Logo of mbcLink to Publisher's site
Mol Biol Cell. 2010 Mar 15; 21(6): 860–870.
PMCID: PMC2836967
A Highlights from MBoC Selection

The RhoA Activator GEF-H1/Lfc Is a Transforming Growth Factor-β Target Gene and Effector That Regulates α-Smooth Muscle Actin Expression and Cell Migration

Keith E. Mostov, Monitoring Editor


Maintenance of the epithelial phenotype is crucial for tissue homeostasis. In the retina, dedifferentiation and loss of integrity of the retinal pigment epithelium (RPE) leads to retinal dysfunction and fibrosis. Transforming growth factor (TGF)-β critically contributes to RPE dedifferentiation and induces various responses, including increased Rho signaling, up-regulation of α-smooth muscle actin (SMA), and cell migration and dedifferentiation. Cellular TGF-β responses are stimulated by different signal transduction pathways: some are Smad dependent and others Smad independent. Alterations in Rho signaling are crucial to both types of TGF-β signaling, but how TGF-β-stimulates Rho signaling is poorly understood. Here, we show that primary RPE cells up-regulated GEF-H1 in response to TGF-β. GEF-H1 was the only detectable Rho exchange factor increased by TGF-β1 in a genome-wide expression analysis. GEF-H1 induction was Smad4-dependant and led to Rho activation. GEF-H1 inhibition counteracted α-SMA up-regulation and cell migration. In patients with retinal detachments and fibrosis, migratory RPE cells exhibited increased GEF-H1 expression, indicating that induction occurs in diseased RPE in vivo. Our data indicate that GEF-H1 is a target and functional effector of TGF-β by orchestrating Rho signaling to regulate gene expression and cell migration, suggesting that it represents a new marker and possible therapeutic target for degenerative and fibrotic diseases.


The retinal pigment epithelium (RPE) underlies the neural retina and is crucial for photoreceptor physiology and survival; hence, various retinopathies originate from changes in RPE function. Retinal detachments due to injury or surgery lead to RPE dysfunction and the development of ocular fibrotic diseases including proliferative vitreoretinopathy (Roberts et al., 2006 blue right-pointing triangle; Saika et al., 2008 blue right-pointing triangle). Major drivers of ocular degenerative and fibrotic diseases are transforming growth factor (TGF)-β and its downstream signaling mechanisms (Connor et al., 1989 blue right-pointing triangle; Hiscott et al., 1999 blue right-pointing triangle; Kon et al., 1999 blue right-pointing triangle; Saika et al., 2004 blue right-pointing triangle). Rho signaling is one of those mechanisms and is activated by TGF-β in fibrotic diseases of different types of epithelia including the RPE (Zheng et al., 2004 blue right-pointing triangle; Nishikimi and Matsuoka, 2006 blue right-pointing triangle). Therefore, identification of regulators of Rho signaling downstream of TGF-β is crucial to understand these pathological changes and to identify novel therapeutic targets.

TGF-β signaling activates different signal transduction mechanisms: they can be Smad dependent or Smad independent and activate different types of cellular responses (Zavadil and Bottinger, 2005 blue right-pointing triangle; Schmierer and Hill, 2007 blue right-pointing triangle; Heldin et al., 2009 blue right-pointing triangle; Zhang, 2009 blue right-pointing triangle). In brief, upon TGF-β binding, the type II receptor kinase activates the type I receptor kinase, leading to phosphorylation of Smad2 and Smad3, which subsequently oligomerize with Smad4 and translocate to the nucleus to regulate gene expression. Smad-dependent signaling is important for cellular responses such as migration (Levy and Hill, 2005 blue right-pointing triangle). TGF-β–stimulated Smad-independent signaling pathways include various branches of mitogen-activated protein kinase pathways (e.g., p38, extracellular signal-regulated kinase 1/2 and c-Jun NH2-terminal kinase) and phosphatidylinositol-3-kinase/AKT pathways depending on the cellular context. Importantly, however, the Smad-dependent and -independent responses cannot always be separated so clearly, because certain signaling mechanisms, such as RhoGTPases, are regulated by both types of responses. Hence, it is important to understand how such Rho signaling mechanisms contribute to specific TGF-β responses.

Modulation of Rho GTPase signaling plays a central role in various TGF-β–induced responses but is only partially understood. TGF-β has opposing temporal effects on RhoA activation, initially inhibition and later activation of Rho signaling. TGF-β induces dissolution of cell–cell adhesion and reorganization of the actin cytoskeleton. During the first phase, RhoA is inactivated by degradation at cell junctions, leading to reduced intercellular adhesion (Ozdamar et al., 2005 blue right-pointing triangle). This initial phase is important for epithelial mesenchymal transition (EMT). In contrast, subsequent cellular responses leading to cytoskeletal reorganization, α-smooth muscle actin (SMA) expression and cell migration require RhoA activation; however, the molecular mechanisms and Rho regulators by which TGF-β induces activation of RhoA signaling are poorly understood (Masszi et al., 2003 blue right-pointing triangle; Fu et al., 2006 blue right-pointing triangle; Kita et al., 2008 blue right-pointing triangle). Activation of Rho GTPases is catalyzed by guanine nucleotide exchange factors (GEFs) and inactivation by GTPase activating proteins (GAPs). Understanding the functional roles of different GEFs and GAPs as well as their regulation of expression and activity in particular signaling pathways is a major challenge, and recent evidence suggests that these proteins may be potential therapeutic targets for developing drugs to treat various diseases (Bos et al., 2007 blue right-pointing triangle).

We now identify GEF-H1 as crucial TGF-β target gene and show that GEF-H1 regulates TGF-β–induced Rho activation, responses in gene expression, and migration in primary RPE cells. GEF-H1 protein expression is also up-regulated in migratory RPE cells of patients with retinal detachments and fibrosis, indicating that the observations in the experimental model reflect processes that occur in human disease. Our data thus indicate that GEF-H1 is a crucial target and mediator of TGF-β signaling and participates in epithelial dysfunction in disease.


Reagents, Cell Culture, and Treatments

Recombinant human TGF-β1 was from (PeproTech Rocky Hill, NJ). SB431542, actinomycin D, and cycloheximide were from Sigma Chemical (Poole, Dorset, United Kingdom). RPE cells were isolated from porcine eyes (Lee et al., 2001 blue right-pointing triangle) and used at passage 1. For TGF-β1 experiments, cells were plated at 3 × 104 cells/cm2, serum starved (0.5% fetal bovine serum) for 24 h, and then stimulated by adding 10 ng/ml TGF-β1 for the indicated times. For inhibitor studies, cells were preincubated with 10 μM SB431542, 50 ng/ml actinomycin D, or 10 μg/ml cycloheximide for 1 h and then treated with TGF-β1 in the continuous presence of the inhibitor for the specified time. For spontaneous transdifferentiation, RPE cells (passage 1) were plated at 0.5 × 104 cells/cm2; in some experiments, cells were cultured in the presence of SB431542 (10 μM; 14 d). The human keratinocyte cell lines HaCaT-TR (stably expressing the tetrcycline [Tet] repressor) and HaCaT-TR-S4 (stably expressing the Tet-inducible Smad4 small interfering RNA (siRNA) in addition to the Tet repressor) have been characterized previously (Levy and Hill, 2005 blue right-pointing triangle). For siRNA induction, the cells were grown for 48 h in the presence of tetracycline (2 μg/ml) and then stimulated by adding 2 ng/ml TGF-β1 for 4 d in the continuous presence of tetracycline. Madin-Darby canine kidney (MDCK) cells allowing the conditional depletion of GEF-H1 were described previously (Benais-Pont et al., 2003 blue right-pointing triangle).

