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PHYSIOLOGY, ENDOCRINOLOGY, AND REPRODUCTION |
Department of Animal Sciences, Ohio Agricultural Research and Development Center, The Ohio State University, Wooster 44691
2 Corresponding author: velleman.1{at}osu.edu
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Key Words: chicken • decorin • extracellular matrix • skeletal muscle • transforming growth factor-β1
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Communication between the extracellular matrix (ECM) and cells plays a pivotal role in the regulation of muscle cell proliferation and differentiation. The connective tissue ECM is defined as a dynamic network of collagens, proteoglycans, and glycoproteins that are secreted outside the cell and required for sustaining cellular behavior (Scott, 1995). In skeletal muscle, the ECM is a major component in the intramuscular connective tissue. There are 3 layers of connective tissue surrounding muscle, including the endomysium, the perimysium, and the epimysium. The endomysium separates individual muscle fibers. The perimysium surrounds bundles of muscle fibers, and the epimysium sheathes the whole muscle. Collagens are the major protein in the ECM present in the connective tissue. Types I, III, IV, V, and VI collagens have been identified in skeletal muscle (Nishimura et al., 1997). The perimysium contains types I and III collagen, whereas the endomysium contains types I, III, IV, and V collagen (Light and Champion, 1984).
The proteoglycans are a diverse family of macromolecules containing a core protein and at least one, but frequently more, covalently attached glycosaminoglycan (GAG) side chains (Hardingham and Fosang 1992). The proteoglycans are involved in the regulation of gene expression, cell proliferation, migration, adhesion, and differentiation, which are all essential for muscle development and growth (Yanagishita, 1993). The presence of the proteoglycans is required for skeletal muscle cells to respond to growth factors. Through interactions with growth factors such as transforming growth factor-β (TGF-β; Yamaguchi et al., 1990), fibroblast growth factor 2 (Rapraeger et al., 1991), and hepatocyte growth factor (Allen et al., 1995), the ECM can regulate the ability of skeletal muscle cells to proliferate or differentiate.
Decorin, a member of the small leucine-rich proteoglycans, is distributed in the ECM of various tissues (Brennan et al., 1984; Day et al., 1987), including skeletal muscle (Eggen et al., 1994; Velleman et al., 1996). Decorin consists of a core protein of approximately 45 kDa and a single covalently attached chondroitin or dermatan sulfate chain (Krusius and Ruoslahti, 1986). Decorin interacts with a variety of proteins that are involved in matrix assembly and regulation of cell attachment, migration, proliferation, and differentiation (Iozzo, 1999). Decorin binds to the collagen fibrils as a spacer during the lateral assembly of collagen fibers, maintaining collagen fibrillogenesis and modifying the morphology of the fibrils (Weber et al., 1996). Decorin is also a regulator of many cellular properties, including cell attachment, migration, proliferation, and differentiation, mediated either through its ability to interact with growth factors or by upregulating cyclin-dependent kinase inhibitors such as p21 and p27 (Iozzo, 1999). The binding of decorin to growth factors modulates the activities of growth factors and subsequently influences cell proliferation and differentiation. Decorin has a high affinity for TGF-β (Hildebrand et al., 1994). The decorin core protein binds to TGF-β, neutralizes TGF-β activity, and modulates TGF-β-dependent cell growth stimulation or inhibition (Yamaguchi et al., 1990; Kresse et al., 1994).
Transforming growth factor-β1, a member of the TGF-β superfamily, is a strong inhibitor of both myoblast proliferation and differentiation (Allen and Boxhorn, 1987). Decorin can bind directly to TGF-β1 and modulate the responsiveness of myoblasts to TGF-β1. In vitro myoblasts not expressing decorin show a decrease in their responsiveness to TGF-β1 (Riquelme et al., 2001). This indicates that the expression of decorin may mediate the ability of myoblasts to respond to TGF-β1 signaling, thus regulating cell proliferation and differentiation.
