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1 Department of Internal Medicine, University of Texas Southwestern Medical Center, Dallas, Texas 75390-8573
2 Department of Pathology, University of Texas Southwestern Medical Center, Dallas, Texas 75390-8573
3 Department of Molecular Biology, University of Texas Southwestern Medical Center, Dallas, Texas 75390-8573
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONDISCLOSURESREFERENCES |
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microarray analysis; biglycan; periostin
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONDISCLOSURESREFERENCES |
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Effective muscle regeneration requires a coordinated repair response involving the inflammatory system, capillary morphogenesis, and myogenic progenitor cell-mediated myogenesis to ultimately restore the architecture of the tissue without fibrosis (i.e., scar formation). This repair process is dependent on an orchestrated response between the inductive signals of cytokines, growth factors, and the extracellular matrix (ECM) (12, 16, 23, 24). Previous studies have established the critical role for ECM components during morphogenesis and tissue regeneration (15, 17, 20). Matrix degradation following injury initially releases growth factors that function as potent mitogenic signals for tissue-specific progenitor cells and the vascular endothelial cells (15, 17, 20). Additionally, at later stages of the regeneration phase, the ECM is capable of scavenging or inactivating these morphogens, resulting in the reestablishment of the tissue architecture. Although considerable progress has been made regarding our understanding of muscle development, the coordinated regenerative response of skeletal muscle is unclear. Applications using emerging technologies to examine the molecular response of muscle regeneration would enhance our understanding of this response and facilitate tissue engineering strategies that would have applications for patients with myopathies.
In the present study, we utilize cellular and molecular technologies to comprehensively examine skeletal muscle regeneration and further focus on the role of periostin (Osf2) and bigylcan (Bgn), which contribute to the ECM. The results of this study highlight the critical role of the ECM in skeletal muscle regeneration, which is a complex process involving thousands of genes, many of which are also expressed during the fetal program.
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EXPERIMENTAL PROCEDURES |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONDISCLOSURESREFERENCES |
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Reverse transcription, real-time, and semi-quantitative PCR.
Total RNA was isolated with TriPure isolation reagent (Roche, Basel, Switzerland) and reverse transcribed with SuperScript II (Invitrogen, Carlsbad, CA) to cDNA. Real-time PCR was performed with the iCycler iQ real-time PCR detection system (Bio-Rad Laboratories, Hercules, CA) and the QuantiTect SYBR Green PCR kit (Qiagen, Valencia, CA) (7, 8). The reaction was carried out in a final volume of 25 µl with 1 x SYBR Green PCR master mix, 0.3 µM each primer, and 20 ng cDNA for 50 cycles (20 s at 94°C to denature, 20 s at 60°C to anneal, and 30 s at 72°C to extend). The 75- to 100-bp extension was detected following each cycle and analyzed with the iCycler iQ software (Bio-Rad) for specificity, using melting curve, and threshold cycle (CT). Data are expressed with comparative CT method as an estimate of mRNA of target to reference control. Real-time PCR primers included the following: Ccnd1, forward 5' CGGATGAGAACAAGCAGACC 3' and reverse 5' GCAGGAGAGGAAGTTGTTGG 3'; Fn1, forward 5' TGTAGGAGAACAGTGGCAGAAA 3' and reverse 5' CAGGTCTACGGCAGTTGTCA 3'; Mmp3, forward 5' CCCAGGAAGATAGCTGAGGA 3' and reverse 5' CAACTGCGAAGATCCACTGA 3'; Myf5, forward 5' CTGTCTGGTCCCGAAAGAAC 3' and reverse 5' AAGCAATCCAAGCTGGACAC 3'; Myf6, forward 5' AATTCTTGAGGGTGCGGATT 3' and reverse 5' ATGGAAGAAAGGCGCTGAAG 3'; Pcna, forward 5' GCTTGGCAATGGGAACATTA 3' and reverse 5' CAGTGGAGTGGCTTTTGTGA 3'; and Rasa3, forward 5' ATTGATGGGGACCGTGAAAC 3' and reverse 5' GGGCCGTCATACACAGACTT 3'.
