help button home button Endocrine Society JCEM
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Journal of Clinical Endocrinology & Metabolism , doi:10.1210/jc.2005-1877
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Submit a related Letter to the Editor
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow Request Copyright Permission
Citing Articles
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Shih, M.-J.
Right arrow Articles by Chuang, L.-M.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Shih, M.-J.
Right arrow Articles by Chuang, L.-M.
Related Collections
Right arrow Thyroid
Right arrow Autoimmunity
The Journal of Clinical Endocrinology & Metabolism Vol. 91, No. 3 1159-1167
Copyright © 2006 by The Endocrine Society

Molecular Pathology of Müller’s Muscle in Graves’ Ophthalmopathy

Mei-Ju Shih1, Shu-Lang Liao1, 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

Address all correspondence and requests for reprints to: Dr. Lee-Ming Chuang, Department of Medicine, National Taiwan University Hospital, No. 7, Chung-Shan South Road, Taipei, Taiwan. E-mail: leeming{at}ha.mc.ntu.edu.tw.


   Abstract
Top
 Abstract
Introduction
 Subjects and Methods
 Results
 Discussion
 References
 
Context: Upper lid retraction is a common sign in Graves’ ophthalmopathy (GO). Whether Müller’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 Müller’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 Müller’s muscle in GO were measured.

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-{gamma} 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 Müller’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.


   Introduction
Top
 Abstract
 Introduction
Subjects and Methods
 Results
 Discussion
 References
 
MANIFESTATIONS 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 Müller’s muscle (8). However, upper eyelid retraction has also been observed in consistently euthyroid patients and following normalization of thyroid function. Moreover, it is difficult to demonstrate increased Müller’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 Müller’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 Müller’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 Müller’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
Top
 Abstract
 Introduction
 Subjects and Methods
Results
 Discussion
 References
 
Subject recruitment and specimen collection

Thirty-one euthyroid subjects (two males and 29 females) with GO who underwent Müllerectomy were consecutively recruited. Clinical characteristics are given in Table 1Go. 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).


View this table:
[in this window]
[in a new window]
  		TABLE 1. Clinical characteristics of patients with thyroid lid retraction (n = 31)

 
Müller’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 Müller’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 Müller’s muscles were fixed immediately in 10% buffered formalin overnight at 4 C. After gradient dehydration, tissue was embedded in paraffin. Sections of 3 µm 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 x200 high-power fields (HPF) by two independent pathologists.

Quantitative histopathological evaluation of Müller’s muscle

Using Masson’s trichrome stain, three major pathological components can be distinguished, i.e. red to purple defines muscle, blue suggests fibrosis, 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 Müller’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 x4 objective lens and a x0.45 wide-angle lens. These images were then merged and proportionally reconstructed using Adobe Photoshop, version 7.0.1 [390 x 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. 1Go. 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.


Figure 1
View larger version (60K):
[in this window]
[in a new window]
  		FIG. 1. Reconstruction of the pathological alterations in Müller’s muscle. All images of the tissue sections were obtained under a x4 objective lens and a x0.45 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.

 
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-µl reaction volume containing 0.1 M dithiothreitol, 10 µM deoxynucleotide triphosphate, 50 µM (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-µl volume with final concentrations of 4 mM MgCl2, 2 µl LightCycler FastStart DNA Master SYBR green I (Roche Applied Science, Penzberg, Germany), 5 µl diluted cDNA, and 0.5 µM 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)-{gamma} (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'-AGGTAGTCGAGCGTTT-3'). 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 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
Top
 Abstract
 Introduction
 Subjects and Methods
 Results
Discussion
 References
 
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 Müller’s muscle specimens underwent pathological analysis only.

Quantitative pathological analyses of Müller’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. 1Go. Normal Müller’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. 2AGo). No fibrosis was found in the normal Müller’s muscle, but collagen fibers are noted in the perimysium around the muscle bundles (Fig. 2AGo). In contrast, decreased muscle volume together with variable degrees of fat and fibrosis were observed in diseased Müller’s muscle (Fig. 2BGo). 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 Müller’s muscle when compared with normal specimens. Of note, no remarkable difference in the histopathological appearance was found in two noncontiguous sections (Fig. 2Go, C and D).