It is clear that the expression of TGF-β1 and decorin is involved in the regulation of muscle growth and development. However, how TGF-β1 affects decorin and regulates skeletal muscle development and growth is not well understood at this time. The present study focused on the effect of TGF-β1 on decorin expression and skeletal muscle growth and development in vivo during both embryonic and posthatch development.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Specific-pathogen-free chickens used in the present study were from flocks maintained by the Ohio Agricultural Research and Development Center at The Ohio State University. The chicken fertile eggs at embryonic day (ED) 3 were injected with 100 ng of recombinant TGF-β1 (Pepro Tech Inc., Rocky Hill, NY) brought to a 100-µL volume in PBS or only PBS as a control. Embryonic d 3 represents a developmental time critical for cell migration and tissue formation and is also a period of time that allows the incorporation of material through the chorioallantoic membrane (Zwilling 1959). The eggs were incubated in a 37.5°C and 60% humidity egg incubator (NatureForm Hatchery Systems Inc., Jacksonville, FL). In brief, the eggshells were drilled with a Dremel tool (Dremel, Racine, WI), resulting in a 1-mm-diameter hole, and then 100 ng of TGF-β1 in 100 µL of PBS or only the PBS control was injected into the air sack area of the eggs at 3 ED with a 26-gauge 1/2 needle attached to a 1-mL syringe, adapted from the method of Zwilling (1959). A total of 150 eggs were randomly split into 5 groups corresponding to the sampling times: 60 eggs for 10 ED, 30 eggs for 17 ED, 20 eggs for 1 d posthatch, 20 eggs for 1 wk posthatch, and 20 eggs for 6 wk posthatch. Each group was randomly split in half for either the control or TGF-β1 treatment. The pectoralis (p. major) muscle from both the control and the TGF-β1-treated group was harvested at 10 ED, 17 ED, 1 d, 1 wk, and 6 wk posthatch.
Total RNA Extraction and cDNA Synthesis
The p. major muscle from each bird was removed, quick-frozen in liquid nitrogen, and stored at –70°C until use. Total RNA was extracted from the tissue samples by using Trizol (Invitrogen, Carlsbad, CA) according to the protocol of the manufacturer. The samples were digested with DNase (Promega, Madison, WI) at 37°C for 30 min before the reverse-transcription reaction. Reverse transcription of total RNA to a cDNA was conducted by using Moloney murine leukemia virus reverse transcriptase (M-MLV, Promega). In brief, an RNA-primer mix [1 µg of total RNA, 1 µL of 50 µM Oligo d(T)20, and nuclease-free water up to 13.5 µL] was incubated at 80°C for 3 min, followed immediately by an incubation on ice. The reaction mix [5 µL of 5x First-Strand buffer, 1.25 µL of 10 mM deoxynucleotide triphosphate mix, 0.5 µL of RNasin (40 units/µL), 1 µL of Moloney murine leukemia virus reverse transcriptase (200 units/µL), and nuclease-free water up to 11.5 µL] was added to the mixture. The complete reaction mixture was incubated at 55°C for 60 min, and then heated at 90°C for 10 min for inactivation.
Real-Time Quantitative PCR
Real-time quantitative PCR was performed by using the DyNAmo Hot Start SYBR Green qPCR kit (New England Biolabs, Beverly, MA). The PCR reaction consisted of 2 µL of the reverse-transcription reaction mixture diluted with 25 µL of nuclease-free water, 10 µL of 2x master mix provided by the manufacturer, 250 nM each of the forward and reverse primers, and nuclease-free water up to 20 µL. The decorin forward primer was 5'-AAGGTTCTGCCTGGAGTTGA-3' and the reverse primer was 5'-TTGGCACTCTTTCCAGACCT-3' (GenBank accession no. X63797). The glyceraldehyde-3-phosphate dehydrogenase (GAPDH) forward primer was 5'-GAGGGTAGTGAAGGCTGCTG-3' and the backward primer was 5'-CCACAACACGGTTGCTGTAT-3' (GenBank accession no. U94327). The specificity of the primers was confirmed by DNA sequence analysis of the amplified PCR product. The real-time PCR amplification was done in a DNA Engine Opticon 2 real-time system (MJ Research, Reno, NV). For all the amplified genes, the cycling program was initiated with a hot-start step at 95°C for 15 min. The cycling program for GAPDH and decorin was 94°C for 30 s, 55°C for 30 s, and an extension at 72°C for 30 s for 34 cycles, with a final elongation of 72°C for 5 min. The PCR products were purified by agarose gel electrophoresis by using a QIAquick Gel Extraction Kit (Qiagen, Valencia, CA). All the sample concentrations fell within the values of the standard curves. The amount of sample cDNA for each gene was interpolated from the corresponding standard curve. The expression of decorin was normalized to GAPDH expression.