Semi-quantitative RT-PCR was performed with a final volume of 20 µl with variable cycle lengths as previously described for quantitative analyses. Conditions for these reactions include 20 s at 94°C to denature, 20 s at 60°C to anneal, and 30 s at 72°C to extend. Following the reaction, the amplicon (periostin, 92 bp; biglycan, 144 bp; 18S rRNA, 149 bp) was observed by using 1% agarose gel electrophoresis. Primer sequences are as follows: periostin, forward 5' CCCTTCCATTCTCATATA 3' and reverse 5' CCCTTCCATTCTCATATA 3'; biglycan, forward 5' CACCTGGACCACAACAAAA 3' and reverse 5' TCCGAATCTGATTGTGACCTA 3'; and 18S rRNA, forward 5' GGACCAGAGCGAAAGCATTTA 3' and reverse 5' TGCCAGAGTCTCGTTCGTTAT 3'.
Probe labeling and Affymetrix array hybridization.
Oligonucleotide array hybridizations were carried out according to the Affymetrix protocol summarized as follows. Total RNA was isolated from uninjured or cardiotoxin-injured gastrocnemius skeletal muscle at defined intervals and pooled from three animals as previously described. Five micrograms of total RNA was reverse transcribed using an oligonucleotide containing a T7 promoter with a poly-dT tail [T7(dT)24]. Ligase and polymerases were then added to produce the second strand of the single-stranded cDNA. Following precipitation, the double-stranded cDNA was converted to biotin-labeled cRNA by using the Enzo BioArray high-yield RNA transcript labeling kit (Enzo Biochem, New York, NY). The purified biotin-labeled cRNA was then fragmented by using Affymetrix fragmentation buffer for 35 min at 95°C. Labeled fragmented cRNA (15 µg) was then hybridized to the high-density oligonucleotide Murine Genome Array U74Av2 GeneChip (Affymetrix, Santa Clara, CA) (pooled RNA isolated from 3 animals/array). After 16 h of hybridization, the array was washed, stained, and scanned according to the manufacturer’s protocol.
Array analysis.
Array quality assessment and expression values were acquired with DNA-Chip Analyzer (dChip) model-based analysis of multiple arrays (n = 18 for modeling). This method utilizes invariant-set normalization and model-based expression indexes (MBEI) with standard error (SE) to measure accuracy. Comparative analysis was performed by using MBEI and SE to construct a 90% confidence interval (CI) of fold change (3, 4). Change was specified as significant if the lower 90% confidence bound of fold change was greater than or equal to 2 and the absolute difference of the group mean expression was greater than 100. Values were exported to GeneCluster for self-organizing map (SOM) clustering to determine common and unique expression profiles for sets of genes during muscle regeneration (27) (see the Supplemental Material, available at the Physiological Genomics web site).1
Gene ontology annotation as defined by the Gene Ontology Consortium (http://www.geneontology.org), either as biological process, molecular function, or cellular component, was performed with dChip.
Riboprobe synthesis.
IMAGE clones (403071, 425785) were sequence-verified and prepared for in vitro transcription following restriction enzyme digestion and gel isolation. Linearized template (500 ng) was transcribed by using either the T7 or SP6 RNA polymerase (Ambion, Austin, TX) with 7.0 µM [
-35S]UTP (1,000 Ci/mmol; Amersham, Piscataway, NJ) to produce the respective sense and antisense riboprobes (6, 25). The periostin riboprobe was 632 bp in length, and the biglycan riboprobe was 3.0 kb in length, for the sense and antisense probes (6, 25). The probes were purified with MicroSpin G-50 columns (Amersham), analyzed for integrity, and stored overnight at -80°C as previously described (25).
In situ hybridization.
In situ hybridization was performed as previously described (6, 25). Briefly, 5-µm paraffin sections of staged embryos or selected tissues (i.e., cardiotoxin-injured skeletal muscle or the mdx heart, diaphragm, and skeletal muscles) were cut, mounted on Vectabond-coated slides, dewaxed, permeabilized, acetylated, and hybridized at 70°C. Riboprobes were diluted in 50% formamide, 0.07 M NaCl, 20 mM Tris·HCl (pH 8.0), 5 nM EDTA (pH 8.0), 10 mM NaPO4 (pH 8.0), 10% dextran sulfate, 1x Denhardt’s, and 0.5 mg/ml tRNA (25). Following hybridization, the slides were rinsed with washes of increasing stringency, treated with RNase A (2 µg/ml, 30 min at 37°C), dehydrated, and dipped in K5 nuclear emulsion gel (Ilford). Autoradiographic exposure was undertaken for a 21-day period. In all cases, sections hybridized with the sense probe resulted in the absence of signal. Reference to all expression data presented in these studies pertains to the respective mRNA transcripts.