Figure 2
View larger version (122K):
[in this window]
[in a new window]
  		FIG. 2. Histopathology of normal and diseased Müller’s muscles. Undulating bundles of smooth muscle fibers as well as scant fatty tissue infiltration were found in normal Müller’s muscle (A). A variable degree of fat and fibrosis and decreased muscle volume were found in the diseased muscle (B). Magnification, x200. There is no remarkable difference in pathological fraction, even in two noncontiguous sections (C and D) after reconstruction of images.

 
Immunohistochemical studies of Müller’s muscle

Two major immunocompetent cell types, mast cells and macrophages, infiltrate normal Müller’s muscle (Fig. 3Go, 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. 3Go, B and D). Remarkably, numerous macrophages surround the extensively fibrotic area of the muscle specimens (Fig. 3EGo).


Figure 3
View larger version (100K):
[in this window]
[in a new window]
  		FIG. 3. Cellular infiltration in Müller’s muscle and staining for TSHr in diseased Müller’s muscle. Abundant mast cells (A) and infrequent macrophages (C) infiltrate normal Müller’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 Müller’s muscle (G). Original magnification, x100 (A, B, C, D, and G); x200 (F); x400 (E).

 
We examined other cell types present in Müller’s muscle specimens. Cells positive for LCA were more abundant in normal Müller’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. 3GGo), 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 Müller’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. 4AGo). A positive correlation exists between macrophage counts and fibrosis (r = 0.503, P = 0.040) (Fig. 4BGo), consistent with increased macrophage infiltration frequently observed in areas of fibrosis (Fig. 3EGo). Moreover, we found a positive correlation between macrophage counts and severity of upper lid retraction (r = 0.512, P = 0.036, Fig. 4CGo).


Figure 4
View larger version (16K):
[in this window]
[in a new window]
  		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.

 
Cell lineage-specific gene expression correlates with pathological changes in Müller’s muscle

To understand the molecular basis for the pathological changes in Müller’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 macrophages. These include PPAR{gamma}, myocardin, c-kit, and MCSF, respectively. Using real-time PCR, we found a remarkable increase in PPAR{gamma} mRNA expression in diseased Müller’s muscle when compared with that in normal tissue (5.73 ± 5.40 vs. 0.38 ± 0.22, P = 0.001) (Fig. 5AGo). The PPAR{gamma} mRNA levels were positively correlated with the fractional fat composition by regression analysis (r = 0.75, P = 0.001). As depicted in Fig. 5AGo, a significant difference in PPAR{gamma} 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. 5BGo) and correlated with fractional muscle mass (r = 0.503, P = 0.047) (data not shown).


Figure 5
View larger version (28K):
[in this window]
[in a new window]
  		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{gamma} mRNA levels are increased in diseased compared with normal muscle (P = 0.001). A significant difference exists between PPAR{gamma} 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 ≤ 0.05; **, P ≤ 0.001 by Wilcoxon’s rank-sum test; #, P = 0.004 by Kruskal-Wallis test.

 
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. 5CGo). In contrast, c-kit mRNA was substantially decreased in diseased tissue (1.67 ± 1.11 vs. 8.73 ± 6.15, P = 0.007, Fig. 5DGo). 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 analyses. These findings suggest a potential role for these factors in the fibrosis of Müller’s muscle in GO.

We were unable to detect TSHr protein by IHC in either diseased Müller’s muscle (Fig. 3GGo), although the positive control (thyroid) was markedly positive (Fig. 3FGo). In contrast, TSHr mRNA was detected in muscle with real-time PCR. As shown in Fig. 5EGo, TSHr mRNA was found in both normal and diseased Müller’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. 5EGo).