Western Blot Analysis
Protein was extracted from the p. major muscle tissue samples by homogenization in Cytobuster protein extraction reagent (Novagen, San Diego, CA). The samples were digested with 1 unit/mL of chondroitinase ABC (Sigma-Aldrich, St. Louis, MO) in 200 µL of 250 mM Tris-HCl, 350 mM NaCl, and 0.05% BSA (pH 8.0) at 37°C overnight to remove the chondroitin-4 and chondroitin-6 sulfate chains, and dermatan sulfate chains attached to the decorin core protein. The reaction was stopped by freezing the samples at –20°C. The undigested samples were directly frozen and stored at –70°C. The protein concentration of each sample was measured by the method of Bradford (1976). The samples were then resuspended in 50 µL of 1% SDS, 10% glycerol, 1% β-mercaptoethanol, 0.1 M Tris-HCl, and 0.1% (wt/vol) bromophenol blue (pH 6.8), and heated at 95°C for 5 min. Eighty micrograms of the protein samples were applied to a 4 to 12% gradient SDS-polyacrylamide gel with a 3.2% polyacrylamide in the stacking gel (Laemmli, 1970). Electrophoresis was run in a 25 mM Tris-HCl, 1.90 mM glycine, and 0.1% SDS (pH 8.4) running buffer at a 20-mA constant current while the samples were in the stacking gel and at 30 mA during the separation phase. After electrophoresis, the proteins were electrophoretically transferred to a polyvinylidene difluoride (Millipore, Bedford, MA) membrane at a 25-mA constant current for 2 h at room temperature in transfer buffer [25 mM Tris-HCl, 192 mM glycine, and 10% (vol/vol) methanol (pH 8.3)] according to the method of Towbin et al. (1979). After the transfer, the membrane was blocked in 5% nonfat dry milk in Tris-buffered saline (20 mM Tris-HCl and 500 mM NaCl, pH 7.5; TBS) for 1 h. After blocking, the membrane was incubated in chicken decorin monoclonal antibody CB-1 (Developmental Studies Hybridoma Bank at the University of Iowa; 1:3,000 in 5% nonfat dry milk in TBS) for 2 h at room temperature and then washed 3x in 1x TBS and 0.05% Tween 20 (TBS-T) for 5 min each. The blot was then incubated in alkaline phosphatase conjugated goat anti-mouse IgG (1:10,000 in 5% nonfat dry milk in TBS-T) for 2 h at room temperature. The membrane was washed 2x in TBS-T and 1x in TBS, with 5 min per wash. The membrane was then incubated with a chemiluminescent alkaline phosphatase substrate detection reagent (Millipore) for 5 min in a dark room, and placed in an x-ray film cassette containing chemiluminescent BioMax film (Kodak, Rochester, NY). The film was developed by using a Konica Medical Film Processor (Konica, Nishishinjuku 1-Chome, Shinjuku-ku, Tokyo, Japan).