Microscopy and photomicrography.
Periostin and biglycan mRNA expression were visualized with a Leitz Laborlux-S microscope equipped with plan-apochromatic optics, standard bright-field condenser, and a Mears low-magnification dark-field condenser. Photomicrographs were obtained with an Optronics VI-470 CCD camera and Power Macintosh G3 with Scion Image 1.62 software.
Cell culture.
C2C12 myogenic cells (derived from satellite cells) were grown as monolayers in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 20% fetal bovine serum and antibiotics. Myotube differentiation was promoted by exposing 80% confluent myoblast cultures to differentiation medium [DMEM supplemented with 2% heat-inactivated horse serum, antibiotics, insulin, and transferrin as previously described (1, 9)]. Differentiation was assessed morphologically by the appearance of multinucleated myotubes. Monolayers of NIH 3T3 fibroblast cells were grown in DMEM supplemented with 10% fetal bovine serum. C2C12 myoblasts, myotubes, and NIH 3T3 fibroblasts were cultured in the presence and absence of supplemented transforming growth factor-ß1 (TGFB1, 5 ng/ml; Sigma, St. Louis, MO). At specified time periods, medium was removed from the respective cell populations, the monolayers of cells were rinsed, and RNA was harvested for RT-PCR analysis of gene expression.
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RESULTS |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONDISCLOSURESREFERENCES |
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75–90% of the muscle is destroyed in the mouse model (7, 8, 12). Following this chemical-induced injury, a well-orchestrated cellular response may be observed by using standard histological techniques. During the acute postinjury period (
6 h), myofibers become hyalinized and vacuolated and their nuclei are pyknotic or lysed (Fig. 1, A and B). Interstitial edema, myonuclear dropout, and the presence of a neutrophilic (PMN) infiltrate further characterize this initial phase (Fig. 1A). Microvascular thromboses are evident (Fig. 1B), and myogenic progenitor cells assume an activated or primed state (not shown) (7, 8). Twelve hours following injury, myofibers lyse, resulting in the production of a protein-rich edema (Fig. 1C). The inflammatory response consists initially of PMNs (Fig. 1D), and at later stages macrophages that phagocytose necrotic myofibers (2–5 days following injury). This chemical-induced injury lacks a prominent lymphocytic response using standard histological techniques. Following activation, myogenic progenitor cells proliferate (days 2–4) and characterize the regenerative phase, forming crescent-like structures around necrotic or damaged myofibers (Fig. 1E). Newly regenerated myofibers are easily identified as small basophilic, centronucleated myofibers (Fig. 1, E–G). A prominent remodeling of the ECM is clearly evident early during the repair process, which persists and matures throughout the repair period (Fig. 1, E–G). Ultimately, the biomatrix organizes and matures corresponding to the completion of muscle regeneration resulting in preservation of the muscle architecture (Fig. 1, G and H). No evidence of residual scar formation or persistent fibrosis is observed with this extensive injury.
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The analysis of the transcriptional profiling experiments revealed discrete molecular responses associated with sequential stages following cardiotoxin-induced muscle injury (Fig. 2). Although the molecular response involving the greatest number of genes occurred within 2 days of injury (1,225 genes dysregulated 
2 fold), the acute period following injury was associated with an induction of inflammatory and signal transduction genes (Fig. 2). Transcripts that encode nuclear proteins were increased within 0.5 days of injury followed by an induction of cell cycle and signal transduction genes at 2 days (Fig. 2). The expression profile of cell cycle genes corresponds to a marked increase in the proliferative capacity of the myogenic progenitor cell population at this time (i.e., Pcna expression, see Table 1 and clusters 10, 11, 15–17, and 21–23 in the Supplemental Material). Increased ECM/cell adhesion-associated gene expression observed at day 5 and thereafter (days 7 and 14) corresponds to the reorganization/maturation of the ECM and the differentiation of the myogenic progenitor cells (Fig. 1, C–H) (Table 1 and Supplemental Material). This differentiation phase, corresponding to the maturation of both the ECM and the newly regenerated myofibers, ultimately results in regenerated skeletal muscle lacking any evidence of residual scarring or fibrosis.
