each muscle on the basis of clinically indicated T1-weighted MRI or CT arthrogram images in the coronal and sagittal-oblique planes.

Using an anterosuperior approach, biopsy samples (ranging from 30 to 500 mg; mean diameter [and standard deviation] of 5.3 ± 2.0 mm) were collected under direct visualization from the scapular fossae, in a region defined by the lateral edge of the glenoid and no more than 2 cm medial from that landmark. Muscle that was macroscopically defined as pink-red organized fascicular tissue was specifically targeted; samples were not obtained from muscles without such tissue. Obvious adipose or tendon tissue was avoided (Fig. 1). The supraspinatus was always evaluated and a biopsy sample obtained if it met the above-mentioned criteria, and in multiple cases, the infraspinatus (including the infraspinatus-teres minor junction) was evaluated and a sample obtained. Immediately after biopsy, samples were pinned under tension along the medial-lateral axis, flash-frozen in liquid nitrogen-cooled isopentane, transported on dry ice, and stored at −80°C.

Ten-micron sections embedded in OCT (optimum cutting temperature) compound were generated on a cryostat (CM3050 S; Leica). For each measurement, the entire area of a single cross-section (6 to 20 fields) was quantified by a single observer with 4 years of muscle histology experience who was blinded to the Goutallier score and the intraoperative appearance (Table II). All quantification was performed using automated procedures in ImageJ18, which required only minimal input from the operator. Hematoxylin and eosin (H & E) and Gomori trichrome stains were used in evaluating the overall tissue composition and structure19. The relative fraction of organized, dense connective tissue; loose, disorganized tissue similar in appearance to granulation tissue; and muscle fibers was quantified on the basis of manual thresholding of staining color and intensity, where loose connective tissue was similar in color but lighter in shade and intensity than dense connective tissue because of the relative packing of the extracellular matrix (ECM). The same program was also used to quantify areas of fat within the sections.

To quantify the prevalence of degeneration and intrafascicular fat accumulation, each fascicle in an H & E-stained cross-section was scored for evidence of degenerating fibers and intrafascicular and perifascicular fat, with the results reported as a percentage of total fascicles per sample. In this analysis, muscle degeneration was defined as the presence of hypercellular infiltration of muscle fibers (i.e., myophagocytosis), the disrupted integrity of muscle fiber membranes, or split muscle fibers6,9. Intrafascicular fat was defined as large, eosin-negative cells residing within the perimysium, while perifascicular fat was defined as similarly eosin-negative cells in contact with, but not clearly within, the fascicular border. Immunostaining for the adipocyte lipid droplet protein perilipin (Abcam) was used to confirm that such eosin-negative cells contained lipid.

The number and size of noncapillary blood vessels (alpha-smooth muscle actin [α-SMA]; Diagnostic Biosystems) and the number of macrophages (CD68; Leica Biosystems/Novocastra) were quantified for whole biopsy cross-sections in ImageJ20,21. Samples were counterstained with laminin-111 or laminin-211 (LAMA1; Sigma, or LAMA2; Vector Laboratories) and 4',6-diamidino-2-phenylindole (DAPI).

The muscle fiber area and centralized nuclei were quantified via laminin and DAPI overlay using ImageJ18. The average muscle fiber area was weighted for total fiber area within each sample in order to account for varying amounts of muscle between samples. As a control group for muscle fiber area and central nuclei, 4 cadaveric supraspinatus muscles with no sign of rotator cuff disease were stained with H & E and quantified manually.

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Statistical Analysis

Data are reported as the mean and standard deviation. Due to the differences in disease state between muscles, no pairwise correction was applied when multiple muscles from the same individual were biopsied. One or 2-way analyses of variance (ANOVAs) were used as appropriate to compare measurements between Goutallier scores, and Student t tests were used to compare patient biopsies with cadaveric data as well as muscle versus nonmuscle regions within the same biopsy samples. Linear regression was used to assess correlations between compositional parameters. Significance thresholds were set at an alpha level of 0.05.