Histochemistry
Pectoralis major muscle tissue was dissected from 17 ED embryos, 1 d, and 6 wk posthatch birds. The sample was obtained by carefully dissecting approximately 0.5 cm of the muscle, following the orientation of the muscle fibers for a length of 3 cm. The ends of the muscle sample were tied to wooden applicator sticks by surgical thread before removal to prevent muscle contraction. Each p. major sample was placed in 10% (vol/vol) buffered formalin fixative (pH 7.0) at 4°C for at least 17 h. After fixation, the sample was dehydrated through a graded series of alcohols, as described previously by Jarrold et al. (1999), and cleared in Pro-par Clearant (Anatech Ltd., Battle Creek, MI) for 1 h with one change at 30 min, and infiltrated with paraffin at 55°C for 4 h with one change at 30 min by using a Leica TP1020 Tissue Processor (Leica, Nussloch, Germany). The samples were then embedded in paraffin, cross-sectioned into 5-µm sections, and mounted on Superfrost Plus slides (Fisher, Pittsburgh, PA). Before staining with hematoxylin and eosin, the muscle tissue sections were incubated at 55°C for 30 min and then rehydrated for 10 min in Pro-par Clearant, 2 min in 100% ethanol, 2 min in 95% ethanol, 2 min in 70% ethanol, 2 min in 50% ethanol, and 2 min in distilled water. After rehydration, the slides were placed in hematoxylin (Fisher) for 4 min. The sections were then rinsed in gentle running tap water for 10 min and transferred to eosin (0.5 g of eosin and 2.5 mL of glacial acetic acid in 500 mL of 70% ethanol) for 2 min. After staining, the slides were dehydrated back through the graded series of alcohols and Pro-par Clearant. The stained sections were analyzed for muscle morphology with an Olympus XI 70 microscope (Olympus America, Inc., Melville, NY) equipped with an Olympus Optronics camera (Olympus America, Inc.). The stained sections were analyzed for the width of the perimysium, and number of myofibers within a 52-µm2 area at 1 d posthatch or within a 35-µm2 area at 6 wk posthatch by using Image Pro software (Media Cybernetics, Silver Spring, MD). Each slide from each bird contained a minimum of 4 sections. From each slide, 20 measurements were taken for each trait analyzed.
Immunohistochemistry
Pectoralis major muscle tissue sections were prepared as described previously. After rehydrating, the sections were digested with 200 µL of chondroitinase ABC (0.425 units in 250 mM Tris-HCl, 350 mM NaCl, 0.05% BSA, pH 8.0) per slide at 37°C for 4 h in a slide moat (Boekel Scientific, Feasterville, PA). The slides were washed with 1x TBS for 5 min. After washing, the sections were covered with 10% goat serum (Gibco BRL, Grand Island, NY) in 1x TBS with 1% BSA and incubated for 1 h at room temperature. At the end of the incubation, the goat serum was removed and the slides were then covered with 200 µL of CB-1 (1:100 in 1x TBS containing 1% BSA; Lennon et al., 1991) at 37°C for 2 h. Negative control slides that did not include the CB-1 antibody were also processed in the 1x TBS solution containing 1% BSA. The slides were then washed 2x for 10 min each in 1x TBS and once for 10 min in 1x TBS containing 0.5% BSA. After washing, the slides were incubated at room temperature for 2 h in a 1:200 dilution of goat anti-mouse rhodamine (Chemicon International Inc., Temecula, CA) in the 1x TBS containing 1% BSA and incubated at room temperature in dark for 2 h. The slides were washed 2x for 5 min in 1x TBS and then mounted in aqueous Gel/Mount mounting media (Biomeda, Foster City, CA) and covered with coverslips (Fisher). The immunolocalization of decorin was observed with an Olympus XI 70 microscope and recorded with an Optronics digital camera.
Statistical Analysis
Statistical analysis was performed by using Student’s t-test to determine the difference between the control and TGF-β1-treated groups in mRNA expression, the perimysium space at each sampling time, and the number of myofibers in a 52-µm2 area at 1 d and in a 35-µm2 area at 6 wk posthatch. Differences were considered significant at P < 0.05.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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At 17 ED, the muscle fiber bundles were surrounded by the perimysium, which was observed as large gaps between the bundles. The endomysium separated the individual muscle fibers and was observed as small spaces surrounding single muscle fibers (Figure 1A
). In the p. major muscle from 1 d posthatch birds, the muscle fiber bundles were distinct, with a well-defined perimysium (Figure 1C). In the 6 wk p. major muscle, the muscle fibers were in tight proximity to each other within a muscle fiber bundle (Figure 1E). The width of the perimysium was increased from 17 ED to 6 wk posthatch (Figure 2A).
