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Research Article
TGFβ stimulated re-epithelialisation is regulated by CTGF and Ras/MEK/ERK signalling Genevieve A. Secker a , Alex J. Shortt a , Edith Sampson b , Quenten P. Schwarz c , Gregory S. Schultz b , Julie T. Daniels a,⁎ a
Department of Pathology, Cells for Sight Transplantation and Research Programme, UCL Institute of Ophthalmology, 11-43 Bath Street, London, EC1V 9EL, UK b Department of ObGyn, The University of Florida, Gainsville, FL 32610, USA c Department of Cellular Therapy, UCL Institute of Ophthalmology, London, EC1V 9EL, UK
ARTICLE INFORMATION
ABS T R AC T
Article Chronology:
The complex mechanisms by which transforming growth factor β (TGFβ) regulate re-
Received 18 May 2007
epithelialisation following injury of stratified epithelia are not fully understood. TGFβ
Revised version received
signals via binding to distinct receptors activating downstream effectors, including Smads
31 August 2007
which initiate transcription of target genes. However, studies have shown that TGFβ can
Accepted 4 September 2007
also signal independently of Smads through MAPK pathways, demonstrating the diversity
Available online 7 September 2007
of TGFβ signalling. Connective tissue growth factor (CTGF) is strongly induced by and acts downstream of TGFβ causing pathophysiology in tissues by inducing matrix deposition,
Keywords:
conversion of fibroblasts into contractile myofibroblasts (e.g. dermis and corneal stroma)
TGFβ
and stimulation of epithelial-to-mesenchymal transition (e.g. kidney and lung) all of which
Re-epithelialisation
are known to cause fibrosis. However, a role for CTGF in epithelial cell function which does
Migration
not involve direct contribution to fibrosis has not been demonstrated. We show for the first
CTGF
time that synthesis of CTGF in cultures of human corneal epithelial cells is induced by TGFβ
Ras/MEK/ERK
through the Ras/MEK/ERK MAPK signalling pathway and that this is required for reepithelialisation to occur through cell migration. These data reveal a novel function for CTGF in the regulation of epithelial tissue repair beyond its established role in fibrosis, and further highlight the complexity of TGFβ regulation of epithelial cell function. © 2007 Elsevier Inc. All rights reserved.
Introduction The pathways involved in TGFβ regulation of re-epithelialisation of stratified epithelia, specifically inhibition of proliferation and promotion of migration of epithelial cells to facilitate wound closure are unclear [1]. Members of the TGFβ family can initiate cellular responses by binding to distinct receptors with
intrinsic serine/threonine kinase activity and activation of specific downstream intracellular effectors termed Smad proteins. The Smads relay the signal from the cell membrane to the nucleus, where they affect transcription of target genes [2]. For example Smad 4 is necessary for TGFβ-induced cell cycle arrest and migration in immortalised keratinocytes (HaCaT) and human pancreatic tumour cells (Colo-357) [3].
⁎ Corresponding author. Fax: +44 2076086887. E-mail address: [email protected] (J.T. Daniels). Abbreviations: TGFβ, transforming growth factor β; SFM, serum free medium; DMEM, Dulbecco's modified Eagle's medium; GAPDH, glyseraldehyde-3-phosphate dehydrogenase; BSA, bovine serum albumin; PBS, phosphate buffered saline; KSFM, keratinocyte serum free medium; HCEC, human corneal epithelial cells; S.E.M., standard error 0014-4827/$ – see front matter © 2007 Elsevier Inc. All rights reserved. doi:10.1016/j.yexcr.2007.09.001
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However, there is also evidence that TGFβ-stimulated epithelial cell migration can occur independently of Smad 4 with signalling via the ERK and JNK MAPK pathways [4,5]. During re-epithelialisation of mouse corneas TGFβ was found to enhance re-epithelialisation by suppressing cell division and enhancing cell migration via the p38 MAPK pathway, independent of Smad signalling [6]. These findings illustrate the complexities of TGFβ signalling and regulation [7]. During early development and tissue repair, TGFβ gene expression in fibroblasts is co-ordinately regulated with that of connective tissue growth factor (CTGF) [8,9]. CTGF is a member of the CCN (CTGF, Cyr61 and Nov) protein family [10] and was first identified in conditioned medium collected from cultured human vascular endothelial cells [11]. CTGF is a cysteine-rich, heparin binding protein,[11] whose gene expression is strongly induced by TGFβ in fibroblasts [8,12]. A 1-h exposure to TGFβ is sufficient to induce CTGF gene transcription for up to 36 h in fibroblasts [13]. A novel TGFβ response element controls transcription in both human and murine CTGF promoters
