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Vol. 20, Issue 8, 2174-2185, April 15, 2009

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Transforming Growth Factor β1-mediated Activation of the Smooth Muscle -Actin Gene in Human Pulmonary Myofibroblasts Is Inhibited by Tumor Necrosis Factor- via Mitogen-activated Protein Kinase Kinase 1-dependent Induction of the Egr-1 Transcriptional Repressor

Xiaoying Liu^*, Robert J. Kelm, Jr., and Arthur R. Strauch^*

*Department of Physiology and Cell Biology and the Dorothy M. Davis Heart and Lung Research Institute, College of Medicine, The Ohio State University, Columbus, OH 43210; and Departments of Medicine and Biochemistry, Cardiovascular Research Institute, University of Vermont, College of Medicine, Burlington, VT 05405

Submitted October 3, 2008; Revised February 10, 2009; Accepted February 19, 2009
Monitoring Editor: Carl-Henrik Heldin

	ABSTRACT

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Transforming growth factor (TGF) β1 is a mediator of myofibroblast differentiation in healing wounds in which it activates transcription of the smooth muscle -actin (SMA) gene via dynamic interplay of nuclear activators and repressors. Targeting components of TGFβ1 signaling may be an effective strategy for controlling myofibroblasts in chronic fibrotic diseases. We examined the ability of proinflammatory tumor necrosis factor (TNF)- to antagonize TGFβ1-mediated human pulmonary myofibroblast differentiation. TNF- abrogated TGFβ1-induced SMA gene expression at the level of transcription without disrupting phosphorylation of regulatory Smads. Intact mitogen-activated protein kinase kinase (Mek)–extracellular signal-regulated kinase (Erk) kinase signaling was required for myofibroblast repression by TNF- via induction of the early growth response factor-1 (Egr-1) DNA-binding protein. Egr-1 bound to the GC-rich SPUR activation element in the SMA promoter and potently suppressed Smad3- and TGFβ1-mediated transcription. Reduction in Smad binding to the SMA promoter in TNF-–treated myofibroblasts was accompanied by an increase in Egr-1 and YB-1 repressor binding, suggesting that the molecular mechanism underlying repression may involve competitive interplay between Egr-1, YB-1, and Smads. The ability of TNF- to attenuate myofibroblast differentiation via modulation of a Mek1/Erk/Egr-1 regulatory axis may be useful in designing new therapeutic targets to offset destructive tissue remodeling in chronic fibrotic disease.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Myofibroblasts are specialized stromal cells that synthesize interstitial collagens and contain a smooth muscle -actin (SMA)-enriched contractile apparatus designed to generate tensile force needed for wound closure and tissue healing (Skalli and Gabbiani, 1988; Ronnov-Jessen and Petersen, 1996; Zalewski and Shi, 1997; Gabbiani, 2003; Desmouliere et al., 2005; Tomasek et al., 2006). Transforming growth factor (TGF) β1) is the principle mediator of myofibroblast differentiation in healing wounds where it accumulates in granulation tissue in activated form during leukocyte infiltration and potently stimulates transcription of the type I2 collagen subunit (COL12) and SMA genes (Becker et al., 2000; Massague and Wotton, 2000; Cogan et al., 2002; Higashi et al., 2003; Grotendorst et al., 2004; Leask and Abraham, 2004; Subramanian et al., 2004; Zhang et al., 2005). Smad proteins 2 and 3 are activated by TGFβ1 receptor engagement and enter the nucleus in which they directly bind SMA and COL12 promoter DNA. Smad3 also reportedly forms additional off-promoter complexes with DNA-binding repressor proteins to govern transcriptional output of the COL12 gene in TGFβ1-activated myofibroblasts (Higashi et al., 2003). Poor termination of TGFβ1-mediated myofibroblast differentiation could contribute to chronic fibroproliferative disease and excessive scar formation in the injured lung, heart, liver, kidney, and skin potentially causing pathobiologic phenomena referred to as "endless healing" (Tomasek et al., 2002). In particular, pulmonary fibrosis typically is associated with chronic myofibroblast activation, deposition of interstitial collagen and SMA, and dysfunctional parenchymal remodeling with a mortality of 50% at 5 y after diagnosis (Zhang et al., 1996; Kunkel et al., 2003; Noble, 2003; Phan, 2003; Selman and Pardo, 2003; Daniels et al., 2004; Marshall et al., 2004; Willis et al., 2006; Wynn, 2007). Similarly, cardiac transplant chronic rejection is associated with pervasive interstitial myofibroblast accumulation and perivascular fibrosis that ultimately impairs heart graft perfusion, performance, and long-term survival (Weber, 1995; Armstrong et al., 1997a,b,c; Subramanian et al., 1998, 2002; Rosenkranz, 2004).

Our laboratory has identified several cis-regulatory elements in SMA promoter that are highly conserved among vertebrate species and function as binding sites for multiple positive and negative trans-acting transcription factors (Foster et al., 1992; Cogan et al., 1995; Sun et al., 1995). Among these factors, Pur, Purβ, and YB-1 show preferential binding to single-stranded DNA and function as transcriptional repressors (Kelm et al., 1999, 2003;Carlini et al., 2002; Zhang et al., 2005; Knapp et al., 2006) whereas others such as transcription enhancer factor (TEF) 1; serum response factor (SRF); Smads 2, 3, and 4; and Sp1/Sp3 represent serum- or TGFβ1-responsive transcriptional activators (Kelm et al., 1999; Cogan et al., 2002; Subramanian et al., 2004). A DNA element referred to as SPUR recently was identified in the mouse as a key component in both basal and TGFβ1-dependent transcriptional activation of the SMA gene through its ability to bind heteromeric complexes of the Sp1/Sp3 activators and Pur/Purβ protein repressors (Subramanian et al., 2004; Knapp et al., 2006). SPUR DNA also encompasses a TGFβ1 control element shown to be an essential mediator of myofibroblast differentiation and dermal wound healing in the mouse (Tomasek et al., 2005). Dynamic physical interplay between transcriptional repressors and activators at SPUR is governed by Smad proteins that are rapidly imported into the nucleus of TGFβ1-activated myofibroblasts in which they seem to neutralize repressor:activator interactions and derepress the SMA promoter during the early stages of the wound healing process.

