0021-972X/06/$15.00/0 Printed in U.S.A.

The Journal of Clinical Endocrinology & Metabolism 91(3):1159 –1167 Copyright © 2006 by The Endocrine Society doi: 10.1210/jc.2005-1877

Molecular Pathology of Mu ¨ ller’s Muscle in Graves’ Ophthalmopathy Mei-Ju Shih,* Shu-Lang Liao,* Kuan-Ting Kuo, Terry J. Smith, and Lee-Ming Chuang Graduate Institute of Clinical Medicine (M.-J.S., L.-M.C.), National Taiwan University School of Medicine, Taipei 10494, Taiwan; Departments of Ophthalmology (S.-L.L.), Pathology (K.-T.K.), and Internal Medicine (L.-M.C.), National Taiwan University Hospital, Taipei 10494, Taiwan; Department of Ophthalmology (M.-J.S.), Shin Kong Memorial Hospital, Taipei 111, Taiwan; Department of Medicine (M.-J.S.), College of Medicine, Hsinchuang, Fu-Jen Catholic University, Taipei Hsien 24205, Taiwan; Division of Molecular Medicine (T.J.S.), Harbor-University of California, Los Angeles Medical Center, Torrance, California 90502; and David Geffen School of Medicine (T.J.S.), University of California, Los Angeles, California 90095 Context: Upper lid retraction is a common sign in Graves’ ophthalmopathy (GO). Whether Mu¨ller’s muscle is involved in upper lid retraction has not been fully elucidated. Objective: The objective of the study was to understand the molecular pathology of Mu¨ller’s muscle in GO. Design/Setting/Participants: A method for measurement of histological changes was developed and used to correlate severity and expression of cell-specific genes in GO. Main Outcome Measures: Histological changes, clinical severity of upper lid retraction, and mRNA expression in Mu¨ller’s muscle in GO were measured.

M

ANIFESTATIONS OF GRAVES’ disease include thyrotoxicosis and several extrathyroidal signs including ophthalmopathy, dermopathy, and acropachy (1). Approximately 25–50% of patients may exhibit ocular complications, termed Graves’ ophthalmopathy (GO) or thyroid eye disease. These include lid retraction, proptosis, soft tissue swelling, strabismus, and compressive optic neuropathy (2). Although it is widely accepted that Graves’ disease is an autoimmune process and that the primary antigen in the thyroid is the TSH receptor (TSHr) (3, 4), the pathogenesis of GO remains poorly understood. Most believe that it is also autoimmune and involves an as-yet-unidentified shared autoantigen(s) in thyroid and orbital tissues (5, 6). Nearly 90% of patients with Graves’ disease manifest upper lid retraction, which is the most common ocular sign (7). Possible mechanisms for upper lid retraction include excessive sympathetic activity, which may overstimulate contractility of Mu¨ller’s muscle (8). However, upper eyelid retraction has also been observed in consistently euthyroid patients and First Published Online December 29, 2005 * M.-J.S. and S.-L.L. contributed equally to this work. Abbreviations: GAPDH, Glyceraldehyde 3-phosphate dehydrogenase; GO, Graves’ ophthalmopathy; HLA, human leukocyte antigen; HPF, high-power fields; IHC, immunohistochemistry; LCA, leukocyte common antigen; MRD1, marginal reflex distance; PPAR, peroxisome proliferator-activated receptor; Q-PCR, quantitative real-time PCR; TSHr, TSH receptor. JCEM is published monthly by The Endocrine Society (http://www. endo-society.org), the foremost professional society serving the endocrine community.

Results: The degree of fibrosis correlates with severity of upper lid retraction. Macrophage infiltration was increased in fibrotic areas, consistent with higher levels of macrophage-colony stimulating factor mRNA. Levels of peroxisome proliferator-activated receptor-␥ mRNA were up-regulated and correlated with fat infiltration. Decreased muscle mass correlated with lower myocardin mRNA expression. The expression of c-kit levels was decreased in diseased muscles, consistent with diminished mast cell numbers. Conclusion: The pathological changes of Mu¨ller’s muscle correlate with clinical severity of upper lid retraction in GO. Patterns of gene expression appear to correlate with the histopathological changes in this disease process. (J Clin Endocrinol Metab 91: 1159 –1167, 2006)

following normalization of thyroid function. Moreover, it is difficult to demonstrate increased Mu¨ller’s muscle contractility when experimental preparations are exposed to exogenous phenylephrine in hyperthyroid rats (9). Therefore, it is possible that intrinsic changes in the muscle might underlie manifestations seen in GO. Some studies have suggested that increased activity of upper lid retractors, including Mu¨ller’s and superior rectus muscles, compensates for the restricted movement of the inferior rectus muscle (10). However, this explanation is not fully accepted because some patients display upper lid retraction without evidence of restriction or enlargement of the inferior rectus. Much debate exists concerning whether pathological changes of Mu¨ller’s muscle are a consistent finding in GO patients. Reports range from no change (11) to an increased infiltration of macrophages and mast cells (12, 13) and muscle hypertrophy (14). Although fatty changes and fibrosis have been found, the correlation between pathological changes and severity of upper lid retraction is lacking (13). Therefore, in this study, we aimed to elucidate the molecular pathology occurring in Mu¨ller’s muscle in GO and address the potential relationship between histopathological changes, severity of upper lid retraction, and mRNA expression of certain cell lineage-specific genes. Subjects and Methods Subject recruitment and specimen collection Thirty-one euthyroid subjects (two males and 29 females) with GO who underwent Mu¨llerectomy were consecutively recruited. Clinical

1159

The Endocrine Society. Downloaded from press.endocrine.org by [${individualUser.displayName}] on 15 January 2017. at 07:28 For personal use only. No other uses without permission. . All rights reserved.