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Results

Of the total sample pool (n = 31), only 25.8% contained histologically identifiable muscle fibers. On average, the samples consisted of 48.2% ± 19.1% dense connective tissue, 36.9% ± 15.9% disorganized and loose connective tissue similar to granulation tissue, and only 10.4% ± 22.0% muscle overall (Fig. 2). Of the samples with any identifiable muscle tissue, the composition was 30.3% ± 11.8% dense connective tissue, 26.7% ± 17.4% loose connective tissue, and 40.1% ± 26.5% muscle. There was a significant interaction between biopsy sample composition and Goutallier score (p = 0.0027), with samples from muscles with lower Goutallier scores containing more muscle tissue on average, and with muscle tissue nearly absent in samples from muscles with a Goutallier score of 4. The average fat composition in all samples was low (6.5% ± 11.6%), which was likely due to our purposeful avoidance of macroscopic fat at biopsy sample collection.

Altered vascularization was prominent in the majority of samples, with an average vessel density among all samples of 66.6 ± 38.0 vessels/mm2 and an average area per vessel of 448.7 ± 204.3 μm2 (Fig. 3). This represents a 4-fold20 to 130-fold22 increase in the normal vessel density of skeletal muscle (dashed line in Figure 3-C, as measured in adult rat hindlimb muscle). No significant difference in vessel density was observed between samples with or without identifiable muscle tissue or on the basis of Goutallier scores.

Profound inflammation was observed (Fig. 4), with an average of 111.3 ± 81.5 macrophages/mm2 across all samples and 103.3 ± 88.8 macrophages/mm2 among the muscle-containing samples. This is 10-fold to 100-fold higher macrophage density than has been reported for either healthy muscle23 or tendon24 (1 to 10 macrophages/mm2, dashed line in Figure 4-C). No significant difference was found in macrophage density across Goutallier scores or between samples with or without histological evidence of muscle. Furthermore, macrophage density was not significantly different in biopsy regions composed primarily of muscle compared with the cumulative macrophage density of the total sample cross-section.

Within regions containing identifiable muscle fibers, signs of muscle degeneration were observed in 90.0% ± 15.6% of all muscle fascicles (Fig. 5). These features qualitatively occurred more frequently around muscle-nonmuscle interfaces and within regions of high cellularity, although precise quantification of this phenomenon proved difficult. The percentage of centrally nucleated muscle fibers was also significantly elevated (11.3% ± 6.3%; p = 0.0063) compared both with the cadaveric controls (2.74% ± 1.0%) and the clinical threshold for abnormal central nuclei (3%)25 (Fig. 6). Interestingly, no difference was found between biopsy samples and cadaveric controls in terms of the average muscle fiber area (Fig. 5-F), although this measurement was mildly underpowered (β = 0.62).

On average, adipose tissue was dispersed throughout the biopsy samples and constituted only a small fraction (6.5% ± 11.6%) of the total area sampled. In samples containing muscle fibers, although the total fat content remained low, fat was consistently concentrated at fascicular borders and between fascicles (perifascicular fat, 60.7% ± 41.4% of fascicles) as well as within fascicles (intrafascicular fat, 42.2% ± 33.6% of fascicles). Interestingly, intrafascicular fat was often seen in combination with degenerative signs, yet the endomysial and perimysial structures appeared intact (Figs. 7 and 8-A). In some cases, perilipin staining appeared to penetrate discontinuous laminin borders of muscle fibers (Fig. 8-B), although this phenomenon of fat infiltration of individual muscle fibers was rare and therefore difficult to quantify.

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Discussion

In the presence of chronic and severe rotator cuff disease, the nonfatty tissues of the normally muscular supraspinatus and infraspinatus fossae were composed primarily of vascularized connective tissue. A large fraction of this tissue was similar in appearance to granulation tissue that forms in response to severe tissue damage, in that it was composed of vascular, loose connective tissue that has a high macrophage density26. This finding is not unprecedented given that chronic inflammation and muscle degeneration cause neovascularization while diminishing muscle volume27,28. Although tissue composition was related to Goutallier score, the assumptions about tissue type and quality that are associated with each scoring level are called into question. Because our biopsy procedure targeted only nonfatty tissue, low fat content relative to clinical images was expected. However, lack of muscle relative to what was observed on imaging and intraoperatively may relate to the hypervascularity of the disorganized connective tissue. Additional research (high-resolution, quantitative MRI and histology) on isolated biopsy samples is needed to test this idea. Combined with the large number of actively degenerating muscle fibers observed, these data suggest that imaging may create considerable overestimation of the fraction of viable contractile tissue within the fossa.