[13]. These observations suggest that CTGF is a downstream mediator of TGFβ activity in fibroblasts [13]. Indeed, CTGF is known to mediate key cellular events in response to TGFβ including migration [13], proliferation [14,15], matrix production [16] and contraction and differentiation of fibroblast into myofibroblasts [14,17,18], which are all elements of a fibrotic response. Previous reports have indicated that CTGF induction by TGFβ is specific for connective tissue cells and does not occur in epithelial cells [19,9,13,20,21,22]. Nevertheless, a growing body of evidence supports the notion that a variety of epithelial cell types can produce CTGF during development and in adult tissues [23–27]. However, the functional significance of CTGF production during homeostasis and following epithelial injury of stratified epithelia such as those of the cornea and skin remain unclear. In the present study, we report that CTGF is produced by differentiating human corneal epithelial cells (HCEC) and that CTGF has an additional role for mediating the activity of TGFβ
Fig. 1 – TGFβ stimulates scratch wound closure in primary HCEC and HCE-T through migration. (A,B) A wound was introduced into primary human corneal epithelial cells and HCE-T serum starved monolayers with a pipette tip. Scratch wound closure was determined in cultures incubated in SFM with (0.1, 1, 10) or without (0) TGFβ1 (ng/ml). All three concentrations significantly stimulated primary corneal epithelial cell (A), and HCE-T cell (B), re-epithelialisation after 20 and 8 h respectively. (C) Cell proliferation was determined using WST-1 assay. Subconfluent serum starved HCE-T cells were incubated in SFM with (0.1, 1, 10) or without (0) TGFβ1 (ng/ml) for 24 h. TGFβ1 significantly inhibited HCE-T cell proliferation at concentrations of 0.1 and 10 ng/ml. (D) Confluent HCE-T cells cultured in cloning rings were growth arrested prior to ring removal and addition of treatment groups. Cells were incubated in SFM with (0.1, 1, 10) or without (0) TGFβ1 (ng/ml) for 72 h, with all concentrations stimulating migration. Error bars represents S.E.M. Asterisks indicate values that are significantly different from TGFβ1 (0) (p < 0.05).
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during tissue repair beyond the regulation of fibrosis; namely regulation of re-epithelialisation. Specifically, CTGF is required for TGFβ-stimulated epithelial cell migration. Furthermore we have demonstrated a role for ERK and p38 MAPK signalling in both TGFβ-stimulated CTGF production and migration in epithelial cells derived from stratified epithelium.
Materials and methods
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described previously [19], were used at concentrations 0– 10 ng/ml in serum free media (SFM) consisting of basal media supplemented with 0.1% bovine serum albumin (BSA). ERK 1/2, p38 and Ras pathways were inhibited using PD98059 (100 μM; Cell Signalling Technology, ERK 1/2 inhibitor, SB203580 (10 μM; Promega, Madison, WI) and Farnesylthiosalicylic acid (5 μM; FTS; Biomol International, Exeter, UK). Antisense oligodeoxynucleotides (AS), synthesised as described previously [17], were used at concentration of 1, 10 and 100 μM and the scrambled control was used at 100 μM.
Cells, cell culture and reagents Scratch wound model Human corneas supplied by Moorfields Eye Hospital National Health Service (NHS) Foundation Trust Eye Bank were used in the study, in accordance with the Declaration of Helsinki for the use of human tissue. Human corneal epithelial cells (HCEC) were isolated and cultured in keratinocyte serum free medium (KSFM) with supplements (Invitrogen, Scotland, UK). Corneal segments were dissected into quadrants and placed in 1x Dispase II (Roche Diagnostics Ltd, East Sussex, UK) overnight at 4 °C before epithelial cells were gently scraped off using forceps. Experiments were performed with cells between passages 1 and 3. HCE-T cells (Riken, Tsukuba Institute) were maintained in corneal epithelial cell medium (CECM) consisting of DMEM:F12 (1:1) (Invitrogen) supplemented with 5% FCS (Invitrogen), 10 ng/ml Epidermal Growth Factor (Invitrogen), 0.1 ng/ml Cholera Toxin (Sigma-Aldrich Chemical Co. Ltd., Poole, UK) and 5 ug/ml Insulin (Sigma-Aldrich). HaCaT cells (a kind gift from F. Watt, Cancer Research UK, London, UK) were cultured in DMEM (Invitrogen) supplemented with 10% FCS. All cells were cultured in the presence of 5% CO2 in air at 37 °C and were grown in the presence of antibiotics. Growth factors, TGFβ1 (R&D Systems Europe Ltd., Oxon, UK) and CTGF produced by baculovirus expression in insect cells, as
HCE-T or HCEC cultured to confluence in marked 96-well plates were serum starved in SFM for 24 h. A 200-μl plastic pipette tip was used to scratch the monolayers, the wounded cells were digitally photographed under an inverted phase contrast microscope prior to addition of treatment groups and 8 (HCE-T) or 18 h (HCEC) post wounding. The distance between the scratch wound edges were measured using Image Tool software (UTHSCSA, San Antonio, TX), with average wound areas calculated to determine percentage wound closure (re-epithelialisation) between time points.