1160

J Clin Endocrinol Metab, March 2006, 91(3):1159 –1167

characteristics are given in Table 1. The degree of upper lid retraction was measured by documenting the distance from the corneal light reflex to the upper lid margin, which is referred to as the upper marginal reflex distance (MRD1, millimeters). The degree of proptosis was evaluated by a Hertel exophthalmometer (in millimeters). Mu¨ller’s muscle specimens were collected from surgery performed to correct lid retraction in GO patients who were stable with regard to thyroid and ophthalmic state for at least 3– 6 months before surgery. A control group of normal Mu¨ller’s muscles was collected from seven patients (six females, one male) with no thyroid history, aged 45–73 yr, who underwent correction of involutional ptosis causing by senile degeneration. This study was approved by the Institutional Review Board of the National Taiwan University Hospital. Written informed consent was obtained from each patient.

Special stain and immunohistochemistry (IHC) A total of 31 excised Mu¨ller’s muscles were fixed immediately in 10% buffered formalin overnight at 4 C. After gradient dehydration, tissue was embedded in paraffin. Sections of 3 ␮m thickness were stained with hematoxylin and eosin, toluidine blue, Masson’s trichrome stain, and immunohistochemical analyses for leukocyte common antigen (LCA), CD3, CD20, CD68, human leukocyte antigens (HLA)-DR, and caspase-3 with appropriate dilutions of the above primary antibodies (Dako, Carpinteria, CA). Anti-TSHr antibody (diluted 1:25) was purchased from Novocastra (Newcastle upon Tyne, UK). Endogenous peroxidase was quenched by 0.3% H2O2 containing sodium azide and levamisole (Dako) for 10 min. Slides were incubated for 60 min with primary antibodies following the manufacturers’ recommendations. After washing with PBS three times, the sections were incubated with secondary antibodies (EnVision⫹ dual link system, Dako) for 30 min. All slides were developed using 3,3⬘-diaminobenzidine chromogen for 5 min and counterstained with hematoxylin. Tonsil and lymph nodes were used as positive controls for LCA, CD3, CD20, CD68, caspase-3, and HLA-DR, whereas thyroid tissue from patients with simple goiter served as the positive control for TSHr. Each sample was also stained with a secondary antibody in the absence of primary antibodies as a negative control. To quantify the inflammatory cells, positive staining was counted manually using a light microscope at ⫻200 high-power fields (HPF) by two independent pathologists.

Quantitative histopathological evaluation of Mu¨ller’s muscle Using Masson’s trichrome stain, three major pathological components can be distinguished, i.e. red to purple defines muscle, blue suggests fiTABLE 1. Clinical characteristics of patients with thyroid lid retraction (n ⫽ 31) Characteristic

Value

Age (yr) Sex Female Male MRD1 (mm) Proptosis (mm) Pretreatment thyroid status Normal Hyper TBII (n ⫽ 19) Normal Abnormal Duration of hyperthyroidism (month) Duration of GO (month) Drug history ATD Steroid Thyroidectomy Sandostatin Smoking Family history of Graves’ disease

39.1 ⫾ 9.4 (23.0 ⬃ 64.0) 29 (93.6%) 2 (6.5%) 7.6 ⫾ 0.8 (6.0 ⬃ 9.0) 19.7 ⫾ 2.6 (15.0 ⬃ 26.0) 3 (9.7%) 28 (90.3%) 6 (31.6%) 13 (68.4%) 68.6 ⫾ 77.0 (12.0 ⬃ 336.0) 43.3 ⫾ 33.2 (3.0 ⬃ 120.0) 27 (87.1%) 16 (51.6%) 5 (16.1%) 1 (3.2%) 4 (12.9%) 3 (9.7%)

TBII, TSH binding inhibitor immunoglobulin; ATD, antithyroid drug.

Shih et al. • Mu¨ller’s Muscle in Thyroid Lid Retraction

brosis, and white indicates fat under standard light microscopy. Because there is a striking variation in composition of the pathological changes observed in different areas of the same specimen, we developed an image reconstruction method for quantifying the changes found in Mu¨ller’s muscle. Using an Eclipse E600 microscope (Nikon, Kanagawa, Japan) and a cooled charge-coupled device of ProgRes C14 (Jenoptik Group, Munich, Germany), all images were captured with a ⫻4 objective lens and a ⫻0.45 wide-angle lens. These images were then merged and proportionally reconstructed using Adobe Photoshop, version 7.0.1 [390 ⫻ 3090 pixels (Adobe Systems, San Jose, CA)] (15). Different pathological changes could be quantified based on color for each component. For more precise quantification, we used the magic wand tool under selection of tolerance values less than 50, in a noncontiguous, non antialiasing condition. Each pathological component was identified and saved as a new layer following summation (16). The total numbers of pixels identified using the histogram function in those pathological components constituted 100% opacity of the foreground color. These data were then transformed into percentages of the total pixels representing each area. Pre- and postreconstruction of images are shown in Fig. 1. To avoid possible bias resulting from uneven distribution in tissue specimens, another, noncontiguous section was also analyzed. Data from the two sets were used for the final calculations.