While understanding tissue composition is critical to interpreting clinical images and making surgical decisions, elucidating the source of muscle loss is potentially a more important issue. Specifically, the distinction between the terms fatty degeneration, fatty atrophy, and fatty infiltration is rarely addressed in the literature. Although previously underappreciated, the implications of atrophy versus degeneration and fatty infiltration versus replacement are, in fact, critical in understanding disease progression on a cellular level.

The most appropriate term to describe the findings of previous studies in this area appears to be fatty atrophy. In evaluating the muscle of early-stage rotator cuff disease, Lundgreen et al. reported decreased muscle fiber diameter and increased perifascicular fat but found no evidence of degeneration-regeneration cycling as evidenced by a normal percentage of centrally nucleated muscle fibers15. Steinbacher et al. reported a similar increase in overall fat and perifascicular ECM, with increased lipid content and decreased myofibrillar organization in affected muscle fibers17. Mendias et al. corroborated these findings, and further showed diminished force production in muscle fibers isolated from chronic, full-thickness tears16. Animal models of rotator cuff disease, particularly large animal models, are relatively effective in recapitulating this atrophic, fat-accumulating phenotype of early and mid-stage disease29-35. However, to our knowledge, no previous study involving humans or animals has addressed the existence of degeneration or the tissue-level pattern of fat accumulation within the muscle in the most chronic and advanced stage of rotator cuff disease.

The data presented here provide clear evidence that muscle degeneration and regeneration occur in advanced rotator cuff disease. The level of muscle degeneration was profound, where 90% of fascicles displayed some degenerative features and 74% of the biopsy samples contained no muscle tissue at all, despite intraoperative targeting of muscle tissue. A previous study noted significant reductions in muscle fiber area between control samples and large tears17, and although our mean muscle fiber areas were also lower on average, they were not significantly different from cadaveric controls. Because our study was underpowered, we do not claim that atrophy was absent in these biopsy samples.

Cumulatively these data suggest that imbalanced degeneration-regeneration, possibly exacerbated by high levels of inflammation, is the primary driver of muscle loss in later-stage rotator cuff disease. We speculate that this degenerative mechanism of muscle loss is somewhat similar to that of the inflammatory myopathies, and contrasts with the atrophy-driven muscle loss reported in early-stage disease15. Functionally, muscle loss from atrophy and degeneration will both result in impaired force production. The distinction between atrophy and degeneration has critical implications for recovery. An atrophic muscle may be rescued with renewed activation and mechanical reloading (i.e., tendon repair and exercise) even in older patients12. Conversely, a chronically degenerated muscle with cell loss and likely impaired regenerative capacity may be further damaged by the same treatment36, with limited capacity to increase muscle fiber number. In the latter case, treatment with anti-inflammatory and/or pro-myogenic (pharmacological or cellular) therapies may be appropriate.

Of interest was the presence of fat in structures that appeared to be previously filled with contractile tissue, indicative of a more chronic and possibly terminal stage of degeneration. In addition to the fatty infiltration described ubiquitously in the literature, we demonstrated the proximity of degenerative fibers and large, lipid-filled structures within architecturally preserved fascicles in >40% of all fascicles. The origin of this lipid remains to be determined, but may be related to the lipid-laden macrophages described by Mendias et al.16, as it is possible that such macrophages could differentiate into adipocytes37. More detailed analysis demonstrated perilipin-positive membranes surrounded by or crossing laminin borders, suggesting that the lipid deposit had replaced the contractile apparatus while maintaining the original muscle fiber structure. To our knowledge, this is the first human example of a muscle-to-fat transition that preserved the underlying muscle matrix architecture in what could be considered “fatty replacement.” This phenomenon is not entirely unprecedented, as previous studies have shown adipogenic transition of both human myeloid cells37 and resident muscle stem cells in rodent systems38,39. Additional studies that specifically trace cell fate are required to identify the exact cell source or sources of the accumulating fat, as it is possible that the fat in different anatomical locations or in distinct stages of disease is derived from divergent cell depots.