Cell proliferation assay HCE-T proliferation was monitored using the water soluble tetrazolium-1 (WST-1) reagent (Roche Diagnostics Ltd) according to the manufacturer's instructions. Briefly, HCE-T cells were seeded into 96-well tissue culture plates, at a density of 5 × 103 cells per well in CECM and cultured for 18 h. CECM was removed, and the cells were washed three times prior to the addition of SFM for a 24-h serum starvation period. Treatment groups and serum free baseline control
Fig. 2 – TGFβ utilises ERK 1/2 and p38 pathway signalling for re-epithelialisation. (A) Phosphorylated ERK 1/2 and p38 expression over time was determined by immunoblotting whole cell lysates stimulated with TGFβ1 (10 ng/ml) using specific antibodies to phosphorylated ERK 1/2, p38 and GAPDH. ERK 1/2 phosphorylation occurred at 1 min and was most prominent at 10 min with p38 phosphorylation occurring at 3 min and then becoming most prominent at 10 min. GAPDH was used as a loading control. (B) A wound was introduced into HCE-T serum starved monolayers with a pipette tip. Scratch wound closure was determined in cultures incubated with (+) or without (−) TGFβ1 (10 ng/ml) in presence and absence (control) of MEK1 (PD98059) and p38 (SB203580) inhibitors. Inhibition of both pathways prevented TGFβ1 stimulated wound closure. Error bars represents S.E.M. Asterisks indicate values that are significantly different from TGFβ1 (+) control (p < 0.05).
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medium (100 μl) were added to the cells (eight replicates) and incubated for 22 h. Ten microlitres of WST-1 reagent was added to each well and the plate was incubated at 37 °C for 2 h. The absorbance, related to the number of viable cells converting the reagent to coloured formazan crystals, was read at 450 nm.
Collection of conditioned medium
Colony dispersion
Immunoblot analysis
Cell migration was assessed using a colony dispersion assay as described by Pilcher et al. [28] Briefly, HCE-T or HaCaT cells were seeded and cultured to confluence within cloning rings. The cells were cultured for a further 24 h in the presence of 1 mM hydroxyurea (Sigma-Aldrich) to induce growth arrest. After removal of the rings, the cells were thoroughly washed with PBS prior to the addition of treatment groups. Cells were allowed to migrate for 72 h, then washed with PBS three times, fixed with 100% chilled methanol for 10 min and stained with Harris hematoxylin (Shandon Life Sciences International, Europe, Ltd., UK) for 30 min prior to washing with water. Dispersion areas were digitally photographed and measured in pixels with Image Tool analysis software.
Sub confluent HCE-T were serum starved in SFM for 48 h prior to addition of treatment groups. Cell monolayers were collected at time points 0 to 1 h using radioimmune precipitation assay (RIPA) buffer (1% Nonidet P-40/0.5% deoxycholic acid/ 0.1% SDS/158 mM NaCl/50 mM Tris/pH 7.2) and centrifugated at 13,000 rpm for 5 min prior to supernatant collection. Proteins were resolved by SDS/10% PAGE (NuPAGE; Invitrogen, Groningen, The Netherlands) for 60 min at 150 V in Tris-acetate running buffer (Invitrogen) and transferred to nitrocellulose for 2 h at 30 V with a NOVEX transfer apparatus and Transfer Buffer (Invitrogen). Nitrocellulose was blocked with TBSTT (TBS/0.1% Tween-20/0.2% Triton-X) with 1% BSA for 1 hr, probed with antiACTIVE® MAPK (Promega, Southampton, UK; 1:2000) and anti-
Cells were seeded at 60% confluence and allowed to adhere overnight prior to serum starvation for 24 h. Conditioned media was collected following addition of treatment groups for 24 h or directly following serum starvation period.