Clinical therapeutic strategies for managing myofibroblast differentiation by specifically targeting TGFβ1 signal transduction may help alleviate dysfunctional tissue remodeling in chronic fibrotic disease (Daniels et al., 2004; Yingling et al., 2004; Wang et al., 2005). Although comparatively little is known about mechanisms of myofibroblast attenuation, one approach under study is based on the ability of proinflammatory cytokines such as interferon (IFN) and TNF- to antagonize profibrotic TGFβ1 signaling (Verrecchia and Mauviel, 2004; Breitkopf et al., 2006; Dooley et al., 2006). TNF- is a 17.5-kDa polypeptide produced by activated macrophages as well as injured stromal and epithelial cells (Verrecchia and Mauviel, 2004; Saika et al., 2006). TNF- mediates host immune responses to microbial infection, tissue injury, and inflammation and has been implicated in etiology of chronic diseases such as rheumatoid arthritis, osteoarthritis, viral myocarditis, and heart failure (Tracey and Cerami, 1993). In these pathological contexts, TNF- fosters degradation of the extracellular matrix by inducing the production of stromal collagenases or by direct inhibition of type I collagen biosynthesis. TNF- binding to its two cognate receptors TNFR-1 and TNFR-2 variably activates the nuclear factor-B (NF-B), c-Jun NH₂-terminal kinase (JNK), and the p38 or Ras–Raf–mitogen-activated protein kinase kinase (Mek)–extracellular signal-regulated kinase (Erk) mitogen-activated protein (MAP) kinase signaling pathways (Verrecchia et al., 2002; Wajant et al., 2003). The antifibrotic property of TNF- was evident in transgenic mice where overexpression of this inflammatory cytokine was observed to markedly diminish pulmonary fibrosis induced by either TGFβ1 or bleomycin instillation. TGFβ1 and TNF- reportedly exhibit opposite effects on nitric oxide production in cardiac myocytes, cytotoxic T cell survival, and transcription of the type I collagen gene expression required for tissue repair and remodeling. Last, TNF- augments fibroblast and smooth muscle cell migration (Goetze et al., 1999), a process that has been definitively linked to reduced expression and deployment of SMA-enriched microfilaments (Ronnov-Jessen and Petersen, 1996).

In this report, we examined the hypothesis that proinflammatory TNF- elicits antifibrotic responses that specifically block TGFβ1-dependent SMA gene expression in differentiated human pulmonary myofibroblasts. We discovered that TNF- inhibits TGFβ1 signaling by activating Mek1 and Erk1/2 kinases needed for induction of the DNA-binding protein early growth response factor-1 (Egr-1). TNF-–inducible Egr-1 was observed to bind the GC-rich SPUR element in the SMA promoter and potently suppressed Smad-mediated transcription of this gene in TGFβ1-activated myofibroblasts. The data suggest that Egr-1 functionally resembles the Pur protein repressors that also target SPUR sequences in quiescent stromal fibroblasts, rendering them less efficient in the expression of SMA protein needed for myofibroblast contractility. TNF- also offsets SMA gene activation in myofibroblasts by enhancing interaction of the YB-1 transcriptional repressor with promoter DNA resulting in partial displacement of profibrotic Smad activators. A better understanding of dynamic interplay between transcriptional activators and repressors that control myofibroblast differentiation may facilitate development of novel antifibrosis therapeutic agents.

	MATERIALS AND METHODS

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Human Fibroblast Culture and Protein Extraction Methods
Human pulmonary fibroblasts (hPFBs) were established in primary culture from enzyme-dispersed tissue fragments of human neonatal lung tissue obtained at autopsy and were the kind gift of Dr. Daren L. Knoell (Departments of Pharmacy and Internal Medicine, The Ohio State University, Columbus, OH). Fibroblasts were maintained up to passage 7 with little observed change in growth rate, morphology, or growth factor responsiveness in a 1:1 mixture of Ham's F-12 and DMEM (1.0 g/l D-glucose) supplemented with penicillin-streptomycin-antimycotic gentamicin (50 µg/ml), and 10% heat-inactivated fetal bovine serum (Zhang et al., 2005). All culture medium reagents were obtained from Invitrogen (Carlsbad, CA). Recombinant human TGFβ1 and human TNF- (R&D Systems, Minneapolis, MN) were added to cell cultures for varying periods and doses as noted in the text. The metabolic inhibitors U0126, PD98059 (Cell Signaling Technology, Danvers, MA), SP600125 (A.G. Scientific, San Diego, CA), and SB203580 (Calbiochem, San Diego, CA) were used as noted in the text. To prepare whole cell protein extracts we followed our previously published methods (Cogan et al., 2002; Subramanian et al., 2004; Zhang et al., 2005). In brief, cell monolayers were rinsed twice with Dulbecco's phosphate-buffered saline (PBS) and then scraped into 0.5 ml of radioimmunoprecipitation assay buffer (PBS containing 1% NP-40, 0.5% sodium deoxycholate, 0.1% SDS, protease inhibitor cocktail, 0.2 mM phenylmethylsulfonyl fluoride [PMSF], and 0.5 mM dithiothreitol [DTT]). After gentle rocking for 15 min at 4°C, the cell lysate was collected at 10,000 x g for 10 min at 4°C. To prepare nuclear protein extracts, cell monolayers were washed twice with PBS, scraped into fresh PBS, sedimented at 3000 rpm, washed once more in PBS, and resuspended in 8 packed cell volumes of a hypotonic buffer containing 10 mM HEPES, pH 7.9, 1.5 mM MgCl₂, 10 mM KCl, 0.2 mM PMSF, and 0.5 mM DTT. Cells were allowed to swell in hypotonic buffer for 10 min on ice before transfer to a Dounce homogenizer for processing with a type B pestle. After centrifugation for 15 min at 4000 rpm, supernatants were removed and retained as the cytosolic protein fraction. The nuclei pellet was resuspended in packed pellet volume of ice-cold, low-salt buffer containing 20 mM HEPES, pH 7.9, 25% glycerol, 1.5 mM MgCl₂, 20 mM KCl, 0.2 mM EDTA, 0.2 mM PMSF, 0.5 mM DTT. An equal volume of high salt buffer containing 20 mM HEPES, pH 7.9, 25% glycerol, 1.5 mM MgCl₂, 1.2 M KCl, 0.2 mM EDTA, 0.2 mM PMSF, and 0.5 mM DTT was added, and the preparation was gently mixed for 30 min at 4°C. Nuclear remnants were removed by centrifugation at 14,500 rpm for 30 min, and the supernatant was dialyzed at 4°C for 3 h against 50 volumes of dialysis buffer containing 20 mM HEPES, pH 7.9, 20% glycerol, 100 mM KCl, 0.2 mM EDTA, 0.2 mM PMSF, and 0.5 mM DTT. Dialyzed nuclear protein was clarified by centrifugation at 14,500 rpm for 20 min, divided into aliquots, and stored at –80°C for use in biochemical assays.