As with many studies of this nature, obtaining the appropriate control data proved difficult. Rotator cuff muscle-specific control data are absent in the literature, and due to limitations inherent to fixed tissue—specifically that extracellular and nonmuscle tissues (especially loose connective and adipose tissue) are severely disrupted and often destroyed in processing—and that immunohistochemical assays are largely ineffective on fixed tissue, we could not obtain meaningful control data for many of our assays.

Another limitation of this study was the relatively low number of biopsy samples that actually contained identifiable muscle fibers, particularly in the supraspinatus. This low muscle fraction was startling, given the MRI and intraoperative appearance of muscle-like tissue in the biopsy region. Although we can associate a biopsy region with an imaging-based region, we did not perform stereotactic biopsy (exact position). Nevertheless, the severe muscle degeneration measured here suggests that tissue quality is not well represented by current clinical imaging modalities. This in turn may help to explain the high number of unsatisfactory clinical outcomes. If the extent of muscle degeneration and replacement by noncontractile tissue is systematically underestimated, which our data emphatically suggest, then standard treatments may insufficiently address the underlying pathology and functional deficits may persist even in the presence of an anatomically repaired cuff.

In summary, we demonstrated that rotator cuff muscle exhibits an active cycle of degeneration and regeneration in advanced rotator cuff disease, with degeneration potentially exacerbated by high levels of inflammation. Muscle loss in these samples was so profound that, in a large majority of the samples, muscle tissue was completely replaced by a disorganized, vascular connective-tissue network with high macrophage density. It is possible that such tissue appears similar to muscle in clinical imaging, leading to gross underestimation of muscle degeneration. We also provided evidence for the existence of a process of “fatty replacement,” whereby degenerating muscle is replaced by fat while maintaining the structure of the native muscle fiber ECM. Clinically, these results indicate that more careful staging of muscle quality may be important for surgical decision-making, as the most severe forms of degeneration may not respond well to muscle reloading.

Investigation performed at the University of California, San Diego, San Diego, California

Disclosure: Financial support for this work was provided by the National Institutes of Health (NIH) R01 HD073180 (S.R.W.); the Orthopaedic Research and Education Foundation (OREF) Resident Research Grant (T.C.); and the University of California, San Diego Frontiers of Innovation Scholars Program Fellowship (M.C.G.). The Disclosure of Potential Conflicts of Interest forms are provided with the online version of the article.