RNA extraction and reverse transcription Muscle specimens from another group of 21 subjects (16 GO and five normal) were divided. One fragment was subjected to quantitative pathological examination, and the other was snap frozen in liquid nitrogen, followed by RNA extraction using TRI REGENTBD (Molecular Research Center, Inc., Cincinnati, OH). Total RNA was reverse transcribed into cDNA in a 20-␮l reaction volume containing 0.1 m dithiothreitol, 10 ␮m deoxynucleotide triphosphate, 50 ␮m (dT)20, 40 U RNase inhibitor, and 200 U SuperScript III reverse transcriptase (Invitrogen Corp., Carlsbad, CA). Reactions were carried out for 42 min at 50 C, 15 min at 70 C, and 5 min at 95 C, followed by cooling to 4 C.

Quantitative real-time PCR (Q-PCR) Real-time or Q-PCR was performed with a LightCycler rapid thermal cycler system (Roche Diagnostics GmbH, Mannheim, Germany) according to the manufacturer’s instructions. The reaction was performed in a 20-␮l volume with final concentrations of 4 mm MgCl2, 2 ␮l LightCycler FastStart DNA Master SYBR green I (Roche Applied Science, Penzberg, Germany), 5 ␮l diluted cDNA, and 0.5 ␮m of both forward and reverse primers. PCR primers used were as follow: TSHr (forward, 5⬘-TGGGTGCAACACGGCT-3⬘; reverse, 5⬘-AGGGGTCTCGGTGTCC-3⬘); peroxisome proliferator-activated receptor (PPAR)-␥ (forward, 5⬘-GCCAGTTTCGCTCCGT-3⬘; reverse, 5⬘-CAAACCTGGGCGGTCT-3⬘); myocardin (forward, 5⬘-ACCCAACAACCCTCAC-3⬘; reverse, 5⬘-CTCCGGGTCATTTGCT-3⬘); macrophage-colony stimulation factor (MCSF) (forward, 5⬘-CCTGATTTCCCGTAAAGG-3⬘; reverse, 5⬘-GTTCACAGTGCAATGGC-3⬘); and c-kit (stem cell factor receptor) (forward, 5⬘GATGACGAGTTGGCCC-3⬘; reverse, 5⬘-AGGTAGTCGAGCGTTT3⬘). Glyceraldehyde 3-phosphate dehydrogenase (GAPDH) (forward, 5⬘-CATCACCATCTTCCAGGAGC-3⬘; reverse, 5⬘-GGATGATGTTCTGGGCTGCC-3⬘) was used as an internal control (Operon Biotechnologies Inc., Huntsville, AL). They were cycled with denaturation for one cycle at 95 C for 5 min and then for 40 cycles of 95 C for 5 sec, 55 or 52 C for 5 sec, and 72 C for 15 sec. The relative mRNA level of each gene was corrected with the corresponding GAPDH level.

Statistical analyses Data are presented as the mean ⫾ sd. The correlation between continuous variables, including pathological changes, severity of upper lid retraction, and mRNA levels of target genes were evaluated using Pearson’s correlation. Differences in MRD1 between clinical categorical data as well as the evaluation of differences in pathological composition and gene expression levels between normal and diseased groups were analyzed by Wilcoxon’s rank-sum test or Kruskal-Wallis test, as indicated. Differences in histopathological parameters within the same group were

The Endocrine Society. Downloaded from press.endocrine.org by [${individualUser.displayName}] on 15 January 2017. at 07:28 For personal use only. No other uses without permission. . All rights reserved.

Shih et al. • Mu¨ller’s Muscle in Thyroid Lid Retraction

J Clin Endocrinol Metab, March 2006, 91(3):1159 –1167

1161

FIG. 1. Reconstruction of the pathological alterations in Mu¨ller’s muscle. All images of the tissue sections were obtained under a ⫻4 objective lens and a ⫻0.45 wide-angle lens. The images were then reconstructed into their original size (central colored figure). Using the Photoshop magic wand, images of the selected area color differences are collected, indicative of total fat, muscle, blood, and fibrosis.

compared using Wilcoxon’s signed-rank test. Multivariate linear regression analysis was performed using variables with P ⬍ 0.05 from univariate analysis. All statistical analyses were performed with SAS software (SAS Institute, Cary, NC). P ⬍ 0.05 was considered statistically significant.

Results Clinical features and severity of upper lid retraction

The severity of upper lid retraction correlated significantly with age (r ⫽ 0.407, P ⫽ 0.020), initial pretreatment thyroid state (P ⫽ 0.045), and the period of hyperthyroidism (r ⫽ 0.501, P ⫽ 0.004) in univariate analysis. There is no significant relation between severity of upper lid retraction and other clinical variables, such as the titer of TBII, drug, smoking, and family history of Graves’ disease. In multivariate analysis after exclusion of colinearity, no correlation could be found between patient age (P ⫽ 0.608) or pretreatment thyroid status (P ⫽ 0.146) and severity of upper lid retraction. The duration of hyperthyroidism is the only independent variable correlated with clinical severity of upper lid retraction (P ⫽ 0.016). We recruited another group of 16 patients for both RNA extraction and pathological examination. Their clinical characteristics were similar to those of 31 patients

whose Mu¨ller’s muscle specimens underwent pathological analysis only. Quantitative pathological analyses of Mu¨ller’s muscle in normals and patients with GO