Immunoblotting and Immunofluorescence Microscopy

Total cell extracts were prepared in SDS-polyacrylamide gel electrophoresis (PAGE) sample buffer and western blotting was performed using standard procedures. For immunofluorescence, cells were fixed in ice-cold methanol and processed for immunostaining as described previously (Benais-Pont et al., 2003 blue right-pointing triangle). Photographs were obtained with an LSM510 confocal microscope (Carl Zeiss, Jena, Germany) using a 63× objective, and the manufacturer's image acquisition software. Brightness and contrast of the images were adjusted with Photoshop (Adobe Systems Mountain View, CA). Antibodies used were as follows: zona occludens-1 (Benais-Pont et al., 2003 blue right-pointing triangle), occludin (mouse; Zymed Laboratories, South San Francisco, CA), cingulin (rabbit; Zymed Laboratories), GEF-H1 (Benais-Pont et al., 2003 blue right-pointing triangle), α-SMA (1A4; Sigma Chemical), α-tubulin (1A2; Kreis, 1987 blue right-pointing triangle), Smad4 (B8; Santa Cruz Biotechnology, Santa Cruz, CA), Slug (Santa Cruz Biotechnology), myosin-IIA (rabbit [Sigma Chemical] and mouse, 3/36), fibronectin (monoclonal F0791; Sigma Chemical) myosin light chain phosphates (MYPT1) and phosphorylated (T696) MYPT1 (Millipore, Billerica, MA). Secondary antibodies conjugated to horseradish peroxidase, fluorescein isothiocyanate, and cyanine 3 were from Jackson ImmunoResearch Laboratories (West Grove, PA). IRDye-680- and IRDye 80-conjugated secondary antibodies were from Li-COR Biosciences (Lincoln, NE) and were used in combination with an Odyssey fluorescence reader.


On approval of the ethics committee of the local health authority (REC 05/Q0504/17), eyes consented for research were obtained from Moorfields Hospital Eye Bank (London, United Kingdom). Nine evisceration specimens and one enucleation specimen were examined. In all eviscerations, there was relatively extensive disorganization of intraocular contents, generally with at least partial retinal detachment and changes of proliferative vitreoretinopathy. Trauma was the most common underlying pathology with two postinfection cases and one patient with retinopathy of prematurity. The enucleation was carried out because of a choroidal malignant melanoma. Routine, buffered Formalin-fixed, paraffin-embedded sections were cut at 4 to 5 μm in thickness. Hybridoma supernatant with anti-GEF-H1 monoclonal antibody (mAb) was diluted 1 in 3 and incubated overnight followed by washing and alkaline phosphatase-conjugated secondary antibody. Immunoreaction product was visualized using a red alkaline phosphatase-based technique and an Autostainer (Dako, Ely, Cambridgeshire, United Kingdom) after pretreatment in citrate buffer, pH 6.0, in a Pascal pressure cooker (Dako) according to manufacturer's instructions.

Semiquantitative Reverse Transcription-Polymerase Chain Reaction (RT-PCR)

Total RNA was reverse transcribed with specific anti-sense primers using avian myeloblastosis virus reverse transcriptase (Promega, Madison, WI) for 1 h. PCR was carried out in the exponential phase (25 cycles) to allow comparison of PCR product levels. This was achieved by performing initial reactions with different amounts of template to determine optimal amounts of input. For reverse transcription, total RNA (0.5 μg in 15-μl RT reaction) was incubated for 1h at 50°C for GEF-H1 primer 5′-acatctgtcatcagcagga-3′. For PCR 1 μl of RT reaction was used: primers 5′-TTCTCATCACCCAGTTCTCA-3′ (forward) and 5′-acatctgtcatcagcagga-3′ (reverse) and an annealing temperature of 56°C. Other primers used were myosin-IIA (forward 5′-AAGCTGCAGGAGATGGA GGGC-3′; reverse 5′-AAAAAAGAATTCCGGCCTGGAGCT CCTCCTCTTT-3′) and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) (forward 5′-ATCACTGCCACCCAGAAGAC-3′; reverse ATGAGGTCCACCACCCTGTT-3′).

Microarray Analysis

RPE cells were incubated for 3 d in the absence or presence of TGF-β1 (10 ng/ml), and RNA was isolated. Three samples for each condition were obtained and the RNA quality analyzed using a Bioanalyzer 2100 (Agilent Technologies, Palo Alto, CA). cDNA and subsequent cRNA were prepared as described previously (Chambers et al., 2003 blue right-pointing triangle) and then hybridized to porcine GeneChip arrays according to Affymetrix (Santa Clara, CA) standard protocols (http://www.affymetrix.com) at the University College London, Institute of Child Health Gene Microarray Centre (London, United Kingdom). Labeled GeneChips were scanned, using a confocal argon ion laser (Agilent Technologies). The data were analyzed using Gene Spring 7.2 software (Agilent Technologies). Genes were excluded if the signal strength did not significantly exceed background values and if expression did not reach a threshold value for reliable detection (based on the relaxed Affymetrix MAS 5.0 probability of detection (p ≤ 0.1) in each of the samples (Seo et al., 2004 blue right-pointing triangle).

Reporter Gene Assays

RPE cells were transfected using Lipofectamine 2000 (Invitrogen, Paisley, United Kingdom) with the indicated reporter promoter constructs driving firefly luciferase expression and an expression construct for GEF-H1 (pCB6-GEF-H1) or empty vector (pCB6), a reference promoter driving Renilla luciferase was used to normalized the data. Reporters genes used were as follows: serum response element (SRE) (SRE containing promoter; Clontech, Mountain View, CA), α-SMA-fl (full-length α-SMA promoter), α-SMA-155 (155-base pair α-SMA promoter construct), and α-SMA-BmAm (155-base pair α-SMA promoter construct with mutated SRE elements; Liu et al., 2003 blue right-pointing triangle). After 26 h, firefly and Renilla luciferase were measured. Where indicated, cells were incubated with 0.5 μM TAT-C3 transferase, a membrane permeable C3 transferase (Coleman et al., 2001 blue right-pointing triangle) for 24 h.

Transfection of siRNAs and Determination of Active RhoA

HaCaT cells were plated into 12-well plates, to determine Rho activation, or 24-well plates, to analyze protein expression. Cells were then transfected with nontargeting control siRNA pools or siRNAs specific for GEF-H1, Snail, and Slug (Thermo Fisher Scientific, Waltham, MA; and Dharmacon RNA Technologies, Lafayette, CO), using Interferin transfection reagent (Polyplus Transfection, Calne, Wilts, United Kingdom), and a total final siRNA concentration was 100 nM (Steed et al., 2009 blue right-pointing triangle). Twenty-four hours after the transfection, TGF-β1 (20 ng/ml) was added, and the cells were analyzed after another 3 d of culture. The cells were then rinsed with phosphate-buffered saline (PBS) and lysed in SDS-PAGE sample buffer for protein analysis, or levels of active RhoA were measured with the G-LISA assay kit (Cytoskeleton, Denver, CO). The assay was performed as instructed by the manufacturer but avoiding the freezing step in all samples.

Inhibition of α-SMA Expression by DN-GEF-H1

RPE cells were transiently transfected with pcDNA4/TO-CTD-VSV construct (DN-GEF-H1 (Aijaz et al., 2005 blue right-pointing triangle) using Lipofectamine 2000 and then treated with TGF-β1 for 3 d. Samples were fixed and stained for α-SMA, vesicular stomatitis virus (VSV; transfected cells), and DNA. Random fields were photographed and the percentages of α-SMA positive cells in the control (VSV-negative) and DN-GEF-H1 expressing (VSV-positive) cells were calculated (a total of 600 cells were counted for each condition, shown are means ± 1 SD of 3 determinations). For the lentiviral constructs, either the CTD-VSV sequence or the VSV epitope cassette were cloned into the lentiviral pHR'IN plasmid (Bainbridge et al., 2001 blue right-pointing triangle), giving rise to LNT-DN-GEF-H1 or LNT-control, respectively. RPE cells were infected with control (LNT-VSV) or DN-GEF-H1 (LNT-VSV-DN-GEF-H1) lentivirus at multiplicity of infection (MOI) 100 and stimulated with TGF-β1 for 3 d (experiments were performed in triplicates). Cells lysates were analyzed for α-SMA and fibronectin expression; α-tubulin was used as loading control.