   Discussion
Top
 Abstract
 Introduction
 Subjects and Methods
 Results
 Discussion
References
 
We demonstrate here for the first time that the degree of inflammatory cells infiltration and pathological changes of Müller’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{gamma}, myocardin, MCSF, and c-kit. Our current findings provide the rationale for the pathological changes observed in Müller’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 Müller’s muscle. Thus, although increased sympathetic activity, hypertrophy, and/or excessive contractility of Müller’s muscle have been reported as causes of upper lid retraction, we provide very strong evidence of pathological changes in Müller’s muscle that also contribute.

PPAR{gamma} 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 Müller’s muscle in GO. The steady-state level of PPAR{gamma} mRNA correlates well with the fractional fat composition in Müller’s muscle specimens. The function of increased PPAR{gamma} in Müller’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{gamma} (20, 21, 22). Thus, we propose that the increased adipose composition found in diseased Müller’s muscle might originate from two pathways. Adipocytes differentiation from fibroblasts, or transdifferentiation from myoblasts into adipocytes might occur. A mechanism involving PPAR{gamma} activation could explain either. Indeed, the expression of adipocyte fatty acid binding protein was also highly expressed in diseased Müller’s muscle corresponding to the expression pattern of PPAR{gamma} (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 whether they are activated by TGFß or a PPAR{gamma} agonist (23, 24).

To understand the mechanisms involved in reduced contractile elements in diseased Müller’s muscle, we studied the expression of the myocardin gene. This protein is critical for the transcriptional regulation of smooth muscle cell differentiation (25, 26, 27). We found that the level of myocardin expression was decreased in diseased tissue, suggesting that a reduction might result in attenuated Müller’s muscle differentiation in GO. We found no evidence of caspase-3-dependent apoptosis in diseased Müller’s muscle. Taken together, the reduced mass of Müller’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 Müller’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-1ß, TNF{alpha}, and IL-10, levels of which are increased in orbital adipose tissue, (30) promote the pathological features in Müller’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 cell-specific 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 Müller’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 Müller’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 Müller’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, 38, 39). In addition, there was no correlation between TSHr levels and any of the pathological changes or PPAR{gamma} mRNA levels. Our data suggest that TSHr may not play a role for the pathological changes occurring in diseased Müller’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 also prominent in these samples, suggesting an immune-mediated process in diseased Müller’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 Müller’s muscle in GO. Expression of PPAR{gamma}, 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, Müller’s muscle may also be considered an important target of autoimmunity in GO.


   Footnotes
 
This work was supported in part by Grants NSC 93-2752-B-002-009-PAE 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.).

First Published Online December 29, 2005

1 M.-J.S. and S.-L.L. contributed equally to this work. Back

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.

Received August 19, 2005.

Accepted December 20, 2005.


   References
Top
 Abstract
 Introduction
 Subjects and Methods
 Results
 Discussion
 References
 