Using Masson’s trichrome staining, we determined the percentage of fat, muscle, or fibrosis composition in muscle specimens subjected to quantitative image analyses, as depicted in Fig. 1. Normal Mu¨ller’s muscle consists of undulating bundles of smooth muscle fibers, which are coated with a thin layer of connective tissue as well as small deposits of fat (Fig. 2A). No fibrosis was found in the normal Mu¨ller’s muscle, but collagen fibers are noted in the perimysium around the muscle bundles (Fig. 2A). In contrast, decreased muscle volume together with variable degrees of fat and fibrosis were observed in diseased Mu¨ller’s muscle (Fig. 2B). Significant fractional increases in fat (22.02 ⫾ 12.10 vs. 11.26 ⫾ 4.38%, P ⫽ 0.032) and fibrosis (33.74 ⫾ 13.96 vs. 20.10 ⫾ 3.93%, P ⫽ 0.006) and decreased muscle (44.23 ⫾ 12.56 vs. 68.64 ⫾ 1.86%, P ⬍ 0.001) were found in diseased Mu¨ller’s muscle when compared with normal specimens. Of note, no remarkable difference in

The Endocrine Society. Downloaded from press.endocrine.org by [${individualUser.displayName}] on 15 January 2017. at 07:28 For personal use only. No other uses without permission. . All rights reserved.

1162

J Clin Endocrinol Metab, March 2006, 91(3):1159 –1167

Shih et al. • Mu¨ller’s Muscle in Thyroid Lid Retraction

FIG. 2. Histopathology of normal and diseased Mu¨ller’s muscles. Undulating bundles of smooth muscle fibers as well as scant fatty tissue infiltration were found in normal Mu¨ller’s muscle (A). A variable degree of fat and fibrosis and decreased muscle volume were found in the diseased muscle (B). Magnification, ⫻200. There is no remarkable difference in pathological fraction, even in two noncontiguous sections (C and D) after reconstruction of images.

the histopathological appearance was found in two noncontiguous sections (Fig. 2, C and D). Immunohistochemical studies of Mu¨ller’s muscle

Two major immunocompetent cell types, mast cells and macrophages, infiltrate normal Mu¨ller’s muscle (Fig. 3, A and C). Mast cell numbers were decreased [29.6 ⫾ 14.0 vs. 60.0 ⫾ 18.8 cells per 5 HPF, P ⬍ 0.001], whereas increased numbers of macrophages infiltrate the muscle in GO (30.7 ⫾ 15.0 vs. 14.6 ⫾ 3.6 cells per 5 HPF, P ⫽ 0.004) (Fig. 3, B and D). Remarkably, numerous macrophages surround the extensively fibrotic area of the muscle specimens (Fig. 3E). We examined other cell types present in Mu¨ller’s muscle specimens. Cells positive for LCA were more abundant in normal Mu¨ller’s muscle than in diseased specimens (54.1 ⫾ 17.2 vs. 36.3 ⫾ 14.0 cells per 5 HPF, P ⫽ 0.023). T cells were also more numerous in normal muscle than GO, but differences failed to reach statistical significance (10.1 ⫾ 2.0 vs. 8.2 ⫾ 6.4 cells per 5 HPF, P ⫽ 0.161). Neither cell type stained positively for CD20, TSHr (Fig. 3G), or HLA-DR in both groups.

To determine whether the decrease in muscle results from greater apoptosis, we immunohistochemical stained for caspase-3. This staining was absent in all samples, suggesting that apoptosis is not enhanced in decreased muscle (data not shown). Relationship among infiltrating inflammatory cells, histopathology, and severity of upper lid retraction

To study the potential relationship among infiltrating immunocompetent cells, pathological changes, and severity of upper lid retraction, measurements were analyzed in regression models. Mast cell counts infiltrating in Mu¨ller’s muscle correlated positively with fractional fat composition (r ⫽ 0.56, P ⫽ 0.019), whereas they correlated negatively with fractional fibrosis (r ⫽ ⫺0.727, P ⫽ 0.001) (Fig. 4A). A positive correlation exists between macrophage counts and fibrosis (r ⫽ 0.503, P ⫽ 0.040) (Fig. 4B), consistent with increased macrophage infiltration frequently observed in areas of fibrosis (Fig. 3E). Moreover, we found a positive correlation between macrophage counts and severity of upper lid retraction (r ⫽ 0.512, P ⫽ 0.036, Fig. 4C).

The Endocrine Society. Downloaded from press.endocrine.org by [${individualUser.displayName}] on 15 January 2017. at 07:28 For personal use only. No other uses without permission. . All rights reserved.

Shih et al. • Mu¨ller’s Muscle in Thyroid Lid Retraction

J Clin Endocrinol Metab, March 2006, 91(3):1159 –1167

1163

FIG. 3. Cellular infiltration in Mu¨ller’s muscle and staining for TSHr in diseased Mu¨ller’s muscle. Abundant mast cells (A) and infrequent macrophages (C) infiltrate normal Mu¨ller’s muscle. In contrast, decreased mast cells (B) and increased macrophages (D) infiltrate diseased muscle. Numerous macrophages surround extensive fibrosis (E). Thyroid tissue from a simple goiter was used as positive control for TSHr IHC (F). There is negative staining in diseased Mu¨ller’s muscle (G). Original magnification, ⫻100 (A, B, C, D, and G); ⫻200 (F); ⫻400 (E).