Wound-Healing and Morphological Assays

The Electric Cell-Substrate Impedance Sensing (ECIS) Model 1600R (Applied BioPhysics, Troy, NY) was used to monitor cell migration. RPE cells were plated in ECIS electrode array (8W1E) (Applied BioPhysics). The following day, they were infected with LNT-control or LN T-DN-GEF-H1 at MOI 100 and 24 h later treated with TGF-β1 (50 ng/ml) for 2 d, and electrical wounds were inflicted as described previously (Keese et al., 2004 blue right-pointing triangle). In another type of wound-healing assays, manual wounds were inflicted with a pipette yellow tip, pictures were then taken after 0, 16, 24, 48 and 72 h, and the wound area was measured. The wound areas were normalized to the ones obtained at 0 h that are referred as 1 and all other areas were then expressed as fractions of the initial wound. For MDCK cells, the same wounding and impendence assay was used. GEF-H1 depletion was induced for 4 d before the wounding assay using tetracycline and was confirmed in parallel experiments as described previously (Benais-Pont et al., 2003 blue right-pointing triangle).

To follow morphological changes, RPE cells confluent for 3 wk were split and plated on glass coverslips coated with fibronectin (6 × 104 cells/cm2), cells were infected with LNT-control or LN T-DN-GEF-H1 at 200 MOI, and reached confluence after 1 d. Monolayer detachment/contraction was then followed for up to 5 d. Quantification of monolayer detachment/contraction was performed by measuring cell-free areas using ImageJ (National Institutes of Health, Bethesda, MD.


TGF-β–induced Disorganization of Cell–Cell Adhesion Correlates with Up-Regulation of GEF-H1

We used primary porcine RPE cells as a model to analyze TGF-β signaling because they form well-differentiated monolayers in culture and respond to TGF-β (Lee et al., 2001 blue right-pointing triangle; Ablonczy and Crosson, 2007 blue right-pointing triangle). As expected, addition of TGF-β stimulated dissolution of cell–cell adhesion structures, such as adherens and tight junctions, correlating with altered cell morphology and reduced expression of junctional proteins, such as ZO-1 and occludin (Figure 1, A–F).

Figure 1.
TGF-β1 induces junctional disruption and GEF-H1 up-regulation in RPE cells. RPE cells were stimulated with TGF-β1 (A–E for 3 d; F–I as indicated) and processed for immunofluorescence (A–E) or immunoblot (F–H) ...

RhoA is a key player in the control of the actin cytoskeleton, cell–cell adhesion and gene expression (Fujita and Braga, 2005 blue right-pointing triangle; Hall, 2005 blue right-pointing triangle; Posern and Treisman, 2006 blue right-pointing triangle; Heasman and Ridley, 2008 blue right-pointing triangle; Nelson, 2008 blue right-pointing triangle). To identify the Rho activators that transmit the TGF-β stimulus, we performed a genome-wide expression analysis using microarrays. Total RNA was isolated from triplicate samples of control and TGF-β–treated RPE cells and used to probe Affymetrix porcine arrays. GEF-H1 was the only detectable Rho exchange factor that was up-regulated in response to TGF-β1 (Table 1), suggesting that induction of GEF-H1 expression is likely to be of functional relevance for TGF-β-induced responses in RPE cells.

Table 1.
Guanine nucleotide exchange factor mRNA expression profile of control and TGF-β 1–treated porcine RPE cells

GEF-H1/Lfc is a guanine nucleotide exchange factor for RhoA (Benais-Pont et al., 2003 blue right-pointing triangle; Aijaz et al., 2005 blue right-pointing triangle; Birkenfeld et al., 2008 blue right-pointing triangle). In contrast to permanent cell lines, GEF-H1 is expressed at very low levels in primary cultures of differentiated RPE cells (Figure 1G), similar to the levels previously reported for adult epithelial tissues (Ryan et al., 2005 blue right-pointing triangle). Stimulation with TGF-β1, however, up-regulated GEF-H1 expression (Figure 1G). Similar results were obtained with GEF-H1 antibodies recognizing different epitopes (data not shown). Immunofluorescence also revealed increased expression of GEF-H1 and accumulation in areas of cell protrusions (Figure 1D). Up-regulation of GEF-H1 not only correlated with increased expression of α-SMA (Figure 1G) but also enhanced phosphorylation of myosin phosphatase (Figure 1I), suggesting increased activity of the Rho–Rho kinase pathway. Thus, up-regulation of GEF-H1 by TGF-β correlates with activation of Rho signaling and α-SMA expression.

We next analyzed the importance of GEF-H1 for the activation of RhoA in response to TGF-β treatment. We used the HaCaT cells, a keratinocyte cell line, for this purpose as we could down-regulate GEF-H1 in these cells effectively with commercially available siRNAs, and control siRNAs did not cause nonspecific effects as in primary porcine RPE cells. Figure 2A shows that TGF-β treatment also resulted in increased GEF-H1 expression in HaCaT cells. Up-regulation was inhibited by transfection of GEF-H1-specific but not nontargeting control siRNAs (Figure 2B). When the levels of active RhoA was measured in identically treated cells, we found that active RhoA levels increased in response to TGF-β and that this was inhibited if up-regulation of GEF-H1 was blocked by RNA interference. These data thus indicate that the increased levels of GEF-H1 expression contribute to Rho activation in response to TGF-β.

Figure 2.
Inhibition of TGF-β–induced Rho activity by GEF-H1 depletion. (A) HaCaT cells were stimulated for 3 d with TGF-β and were then analyzed for expression of GEF-H1 and α-tubulin. Expression of GEF-H1 was monitored with two ...

TGF-β Transcriptionally Up-Regulates GEF-H1 Expression through a Smad4-dependent Pathway

In response to TGF-β, activated Smad2 and Smad3 form complexes with Smad4 and accumulate in the nucleus, where they regulate expression of TGF-β target genes (Ross and Hill, 2008 blue right-pointing triangle). Therefore, we next analyzed whether the TGF-β–induced up-regulation of GEF-H1 at the protein level and the increased mRNA levels observed by microarray analysis were due to changes at the transcriptional level and whether up-regulation depended on Smad4.

Figure 3A shows that increased GEF-H1 mRNA levels in response to TGF-β were also observed if analyzed by semiquantitative RT-PCR instead of microarrays: in both types of assays, an approximately twofold up-regulation of GEF-H1 mRNA was observed. To determine whether transcription was required for up-regulation of GEF-H1 protein, we treated the cells with actinomycin D or cycloheximide. Both drugs inhibited induction of GEF-H1, indicating that transcription is required (Figure 3B).

Figure 3.
TGF-β1-induced GEF-H1 up-regulation is Smad4 dependent. (A) RT-PCR analysis for GEF-H1 mRNA levels in control and TGF-β1–treated (3-d) samples; GAPDH was used as a loading control. Note that the increase observed by RT-PCR (>2-fold) ...

We next tested whether up-regulation of GEF-H1 involves the canonical Smad pathway (Derynck and Zhang, 2003 blue right-pointing triangle). Treatment of RPE cells with the ALK5 kinase inhibitor SB431542 abrogated GEF-H1 expression (Figure 3C), indicating that TGF-β type I receptor kinase activity is necessary for GEF-H1 expression. To test involvement of the Smad pathway directly, we used again HaCaT cells that stably express a tetracycline-inducible shRNA targeting Smad4 (HaCaT-TR-S4 cells) (Levy and Hill, 2005 blue right-pointing triangle). TGF-β1 induced the expression of GEF-H1 in control HaCaT cells, and Smad4 depletion inhibited TGF-β1 induced GEF-H1 up-regulation (Figure 3D), revealing that GEF-H1 induction by TGF-β1 requires Smad4. These observations thus indicate that up-regulation of GEF-H1 involves activation of the TGF-β type I receptor kinase and the Smad pathway.