Fig. 3 – CTGF is produced during HCEC differentiation and TGFβ stimulates CTGF production in epithelial cells via the ERK 1/2 and p38 pathways. (A) Conditioned media collected from serum starved primary human corneal epithelial cells analysed by CTGF ELISA demonstrated a temporal production of CTGF during differentiation. (B) CTGF ELISA was performed on conditioned media collected from serum starved HCE-T incubated with (0.1, 1, 10) or without (0) TGFβ1 (ng/ml) for 24 h. All concentration of TGFβ1 stimulate CTGF protein production, (C) HCE-T cells were incubated with (+) or without (−) TGFβ1 (10 ng/ml) in the presence or absence (control) of MEK1 (PD98059) and p38 (SB203580) inhibitors demonstrated reduced CTGF protein production. Representative images of differentiating HCEC (D) Day 4, (E) Day 7 and (F) Day 14. Error bars represent S.E.M. Asterisks indicate values that are significantly different from TGFβ1 (0) (p < 0.05).
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ACTIVE® p38 (Promega; 1:5000) primary antibodies in block for 1 h, then washed 3 ×30 min prior to addition of Donkey antiRabbit-HRP (Jackson Laboratories) secondary antibody in TBSTT+ 0.1% BSA for 1 h followed by 3×30 min washes. Immunocomplexes were detected by enhanced chemiluminescence (ECL) (Amersham, Amersham, UK) and images analyzed and processed using an image reader (LAS-1000 Pro, ver.2.1; Fuji, Bedford, UK).
CTGF enzyme linked immunosorbant assay CTGF was measured using a capture sandwich ELISA with biotinylated and non-biotinylated affinity purified goat polyclonal antibodies to human CTGF as described [26]. Briefly, a flat-bottom ELISA plate was coated with 50 μl of goat antihuman CTGF antibody at a concentration of 10 μg/ml in PBS/ 0.02% sodium azide for 18 h at 4 °C. Wells were washed three times and incubated with 300 μl of blocking buffer (PBS/0·02%
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sodium azide/1% BSA) for 1 h at room temperature. The wells were washed three times and 50 μl of recombinant human CTGF protein (from 0.156 to 100 ng/ml) or sample was added and incubated at room temperature for 2 h. After washing three times, 50 μl of biotinylated goat anti-human CTGF (2 μg/ml) was added and incubated at room temperature in the dark for 1 h, then washed three times, and 50 μl of alkaline phosphatase-conjugated streptavidin (1·5 μg/ml, Zymed, South San Francisco, CA, USA) was added and incubated at room temperature for 1 h. The wells were washed again three times and incubated with 100 μl of alkaline phosphatase substrate solution (1 mg/ml p-nitrophenyl phosphate, Sigma Chemicals, St. Louis, MO, USA) in sodium carbonate/bicarbonate buffer, pH = 9.6. Absorbance at 405 nm was measured using a microplate reader (Molecular Devices, Sunnyvale, CA). CTGF levels were expressed as ng/ml for three replicate samples for each condition.
Fig. 4 – CTGF stimulates scratch wound closure of HCEC and HCE-T through migration. (A, B) A wound was introduced into primary human corneal epithelial cells and HCE-T serum starved monolayers with a pipette tip. Scratch wound closure was determined in cultures incubated in SFM with (0.1, 1, 10) or without (0) CTGF (ng/ml). All three concentrations significantly stimulated re-epithelialisation of primary corneal epithelial cell (A) with (1 and 10 ng/ml) stimulating HCE-T cells (B) after 20 and 8 h respectively. (C) Cell proliferation was determined using WST-1 assay. Subconfluent serum starved HCE-T cells were incubated in SFM with (0.1, 1, 10) or without (0) CTGF (ng/ml) for 24 h. CTGF significantly inhibited HCE-T cell proliferation at concentrations of 0.1 and 1 ng/ml. (D) Confluent HCE-T cells cultured in cloning rings were growth arrested prior to ring removal and addition of treatment groups. Cells were incubated in SFM with (0.1, 1, 10) or without (0) CTGF (ng/ml) for 72 h, with all concentrations stimulating migration. Error bars represent S.E.M. Asterisks indicates values that are significantly different from CTGF (0) (p < 0.05).
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Statistical analysis One-way analysis of variance (ANOVA) was used to compare data. A difference between means of p < 0.05 was considered to be statistically significant.