Cell lineage-specific gene expression correlates with pathological changes in Mu¨ller’s muscle

To understand the molecular basis for the pathological changes in Mu¨ller’s muscle in GO, we studied genes that have been found previously involved in the regulation of differentiation, such as those implicated in adipogenesis, myogenesis, and differentiation of mast cells and macro-

phages. These include PPAR␥, myocardin, c-kit, and MCSF, respectively. Using real-time PCR, we found a remarkable increase in PPAR␥ mRNA expression in diseased Mu¨ller’s muscle when compared with that in normal tissue (5.73 ⫾ 5.40 vs. 0.38 ⫾ 0.22, P ⫽ 0.001) (Fig. 5A). The PPAR␥ mRNA levels were positively correlated with the fractional fat composition by regression analysis (r ⫽ 0.75, P ⫽ 0.001). As

The Endocrine Society. Downloaded from press.endocrine.org by [${individualUser.displayName}] on 15 January 2017. at 07:28 For personal use only. No other uses without permission. . All rights reserved.

1164

J Clin Endocrinol Metab, March 2006, 91(3):1159 –1167

Shih et al. • Mu¨ller’s Muscle in Thyroid Lid Retraction

analyses. These findings suggest a potential role for these factors in the fibrosis of Mu¨ller’s muscle in GO. We were unable to detect TSHr protein by IHC in either diseased Mu¨ller’s muscle (Fig. 3G), although the positive control (thyroid) was markedly positive (Fig. 3F). In contrast, TSHr mRNA was detected in muscle with real-time PCR. As shown in Fig. 5E, TSHr mRNA was found in both normal and diseased Mu¨ller’s muscle specimens. No significant difference of the expression level was found between the two groups (1.63 ⫾ 1.05 vs. 1.44 ⫾ 0.89, P ⫽ 0.493, Fig. 5E). Discussion

FIG. 4. Correlation among cellular infiltration, pathological changes, and severity of upper lid retraction. A, Mast cell counts were positively correlated with fractional fat composition and negatively correlated with fibrosis. B, Significant correlation between macrophage infiltration and fibrosis. C, Positive correlation between macrophage infiltration and severity of upper lid retraction.

depicted in Fig. 5A, a significant difference in PPAR␥ mRNA levels was observed in groups differing in fat composition, i.e. less than 10%, 10 – 40%, and more than 40% (P ⫽ 0.004), respectively. Myocardin mRNA, a crucial smooth muscle transcriptional factor, was diminished in diseased muscle (0.99 ⫾ 0.36 vs. 2.98 ⫾ 0.97, P ⫽ 0.001, Fig. 5B) and correlated with fractional muscle mass (r ⫽ 0.503, P ⫽ 0.047) (data not shown). MCSF mRNA levels were increased in diseased muscle (1.60 ⫾ 0.62 vs. 1.00 ⫾ 0.13, P ⫽ 0.004), especially in the specimens in which fibrosis was greater than 40% (Fig. 5C). In contrast, c-kit mRNA was substantially decreased in diseased tissue (1.67 ⫾ 1.11 vs. 8.73 ⫾ 6.15, P ⫽ 0.007, Fig. 5D). Interestingly, MCSF mRNA correlated positively with fractional fibrosis (r ⫽ 0.609, P ⫽ 0.012), whereas c-kit mRNA correlated negatively (r ⫽ ⫺0.496, P ⫽ 0.051) in the regression

We demonstrate here for the first time that the degree of inflammatory cells infiltration and pathological changes of Mu¨ller’s muscle correlate with clinical severity of upper eyelid retraction in GO. We also provide a putative molecular basis for differential gene expression of several key factors, including PPAR␥, myocardin, MCSF, and c-kit. Our current findings provide the rationale for the pathological changes observed in Mu¨ller’s muscle. Among the clinical variables from our studies, age, pretreatment thyroid function status, and duration of hyperthyroidism are considered risk factors of developing upper lid retraction. However, duration of hyperthyroidism proved to be the only independent predictor in the multivariate regression models. With the image processing system presented in this report, we demonstrated that severity of upper lid retraction is positively correlated with the degree of fibrosis and macrophage infiltration the Mu¨ller’s muscle. Thus, although increased sympathetic activity, hypertrophy, and/or excessive contractility of Mu¨ller’s muscle have been reported as causes of upper lid retraction, we provide very strong evidence of pathological changes in Mu¨ller’s muscle that also contribute. PPAR␥ is a nuclear hormone receptor that plays a crucial role in the complex process for adipogenesis (17, 18). Its expression is increased in orbital adipose tissues and extraocular muscles (19). Here we report that its expression is also higher in Mu¨ller’s muscle in GO. The steady-state level of PPAR␥ mRNA correlates well with the fractional fat composition in Mu¨ller’s muscle specimens. The function of increased PPAR␥ in Mu¨ller’s muscle is currently unknown. However, a reciprocal relationship was found in the percentages of skeletal muscle and adipose tissue reported in myodystrophic states, raising the possibility that myoblasts might transdifferentiate into adipocytes through the activation of PPAR␥ (20 –22). Thus, we propose that the increased adipose composition found in diseased Mu¨ller’s muscle might originate from two pathways. Adipocytes differentiation from fibroblasts, or transdifferentiation from myoblasts into adipocytes might occur. A mechanism involving PPAR␥ activation could explain either. Indeed, the expression of adipocyte fatty acid binding protein was also highly expressed in diseased Mu¨ller’s muscle corresponding to the expression pattern of PPAR␥ (Chuang, L.-M., M.-J. Shih, and S.-L. Liao, unpublished observations). It should be noted that fibroblasts from orbital fat are heterogeneous with regard to Thy-1 display and can be differentiated into either adipocytes or myofibroblasts, depending on their lineage and