DNA Binding Assay
Synthetic oligonucleotide probes used in this report were derived from selected sequences present in the human SMA promoter (Cogan et al., 1995; Sun et al., 1995; Kelm et al., 1996; Subramanian et al., 2004; Zhang et al., 2005). Reaction mixtures containing protein extract (100 µg of protein) and 3'-biotinylated oligonucleotides (100 pmol; Integrated DNA Technologies, Coralville, IA) were incubated in a buffer containing poly(dI-dC), 10 mM Tris, pH 7.5, 50 mM NaCl, 0.5 mM DTT, 0.5 mM EDTA, 0.12 mM PMSF, and 4% glycerol. Biotinylated DNA:protein complexes were incubated for 30 min with a 0.6-ml aliquot of streptavidin-coated paramagnetic particles (Promega, Madison, WI). After washing four times with buffer containing 25 mM Tris-HCl, pH 7.5, 1 mM EDTA, and 100 mM NaCl, the bound protein was eluted using 2x SDS protein denaturing buffer, size fractionated on 10% SDS-polyacrylamide gels, and then electrophoretically transferred to nitrocellulose membranes (Whatman Schleicher and Schuell, Keene, NH) for immunoblot evaluation. Protein blots were blocked at 4°C in Tris-buffered saline (TBS) containing 25 mM Tris-HCl, pH 7.5, 150 mM NaCl, 3% (wt/vol) nonfat dry milk, and 0.5% bovine serum albumin and incubated with the indicated antibodies for 90 min at room temperature with gentle rocking. For each of the primary antibodies used in our analysis, preimmune serum or antibodies presaturated with blocking peptides (in the case of peptide epitope antibodies for YB-1) were used as negative controls and protein extracts prepared from mouse heart or AKR-2B mouse embryonic fibroblasts known to contain SMA gene repressors and activators (Cogan et al., 1995; Sun et al., 1995; Kelm et al., 1999) were used as positive controls and to monitor antibody binding specificity. Blots were washed four times at room temperature over a 20-min period in TBS containing Tween 20 (0.05% vol/vol) and then incubated for 45 min with a species-appropriate, horseradish peroxidase (HRP)-conjugated secondary antibody (1:2000; GE Healthcare, Chalfont St. Giles, Buckinghamshire, United Kingdom). The blots were washed as described above, processed for antibody visualization by chemiluminescence (GE Healthcare), and imaged onto Biomax film (Eastman Kodak, Rochester, NY). Commercial antibodies used in this study were specific for Smad2/3 (rabbit polyclonal antibody, 1:1000; Cell Signaling Technology), Egr-1 (clone 44D5 rabbit monoclonal antibody [mAb], 1:1000; Cell Signaling Technology; or C-19 rabbit polyclonal antibody, 1:200; Santa Cruz Biotechnology, Santa Cruz, CA), Smad7 (goat polyclonal antibody, 1:250; Abcam, Cambridge, MA), phosphorylated Smad2 (Ser 465/467-specific, rabbit polyclonal antibody, 1:1000; Cell Signaling Technology), SMA (EPOS anti-human smooth muscle -actin/HRP, clone 1A4, 1:200; Dako North America, Carpiteria, CA), Erk1/2 (anti-MAP kinase p44/p42, rabbit polyclonal antibody, 1:1000; Cell Signaling Technology), phosphorylated Erk1/2 (anti-MAP kinase phospho-p44/p42, T202/Y204-specific, clone E10 mouse mAb, 1:1000; Cell Signaling Technology), and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) (anti-GAPDH/HRP, 1:200; Santa Cruz Biotechnology). Rabbit polyclonal antibodies specific for YB-1 (anti-YB1 M85-110 and anti-YB1 M276-302, 1:2000) were described in our previous reports (Kelm et al., 1999). The two Egr-1–specific antibodies gave slightly different results on immunoblots owing to Egr-1 size variation, possibly because of protein phosphorylation. However, antibodies from both vendors reacted with a dominant protein band that migrated on SDS gels with a molecular size expected for authentic Egr-1.

Mammalian Protein Overexpression Plasmids, Cell Transfection, and Reporter Gene Assay
Monolayers of hPFBs maintained at 40–50% confluence were transfected as described previously (Zhang et al., 2005). Optimized mixtures of the SMA promoter: reporter fusion plasmids (VSMP4) with or without mammalian expression plasmids encoding various transcriptional regulatory proteins were transfected using the Mirus (Invitrogen) transfection reagent and protocol provided by the manufacturer. Transfection efficiencies using these methods and reagents typically were in the range of 40–50% (Zhang et al., 2008). All plasmids used in this study were purified using QIAGEN preparative resin and a protocol provided by the manufacturer (QIAGEN, Valencia, CA). The Mek1-SSED and Mek1-K97M expression plasmids were kindly provided by Dr. Natalie Ahn (University of Colorado, Boulder, CO), the Egr-1 expression plasmid was a gift from Dr. Vikas P. Sukhatme (Harvard Medical School, Boston, MA), and the Smad2 and Smad3 expression plasmids were kindly provided by Dr. J. Horowitz (North Carolina State University, Raleigh, NC). Forty-eight hours after transfection, cells were washed three times with cold PBS and then lysed using chloramphenicol acetyltransferase (CAT) enzyme-linked immunosorbent assay (ELISA) lysis buffer (Roche Applied Science, Indianapolis, IN). Lysates were clarified by centrifugation at 14,000 x g for 10 min at 4°C, and the protein concentration of CAT protein-enriched supernatants determined by the bicinchoninic acid assay (Pierce Chemical, Rockford, IL). Equivalent amounts of supernatants were analyzed on immunoblot to verify overexpression of the various plasmid-encoded transcriptional regulatory proteins in transfected cells and expression of the SMA promoter-driven CAT reporter gene was quantified using a commercial CAT-ELISA kit (Roche Applied Science). Reporter gene expression was normalized with respect to total cell protein and transfections were performed in triplicate and repeated three to five times. We did not observe discernible differences in cell size, growth rate, or yield of protein extract between cell preparations transfected with the various overexpression plasmids used in the study. Data sets were subjected to analysis of variance to assess statistical significance set at p < 0.05.