Microarray analysis previously identified two populations of TGF-target genes: Smad-dependent and -independent genes (Levy and Hill, 2005 blue right-pointing triangle). Because the Smad-dependence groups TGF-β–responsive genes are into different functional groups, the observed Smad-dependence for GEF-H1 suggests that it may function in Smad-dependent processes such as cell migration. Certain other TGF-β–stimulated processes, such as EMT, are Smad-independent and require the up-regulation of other transcription factors, such as Snail and Slug (Levy and Hill, 2005 blue right-pointing triangle). Hence, we tested whether Snail and Slug are involved in GEF-H1 up-regulation by transfecting cells with siRNAs targeting the two transcription factors before TGF-β stimulation. Figure 3E shows that up-regulation of GEF-H1 was not prevented by down-regulation of Slug. We were not able to detect Snail in HaCaT cells using two different antibodies, suggesting that Snail does not become up-regulated in these cells. This is in agreement with previous observations (Levy and Hill, 2005 blue right-pointing triangle). Thus, TGF-β1 induced GEF-H1 expression is smad4 dependent and does not require up-regulation of Slug.

Primary RPE cells in culture transdifferentiate into myofibroblast-like cells not only when treated with TGF-β but also spontaneously when plated at low density (Grisanti and Guidry, 1995 blue right-pointing triangle; Lee et al., 2001 blue right-pointing triangle; Wiencke et al., 2003 blue right-pointing triangle). Transdifferentiated RPE cells up-regulate both α-SMA and GEF-H1, supporting a myofibroblast-like phenotype (Figure 1G). Strikingly, treatment of low-density cultures with the ALK5 inhibitor prevented morphological changes (Figure 3F) as well as α-SMA and GEF-H1 up-regulation (Figure 3G), further supporting the correlation between TGF-β signaling and expression of α-SMA and GEF-H1.

TGF-β modulates cellular phenotypes not only by regulating α-SMA expression but also of nonmuscle myosin isoforms (Sinha et al., 2004 blue right-pointing triangle; Obara et al., 2005 blue right-pointing triangle). Therefore, we examined whether TGF-β stimulation affects myosin-IIA expression. Indeed, TGF-β increased myosin-IIA expression with similar kinetics as expression of GEF-H1 (Figure 4A). Although the ALK5 kinase inhibitor abrogated myosin-IIA up-regulation (Figure 4A), depletion of Smad4 only partially counteracted the increase (Figure 4B) and mRNA levels did not significantly change in response to TGF-β (Figure 4C), indicating posttranscriptional regulation by a mechanism at least partially distinct from the one that targets GEF-H1.

Figure 4.
TGF-β1–induced myosin-IIA up-regulation is Smad4 independent. (A) RPE cultures in the absence or presence of the ALK45 kinase inhibitor SB431542 were stimulated TGF-β1 as indicated. Immunoblots of total cell extracts are shown ...

GEF-H1 Regulates α-SMA Expression Induced by TGF-β1

Induction of α-SMA expression has been suggested to be of functional relevance for the pathologies of retinopathies such as retinal detachments or proliferative vitreoretinal disorders (Grisanti and Guidry, 1995 blue right-pointing triangle). Transcription of α-SMA is regulated by actin reorganization induced by Rho activation through serum response factor (SRF) (Hill et al., 1995 blue right-pointing triangle; Wamhoff et al., 2006 blue right-pointing triangle). Because GEF-H1 is an activator of RhoA and can activate an SRE-specific reporter gene construct in MDCK (Aijaz et al., 2005 blue right-pointing triangle) and RPE cells (Supplemental Figure 1), we next asked whether GEF-H1 stimulates α-SMA expression in response to TGF-β.

We first used a reporter gene assay to determine whether GEF-H1 is able to stimulate transcription of the α-SMA promoter and, if yes, whether this involves the SREs, the binding sites of the transcription factor SRF (Mack and Owens, 1999 blue right-pointing triangle; Miano et al., 2007 blue right-pointing triangle). Figure 5A shows that cotransfection of GEF-H1 stimulated the full-length promoter (α-SMA-fl) and a shorter promoter (α-SMA-155) in a manner that depended on the two SRE elements (α-155-BmAm) (Figure 5B). As expected, the short promoter responded more strongly to Rho activation as it lacks the repressing upstream region of the promoter. Inhibition of Rho with C3 transferase counteracted stimulation of the α-SMA promoter by GEF-H1 (Figure 5C), confirming the Rho dependence. Thus, GEF-H1 regulates α-SMA promoter activity in an SRE- and Rho-dependent manner.

Figure 5.
GEF-H1 regulates the α-SMA promoter in a Rho-dependent manner. (A) Schematic representation of the α-SMA promoter constructs used. α-SMA-fl is a 2.8-kb full-length α-SMA promoter; α-SMA-155 is a minimal promoter ...

We next tested whether GEF-H1 and Rho signaling regulate α-SMA expression during TGF-β stimulation. First, we incubated control and TGF-β–treated RPE cells with membrane-permeable C3 transferase. Figure 6A shows that this resulted in an efficient repression of α-SMA induction, indicating up-regulation requires Rho signaling.

Figure 6.
Rho signaling and GEF-H1 regulate α-SMA expression induced by TGF-β1. (A) RPE cells were incubated for 3 d with or without TGF-β1. During the last 2 d, membrane-permeable C3 transferase was added as indicated. Expression of GEF-H1 ...

We next used RNA interference to down-regulate GEF-H1 expression in RPE cells. However, various control siRNAs already repressed α-SMA levels, indicating an unspecific effect of siRNAs in RPE cells. Therefore, we made use of a dominant-negative (DN) construct containing the C-terminal domain (CTD) of GEF-H1and a C-terminal VSV epitope as a tag (DN-GEF-H1). SRE reporter assays confirmed that DN-GEF-H1 is able to suppress SRE-driven transcription (Supplemental Figure 2).

We next used a transient transfection assay to determine whether DN-GEF-H1 is able to counteract α-SMA-induction by TGF-β1. Double immunofluorescence revealed that most DN-GEF-H1–expressing cells failed to up-regulate α-SMA (Figure 6B). Quantification demonstrated that only 25% of the DN-GEF-H1–expressing cells were positive for α-SMA, whereas 60% of the control cells expressed the EMT marker (Figure 6C). For biochemical quantification, we repeated the experiment with lentiviral vectors to transduce RPE cells with DN-GEF-H1 (LNT-DN-GEF-H1) or a control lentivirus (LNT-control) and then stimulated with TGF-β1. Immunoblot analysis showed that LNT-DN-GEF-H1 transduction resulted in a 2.7-fold decrease in the α-SMA expression compared with LNT-control (Figure 6D). If the samples were probed for expression of fibronectin, a TGF-β target gene that is up-regulated in a Smad-independent manner (Tsuchida et al., 2003 blue right-pointing triangle), no inhibition of up-regulation was observed. These results show that expression of DN-GEF-H1 counteracts the TGF-β1–induced increase in α-SMA expression.

Treatment of cultures plated at low-density (0.5 × 104 cells/cm2) with the ALK5 inhibitor prevented morphological changes (Figure 3F) as well as α-SMA and GEF-H1 up-regulation (Figure 3G). ALK5 inhibitor also prevented the generation of gaps and monolayer detachment and contraction of older primary cultures (>3 wk) that were plated at high density (6 × 104 cells/cm2) on fibronectin after they had reached confluence, indicating that it was also caused by endogenous TGF-β production (Figure 7A). Because inhibition of GEF-H1 counteracts up-regulation of α-SMA expression, we next tested whether it also inhibits monolayer contraction and cell detachment.

Figure 7.
GEF-H1 inhibition counteracts TGF-β–dependent monolayer contraction and detachment. (A) Three-week-old primary cultures of porcine RPE cells were plated at high density (6 × 104 cells/cm2) on fibronectin-coated coverslips. The ...

Figure 7B shows that monolayers formed by RPE cells infected with a control lentivirus (LNT-control) started to detach and contract, whereas those infected with a virus encoding DN-GEF-H1 (LNT-DN-GEF-H1) did not. Quantification of such images confirmed that expression of dominant-negative GEF-H1 counteracted the appearance of cell-free areas even after 4 d of culture (Figure 7C). Thus, these results indicate that GEF-H1 drives morphological changes such as cell contraction and detachment induced by TGF-β1. Because contraction and detachment were measured by quantification of cell-free areas (Figure 7), further analysis will be necessary to identify whether cell-free areas are due to contraction only or also to reduced adhesion and/or increased cell death.