The Endocrine Society. Downloaded from press.endocrine.org by [${individualUser.displayName}] on 15 January 2017. at 07:28 For personal use only. No other uses without permission. . All rights reserved.

Shih et al. • Mu¨ller’s Muscle in Thyroid Lid Retraction

J Clin Endocrinol Metab, March 2006, 91(3):1159 –1167

1165

FIG. 5. Expression levels of cell lineage-specific mRNAs in normal and diseased muscle. mRNA levels were determined with Q-PCR and normalized to corresponding GAPDH mRNA levels. A, PPAR␥ mRNA levels are increased in diseased compared with normal muscle (P ⫽ 0.001). A significant difference exists between PPAR␥ levels and various degrees of fat infiltration, i.e. less than 10%, 10 – 40%, and more than 40% (P ⫽ 0.004). B, Myocardin mRNA levels are diminished in diseased muscle compared with normal specimens (P ⫽ 0.001). C, MCSF mRNA levels are increased in diseased compared with normal muscle (P ⫽ 0.004), and a significant difference exists between more than 40% and less than 40% fibrosis (P ⫽ 0.017). D, c-kit mRNA levels are reduced in diseased muscle (P ⫽ 0.007 vs. normal). E, TSHr mRNA levels are similar in normal and diseased muscle. *, P ⱕ 0.05; **, P ⱕ 0.001 by Wilcoxon’s rank-sum test; #, P ⫽ 0.004 by Kruskal-Wallis test.

whether they are activated by TGF␤ or a PPAR␥ agonist (23, 24). To understand the mechanisms involved in reduced contractile elements in diseased Mu¨ller’s muscle, we studied the

expression of the myocardin gene. This protein is critical for the transcriptional regulation of smooth muscle cell differentiation (25–27). We found that the level of myocardin expression was decreased in diseased tissue, suggesting that a

The Endocrine Society. Downloaded from press.endocrine.org by [${individualUser.displayName}] on 15 January 2017. at 07:28 For personal use only. No other uses without permission. . All rights reserved.

1166

J Clin Endocrinol Metab, March 2006, 91(3):1159 –1167

reduction might result in attenuated Mu¨ller’s muscle differentiation in GO. We found no evidence of caspase-3-dependent apoptosis in diseased Mu¨ller’s muscle. Taken together, the reduced mass of Mu¨ller’s muscle in GO might result from attenuated myogenesis, increased adipogenesis, and the potential transdifferentiation of myoblasts into adipocytes. Clearly future studies in vitro might shed additional light on these remaining mechanistic questions. In our study, the numbers of infiltrating macrophages were found increased in the majority of diseased Mu¨ller’s muscle specimens examined. This was especially true in specimens with prominent fibrosis. Consistently, we found that the expression of MCSF, a growth factor promoting survival, proliferation, differentiation, and activation of mononuclear phagocytes (28, 29) was increased in diseased muscle. In addition, MCSF mRNA levels were positively correlated with the fractional fibrosis, implying a role for macrophage propagation in fibrosis muscle fibrosis in GO. Whether locally generated macrophage-derived cytokines, such as IL-1␤, TNF␣, and IL-10, levels of which are increased in orbital adipose tissue, (30) promote the pathological features in Mu¨ller’s muscle remains to be investigated. We found a negative correlation between the c-kit and fractional fibrosis in diseased muscle. c-kit is a mast cellspecific gene (31, 32). Mast cells proliferate in response to c-kit activation. Our observation of low c-kit levels coupled with finding relatively few mast cells in diseased muscle is somewhat surprising. Previous studies have shown that they not only induce fibrosis by stimulating fibroblasts to proliferate in the skin and lung (33, 34) but also participate in tissue remodeling possibly through increased production of proinflammatory prostaglandin E2 and hyaluronan in GO (35, 36). Further study will be required to clarify the role of mast cell and the expression the c-kit in the fibrosis in Mu¨ller’s muscle in GO. Kumar et al. (19) demonstrated a parallel increase in TSHr expression and adipocyte-specific genes in the orbital tissues of GO patients. They interpreted their findings as suggesting a role of TSHr in the pathogenesis of GO. Although we found expression of TSHr mRNA level in normal and diseased Mu¨ller’s muscle, the TSHr protein could not be detected with IHC. Moreover, we did not find a difference in the expression of TSHr mRNA level in normal and diseased Mu¨ller’s muscle. The underlying mechanism for explaining the discrepancy between mRNA and protein expression is not clear; nonetheless, a possible illegitimate transcription of the TSHr mRNA in extrathyroid tissues has been reported (37–39). In addition, there was no correlation between TSHr levels and any of the pathological changes or PPAR␥ mRNA levels. Our data suggest that TSHr may not play a role for the pathological changes occurring in diseased Mu¨ller’s muscle. These studies have certain limitations. We could study only samples from surgical resection in patients who failed medical treatment. This may have biased our findings. Rarely does opportunity occur for examining tissues early in the disease or specimens from patients in whom medical therapies prove effective. We did examine tissues from two patients after relatively short disease duration (3 and 5 months after onset of hyperthyroidism). Surprisingly, IHC studies revealed substantial fibrosis. T cell infiltration was