RNA Extraction, Real-Time Quantitative Polymerase Chain Reaction (PCR), and Chromatin Immunoprecipitation (ChIP)
Total RNA was extracted from hPFBs by using TRIzol reagent (Invitrogen) according to the manufacturer's protocol, and 1-µg aliquots were reverse-transcribed using SuperScript First Strand Synthesis System for reverse transcription (RT)-PCR kit (Invitrogen). PCR amplification was performed in a total volume of 50 µl by using TaqMan PCR core reagent kit (Applied Biosystems, Foster City, CA) with primers and probes specific to the human SMA promoter as follows: forward primer, 5'-CCCCTGCTCTGCCTCTAGC-3'; reverse primer, 5'-GAACTGGAGGCGCTGATCC-3'; and TaqMan probe, 5'-(FAM)-ACAACTGTGAACGTTTTG-(MGBNFQ)-3'. A probe specific for 18S rRNA also was used as a control. Samples were processed for real-time quantitative PCR by using the ABI PRISM 7700 sequence detection system (Applied Biosystems). ChIP was performed using a commercial ChIP kit (Millipore, Billerica, MA) following the manufacturer's protocol. Briefly, monolayers of subconfluent hPFBs were exposed to 1% formaldehyde in PBS for 10 min, rinsed in PBS, and the fixed cells were collected into 200 µl of SDS lysis buffer supplemented with proteases inhibitors. After incubation on ice for 10 min, the lysates were sonicated using optimized conditions to obtain sufficient DNA shearing, diluted 10-fold into ChIP dilution buffer, precleared with salmon sperm DNA:protein A beads for 30 min at 4°C, and incubated overnight at 4°C with anti-Egr-1 antibody. The material immunoprecipitated with anti-Egr-1 antibody was then collected on salmon sperm DNA:protein A beads during a 1-h incubation, washed with low-salt buffer followed by high-salt buffer, LiCl buffer, and TE. Protein:DNA complexes were eluted from the protein A beads by exposure to 5 M NaCl for 4 h at 65°C and treated with protease K, EDTA, and Tris-HCl for 1 h at 45°C. DNA was extracted using phenol:chloroform and precipitated with isopropanol. PCR was performed using primers specific for the SMA promoter (forward strand, 5'-ACAGGCTCAAGTCTGTCTTTGCTC-3' and reverse strand, 5'-CCAAATCCCTGCAGATGGTGTCTT-3') by using the following conditions: 95°C for 4 min, 40 cycles at 94°C for 1 min, 55°C for 1 min and 72°C for 30 s, and a final extension phase of 72°C for 2 min. The appearance of an expected 230 base pairs PCR product in the various ChIP preparations was assessed by gel electrophoresis on 1.5% agarose.

	RESULTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
TNF- Inhibits TGFβ1-inducible SMA Gene Activity in Human Pulmonary Fibroblasts without Inducing Smad7 or Blocking Smad2 Phosphorylation
We previously showed by immunoblot and immunofluorescence microscopy that TGFβ1 substantially increased SMA mRNA and protein expression in early passage human pulmonary fibroblasts (Zhang et al., 2005). As shown in Figure 1a, TNF- (10 ng/ml) provided to hPFBs 2 h before treatment with TGFβ1 (5 ng/ml) markedly reduced the accumulation of SMA protein relative to the level of expression noted in cells treated with TGFβ1 alone. Inhibition also was observed if TNF- was added concurrently with exposure of the cells to TGFβ1 (data not shown). Immunofluorescence microscopic assessments of untreated control preparations of hPFBs incubated with the 1A4 SMA-specific antibody revealed that only 3% of these cells exhibited discernible expression of SMA protein above that seen in control preparations that omitted the primary antibody. Supportive of immunoblot data shown in Figure 1a, exposure of the cells to a 48-h treatment with TGFβ1 resulted in strong induction of SMA in 90% of the cells, whereas dual treatment with both TGFβ1 and TNF- reduced SMA expression in these cells to background levels comparable to that observed in untreated control preparations (data not shown). Moreover, evaluation of cell number by using 4,6-diamidino-2-phenylindole nuclear staining showed that neither TGFβ1 nor TNF- had substantial effect on cell abundance or survival at the doses used in this study over the course of a 48-h observation period. RT-PCR analysis revealed that accumulation of SMA mRNA in TGFβ1-treated hPFBs was significantly blunted by TNF- (Figure 1b). TGFβ1 also stimulated a fivefold increase in transcription from a VSMP4 SMA promoter:CAT reporter fusion gene construct in transfected hPFBs, but this increase was significantly attenuated in the presence of TNF- (Figure 1c). This observation suggested that the DNA sequence determinants for TNF-–mediated repression of TGFβ1 response in hPFBs were contained within the same region of the SMA core promoter previously shown to contain two TGFβ1-inducible elements referred to as SPUR and TGFβ1-hypersensitive region (THR) that are located between –59 and –28 and –170 and –150, respectively. Together, the observed effects of TNF- on hPFBs was suggestive of a rapid and dominantly acting inhibitory mechanism capable of neutralizing de novo transcriptional activation of the SMA promoter by TGFβ1.

View larger version (27K): [in this window] [in a new window]   Figure 1. TNF- suppresses TGFβ1-dependent production of SMA protein and mRNA in human pulmonary myofibroblasts. (a) hPFBs were pretreated with TNF- (10 ng/ml) for 2 h followed by exposure to TGFβ1 (5 ng/ml) for another 48 h after which SMA expression in cell extracts was examined by immunoblot using an anti-SMA antibody. Blots were reprobed with anti-GAPDH antibody to verify equivalent protein loading. (b) hPFBs were similarly pretreated with TNF- but harvested after a 22-h exposure to TGFβ1 for isolation of total RNA. SMA mRNA levels were quantitated by real-time PCR analysis using primers and TaqMan probe specific to the SMA sequence. (c) CAT ELISA was performed on hPFBs transfected with VSMP4 after a 24-h exposure to TGFβ1 alone or TGFβ1 plus a 2-h pretreatment with TNF-. Asterisks denote statistically significant differences between TGFβ1 treatments in the absence and presence of TNF- (p < 0.05).