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In addition, the confirmation of the microarray results of selected candidate genes was undertaken by using quantitative real-time RT-PCR analysis (Fig. 3). As shown in Fig. 3, the real-time RT-PCR analysis confirmed the array analyses and further verified the stage-specific expression pattern for transcripts associated with the ECM remodeling (Mmp3 and Fn1), cell cycle regulation (Ccnd1 and Pcna), myogenesis (Myf6 and Myf5), and cellular signaling (Rasa3). These results, in combination with previously published data, provide further support for the molecular analysis undertaken in the present study (11, 18, 19).
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Periostin was expressed early during murine embryogenesis. In the E9.5 embryo, periostin was restricted to the cardiac cushion, vascular outflow tract, amnion, vitelline artery, and vein (Fig. 4A). Persistent expression was observed in the cardiac cushion, outflow tract, peripheral nerves (but not dorsal root ganglia), head-fold mesenchyme, and the periaortic mesenchyme at E11.5 (Fig. 4B). The expression pattern broadens during the midgestational period (E13.5–E15.5) to primarily include the mesenchyme as well as the skin, perichondrium, endocardium, perinephric stroma, choroid plexus, and diaphragm (Fig. 4, C–G). These results support the conclusion that periostin is expressed principally in mesenchymal tissues.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONDISCLOSURESREFERENCES |
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In the present study, we pursued a comprehensive cellular and molecular analysis of the remodeling process following chemical-induced muscle injury. These results clearly establish the presence of phases associated with the inflammatory process, myogenic progenitor cell activation, and proliferation, myogenic differentiation, growth factor, or mitogenic stimuli and the contribution of the ECM. We further examined two candidate genes in detail to define the contribution of the biomatrix during muscle development and injury/regeneration.
Molecular mechanisms of muscle injury and regeneration.
Analysis of other vertebrate tissues that display increased regenerative capacity emphasizes common or shared molecular and cellular programs that characterize the injury-repair process and underscores the importance of the ECM in this process (15, 17, 20). Regeneration of hepatic parenchymal tissue following an extensive injury or partial hepatectomy highlights stages of the repair process, which occurs over a 2- to 3-wk period. The acute phase following an extensive hepatic injury is characterized by the induction of an immediate early gene program (Fos, Jun, JAK/STAT, NF-
B, etc.), a pronounced inflammatory response with the release of cytokines and mitogenic signals, which concludes with the activation and increased proliferative capacity of hepatic precursor cell populations (i.e., oval cells). Growth factors (Igf1, Igf2, Hgf, Fgf1, Tgfb1) in combination with cytokines (Il1, Il6, Tnf) coordinately promote cellular proliferation and the remodeling of the hepatic matrix composition which characterize the second stage of hepatic repair (17). Hepatic differentiation and matrix maturation resulting in preserved architecture characterize the final stages of the repair process. Notably, regeneration of hepatic tissue undergoes similar discrete phases of regeneration as observed in the present study associated with regenerating skeletal muscle.
Intense interest has focused on the regulatory mechanisms of the hepatic precursor cell population and the interaction of key signals (i.e., Hgf, Fgf, and Tgfb1) during these stages, but the signals that regulate the termination of the regenerative response are incompletely defined (15, 17, 20). One candidate factor that may initiate the termination signal for liver regeneration is Tgfb1, as it has been previously shown to inhibit hepatocyte proliferation and modulate the ECM of the regenerating liver. Although limited information is available regarding the termination signal for regeneration, it is highly probable that multiple factors (growth factors, cytokines, and ECM restoration) may deliver in aggregate a set of signals resulting in the termination of the repair process (17). A recent report further examined the molecular response during bone repair and complements the studies of hepatic regeneration and our studies of muscle regeneration. Specifically, transcriptional changes associated with bone formation and the ECM follow a similar expression profile observed in the present study of muscle regeneration, illustrating the stage-specific and essential role of the ECM during regeneration of different tissues. Although the molecular analysis utilized in the present study was performed on whole skeletal muscle tissue, the expression profiles of cell cycle, ECM, and muscle differentiation genes are consistent with array analysis performed on differentiating C2C12 myoblasts (18). The current study further focused on two ECM proteins that were robustly expressed during the muscle repair process.
Remodeling of the ECM during muscle injury and regeneration.