Shih et al. • Mu¨ller’s Muscle in Thyroid Lid Retraction

also prominent in these samples, suggesting an immunemediated process in diseased Mu¨ller’s muscle (Shih, M.-J., S.-L. Liao, and L.-M. Chuang, unpublished observations). We suspect that a transient, early inflammatory phase gives way to fibrosis. In conclusion, we provide for the first time the potential mechanistic explanation for pathological changes in Mu¨ller’s muscle in GO. Expression of PPAR␥, myocardin, MCSF, and c-kit appear to be altered. We propose that these molecular derangements underlie the muscle pathology. In addition to the orbital adipose tissues and extraocular muscles, Mu¨ller’s muscle may also be considered an important target of autoimmunity in GO. Acknowledgments Received August 19, 2005. Accepted December 20, 2005. Address all correspondence and requests for reprints to: Dr. LeeMing Chuang, Department of Medicine, National Taiwan University Hospital, No. 7, Chung-Shan South Road, Taipei, Taiwan. E-mail: [email protected]. This work was supported in part by Grants NSC 93-2752-B-002-009PAE and NSC 94-2752-B-002-008-PAE from the National Science Council of the Republic of China (to L.-M.C.) and EY08976, EY07118, and DK063121 from the National Institutes of Health (to T.J.S.).

References 1. Taylor S 1986 Graves of Graves’ disease. J R Coll Phys London 20:298 –300 2. Bartley GB, Gorman CA 1995 Diagnostic criteria for Graves’ ophthalmopathy. Am J Ophthalmol 119:792–795 3. Paschke R, Ludgate M 1997 The thyrotropin receptor in thyroid disease. N Engl J Med 337:1675–1681 4. Kohn LD, Shimojo N, Kohno Y, Suzuki K 2000 An animal model of Graves’ disease: understanding the cause of autoimmune hyperthyroidism. Rev Endocrinol Metab Dis 1:59 – 67 5. Bahn RS, Heufelfelder AE 1993 Pathogenesis of Graves’ ophthalmopathy. N Engl J Med 329:1468 –1475 6. Bahn RS 2003 Pathophysiology of Graves’ ophthalmopathy: the cycle of disease. J Clin Endocrinol Metab 88:1939 –1946 7. Bartley GB, Fatourechi V, Kadrmas EF, Jacobsen SJ, Ilstrup DM, Garrity JA, Gorman CA 1996 Clinical features of Graves’ ophthalmopathy in an incidence cohort. Am J Ophthalmol 121:284 –290 8. Putterman AM, Urist M 1972 Surgical treatment of upper eyelid retraction. Arch Ophthalmol 87:401– 405 9. Bodker FS, Putterman AM, Laris A, Miletich DA, Volgel SM, Viana MAG 1997 The effect of hyperthyroidism on Mu¨ller’s muscle contractility in rats. Ophthal Plast Reconstr Surg 13:161–167 10. Jampolsky A 1970 What can electromyography do for the ophthalmologist? Invest Ophthalmol 9:570 –599 11. Rootman J, Patel S, Berry K, Nugent R 1987 Pathological and clinical study of the Mu¨ller’s muscle in Graves’ ophthalmology. Can J Ophthalmol 22:32–36 12. Gaag RVD, Schmidt D, Koornneef L 1993 Retrobulbar histology and immunohistochemistry in endocrine ophthalmopathy. Dev Ophthalmol 25:1–10 13. Cockerham KP, Hidayat AA, Brown HG, Cockerham GC, Graner SR 2002 Clinicopathologic evaluation of the Mueller muscle in thyroid-associated orbitopathy. Ophthal Plast Reconstr Surg 18:11–17 14. Kagoshima T, Hori S, Inoue Y 1987 Qualitative and quantitative analyses of Mu¨ller’s muscle in dysthyroid ophthalmopathy. Jpn J Ophthalmol 31:646 – 654 15. Caruso RD, Postel GC 2002 Image annotation with Adobe Photoshop. J Digit Imaging 15:197–202 16. Lahm A, Uhl M, Lehr HA, Ihling C, Kreuz PC, Haberstroh J 2004 Photoshopbased image analysis of canine articular cartilage after subchondral damage. Arch Orthop Trauma Surg 124:431– 436 17. Wright HM, Clish CB, Mikami T, Hauser S, Yanagi K, Hiramatsu R, Serhan CN, Spiegelman BM 2000 A synthetic antagonist for the peroxisome proliferator-activated receptor ␥ inhibits adipocyte differentiation. J Biol Chem 275:1873–1877 18. Tontonoz P, Hu E, Spiegelman BM 1994 Stimulation of adipogenesis in fibroblasts by PPAR␥2, a lipid-activated transcription factor. Cell 79:1147–1156 19. Kumar S, Coenen MJ, Scherer PE, Bahn RS 2004 Evidence for enhanced adipogenesis in the orbits of patients with Graves’ ophthalmopathy. J Clin Endocrinol Metab 89:930 –935 20. Hu E, Tontonoz P, Spiegelman BM 1995 Transdifferentiation of myoblasts by

The Endocrine Society. Downloaded from press.endocrine.org by [${individualUser.displayName}] on 15 January 2017. at 07:28 For personal use only. No other uses without permission. . All rights reserved.