GEF-H1 Is Up-Regulated in RPE from Patients with Disorganized Retina and Pigment Epithelium

TGF-β signaling and expression of α-SMA have been related to the ability of RPE cells to form periretinal membranes and are thought to contribute to retinal detachments in proliferative vitreoretinopathy (PVR) and in response to trauma (Fuchs et al., 1991 blue right-pointing triangle; Saika et al., 2004 blue right-pointing triangle; Zheng et al., 2004 blue right-pointing triangle). Therefore, we next studied the expression of GEF-H1 in eye sections from patients with retinal detachments due to different types of insults.

In control RPE cells, there was little or no immunoreactivity for GEF-H1 (Figure 8A1), confirming the observations we made in nonstimulated primary porcine cultures and further supporting the conclusion that expression of high levels of GEF-H1 requires a stimulus in most adult tissues. In contrast, in pathological specimens, there was consistent GEF-H1 immunoreactivity in subsets of RPE cells that had migrated away from their normal location between photoreceptor outer segments and Bruch's membrane in nine of the ten investigated samples (Figure 8). There were four pigmented RPE cell phenotypes associated with this staining: migratory cells that remained configured as a monolayer, RPE cells around blood vessels, individual migratory cells, or apex to apex islands of RPE cells (Fig. 8, A2–A5). RPE cells were identified on the basis of intense pigmentation and a side-to-side arrangement typical of epithelia except for when arranged as individual cells. Furthermore, their cytoarchitecture was generally cuboidal or polygonal rather than rounded, as would be expected for macrophages that had engulfed uveal pigment. Clusters of CD68 expressing macrophages were, however, identified and they were also strongly immunoreactive (data not shown). These observations indicate that up-regulation of GEF-H1 occurs in response to ocular insults and can be observed in migratory RPE cells in vivo.

Figure 8.
GEF-H1 is up-regulated in migratory RPE cells in vivo and regulates cell migration. (A1) Negative GEF-H1 staining was observed in RPE cells from a normal-looking area adjacent to a choroidal malignant melanoma (case 1, 40×). (A2) Positive GEF-H1 ...

GEF-H1 Regulates Cell Migration

The observed up-regulation of GEF-H1 in migratory pigmented RPE cells in vivo, suggests that the exchange factor plays a role in TGF-β-stimulated migration, a process that involves RhoA activation and that is thought to be one of the underlying reasons for failure of retinal detachment surgery due to PVR (Kon et al., 1999 blue right-pointing triangle; Kim et al., 2006 blue right-pointing triangle). As TGF-β-induced migration is abolished after silencing of Smad4 in HaCaT cells (Levy and Hill, 2005 blue right-pointing triangle) as is up-regulation of GEF-H1 (Figure 3D), we next tested whether GEF-H1 contributes to TGF-β-induced RPE migration using manual and electrical wound-healing assays.

RPE cells were infected with LNT-DN-GEF-H1 or LNT-control, pre-stimulated with TGF-β and then wounded either manually or with a strong electrical field. Wound closure was then followed microscopically or by measuring impedance of the monolayer. Figure 8, B and C, shows that expression of DN-GEF-H1 impaired wound closure in both assays. This indicates that the exchange factor indeed regulates TGF-β-induced RPE cell migration.

To test the importance of GEF-H1 for cell migration with a different cell type, we used spontaneously immortalized MDCK cells that constitutively express high levels of the exchange factor (Benais-Pont et al., 2003 blue right-pointing triangle; Aijaz et al., 2005 blue right-pointing triangle). We took advantage of previously generated cell lines that permit the tetracycline-induced depletion of GEF-H1 by RNA interference (Benais-Pont et al., 2003 blue right-pointing triangle; Aijaz et al., 2005 blue right-pointing triangle). Figure 8D shows that depletion of GEF-H1 resulted in a strong retardation of wound closure in the electrical wound healing assay. Visual inspection of the slides confirmed that the failure in wound healing was due to reduced migration of cells into the induced wound as compared with control RNA interference cells. Thus, GEF-H1 regulates migration of different epithelial cell types and may be of general importance for epithelial migration.


TGF-β-induced expression of alpha-smooth muscle actin and cell migration occurs during the development of different tissues and in several diseases including cancer and fibrosis, a common complication after tissue damage and surgery (Liu, 2006 blue right-pointing triangle; Roberts et al., 2006 blue right-pointing triangle). Our results demonstrate that the Rho guanine nucleotide exchange factor GEF-H1 is a novel target gene and functional effector of two crucial TGF-β-driven processes: α-SMA up-regulation, a marker for transdifferentiation, and cell migration.

TGF-β activates Smad-dependent and independent signaling pathways that regulate various cellular responses including cell migration, adhesion, proliferation and EMT (Derynck and Zhang, 2003 blue right-pointing triangle; Ikenouchi et al., 2003 blue right-pointing triangle; Peinado et al., 2007 blue right-pointing triangle; Ross and Hill, 2008 blue right-pointing triangle; Thuault et al., 2008 blue right-pointing triangle; Heldin et al., 2009 blue right-pointing triangle). Via the Smad-independent pathway, TGF-β receptor II triggers PAR6 mediated down-regulation of RhoA signaling at cell-cell junctions, which initiates dissociation of cell-cell adhesion (Ozdamar et al., 2005 blue right-pointing triangle). However, Smad-dependent and independent processes then require RhoA activation in a spatially and temporally controlled manner. Interestingly, certain processes only require one branch of TGF-β signaling, as, for example, Smad4 is required for TGF-β-induced migration, but not EMT, which is Slug dependent but smad4 independent in HaCaT cells (Levy and Hill, 2005 blue right-pointing triangle). Here, we found that the Smad4-dependent pathway up-regulates GEF-H1 expression induced by TGF-β. Hence, one way by which Smad4-dependent signaling drives the migratory phenotype is by controlling the expression of GEF-H1 and, thereby, Rho activation.

In epithelial cells, GEF-H1, a guanine nucleotide exchange factor for RhoA, associates with tight junctions; and functions in the regulation of paracellular permeability, cell proliferation and junction disassembly (Benais-Pont et al., 2003 blue right-pointing triangle; Aijaz et al., 2005 blue right-pointing triangle; Birukova et al., 2006 blue right-pointing triangle; Samarin et al., 2007 blue right-pointing triangle). We now found that GEF-H1 also supports α-SMA expression and cell migration. The activity of Rho GTPases has to be carefully timed and controlled to guide epithelial proliferation and differentiation (Fujita and Braga, 2005 blue right-pointing triangle; Heasman and Ridley, 2008 blue right-pointing triangle; Nelson, 2008 blue right-pointing triangle; Wheelock et al., 2008 blue right-pointing triangle; Yu et al., 2008 blue right-pointing triangle). In epithelial cells in culture, the endogenous levels of expression of GEF-H1 are generally high; hence, it was previously poorly understood how expression of GEF-H1 is stimulated. In adult epithelial tissues, however, GEF-H1 levels are low. The same is true for the RPE as both primary culture and in vivo experiments indicate that expression of GEF-H1 is low in differentiated cells (Figures 1 and and8).8). Our data now show that TGF-β induces a striking up-regulation of GEF-H1 in Smad4-dependent pathway and in two different epithelial models.

In primary RPE cells in culture, transdifferentiation can be induced when cells are plated at low density, resulting in increased expression of α-SMA (Grisanti and Guidry, 1995 blue right-pointing triangle; Lee et al., 2001 blue right-pointing triangle; Wiencke et al., 2003 blue right-pointing triangle) as well as GEF-H1 (Figure 3). As it has been suggested that at low confluence RPE cell secrete TGF-β, we inhibited the TGF-β receptor I with the ALK5 inhibitor and indeed found that it prevented morphological degeneration as well as α-SMA and GEF-H1 up-regulation. Thus, exogenous as well as autocrine TGF-β induces GEF-H1, indicating that TGF-β is a major driver of GEF-H1 expression in epithelial cells. Although Smad4, but not Slug, is required for GEF-H1 up-regulation, how transcription is induced is not clear yet. The late and sustained expression of GEF-H1 indicates that it may be an indirect target of Smad4-dependent signaling. Recent evidence also shows that GEF-H1 activation is regulated by phosphorylation and TNF-alpha (Zenke et al., 2004 blue right-pointing triangle; Callow et al., 2005 blue right-pointing triangle; Chang and Lee, 2006 blue right-pointing triangle; Fujishiro et al., 2008 blue right-pointing triangle; Kakiashvili et al., 2009 blue right-pointing triangle; Nie et al., 2009 blue right-pointing triangle), suggesting that GEF-H1 regulation occurs at different levels and is target by different signaling pathways. Nevertheless, as most adult tissues express little GEF-H1, up-regulation represents an important step in activation of GEF-H1 signaling.