View larger version (74K): [in this window] [in a new window]   Figure 2. TNF- neither augments expression of Smad7 nor blocks accumulation of phosphorylated Smad2 in human pulmonary myofibroblasts. (a) Immunoblot showing unaltered expression of Smad7 in TNF--treated hPFBs for periods up to 8 h. Similarly, b shows that Smad2 phosphorylation in TGFβ1-activated myofibroblasts was unaffected by TNF-. Cells were pretreated with TNF- (10 ng/ml) for 2 h before the addition of TGFβ1 (5 ng/ml) for an additional 15 min. GAPDH immunoblots are presented to indicate equivalent sample protein loadings on the gels.

Erk1/2 MAP Kinase Is Required for TNF-–mediated SMA Gene Repression

View larger version (26K): [in this window] [in a new window]   Figure 3. Intact Mek/Erk signaling was required for TNF- to repress activation of the SMA gene by TGFβ1 in human pulmonary myofibroblasts. As shown in a, Quiescent hPFBs were pretreated with DMSO (vehicle), SP600125 (10 µM, JNK inhibitor), or SB203580 (20 µM, p38 inhibitor). (b) Cells were pretreated with vehicle, UO126 (10 µM, Mek1/2 inhibitor), or PD98059 (50 µM, Mek1 inhibitor). After 1 h, some preparations were pretreated with TNF- (10 ng/ml) for another 2 h before adding TGFβ1 (5 ng/ml) to all preparations followed by a final 48-h incubation period. SMA expression in whole cell lysates was examined by immunoblot analysis. GAPDH immunoblots are provided to indicate equivalent sample protein loadings on the gels.

View larger version (28K): [in this window] [in a new window]   Figure 4. Immunoblot analysis showing TNF-–mediated activation of Erk 1 and 2 in human pulmonary fibroblasts. Quiescent hPFBs were treated with TNF- (10 ng/ml) for up to 90 min, lysed, and examined for Erk1/2 phosphorylation by using a phospho-epitope–specific anti-Erk1/2 antibody. Total Erk1/2 protein levels in lysed cell preparations also was examined to verify that the increase in Erk1/2 phosphorylation was not simply the result of an increase in Erk1/2 protein availability which remain constant for the duration of the experiment (0 and 15 min time points shown in the bottom panel).

View larger version (33K): [in this window] [in a new window]   Figure 5. Effects of exogenous Mek1 signaling on SMA promoter activity in TGFβ1-treated human pulmonary myofibroblasts. hPFBs were transfected with the SMA promoter: CAT reporter fusion gene (VSMP4) together with expression plasmids encoding either a constitutively active Mek1 mutant (SSED) or dominant-negative Mek1 mutant (K97M). Control cells were transfected with same amount of empty pcDNA3 expression vector. After exposure to TGFβ1 for 24 h (gray-shaded bars), CAT protein levels were analyzed in cell lysates. The black bars denote CAT reporter gene expression in transfected cells treated with vehicle alone. Asterisks denote statistically significant differences between transfections performed using the empty plasmid and Mek1-SSED (p < 0.05).

Egr-1 Is Induced by TNF-–dependent Mek1 and Binds to the SMA Promoter

View larger version (59K): [in this window] [in a new window]   Figure 6. Egr-1 expression in human pulmonary fibroblasts and interaction with a TGFβ1-response element in the SMA promoter. (a) Transiently elevated Egr-1 protein expression in nuclear preparations from TNF-–treated hPFBs. Quiescent hPFBs were exposed to TNF- (10 ng/ml) for various periods up to 180 min, and resulting nuclear protein extracts were analyzed by Egr-1 immunoblot. (b) Egr-1 binds to SPUR DNA but not the binding sites for Smad proteins in either the PAI1 promoter (SBE) or the SMA promoter (THR) nor does it bind the MCAT element in the SMA promoter needed for gene induction by the TEF1 muscle-specific trans-activating protein. The lane labeled ns denotes Egr-1 binding to a nonspecific DNA probe and serves as a negative control. (c) ChIP analysis revealing formation of a complex between the Egr-1 protein and SPUR DNA in native chromatin context. Quiescent hPFBs were treated with TNF- (10 ng/ml) or vehicle for 1 h and then processed for ChIP using either nonspecific (control) or anti-Egr-1 (anti-Egr1) antibodies in combination with SPUR-specific primers as described in Materials and Methods. (d) Overexpression of constitutively active Mek1 (Mek-SSED) in hPFBs enhanced expression of Egr-1 nuclear protein and that inhibition of Mek1 kinase activity in transfected cells by using UO126 abolished both basal and Mek1-inducible expression of Egr-1. hPFBs were transfected with either the Mek1-SSED expression plasmid (3 µg) or identical amounts of empty pcDNA3 vector, and the resulting nuclear extracts were processed 24 h later for evaluation by Egr-1 immunoblot. GAPDH immunoblots in a and d are provided to indicate equivalent sample protein loadings on the gels.

Egr-1 Inhibits TGFβ1- and Smad-mediated Activation of the SMA Promoter

View larger version (18K): [in this window] [in a new window]   Figure 7. Egr-1 overexpression abrogates TGFβ1- and Smad-activated SMA promoter activity in human pulmonary myofibroblasts. hPFBs were transiently cotransfected with the VSMP4 reporter gene plus various amounts (0.1–1 µg) of expression plasmid encoding full-length Egr-1. After 16 h, transfected cells were placed in serum-free medium for 48 h, and TGFβ1 (5 ng/ml) was added to cells for another 24 h. Top, CAT ELISA values for cells transfected with varying amounts of the Egr-1 expression plasmid and then subjected to treatment with TGFβ1. Bottom, CAT ELISA was performed on hPFBs cotransfected with VSMP4 reporter (0.5 µg), a fixed amount of expression plasmid encoding for Smad3 (0.5 µg), and varying amounts of Egr-1 expression plasmid (0.3, 0.6, or 1.0 µg). After 48 h in serum-free medium, cell lysates were prepared and analyzed for CAT activity. In all cases, pcDNA control vector was used to ensure mole equivalent amounts of DNA in each transfection mixture. Transfections were performed in triplicate and asterisks denote statistically significant differences between extracts prepared from cells after TGFβ1 treatment (top) or after Smad3 transfection (bottom) in the absence or presence of Egr-1 expression plasmids (p < 0.05).