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References

1. Yamamoto A, Takagishi K, Osawa T, Yanagawa T, Nakajima D, Shitara H, Kobayashi T. Prevalence and risk factors of a rotator cuff tear in the general population. J Shoulder Elbow Surg. 2010 ;19(1):116–20.
2. Bishop J, Klepps S, Lo IK, Bird J, Gladstone JN, Flatow EL. Cuff integrity after arthroscopic versus open rotator cuff repair: a prospective study. J Shoulder Elbow Surg. 2006 ;15(3):290–9.
3. Longo UG, Franceschi F, Ruzzini L, Rabitti C, Morini S, Maffulli N, Denaro V. Histopathology of the supraspinatus tendon in rotator cuff tears. Am J Sports Med. 2008 ;36(3):533–8. Epub 2007 Nov 15.
4. Riley GP, Goddard MJ, Hazleman BL. Histopathological assessment and pathological significance of matrix degeneration in supraspinatus tendons. Rheumatology (Oxford). 2001 ;40(2):229–30.
5. DE Giorgi S, Saracino M, Castagna A. Degenerative disease in rotator cuff tears: what are the biochemical and histological changes? Joints. 2014 ;2(1):26–8. Epub 2014 May 8.
6. Dubowitz V, Sewry CA, Oldfors A. Muscle biopsy: a practical approach. 4th ed. Philadelphia: Elsevier Health Sciences; 2013.
7. Bonaldo P, Sandri M. Cellular and molecular mechanisms of muscle atrophy. Dis Model Mech. 2013 ;6(1):25–39.
8. Wallace GQ, McNally EM. Mechanisms of muscle degeneration, regeneration, and repair in the muscular dystrophies. Annu Rev Physiol. 2009;71:37–57.
9. Preedy VR, Peters TJ. Skeletal muscle: pathology, diagnosis and management of disease. New York: Cambridge University Press; 2002.
10. Thomason DB, Booth FW. Atrophy of the soleus muscle by hindlimb unweighting. J Appl Physiol (1985). 1990 ;68(1):1–12.
11. Levine S, Nguyen T, Taylor N, Friscia ME, Budak MT, Rothenberg P, Zhu J, Sachdeva R, Sonnad S, Kaiser LR, Rubinstein NA, Powers SK, Shrager JB. Rapid disuse atrophy of diaphragm fibers in mechanically ventilated humans. N Engl J Med. 2008 ;358(13):1327–35.
12. Frontera WR, Meredith CN, O’Reilly KP, Knuttgen HG, Evans WJ. Strength conditioning in older men: skeletal muscle hypertrophy and improved function. J Appl Physiol (1985). 1988 ;64(3):1038–44.
13. Goutallier D, Postel JM, Bernageau J, Lavau L, Voisin MC. Fatty muscle degeneration in cuff ruptures. Pre- and postoperative evaluation by CT scan. Clin Orthop Relat Res. 1994 ;304:78–83.
14. Fuchs B, Weishaupt D, Zanetti M, Hodler J, Gerber C. Fatty degeneration of the muscles of the rotator cuff: assessment by computed tomography versus magnetic resonance imaging. J Shoulder Elbow Surg. 1999 ;8(6):599–605.
15. Lundgreen K, Lian ØB, Engebretsen L, Scott A. Lower muscle regenerative potential in full-thickness supraspinatus tears compared to partial-thickness tears. Acta Orthop. 2013 ;84(6):565–70. Epub 2013 Oct 31.
16. Mendias CL, Roche SM, Harning JA, Davis ME, Lynch EB, Sibilsky Enselman ER, Jacobson JA, Claflin DR, Calve S, Bedi A. Reduced muscle fiber force production and disrupted myofibril architecture in patients with chronic rotator cuff tears. J Shoulder Elbow Surg. 2015 ;24(1):111–9. Epub 2014 Sep 3.
17. Steinbacher P, Tauber M, Kogler S, Stoiber W, Resch H, Sänger AM. Effects of rotator cuff ruptures on the cellular and intracellular composition of the human supraspinatus muscle. Tissue Cell. 2010 ;42(1):37–41. Epub 2009 Aug 25.
18. Abramoff MD, Magalhães PJ, Ram SJ. Image processing with ImageJ. Biophotonics International. 2004 ;11(7):36–42.
19. Miller JL, Watkin KL, Chen MF. Muscle, adipose, and connective tissue variations in intrinsic musculature of the adult human tongue. J Speech Lang Hear Res. 2002 ;45(1):51–65.
20. Hansen-Smith F, Egginton S, Hudlicka O. Growth of arterioles in chronically stimulated adult rat skeletal muscle. Microcirculation. 1998;5(1):49–59.