Regulation of Rho activity has previously been linked to TGF-β stimulation in different cell types (Bhowmick et al., 2001 blue right-pointing triangle; Bakin et al., 2002 blue right-pointing triangle; Edlund et al., 2002 blue right-pointing triangle). TGF-β also enhances the expression of RhoB (Engel et al., 1998 blue right-pointing triangle) as well as NET1, a RhoA-specific guanine exchange factor (Shen et al., 2001 blue right-pointing triangle; Levy and Hill, 2005 blue right-pointing triangle). However, we have not been able to detect NET1 in RPE cells treated with TGF-β1 (not shown). We also failed to detect up-regulation of other Rho exchange factors such as ARHGEF18 by immunoblotting (Supplemental Figure 3) as well as by means of cDNA arrays (Table 1). Hence, RPE cells seem to up-regulate GEF-H1 specifically, indicating that the exchange factor is a major TGF-β target gene in respect to Rho signaling.

As activation of the α-SMA promoter seems to involve Rho signaling in TGF-β-induced transdifferentiation of renal epithelial cells (Masszi et al., 2003 blue right-pointing triangle), we assessed α-SMA promoter activity and protein expression in RPE cells stimulated with TGF-β in the presence or absence of GEF-H1 inhibition. Our results indicate that GEF-H1 mediates Rho stimulation to induce α-SMA expression by activation of its promoter. Therefore, GEF-H1 is not only a target gene of TGF-β, but functionally contributes to the expression of marker genes associated with transdifferentiation and fibrosis. Hence, GEF-H1 represents a possible target to inhibit α-SMA expression for the treatment of fibrosis.

Although Rho signaling is thought to be important for fibrosis, the mechanisms that drive Rho activation in fibrosis had previously not been identified. We have observed strong increases in GEF-H1 expression in RPE cells of patients with retinal detachments due to different types of insults that triggered retinopathies and disorganization of the pigment epithelium (i.e., dislocation from Bruch's membrane). RPE cells have been suggested to contribute to retinal detachments in PVR and in response to trauma (Fuchs et al., 1991 blue right-pointing triangle; Saika et al., 2004 blue right-pointing triangle; Zheng et al., 2004 blue right-pointing triangle), and inhibition of the Rho-kinase pathway suppresses the expression of α-SMA in rabbit RPE cells in culture and attenuates retinal detachment in a rabbit PVR model (Zheng et al., 2004 blue right-pointing triangle; Kita et al., 2008 blue right-pointing triangle). Furthermore, the analysis of expression of GEF-H1 in eye sections from patients with retinal detachments demonstrated that GEF-H1 is up-regulated in migratory RPE cells (Figure 8), suggesting that increased expression of GEF-H1 is an early event in the translocation of RPE from their normal location at the back of the retina and is likely to contribute transdifferentiation in vivo. Thus, GEF-H1 represents a possible therapeutic target to attenuate RPE migration and retinal detachments after injury or surgery.

TGF-β is involved in cell migration in different cell types using Smad-dependent or -independent pathways. Rho also plays a role in cell migration. Our results show that GEF-H1 regulates Rho activation and migration induced TGF-β in primary RPE cells, HaCaT as well as MDCK cells, a spontaneously immortalized cell line that constitutively expresses high levels of GEF-H1. When this article was under revision, a study was published that suggested that GEF-H1 also regulates migration in a tumor cell line (Nalbant et al., 2009 blue right-pointing triangle). Thus, activation of Rho signaling by GEF-H1 seems to be connected to cell migration in different cellular contexts, indicating that GEF-H1 represents a link by which TGF-β stimulates molecular mechanisms of general importance for cell migration and gene expression.

In summary, we have identified a new target and functional effector of TGF-β signaling, the Rho guanine nucleotide exchange factor GEF-H1 that regulates expression genes related to transdifferentiation, such as α-SMA, and epithelial cell migration. Up-regulation of GEF-H1 occurs in migratory RPE in patients with retinal detachments, suggesting that GEF-H1 is a marker and novel therapeutic target for retinal detachments, and may be a crucial signaling protein to be targeted during the manipulation of RPE cells for transplantation and in fibrotic diseases.

Supplementary Material

[Supplemental Materials]


We are thankful to Dr. G. K. Owens (University of Virginia, Charlottesville, VA) for the α-SMA promoter constructs and to Christine Gaughan for the immunocytochemistry of human retinas. This research was supported by Medical Research Council grant G0400678), Association for International Cancer Research grant 06-522), and the Wellcome Trust grant 084678/Z/08/Z.


This article was published online ahead of print in MBC in Press (http://www.molbiolcell.org/cgi/doi/10.1091/mbc.E09-07-0567) on January 20, 2010.