Binding of Egr-1 to the SMA Promoter Has Opposite Effects on the Binding of Smad Activator and YB-1 Repressor to Their Cognate Transcriptional Regulatory Sites

View larger version (57K): [in this window] [in a new window]   Figure 8. Enhanced binding of Egr-1 to the SMA promoter in TNF-–treated human pulmonary myofibroblasts is accompanied by decreased Smad interaction with the promoter but increased binding of YB-1. (a) Although nuclear levels of Smad2 and -3 did not change appreciably when hPFBs were treated with both TGFβ1 and TNF-, the amount of Smad binding to a 191-base pair segment of the SMA promoter containing the TGFβ1-regulatable THR control element decreased significantly (SPUR+THR double-stranded DNA probe; b). Egr-1 protein levels in nuclear extracts increased substantially after treatment with TNF- (a) as did Egr-1 interaction with the double-stranded SPUR+THR DNA probe (b), which was notably higher than binding observed in extracts from cells treated with TGFβ1 alone (b). SMA protein level increased after exposure to TGFβ1 but decreased if cells also were exposed to TNF- (a). The level of GAPDH protein in cytosolic extracts (a) indicates equivalent sample protein loading on the gel. In nuclear extracts prepared from hPFBs treated with TNF- alone (c), binding of YB-1 repressor protein to the reverse strand of DNA encompassing the THR region of the SMA promoter increased by 60% over baseline levels (p < 0.05).

	DISCUSSION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Recent studies indicate that inflammation and fibrosis are distinctly different, perhaps dichotomous phases of the wound healing process (Kaminski et al., 2000; Henson, 2003; Kunkel et al., 2003; Verrecchia and Mauviel, 2004). Profibrotic TGFβ1 is proteolytically released from a latent state during cellular injury and functions as the chief mediator of myofibroblast differentiation and extracellular matrix biosynthesis needed for tissue repair, regeneration, and remodeling (Gabbiani, 2003; Shi and Massague, 2003; Tomasek et al., 2005). The wound provisional matrix also helps organize neovascular elements to restore oxygen and nutrient perfusion to the damaged tissue bed. Secretion of the proinflammatory cytokine TNF- also is elevated after tissue injury to assist in neovasculogenesis but persists during formation of the provisional matrix when it may help balance TGFβ1-mediated matrix biosynthesis in myofibroblasts and prevent progression to hypertrophic scarring (Verrecchia and Mauviel, 2004). The process of matrix equilibration seems quite important in tissue homeostasis since TGFβ1-mediated fibrosis after corneal injury is much more severe in TNF- null mice that lack robust inflammatory responses compared with wild-type mice (Saika et al., 2006). Several pulmonary fibroproliferative diseases seem to involve dysfunctional signal transduction interplay between profibrotic TGFβ1 activated by alveolar phagocytic cells and proinflammatory TNF- released by polarized T cells. If unchecked, faulty signaling could promote cycles of matrix deposition and destruction that could eventually compromise parenchymal cell function (Moller, 2003). Similarly, mononuclear cell release of IFN, a T-helper 1 cytokine that often functions in tandem with TNF-, suppresses fibroblast proliferation and collagen deposition and has long been associated with classic type I, innate-immune responses (Kunkel et al., 2003). Notably, the antifibrotic activity of IFN seems to be partly based on its ability to functionally sequester TGFβ1 receptor-regulated Smad3 protein as a heteromeric complex with the YB-1 transcriptional repressor protein thus preventing binding of activated Smads to their cognate CAGA cis-activation sequence motifs in the type I collagen promoter (Higashi et al., 2003; Dooley et al., 2006). Aside from this one mechanistic detail, the molecular pathways governing termination of TGFβ1-mediated myofibroblast differentiation are not well understood but theoretically important in the search for new therapeutic targets to prevent chronic fibroproliferative diseases of the heart, lung, liver, and kidney. Accordingly, the objectives of this study were to 1) assess the ability of proinflammatory TNF- to block activation of the SMA promoter, the leading early indicator of myofibroblast differentiation and fibrogenesis in healing wounds; and 2) identify the specific molecular mechanism(s) used by antifibrotic TNF- to intercept profibrotic TGFβ1 signaling in differentiated pulmonary myofibroblasts.

The induction of Egr-1 through the Ras–Raf–Mek–Erk MAP kinase signaling cascade has been reported in a variety of cell types, including kidney and intestinal epithelial cells, vascular cells, and astrocytes in the CNS (Morawietz et al., 1999; McCaffrey et al., 2000; Tan et al., 2003; Autieri et al., 2004; Cho et al., 2006; Khachigian, 2006; Harada et al., 2007). In human lung fibroblasts, we observed that overexpression of a constitutively active Mek1 variant was accompanied by increased expression of Egr-1 protein. Accumulation of Egr-1 protein was significantly attenuated by the U0126 Mek1 inhibitor and was observed in human pulmonary fibroblasts that were transfected with the Mek1 variant or exposed directly to TNF-. Data presented in this report also indicated that suppression was dominant over TGFβ1-mediated activation of the SMA promoter in human pulmonary myofibroblasts and dependent on intact Mek1-Erk1/2 MAP kinase signaling. Biochemical inhibition of p38 and JNK signal transduction in neonatal human pulmonary myofibroblasts had no discernible effect on SMA repression by TNF- nor was termination of TGFβ1 signaling by TNF- associated with increased expression of the Smad7 antagonist or reduced efficiency of Smad2 and -3 agonist phosphorylation by the type I receptor kinase (Bitzer et al., 2000; Nagarajan et al., 2000; Verrecchia et al., 2000). In contrast, TNF- reportedly interferes with type I collagen subunit gene expression in various types of fibroblasts via a more combinatorial strategy involving alteration of Smad7 expression (Bitzer et al., 2000; Nagarajan et al., 2000) as well as modulation of Jun NH₂-terminal kinase activity (Verrecchia et al., 2003; Verrecchia and Mauviel, 2004). From a mechanistic standpoint, the regulation of collagen matrix deposition in mammalian fibroblasts may be more complicated than the scheme used to regulate expression of the SMA contractile protein gene and conceivably could involve more than one signal transduction cascade. Indeed, multiple-level control of Smad7 expression as well as JNK and perhaps p38 signaling might be needed to coordinate progression along the collagen fiber assembly arm of matrix biosynthesis with the remodeling arm that drives accumulation of matrix metalloproteinases required for collagen fiber degradation and remodeling. Superimposed on these functional needs may be additional differences between dermal, renal, pulmonary, hepatic, and cardiac fibroblast lineages that could specify how signal transduction operates based on cell fate determination rules laid down during early embryogenesis. Notably, SMA and type I collagen gene transcriptional control mechanisms share a need for ubiquitous Sp1 and TGFβ1-activated Smads, but it is important to realize that the SMA gene additionally is activated by the SRF, myocardin, and TEF1 DNA-binding proteins that are widely viewed as important trans-determinants of smooth and cardiac myogenesis. Conversely, activator protein-1 seems relatively important for activation of the COL12 gene promoter but does not seem to play a significant role in activation of SMA gene transcription. Comparative analysis of SMA and type I collagen gene transcriptional control mechanisms used by various fibroblast subtypes from fetal, neonatal, and adult tissues may be quite informative in this regard and provide much needed insight into the molecular basis of myofibroblast functional plasticity in normal development and chronic fibrotic disease.