Periostin (also named "osteoblast-specific factor 2," or Osf2) was isolated using differential screening techniques and previously believed to be restricted to bone (26). Previous studies reported that periostin was a secreted protein that functioned in osteoblast recruitment, attachment, and spreading (13). Moreover, periostin has been shown to be regulated by Tgfb1 in osteoblast cells (13). More recently, embryological expression studies of periostin revealed that it was also expressed in the cardiac cushion (14). In the present study, we observed periostin to be robustly upregulated during the muscle repair process (>80-fold upregulation) using oligonucleotide array analysis.
Using in situ hybridization techniques, we observed periostin to be more broadly expressed in the head mesenchyme, periaortic mesenchyme, diaphragm, periosteum, skin, and cardiac cushion during mouse embryogenesis. Furthermore, periostin is robustly expressed in the remodeling ECM following muscle injury and in the regenerating diaphragm and heart of the mdx mouse model. In addition, we further establish that periostin is expressed in C2C12 myoblasts and regulated in this cell line by TGFB1. TGFB1 is a multifunctional, contextual cytokine that has previously been reported to function in modulating the tissue repair response through the regulation of the fibroblast cell population and matrix maturation as well as a regulator of cell migration and chemotaxis in regenerating tissues such as the skin (15). Further studies will focus on the complex interaction between growth factors and the biological role of periostin as important regulators of muscle regeneration.
Biglycan is an example of a secreted multifunctional leucine-rich proteoglycan that functions in matrix organization and the modulation of various growth factors (28, 29). Biglycan exhibits predominantly a pericellular location, and via its two glycosaminoglycan chains binds to -dystroglycan located in the skeletal muscle plasmalemma (2). The implications of the interaction of biglycan with -dystroglycan have not been elucidated, but one possibility is that it may serve to localize signaling molecules to the plasmalemma. For example, biglycan has been observed to bind growth factors or cytokines (i.e., Tgfb1, Tnf, etc.) and may modulate the presentation or the sequestration (i.e., inactivation) of these growth factors, thereby affecting muscle and/or matrix remodeling (28, 29). Additionally, biglycan may function in the matrix assembly through the binding of this proteoglycan to the collagen network (i.e., collagen VI and possibly collagen type I) (22, 30) or the elastin microfibril network (i.e., tropoelastin) (21). Furthermore, studies utilizing gene disruption strategies report that biglycan-deficient mice have perturbed bone remodeling as they manifest an osteoporosis-like phenotype characterized by decreased bone mass, suggesting that biglycan is a positive regulator of bone formation (31). Future studies using the biglycan-deficient mouse model will be of interest to further define a broader role for this proteoglycan during muscle regeneration and as a direct or indirect regulator of the myogenic progenitor cell population.
In the present study, we establish that biglycan is expressed in embryonic mesenchymal tissues early during mouse development and has an overlapping expression pattern with periostin in the embryo at midgestational ages. Similarly, biglycan is robustly upregulated during muscle regeneration and in the dystrophic mdx diaphragm and heart consistent with its role in regeneration and muscle remodeling. Using microarray analysis, we observed that Tgfb1 expression precedes that of biglycan and induces expression in the NIH 3T3 fibroblast cell line. These results further establish the dynamic role of the ECM during development and regeneration and underscores the coordinated regulation of myogenesis by growth factors and the ECM for efficient repair during the regenerative process.
In summary, the results of these studies underscore the molecular complexity associated with muscle regeneration and the essential role of the ECM proteins. Furthermore, these studies provide a foundation for future experiments that will enhance our understanding of injury/regenerative mechanisms and have applications for the treatment of chronic debilitating diseases.
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DISCLOSURES |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONDISCLOSURESREFERENCES |
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ACKNOWLEDGMENTS |
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Present address of T. J. Hawke: School of Kinesiology and Health Science, York University, Toronto, Ontario, Canada M3J 1P3.
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FOOTNOTES |
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Address for reprint requests and other correspondence: D. J. Garry, NB11.118A, 5323 Harry Hines Blvd., UT Southwestern Medical Center, Dallas, TX 75390-8573 (E-mail: daniel.garry{at}utsouthwestern.edu).
10.1152/physiolgenomics.00056.2003.
1 The Supplementary Material for this article (a figure and a table) is available online at http://physiolgenomics.physiology.org/cgi/content/full/00056.2003/DC1. 
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REFERENCES |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONDISCLOSURESREFERENCES |
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-dystroglycan and is upregulated in dystrophic muscle. J Cell Biol 148: 801–810, 2000. interacts with biglycan and decorin. FEBS Lett 530: 124–128, 2002.[Medline]
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