21. Pulford KA, Sipos A, Cordell JL, Stross WP, Mason DY. Distribution of the CD68 macrophage/myeloid associated antigen. Int Immunol. 1990;2(10):973–80.
22. Laughlin MH, Cook JD, Tremble R, Ingram D, Colleran PN, Turk JR. Exercise training produces nonuniform increases in arteriolar density of rat soleus and gastrocnemius muscle. Microcirculation. 2006 ;13(3):175–86.
23. Dorph C, Englund P, Nennesmo I, Lundberg IE. Signs of inflammation in both symptomatic and asymptomatic muscles from patients with polymyositis and dermatomyositis. Ann Rheum Dis. 2006 ;65(12):1565–71. Epub 2006 Jul 10.
24. Matthews TJ, Hand GC, Rees JL, Athanasou NA, Carr AJ. Pathology of the torn rotator cuff tendon. Reduction in potential for repair as tear size increases. J Bone Joint Surg Br. 2006 ;88(4):489–95.
25. Sorarù G, D’Ascenzo C, Polo A, Palmieri A, Baggio L, Vergani L, Gellera C, Moretto G, Pegoraro E, Angelini C. Spinal and bulbar muscular atrophy: skeletal muscle pathology in male patients and heterozygous females. J Neurol Sci. 2008 ;264(1-2):100–5. Epub 2007 Sep 12.
26. Clark R. The molecular and cellular biology of wound repair. New York: Springer; 2013.
27. Jackson JR, Seed MP, Kircher CH, Willoughby DA, Winkler JD. The codependence of angiogenesis and chronic inflammation. FASEB J. 1997 ;11(6):457–65.
28. Conn PM. Methods in neurosciences. Vol 8, Neurotoxins. Atlanta: Elsevier Science; 1992.
29. Liu X, Manzano G, Kim HT, Feeley BT. A rat model of massive rotator cuff tears. J Orthop Res. 2011 ;29(4):588–95. Epub 2010 Oct 14.
30. Gerber C, Meyer DC, Schneeberger AG, Hoppeler H, von Rechenberg B. Effect of tendon release and delayed repair on the structure of the muscles of the rotator cuff: an experimental study in sheep. J Bone Joint Surg Am. 2004 ;86(9):1973–82.
31. Gumucio JP, Davis ME, Bradley JR, Stafford PL, Schiffman CJ, Lynch EB, Claflin DR, Bedi A, Mendias CL. Rotator cuff tear reduces muscle fiber specific force production and induces macrophage accumulation and autophagy. J Orthop Res. 2012 ;30(12):1963–70. Epub 2012 Jun 13.
32. Safran O, Derwin KA, Powell K, Iannotti JP. Changes in rotator cuff muscle volume, fat content, and passive mechanics after chronic detachment in a canine model. J Bone Joint Surg Am. 2005 ;87(12):2662–70.
33. Kim HM, Galatz LM, Lim C, Havlioglu N, Thomopoulos S. The effect of tear size and nerve injury on rotator cuff muscle fatty degeneration in a rodent animal model. J Shoulder Elbow Surg. 2012 ;21(7):847–58. Epub 2011 Aug 10.
34. Rowshan K, Hadley S, Pham K, Caiozzo V, Lee TQ, Gupta R. Development of fatty atrophy after neurologic and rotator cuff injuries in an animal model of rotator cuff pathology. J Bone Joint Surg Am. 2010 ;92(13):2270–8.
35. Gerber C, Schneeberger AG, Perren SM, Nyffeler RW. Experimental rotator cuff repair. A preliminary study. J Bone Joint Surg Am. 1999 ;81(9):1281–90.
36. Garrett WE, Kirkendall DT. Exercise and sport science. Philadelphia: Lippincott Williams & Wilkins; 2000.
37. Majka SM, Fox KE, Psilas JC, Helm KM, Childs CR, Acosta AS, Janssen RC, Friedman JE, Woessner BT, Shade TR, Varella-Garcia M, Klemm DJ. De novo generation of white adipocytes from the myeloid lineage via mesenchymal intermediates is age, adipose depot, and gender specific. Proc Natl Acad Sci U S A. 2010 ;107(33):14781–6. Epub 2010 Aug 2.
38. Asakura A, Komaki M, Rudnicki M. Muscle satellite cells are multipotential stem cells that exhibit myogenic, osteogenic, and adipogenic differentiation. Differentiation. 2001 ;68(4-5):245–53.
39. Uezumi A, Ikemoto-Uezumi M, Tsuchida K. Roles of nonmyogenic mesenchymal progenitors in pathogenesis and regeneration of skeletal muscle. Front Physiol. 2014;5:68.
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