We discovered that expression of the Egr-1 transcription factor downstream from Mek-Erk1/2 activation could be a factor in down-regulation of myofibroblast differentiation based on the ability of this DNA-binding protein to block both TGFβ1- and Smad2/3/4-inducible SMA gene transcription. In TNF-–treated human pulmonary fibroblasts, Egr-1 was able to bind the tissue injury-responsive SPUR sequence motif in the SMA promoter. This was an important observation because SPUR was shown previously to be the binding site for a Smad-regulated Sp1/3:Pur transcriptional control complex in a mouse myofibroblast-like cell line (Cogan et al., 2002; Subramanian et al., 2004) as well as an essential determinant in the SMA promoter required for dermal wound healing in the mouse (Tomasek et al., 2005). Importantly, the myofibroblast suppression property of TNF- also could be mediated by activation of Mek1-Erk1/2 signaling because earlier reports indicate that Erk-dependent phosphorylation of the Smad2 or -3 linker regions prevents their nuclear translocation, thus providing a significant impediment to TGFβ1-mediated myofibroblast differentiation (Kretzschmar et al., 1999). Analysis of Smad nuclear translocation in TGFβ1-stimulated fibroblasts overexpressing the constitutively active form of Mek1 may clarify the relative contributions of Erk- and Egr-1–directed forms of myofibroblast repression. The TGFβ1/TNF-/Erk/Egr-1 regulatory axis may provide a physiological rheostat that allows rapid emergence of SMA-positive myofibroblasts yet prevents unchecked expansion of these contractile cells in healing wounds.

Egr-1 is not expressed in normal tissue but rapidly induced by inflammatory cytokines released into damaged heart, lung, and kidney tissue beds as a result of mechanical trauma and hypoxia (McCaffrey et al., 2000; Yan et al., 2000; Okada et al., 2002; Autieri et al., 2004; Khachigian, 2006; Harada et al., 2007). Hypoxia induces Egr-1 expression in the microvascular endothelium and adventitial fibroblasts via protein kinase C-mediated Ras/Raf/Mek/Erk signaling (Khachigian, 2006). Repression of both myofibroblast differentiation and connective tissue biosynthesis by Egr-1 coupled with increased transcription of Egr-1–dependent genes such as interleukin (IL)-1β, monocyte chemoattractant protein-1, intercellular adhesion molecule-1, macrophage inflammatory protein-2, 10-kDa interferon-inducible protein, and (RANTES) protein may improve wound healing efficiency because these proinflammatory cytokines and chemokines seem to collaborate with hypoxia-induced hypoxia-inducible factor and vascular endothelial growth factor in the repair and expansion of microvascular networks needed to restore blood flow to damaged tissues. We recently reported that hypoxia (3% molecular oxygen) significantly repressed SMA gene expression in primary culture preparations of mouse cardiac fibroblasts (Roy et al., 2007). Although the role of Egr-1 in mediating transcriptional response of the SMA gene to hypoxia was not specifically examined in that study, we did report a positive correlation between hypoxia, SMA gene repression, and elevated levels of YB-1 protein in cardiac fibroblasts. As we now show, YB-1 and Egr-1 seem to function as corepressors in antagonizing the human pulmonary fibroblast response to TGFβ1. We also recently proposed that dysfunctional remodeling of syngeneic murine heart grafts was initiated by cardiac myofibroblast differentiation arising as a consequence of mechanical trauma and ischemia-reperfusion injury inflicted by repeated transplant surgery (Zhang et al., 2008). In this alloantigen-independent model of cardiac injury, induction of Egr-1 expression was noted in hearts subjected to one or two rounds of transplant surgery (David and Strauch, unpublished data).