  • Ablonczy Z., Crosson C. E. VEGF modulation of retinal pigment epithelium resistance. Exp. Eye Res. 2007;85:762–771. [PMC free article] [PubMed]
  • Aijaz S., D'Atri F., Citi S., Balda M. S., Matter K. Binding of GEF-H1 to the tight junction-associated adaptor cingulin results in inhibition of Rho signaling and G1/S phase transition. Dev. Cell. 2005;8:777–786. [PubMed]
  • Bainbridge J. W., Stephens C., Parsley K., Demaison C., Halfyard A., Thrasher A. J., Ali R. R. In vivo gene transfer to the mouse eye using an HIV-based lentiviral vector; efficient long-term transduction of corneal endothelium and retinal pigment epithelium. Gene Ther. 2001;8:1665–1668. [PubMed]
  • Bakin A. V., Rinehart C., Tomlinson A. K., Arteaga C. L. p38 mitogen-activated protein kinase is required for TGFbeta-mediated fibroblastic transdifferentiation and cell migration. J. Cell Sci. 2002;115:3193–3206. [PubMed]
  • Benais-Pont G., Punn A., Flores-Maldonado C., Eckert J., Raposo G., Fleming T. P., Cereijido M., Balda M. S., Matter K. Identification of a tight junction-associated guanine nucleotide exchange factor that activates Rho and regulates paracellular permeability. J. Cell Biol. 2003;160:729–740. [PMC free article] [PubMed]
  • Bhowmick N. A., Ghiassi M., Bakin A., Aakre M., Lundquist C. A., Engel M. E., Arteaga C. L., Moses H. L. Transforming growth factor-beta1 mediates epithelial to mesenchymal transdifferentiation through a RhoA-dependent mechanism. Mol. Biol. Cell. 2001;12:27–36. [PMC free article] [PubMed]
  • Birkenfeld J., Nalbant P., Yoon S. H., Bokoch G. M. Cellular functions of GEF-H1, a microtubule-regulated Rho-GEF: is altered GEF-H1 activity a crucial determinant of disease pathogenesis? Trends Cell Biol. 2008;18:210–219. [PubMed]
  • Birukova A. A., Adyshev D., Gorshkov B., Bokoch G. M., Birukov K. G., Verin A. D. GEF-H1 is involved in agonist-induced human pulmonary endothelial barrier dysfunction. Am. J. Physiol. Lung Cell Mol. Physiol. 2006;290:L540–L548. [PubMed]
  • Bos J. L., Rehmann H., Wittinghofer A. GEFs and GAPs: critical elements in the control of small G proteins. Cell. 2007;129:865–877. [PubMed]
  • Callow M. G., Zozulya S., Gishizky M. L., Jallal B., Smeal T. PAK4 mediates morphological changes through the regulation of GEF-H1. J. Cell Sci. 2005;118:1861–1872. [PubMed]
  • Chambers R. C., Leoni P., Kaminski N., Laurent G. J., Heller R. A. Global expression profiling of fibroblast responses to transforming growth factor-beta1 reveals the induction of inhibitor of differentiation-1 and provides evidence of smooth muscle cell phenotypic switching. Am. J. Pathol. 2003;162:533–546. [PMC free article] [PubMed]
  • Chang Z. F., Lee H. H. RhoA signaling in phorbol ester-induced apoptosis. J. Biomed. Sci. 2006;13:173–180. [PubMed]
  • Coleman M. L., Sahai E. A., Yeo M., Bosch M., Dewar A., Olson M. F. Membrane blebbing during apoptosis results from caspase-mediated activation of ROCK I. Nat. Cell Biol. 2001;3:339–345. [PubMed]
  • Connor T. B., Jr, et al. Correlation of fibrosis and transforming growth factor-beta type 2 levels in the eye. J. Clin. Invest. 1989;83:1661–1666. [PMC free article] [PubMed]
  • Derynck R., Zhang Y. E. Smad-dependent and Smad-independent pathways in TGF-beta family signalling. Nature. 2003;425:577–584. [PubMed]
  • Edlund S., Landstrom M., Heldin C. H., Aspenstrom P. Transforming growth factor-beta-induced mobilization of actin cytoskeleton requires signaling by small GTPases Cdc42 and RhoA. Mol. Biol. Cell. 2002;13:902–914. [PMC free article] [PubMed]
  • Engel M. E., Datta P. K., Moses H. L. RhoB is stabilized by transforming growth factor beta and antagonizes transcriptional activation. J. Biol. Chem. 1998;273:9921–9926. [PubMed]
  • Fu P., Liu F., Su S., Wang W., Huang X. R., Entman M. L., Schwartz R. J., Wei L., Lan H. Y. Signaling mechanism of renal fibrosis in unilateral ureteral obstructive kidney disease in ROCK1 knockout mice. J. Am. Soc. Nephrol. 2006;17:3105–3114. [PubMed]
  • Fuchs U., Kivela T., Tarkkanen A. Cytoskeleton in normal and reactive human retinal pigment epithelial cells. Invest. Ophthalmol. Vis. Sci. 1991;32:3178–3186. [PubMed]
  • Fujishiro S. H., Tanimura S., Mure S., Kashimoto Y., Watanabe K., Kohno M. ERK1/2 phosphorylate GEF-H1 to enhance its guanine nucleotide exchange activity toward RhoA. Biochem. Biophys. Res. Commun. 2008;368:162–167. [PubMed]
  • Fujita Y., Braga V. Epithelial cell shape and Rho small GTPases. Novartis Found Symp. 2005;269:144–155. discussion 155–148, 223–230. [PubMed]
  • Grisanti S., Guidry C. Transdifferentiation of retinal pigment epithelial cells from epithelial to mesenchymal phenotype. Invest. Ophthalmol. Vis Sci. 1995;36:391–405. [PubMed]
  • Hall A. Rho GTPases and the control of cell behaviour. Biochem. Soc. Trans. 2005;33:891–895. [PubMed]
  • Heasman S. J., Ridley A. J. Mammalian Rho GTPases: new insights into their functions from in vivo studies. Nat. Rev. Mol. Cell Biol. 2008;9:690–701. [PubMed]
  • Heldin C. H., Landstrom M., Moustakas A. Mechanism of TGF-beta signaling to growth arrest, apoptosis, and epithelial-mesenchymal transition. Curr. Opin. Cell Biol. 2009 [PubMed]
  • Hill C. S., Wynne J., Treisman R. The Rho family GTPases RhoA, Rac1, and CDC42Hs regulate transcriptional activation by SRF. Cell. 1995;81:1159–1170. [PubMed]
  • Hiscott P., Sheridan C., Magee R. M., Grierson I. Matrix and the retinal pigment epithelium in proliferative retinal disease. Prog. Retin. Eye Res. 1999;18:167–190. [PubMed]
  • Ikenouchi J., Matsuda M., Furuse M., Tsukita S. Regulation of tight junctions during the epithelium-mesenchyme transition: direct repression of the gene expression of claudins/occludin by Snail. J. Cell Sci. 2003;116:1959–1967. [PubMed]
  • Kakiashvili E., Speight P., Waheed F., Seth R., Lodyga M., Tanimura S., Kohno M., Rotstein O. D., Kapus A., Szaszi K. GEF-H1 mediates tumor necrosis factor-alpha-induced Rho activation and myosin phosphorylation: role in the regulation of tubular paracellular permeability. J. Biol. Chem. 2009;284:11454–11466. [PMC free article] [PubMed]
  • Keese C. R., Wegener J., Walker S. R., Giaever I. Electrical wound-healing assay for cells in vitro. Proc. Natl. Acad. Sci. USA. 2004;101:1554–1559. [PMC free article] [PubMed]
  • Kim J. S., et al. Transforming growth factor-beta1 regulates macrophage migration via RhoA. Blood. 2006;108:1821–1829. [PubMed]
  • Kita T., et al. Role of TGF-beta in proliferative vitreoretinal diseases and ROCK as a therapeutic target. Proc. Natl. Acad. Sci. USA. 2008;105:17504–17509. [PMC free article] [PubMed]
  • Kon C. H., Occleston N. L., Aylward G. W., Khaw P. T. Expression of vitreous cytokines in proliferative vitreoretinopathy: a prospective study. Invest. Ophthalmol. Vis. Sci. 1999;40:705–712. [PubMed]
  • Kreis T. E. Microtubules containing detyrosinated tubulin are less dynamic. EMBO J. 1987;6:2597–2606. [PMC free article] [PubMed]
  • Lee S. C., Kim S. H., Koh H. J., Kwon O. W. TGF-betas synthesized by RPE cells have autocrine activity on mesenchymal transformation and cell proliferation. Yonsei Med. J. 2001;42:271–277. [PubMed]
  • Levy L., Hill C. S. Smad4 dependency defines two classes of transforming growth factor {beta} (TGF-{beta}) target genes and distinguishes TGF-{beta}-induced epithelial-mesenchymal transition from its antiproliferative and migratory responses. Mol. Cell Biol. 2005;25:8108–8125. [PMC free article] [PubMed]
  • Liu Y. Renal fibrosis: new insights into the pathogenesis and therapeutics. Kidney Int. 2006;69:213–217. [PubMed]
  • Liu Y., Sinha S., Owens G. A transforming growth factor-beta control element required for SM alpha-actin expression in vivo also partially mediates GKLF-dependent transcriptional repression. J. Biol. Chem. 2003;278:48004–48011. [PubMed]
  • Mack C. P., Owens G. K. Regulation of smooth muscle alpha-actin expression in vivo is dependent on CArG elements within the 5′ and first intron promoter regions. Circ. Res. 1999;84:852–861. [PubMed]
  • Masszi A., Di Ciano C., Sirokmany G., Arthur W. T., Rotstein O. D., Wang J., McCulloch C. A., Rosivall L., Mucsi I., Kapus A. Central role for Rho in TGF-beta1-induced alpha-smooth muscle actin expression during epithelial-mesenchymal transition. Am. J. Physiol. Renal Physiol. 2003;284:F911–F924. [PubMed]
  • Miano J. M., Long X., Fujiwara K. Serum response factor: master regulator of the actin cytoskeleton and contractile apparatus. Am. J. Physiol. Cell Physiol. 2007;292:C70–C81. [PubMed]
  • Nalbant P., Chang Y. C., Birkenfeld J., Chang Z. F., Bokoch G. M. Guanine nucleotide exchange factor-H1 regulates cell migration via localized activation of RhoA at the leading edge. Mol. Biol. Cell. 2009;20:4070–4082. [PMC free article] [PubMed]
  • Nelson W. J. Regulation of cell-cell adhesion by the cadherin-catenin complex. Biochem. Soc. Trans. 2008;36:149–155. [PMC free article] [PubMed]
  • Nie M., Aijaz S., Leefa Chong San I. V., Balda M. S., Matter K. The Y-box factor ZONAB/DbpA associates with GEF-H1/Lfc and mediates Rho-stimulated transcription. EMBO Rep. 2009;10:1125–1131. [PMC free article] [PubMed]
  • Nishikimi T., Matsuoka H. Molecular mechanisms and therapeutic strategies of chronic renal injury: renoprotective effect of rho-kinase inhibitor in hypertensive glomerulosclerosis. J. Pharmacol. Sci. 2006;100:22–28. [PubMed]
  • Obara K., Bilim V., Suzuki K., Kobayashi K., Hara N., Kasahara T., Nishiyama T., Takahashi K. Transforming growth factor-beta1 regulates cell growth and causes downregulation of SMemb/non-muscle myosin heavy chain B mRNA in human prostate stromal cells. Scand. J. Urol. Nephrol. 2005;39:366–371. [PubMed]
  • Ozdamar B., Bose R., Barrios-Rodiles M., Wang H. R., Zhang Y., Wrana J. L. Regulation of the polarity protein Par6 by TGFbeta receptors controls epithelial cell plasticity. Science. 2005;307:1603–1609. [PubMed]
  • Peinado H., Olmeda D., Cano A. Snail, Zeb and bHLH factors in tumour progression: an alliance against the epithelial phenotype? Nat. Rev. Cancer. 2007;7:415–428. [PubMed]
  • Posern G., Treisman R. Actin' together: serum response factor, its cofactors and the link to signal transduction. Trends Cell Biol. 2006;16:588–596. [PubMed]
  • Roberts A. B., Tian F., Byfield S. D., Stuelten C., Ooshima A., Saika S., Flanders K. C. Smad3 is key to TGF-beta-mediated epithelial-to-mesenchymal transition, fibrosis, tumor suppression and metastasis. Cytokine Growth Factor Rev. 2006;17:19–27. [PubMed]
  • Ross S., Hill C. S. How the Smads regulate transcription. Int. J. Biochem. Cell Biol. 2008;40:383–408. [PubMed]
  • Ryan X. P., Alldritt J., Svenningsson P., Allen P. B., Wu G. Y., Nairn A. C., Greengard P. The Rho-specific GEF Lfc interacts with neurabin and spinophilin to regulate dendritic spine morphology. Neuron. 2005;47:85–100. [PubMed]
  • Saika S., et al. Smad3 is required for dedifferentiation of retinal pigment epithelium following retinal detachment in mice. Lab. Invest. 2004;84:1245–1258. [PubMed]
  • Saika S., et al. Epithelial-mesenchymal transition as a therapeutic target for prevention of ocular tissue fibrosis. Endocr. Metab. Immune. Disord. Drug Targets. 2008;8:69–76. [PubMed]
  • Samarin S. N., Ivanov A. I., Flatau G., Parkos C. A., Nusrat A. Rho/Rho-associated kinase-II signaling mediates disassembly of epithelial apical junctions. Mol. Biol. Cell. 2007;18:3429–3439. [PMC free article] [PubMed]
  • Schmierer B., Hill C. S. TGFbeta-SMAD signal transduction: molecular specificity and functional flexibility. Nat. Rev. Mol. Cell Biol. 2007;8:970–982. [PubMed]
  • Seo J., Bakay M., Chen Y. W., Hilmer S., Shneiderman B., Hoffman E. P. Interactively optimizing signal-to-noise ratios in expression profiling: project-specific algorithm selection and detection p-value weighting in Affymetrix microarrays. Bioinformatics. 2004;20:2534–2544. [PubMed]
  • Shen X., Li J., Hu P. P., Waddell D., Zhang J., Wang X. F. The activity of guanine exchange factor NET1 is essential for transforming growth factor-beta-mediated stress fiber formation. J. Biol. Chem. 2001;276:15362–15368. [PubMed]
  • Sinha S., Hoofnagle M. H., Kingston P. A., McCanna M. E., Owens G. K. Transforming growth factor-beta1 signaling contributes to development of smooth muscle cells from embryonic stem cells. Am. J. Physiol. Cell Physiol. 2004;287:C1560–C1568. [PubMed]
  • Steed E., Rodrigues N.T.L., Balda M. S., Matter K. Identification of MarvelD3 as a tight junction associated transmembrane protein of the occludin family. BMC Cell Biol. 2009;10:95. [PMC free article] [PubMed]
  • Thuault S., Tan E. J., Peinado H., Cano A., Heldin C. H., Moustakas A. HMGA2 and Smads co-regulate SNAIL1 expression during induction of epithelial-to-mesenchymal transition. J. Biol. Chem. 2008;283:33437–33446. [PMC free article] [PubMed]
  • Tsuchida K., Zhu Y., Siva S., Dunn S. R., Sharma K. Role of Smad4 on TGF-beta-induced extracellular matrix stimulation in mesangial cells. Kidney Int. 2003;63:2000–2009. [PubMed]
  • Wamhoff B. R., Bowles D. K., Owens G. K. Excitation-transcription coupling in arterial smooth muscle. Circ. Res. 2006;98:868–878. [PubMed]
  • Wheelock M. J., Shintani Y., Maeda M., Fukumoto Y., Johnson K. R. Cadherin switching. J. Cell Sci. 2008;121:727–735. [PubMed]
  • Wiencke A. K., Kiilgaard J. F., Nicolini J., Bundgaard M., Ropke C., La Cour M. Growth of cultured porcine retinal pigment epithelial cells. Acta Ophthalmol. Scand. 2003;81:170–176. [PubMed]
  • Yu W., Shewan A. M., Brakeman P., Eastburn D. J., Datta A., Bryant D. M., Fan Q. W., Weiss W. A., Zegers M. M., Mostov K. E. Involvement of RhoA, ROCK I and myosin II in inverted orientation of epithelial polarity. EMBO Rep. 2008;9:923–929. [PMC free article] [PubMed]
  • Zavadil J., Bottinger E. P. TGF-beta and epithelial-to-mesenchymal transitions. Oncogene. 2005;24:5764–5774. [PubMed]
  • Zenke F. T., Krendel M., DerMardirossian C., King C. C., Bohl B. P., Bokoch G. M. p21-activated kinase 1 phosphorylates and regulates 14-3-3 binding to GEF-H1, a microtubule-localized Rho exchange factor. J. Biol. Chem. 2004;279:18392–18400. [PubMed]
  • Zhang Y. E. Non-Smad pathways in TGF-beta signaling. Cell Res. 2009;19:128–139. [PMC free article] [PubMed]
  • Zheng Y., Bando H., Ikuno Y., Oshima Y., Sawa M., Ohji M., Tano Y. Involvement of rho-kinase pathway in contractile activity of rabbit RPE cells in vivo and in vitro. Invest. Ophthalmol. Vis. Sci. 2004;45:668–674. [PubMed]