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). After hatch, the width of the perimysium decreased from 1 d to 6 wk posthatch. At 1 d and 6 wk posthatch, the width of the perimysium was decreased by TGF-β1 in the p. major muscle, compared to the control (Figure 1C, 1D, 1E, and 1F). The width of the perimysium surrounding the muscle fiber bundles was gradually increased from embryonic development through the posthatch stages. The TGF-β1 treatment significantly reduced the perimysium space at 17 ED, 1 d, and 6 wk posthatch (Figure 2A). The number of myofibers was significantly decreased in the TGF-β1-treated p. major muscle at both 1 d and 6 wk posthatch, compared to the control (Figure 2B and 2C). Effect of TGF-β1 on Decorin Expression in Skeletal Muscle
The mRNA expression of decorin was greater in the p. major muscle at 10 ED compared to 17 ED (Figure 3A). After hatch, decorin expression decreased continuously through 6 wk posthatch. At 1 wk posthatch, decorin expression was decreased more than at 6 wk posthatch (Figure 3B). The TGF-β1-treated p. major muscle had a decorin expression pattern similar to the control muscle, except at 10 ED, when the expression of decorin was significantly decreased by TGF-β1 compared to the control (Figure 3A and 3B). No significant difference was detected between the TGF-β1-treated and control groups from 17 ED to 6 wk posthatch (Figure 3A and 3B).
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). No decorin protein was detected in the 10 ED p. major muscle. From 17 ED to 1 d post-hatch, decorin protein was highly synthesized in the p. major muscle. At 1 and 6 wk posthatch, decorin protein was not detected in the p. major muscle (data not shown). The TGF-β1 treatment reduced decorin protein synthesis by 10% in the p. major muscle at 17 ED and by 30% at 1 d posthatch (Figures 4A and 4B). After chondroitinase ABC digestion, the core protein of decorin was approximately 45 kDa at all the developmental stages analyzed (Figure 4C). The β-actin protein levels were decreased at 1 d posthatch, compared to 10 and 17 ED (Figure 4A and 4C).
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In 17 ED p. major muscle, decorin was localized in both the perimysium and endomysium connective tissue layers (Figure 5A). At 1 d posthatch, decorin was mainly localized in the perimysium layer (Figure 6A). In the 6 wk posthatch muscle, the localization of decorin was observed only in the perimysium, but not in the endomysium, and the intensity of decorin staining was much less compared to earlier developmental stages (Figure 7A).
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). The immunostaining of decorin in the TGF-β1-treated muscle was much lower than in the control at 17 ED. At 1 d posthatch, the TGF-β1 treatment reduced the density of decorin in the perimysium, compared to the control. No decorin was detected in the endomyosium in the TGF-β1-treated muscle (Figure 6A and 5C). In the p. major muscle at 6 wk posthatch, the intensity of decorin staining detected in the TGF-β1-treated muscle was less compared to the control muscle (Figure 7A and 7C). |
DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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In addition, the expression of the ECM components changes during muscle growth and development. The mRNA expression of the proteoglycan decorin was initially high at 10 ED in p. major muscle and then decreased dramatically. After hatch, decorin expression decreased from 1 d to 6 wk posthatch. Velleman et al. (1996, 1997) reported that the synthesis of chicken decorin and turkey decorin decreased with skeletal muscle development. This expression pattern is similar to that reported for cattle (Nishimura et al., 2002) and rats (Nishimura et al., 2007). Interestingly, the synthesis of decorin protein was not detected until 17 ED decorin amounts were elevated and then decreased after hatch. The synthesis of the decorin core protein appeared with a pattern similar to the expression of the total decorin protein.
Changes in decorin expression during development indicate a critical role of decorin in the regulation of muscle differentiation and development. The mRNA expression of decorin was greater at 10 ED in the p. major muscle. However, the decorin protein amount was very low, which indicates that decorin mRNA may not be translated into protein in muscle until the later stages of embryonic development. This suggests a developmental change in the ECM environment surrounding the myofibers.
During embryonic development, the intramuscular connective tissue, including the endomysium and the perimysium, develops dramatically with muscle fiber formation (Nishimura et al., 2002). The ECM components, including collagen in the endomysium and the perimysium, are upregulated to support the muscle fibers and fiber bundles. Decorin regulates collagen fibrillogenesis and stabilizes collagen fibers in the connective tissue (Weber et al., 1996; Danielson et al., 1997). Therefore, the increased decorin protein amount may contribute to the regulation of collagen fibrillogenesis during intramuscular connective tissue development.