From the standpoint of therapeutic intervention, approaches that target production of individual cytokines released after tissue injury may not be very effective in managing wound healing outcomes because proinflammatory TNF-, IFN, and IL-1β often exhibit redundant behavior broadly related to their common ability to stimulate NF-B–mediated gene expression (Kunkel et al., 2003). In contrast, identifying downstream negative effectors of myofibroblast differentiation may be a more useful strategy because of their possible rate-limiting roles in the etiology of chronic fibrotic disease. Egr-1 is activated early after tissue injury and directs a family of target genes whose expression is likely to be essential for various aspects of the wound repair process (Yan et al., 2000; Cho et al., 2006; Khachigian, 2006). The development of therapeutic agents that augment Egr-1 expression or mimic its ability to neutralize Smad-based transcription could offer clinical benefits by minimizing unregulated myofibroblast differentiation without the complications presented by targeting functionally redundant and possibly beneficial immune cell-derived cytokines (Harada et al., 2007). Of particular interest would be the characterization of Egr-1–activated gene products such as signaling kinases and phosphatases or identifying peptide domains in the Egr-1 molecule that inactivate receptor-regulated Smads or prevent them from forming transcriptional activation complexes with Sp1/3 or SRF within the SMA promoter. An earlier report also mentions that Egr-1 interferes with Sp1 binding to GC-rich sites in the type I collagen collagen promoter leading us to speculate that Sp1 interaction with SPUR might be similarly disturbed by Egr-1 resulting in loss of SMA gene transcription (Tan et al., 2003). However, preliminary assessment of nuclear extracts prepared from TNF-–treated human pulmonary fibroblasts showed little stoichiometric change in Sp1:SPUR interaction relative to extracts prepared from untreated control cells (Liu and Strauch, unpublished data). Conceivably, Egr-1 may alter Sp3 interaction with SPUR. We previously showed that Sp3 was weaker than Sp1 in the ability to transactivate the SMA promoter in transfected fibroblasts, a finding that was consistent with earlier reports indicating that Sp1 and Sp3 have different, possibly opposite effects on the expression of certain mammalian genes (Cogan et al., 2002). Moreover, Sp3 in fibroblasts and vascular smooth muscle cells exhibits a higher capacity to bind Pur repressor proteins compared with Sp1 (Knapp et al., 2006). Collectively, these data suggest that Egr-1 could attenuate transcriptional output from the SMA promoter in the presence of TNF- by altering dynamic interplay of Sp1 and Sp3 with SPUR DNA, perhaps shifting the balance toward preferential binding of Sp3. Investigation of possible competitive displacement of transcriptional activators mediated by Egr-1 might be accomplished by characterizing the relative DNA binding activity of Sp1 and Sp3 in titration studies using recombinant Egr-1 or by monitoring the level of each DNA-binding protein by chromatin immunoprecipitation using TGFβ1- and TNF-–treated pulmonary myofibroblasts.

Concurrent with altered protein:DNA interactions at SPUR, the observed enhanced loading of YB-1 at the THR element in TNF-–treated fibroblasts expressing Egr-1 may cause widespread changes in DNA structure within the 200-base pair segment of the SMA promoter that directs TGFβ1 responsiveness in myofibroblasts (Figure 9). We reported previously that de novo transcriptional activation of the SMA gene in human pulmonary myofibroblasts by TGFβ1 required nuclear export of the YB-1 repressor (Zhang et al., 2005). However, TNF-–mediated blockade of TGFβ1 activation did not simply reverse nuclear export of YB-1 even though binding of this repressor to SMA promoter DNA was gradually and significantly enhanced by this proinflammatory cytokine. Moreover, as YB-1 binding to the reverse strand of the MCAT/THR region increased after exposure to TNF-, Smad binding to the THR was diminished. The observed displacement of Smads 2 and 3 may occur independently from loading of Egr-1 onto SPUR DNA, given the physical distance separating the SPUR and THR regions, although this remains to be examined experimentally. Currently we cannot rule out the existence of hypothetical "super-repression" protein complex consisting of Egr-1:YB-1 multimers spanning the SPUR-THR segment within the context of native chromatin. However, immunoprecipitation analysis performed on nuclear and cytosolic extracts prepared from human pulmonary myofibroblasts treated with either TGFβ1 or TNF- alone or in various combinations revealed little evidence for the existence of stable Egr-1:YB-1 protein complexes. There is a theoretical possibility that formation and/or biochemical stabilization of this hypothetical super-repressor complex may require promoter DNA substrate to anchor Egr-1:YB-1 protein:protein interactions. This idea currently is under investigation using a DNA-based, solid-phase ELISA analytical approach. The model presented in Figure 9 also predicts that Egr-1 need not bind and sequester Smads to mediate transcriptional repression since direct physical disruption of the Smad binding site at the THR by YB-1 may serve this purpose. YB-1 has well documented duplex-DNA unwinding activity (Gaudreault et al., 2004), and we previously presented evidence showing that YB-1 and Smads compete for binding to the region of the SMA promoter harboring the THR element (Subramanian et al., 2004). YB-1 binds with high-affinity to the reverse strand of the MCAT/THR motif and may maintain this segment of DNA in an unwound state with assistance from the Pur and Purβ corepressor proteins that bind the forward strand of this same region (Strauch et al., 1997; Kelm et al., 1999; Carlini et al., 2002). We are examining duplex MCAT/THR melting in the presence and absence of recombinant YB-1 and Smads to validate various aspects of the working hypothesis presented in Figure 9. Egr-1 could also augment expression of other factors needed to facilitate YB-1–mediated binding and unwinding at THR such as regulatory kinases or phosphatases. We have emerging data showing that Erk1/2 activation is necessary for YB-1 nuclear uptake, oligomerization, and binding to single-strand nucleic acids in human pulmonary fibroblasts (Willis and Strauch, unpublished data). YB-1 is a phosphoprotein whose nuclear translocation in cancer cells was shown to be dependent on serine phosphorylation (Sutherland et al., 2005). Given its central role in controlling both SMA and type I collagen gene expression, exploring ways to stabilize compartmentalization of YB-1 in the cell nucleus or developing cell-permeable peptide derivatives that mimic the localized duplex DNA-unwinding and/or Smad3-sequestering activities of YB-1 (Higashi et al., 2003; Dooley et al., 2006) may lead to the development of myofibroblast-targeted therapy for controlling excessive scar formation.

View larger version (27K): [in this window] [in a new window]   Figure 9. Hypothetical model for dynamic interplay of Egr-1 and YB-1 transcriptional repressors with Smad activators in human pulmonary myofibroblasts. The top illustration depicts a state of SMA gene transcriptional activation in myofibroblasts exposed to profibrotic TGFβ1. Termination of myofibroblast activity by antifibrotic TNF- is shown in the bottom illustration and may involve functional displacement of Smad activators via concerted binding of Egr-1 and YB-1 repressors to their spatially distinct binding sites at SPUR and THR, respectively. Unwinding of duplex DNA at the THR may be fostered by collaboration between the YB-1 and Pur/β repressors that bind to opposite strands of this essential Smad activation site in the SMA promoter.

	ACKNOWLEDGMENTS

 
This work was supported by National Institutes of Health National Heart Lung and Blood Institute grants HL-085109 (to A.R.S.) and HL-054281 (to R.J.K.).

	Footnotes

 
This was published online ahead of print in MBC in Press (http://www.molbiolcell.org/cgi/doi/10.1091/mbc.E08-10-0994) on March 4, 2009.

Address correspondence to: Arthur R. Strauch (strauch.1{at}osu.edu)

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