Articles from Molecular Biology of the Cell are provided here courtesy of American Society for Cell Biology
PubReader format: click here to try


Save items

Related citations in PubMed

See reviews...See all...

Cited by other articles in PMC

See all...


  • Compound
    PubChem chemical compound records that cite the current articles. These references are taken from those provided on submitted PubChem chemical substance records. Multiple substance records may contribute to the PubChem compound record.
  • Gene
    Gene records that cite the current articles. Citations in Gene are added manually by NCBI or imported from outside public resources.
  • Gene (nucleotide)
    Gene (nucleotide)
    Records in Gene identified from shared sequence and PMC links.
  • GEO Profiles
    GEO Profiles
    Gene Expression Omnibus (GEO) Profiles of molecular abundance data. The current articles are references on the Gene record associated with the GEO profile.
  • HomoloGene
    HomoloGene clusters of homologous genes and sequences that cite the current articles. These are references on the Gene and sequence records in the HomoloGene entry.
  • MedGen
    Related information in MedGen
  • Nucleotide
    Primary database (GenBank) nucleotide records reported in the current articles as well as Reference Sequences (RefSeqs) that include the articles as references.
  • Protein
    Protein translation features of primary database (GenBank) nucleotide records reported in the current articles as well as Reference Sequences (RefSeqs) that include the articles as references.
  • PubMed
    PubMed citations for these articles
  • Substance
    PubChem chemical substance records that cite the current articles. These references are taken from those provided on submitted PubChem chemical substance records.

Recent Activity

Your browsing activity is empty.

Activity recording is turned off.

Turn recording back on

See more...