Shih et al. • Mu¨ller’s Muscle in Thyroid Lid Retraction

21. 22. 23. 24. 25. 26. 27.

28. 29. 30.

the adipogenic transcription factors PPAR␥ and C/EBP␣. Proc Natl Acad Sci USA 92:9856 –9860 Grimaldi PA, Teboul L, Inadera H, Gaillard D, Amri EZ 1997 Trans-differentiation of myoblasts to adipoblasts: triggering effects of fatty acids and thiazolidinediones. Prostaglandins Leukot Essent Fatty Acids 57:71–75 Holst D, Luquet S, Kristiansen K, Grimaldi PA 2003 Roles of peroxisome proliferator-activated receptors ␦ and ␥ in myoblast transdifferentiation. Exp Cell Res 288:168 –176 Smith TJ, Koumas L, Gagnon A, Bell A, Sempowski GD, Phipps RP, Sorisky A 2002 Orbital fibroblast heterogeneity may determine the clinical presentation of thyroid-associated ophthalmopathy. J Clin Endocrinol Metab 87:385–392 Koumas, L, Smith TJ, Feldon S, Blumberg N, Phipps RP 2003 Thy-1 expression in human fibroblast subsets defines myofibroblastic or lipofibroblastic phenotypes. Am J Pathol 163:1291–1300 Suzuki T, Nagai R, Yazaki Y 1998 Mechanisms of transcription regulation of gene expression in smooth muscle cells. Circ Res 82:1238 –1242 Wang Z, Wang DZ, Pipes GC, Olson EN 2003 Myocardin is a master regulator of smooth muscle gene expression. Proc Natl Acad Sci USA 100:7129 –7134 Du KL, Ip HS, Li J, Chen M, Dandre F, Yu W, Lu MM, Owens GK, Parmacek MS 2003 Myocardin is a critical serum response factor cofactor in the transcriptional program regulating smooth muscle cell differentiation. Mol Cell Biol 23:2425–2437 Tushinski RJ, Oliver IT, Guilbert LJ, Tynan PW, Warner JR, Stanley ER 1982 Survival of mononuclear phagocytes depends on a lineage-specific growth factor that the differentiated cells selectively destroy. Cell 28:71– 81 Wang JM, Griffin JD, Rambaldi A, Chen ZG, and Mantovani A 1988 Induction of monocyte migration by recombinant macrophage colony-stimulating factor. J Immunol 141:575–579 Kumar S, Bahn RS 2003 Relative overexpression of macrophage-derived cy-

J Clin Endocrinol Metab, March 2006, 91(3):1159 –1167

31. 32. 33.

34.

35.

36. 37.

38.

39.

1167

tokines in orbital adipose tissue from patients with Graves’ ophthalmopathy. J Clin Endocrinol Metab 88:4246 – 4250 Ashman LK 1999 The biology of stem cell factor and receptor c-kit. Int. J Biochem Cell Biol 31:1037–1051 Linnekin D 1999 Early signaling pathways activated by c-kit in hematopoietic cells. Int J Biochem Cell Biol 31:1035–1074 Garbuzenko E, Puxeddu I, Levi-Schaffer F, Berkman N, Kramer M, Nagler A 2004 Mast cells induce activation of human lung fibroblasts in vitro. Exp Lung Res 30:705–721 Garbuzenko E, Nagler A, Pickholtz D, Gillery P, Reich R, Maquart FX, Levi-Schaffer F 2002 Human mast cells stimulate fibroblast proliferation, collagen synthesis and lattice contraction: a direct role for mast cells in skin fibrosis. Clin Exp Allergy 32:237–246 Smith TJ, Parikh SJ 1999 HMC-1 mast cells activate human orbital fibroblasts in coculture: evidence for up-regulation of prostaglandin E2 and hyaluronan synthesis. Endocrinology 140:3518 –3525 Smith TJ 2003 Unique properties of orbital connective tissue underlie its involvement in Graves’ disease. Minerva Endocrinol 28:213–222 Chelly J, Concordet JP, Kaplan JC, Kahn A 1989 Illegitimate transcription: transcription of any gene in any cell types. Proc Natl Acad Sci USA 86:2617– 2621 Aust G, Crisp M, Bosenberg E, Ludgate M, Weetman AP, Paschke R 1998 Transcription of thyroid autoantigens in non-expressing tissues. Exp Clin Endocrinol Diabetes 106:319 –323 Agretti P, Chiovato L, De Marco G, Marcocci C, Mazzi B, Sellari-Franceschini S, Vitti P, Pinchera A, Tonacchera M 2002 Real-time PCR provides evidence for thyrotropin receptor mRNA expression in orbital as well as in extraorbital tissues. Eur J Endocrinol 147:733–739

JCEM is published monthly by The Endocrine Society (http://www.endo-society.org), the foremost professional society serving the endocrine community.

The Endocrine Society. Downloaded from press.endocrine.org by [${individualUser.displayName}] on 15 January 2017. at 07:28 For personal use only. No other uses without permission. . All rights reserved.