  1. Taylor S 1986 Graves of Graves’ disease. J R Coll Phys London 20:298–300[Medline]
  2. Bartley GB, Gorman CA 1995 Diagnostic criteria for Graves’ ophthalmopathy. Am J Ophthalmol 119:792–795[Medline]
  3. Paschke R, Ludgate M 1997 The thyrotropin receptor in thyroid disease. N Engl J Med 337:1675–1681[Free Full Text]
  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[Free Full Text]
  6. Bahn RS 2003 Pathophysiology of Graves’ ophthalmopathy: the cycle of disease. J Clin Endocrinol Metab 88:1939–1946[Free Full Text]
  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[Medline]
  8. Putterman AM, Urist M 1972 Surgical treatment of upper eyelid retraction. Arch Ophthalmol 87:401–405[Abstract/Free Full Text]
  9. Bodker FS, Putterman AM, Laris A, Miletich DA, Volgel SM, Viana MAG 1997 The effect of hyperthyroidism on Müller’s muscle contractility in rats. Ophthal Plast Reconstr Surg 13:161–167[Medline]
  10. Jampolsky A 1970 What can electromyography do for the ophthalmologist? Invest Ophthalmol 9:570–599[Abstract/Free Full Text]
  11. Rootman J, Patel S, Berry K, Nugent R 1987 Pathological and clinical study of the Müller’s muscle in Graves’ ophthalmology. Can J Ophthalmol 22:32–36[Medline]
  12. Gaag RVD, Schmidt D, Koornneef L 1993 Retrobulbar histology and immunohistochemistry in endocrine ophthalmopathy. Dev Ophthalmol 25:1–10[Medline]
  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[CrossRef][Medline]
  14. Kagoshima T, Hori S, Inoue Y 1987 Qualitative and quantitative analyses of Müller’s muscle in dysthyroid ophthalmopathy. Jpn J Ophthalmol 31:646–654[Medline]
  15. Caruso RD, Postel GC 2002 Image annotation with Adobe Photoshop. J Digit Imaging 15:197–202[CrossRef][Medline]
  16. Lahm A, Uhl M, Lehr HA, Ihling C, Kreuz PC, Haberstroh J 2004 Photoshop-based 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 {gamma} inhibits adipocyte differentiation. J Biol Chem 275:1873–1877[Abstract/Free Full Text]
  18. Tontonoz P, Hu E, Spiegelman BM 1994 Stimulation of adipogenesis in fibroblasts by PPAR{gamma}2, a lipid-activated transcription factor. Cell 79:1147–1156[CrossRef][Medline]
  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[Abstract/Free Full Text]
  20. Hu E, Tontonoz P, Spiegelman BM 1995 Transdifferentiation of myoblasts by the adipogenic transcription factors PPAR{gamma} and C/EBP{alpha}. Proc Natl Acad Sci USA 92:9856–9860[Abstract/Free Full Text]
  21. 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[CrossRef][Medline]
  22. Holst D, Luquet S, Kristiansen K, Grimaldi PA 2003 Roles of peroxisome proliferator-activated receptors {delta} and {gamma} in myoblast transdifferentiation. Exp Cell Res 288:168–176[CrossRef][Medline]
  23. 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[Abstract/Free Full Text]
  24. 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[Abstract/Free Full Text]
  25. Suzuki T, Nagai R, Yazaki Y 1998 Mechanisms of transcription regulation of gene expression in smooth muscle cells. Circ Res 82:1238–1242[Free Full Text]
  26. 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[Abstract/Free Full Text]
  27. 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[Abstract/Free Full Text]
  28. 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[CrossRef][Medline]
  29. 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[Abstract]
  30. Kumar S, Bahn RS 2003 Relative overexpression of macrophage-derived cytokines in orbital adipose tissue from patients with Graves’ ophthalmopathy. J Clin Endocrinol Metab 88:4246–4250[Abstract/Free Full Text]
  31. Ashman LK 1999 The biology of stem cell factor and receptor c-kit. Int. J Biochem Cell Biol 31:1037–1051
  32. Linnekin D 1999 Early signaling pathways activated by c-kit in hematopoietic cells. Int J Biochem Cell Biol 31:1035–1074
  33. 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[CrossRef][Medline]
  34. 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[CrossRef][Medline]
  35. 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[Abstract/Free Full Text]
  36. Smith TJ 2003 Unique properties of orbital connective tissue underlie its involvement in Graves’ disease. Minerva Endocrinol 28:213–222[Medline]
  37. 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[Abstract/Free Full Text]
  38. 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[Medline]
  39. 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[Abstract]




This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Submit a related Letter to the Editor
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow Request Copyright Permission
Citing Articles
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Shih, M.-J.
Right arrow Articles by Chuang, L.-M.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Shih, M.-J.
Right arrow Articles by Chuang, L.-M.
Related Collections
Right arrow Thyroid
Right arrow Autoimmunity

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Endocrinology Endocrine Reviews J. Clin. End. & Metab.
Molecular Endocrinology Recent Prog. Horm. Res. All Endocrine Journals