The proteoglycan decorin contains a single chondroitin or dermatan sulfate chain covalently attached to the core protein. The size of the GAG chain varies during development. In the current study, the decorin protein was immunostained by a decorin monoclonal antibody as a diffuse band ranging from 90 to 120 kDa at 17 ED. After hatch, the size of the decorin protein slightly shifted to a lower molecular weight compared to that at 17 ED. It is common for the proteoglycans to appear as diffuse bands in SDS-PAGE because of various GAG chains. The difference in the molecular weight of decorin represents a difference in the molecular weight of the GAG chain, which varies at different developmental stages. Using Western blot analysis, Nishimura et al. (2002) demonstrated that the molecular weight of decorin ranges from 110 to 150 kDa in 2.5-mo-old bovine fetuses and is reduced from 110 to 120 kDa in 9-mo-old fetuses. The variation in molecular weight of decorin suggests a developmental regulation of GAG chain size. The GAG chain attached to decorin core protein has been shown to be involved in the stabilization of the collagen fibril matrix (Scott and Thomlinson, 1998). Nishimura et al. (2002) demonstrated that the longer decorin GAG chain may create space between collagen fibrils, which is likely involved in the development of intramuscular connective tissue. The current study showed that decorin had a molecular size change, which suggests that changes in the decorin GAG chain occur during chicken skeletal muscle development. Decorin may contain 1 or 2 types of GAG chains, chondroitin or dermatan sulfate, depending on different species and type of tissues. However, it is still not clear which type of GAG is present, or whether both types of GAG are present, in the skeletal muscle and why the GAG chain changes in size with muscle development.
The expression of decorin is known to be regulated by TGF-β1 in many cell types (Bassols and Massagué, 1988; Border et al., 1990; Kähäri et al., 1991; Heimer et al., 1995; Li et al., 2006). However, reports on how TGF-β1 regulates decorin expression remain unclear, depending on the cell or tissue type and assay conditions. Border et al. (1990) demonstrated that TGF-β1 increased decorin synthesis in mesangial cells and rat liver cells. In cultured myocardium fibroblasts, TGF-β1 upregulated decorin protein synthesis (Heimer et al., 1995). In contrast, decorin core protein synthesis is markedly decreased by the presence of TGF-β1 in cultured human skin fibroblasts (Kähäri et al., 1991). Li et al. (2006) showed that decorin mRNA expression was reduced by TGF-β1 during chicken myogenic satellite cell differentiation. The current study showed that the TGF-β1-treated muscle had a significant decrease in decorin mRNA expression at 10 ED and that decorin protein amounts were reduced in the TGF-β1-treated muscle at both 17 ED and 1 d posthatch. The immunostaining of decorin in the p. major muscle was decreased by TGF-β1 from 17 ED through 6 wk posthatch. These data suggest that the expression of decorin is regulated by a cell-type-specific TGF-β1-mediated mechanism.
In addition, the perimysium space was dramatically decreased in the TGF-β1-treated p. major muscle compared to the control muscle from embryonic muscle development through the posthatch stages. Decorin was reduced by TGF-β1 in both the endomysium and perimysium from 17 ED to 6 wk posthatch. Danielson et al. (1997) demonstrated that decorin knockout mice had abnormal collagen morphology. Therefore, the reduction in decorin may cause the irregular formation of collagen fibrils, leading to decreased endomysium and perimysium space.
Transforming growth factor-β1 can also regulate collagen expression (Ignotz and Massagué, 1986; Varga et al., 1987). The effect of TGF-β1 on the expression of different types of collagen may contribute to the reduction of intramuscular connective tissue space. Further study is needed to elucidate the changes in the expression of collagens in response to TGF-β1 during skeletal muscle development.
The data from the current study provided evidence that the expression of the ECM proteoglycan decorin and the structure of the ECM changed in response to TGF-β1 during skeletal muscle development and growth. Thus, TGF-β1 likely has multiple functions in the developing skeletal muscle regulating muscle fiber formation, intramuscular connective tissue organization, the proper organization of the muscle fibers, muscle fiber bundles, and skeletal muscle growth.
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FOOTNOTES |
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Received for publication July 3, 2008. Accepted for publication October 1, 2008.
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