Results TGFβ1 stimulates re-epithelialisation through the up-regulation of migration and inhibition of proliferation An in vitro scratch wound assay was used to determine the effect of TGFβ1 on re-epithelialisation. All concentrations of TGFβ1 tested (0.1 to 10 ng/ml) stimulated scratch wound closure of both primary HCEC (Fig. 1A, p < 0.05) and the HCE-T cell line (Fig. 1B, p < 0.05). HCE-T were able to respond more quickly than HCEC, however the response trends were comparable indicating the validity of the cell line in the analysis of corneal epithelial cell behaviour. The process of re-epithelialisation is driven by cell migration and proliferation. Using a colony dispersion assay, in which the cells are growth arrested to eliminate the proliferation component [28], TGFβ1 was found to stimulate HCE-T migration (Fig. 1D, p < 0.05) with an inverse dose response of cellular migratory activity to TGFβ1 concentration. Conversely, all concentrations of TGFβ1 tested, with the exception of 1 μg/ml, significantly inhibited HCE-T proliferation (Fig. 1C, p < 0.05).
TGFβ1 utilises the ERK and p38 MAPK signalling pathways during TGFβ1 stimulated re-epithelialisation To identify intracellular signalling pathways involved in TGFβ1 regulation of epithelial cells, western blotting was performed on cell lysates to detect phosphorylated proteins
of the MAPK pathways known to be involved in cell proliferation and migration. Following stimulation of serumstarved HCE-T with TGFβ1, phosphorylation of ERK 1/2 were detected at 1 min (Fig. 2A). By 3 min phosphorylation of ERK 2 was more pronounced than ERK 1. By 10 min maximal phosphorylation of ERK 1/2 had occurred. De-phosphorylation was evident by 30 min indicating a specific growth factor response. Unlike the ERKs, phosphorylation of p38 was delayed until 3 min post-stimulation with TGFβ1. Subsequent to this time point phosphorylation of p38 followed the same pattern as ERK 1/2 phosphorylation. The functional significance of ERK and p38 MAPK signalling pathway activation was demonstrated by the inhibition of TGFβ1 stimulated scratch wound closure in the presence of the MEK1 inhibitor PD98059 and the p38 inhibitor SB203580 respectively (Fig. 2B, p < 0.05).
Epithelial cells produce CTGF and TGFβ stimulates CTGF production via the ERK 1/2 and p38 MAPK signalling pathways We have previously found CTGF to be an autocrine downstream mediator of TGFβ1 activity in fibroblasts [17,18,15]. To determine if epithelial cells are capable of producing CTGF, conditioned media collected from serum starved primary HCECs exhibiting characteristic morphology of differentiating cells (Figs. 3C, D and F) were analysed by CTGF ELISA [26]. Over a 21-day period increasing levels of CTGF were released from the cells (Fig. 3A) confirming the ability of corneal epithelial cells to produce CTGF. Furthermore, TGFβ1 stimulated the production of CTGF in HCE-T (Fig. 3B) via the MEK1 and p38 (Fig. 3C) MAPK pathways. Inhibition of the MEK1 and p38 MAPK pathways reduced TGFβ1 stimulated CTGF production by 100% and approximately 50% respectively, potentially indicating a secondary cooperative rather than a primary role for p38 signalling in CTGF production.
Fig. 5 – CTGF antisense inhibits TGFβ mediated scratch wound closure in HCEC and HCE-T. (A, B) Antisense (AS) and Scrambled control (Scr) oligodeoxynucleotides targeting CTGF mRNA were used to block CTGF protein production. A wound was introduced into primary human corneal epithelial cells and HCE-T serum starved monolayers with a pipette tip. Scratch wound closure was determined in cultures incubated in SFM with (10 ng/ml) or without (Control) TGFβ1 in the presence (1, 10, 100 μM) and absence (−) of AS along with a control (scr 100μM). (A) 10 μM and 100 μM AS inhibited TGFβ1-stimulated primary human corneal epithelial cell scratch wound closure at 18 h. (B) 100 μM AS inhibited TGFβ1-stimulated HCE-T scratch wound closure at 8 h. (C) Demonstration of antisense oligonucleotides ability to knockdown CTGF expression in HCE-T cells. Error bars represents S.E.M. Asterisks indicate values that are significantly different from AS (−) control (p < 0.05).
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CTGF stimulates re-epithelialisation through the up-regulation of migration and inhibition of proliferation
CTGF activation of ERK MAPK signalling in epithelial cells is required for re-epithelialisation
Scratch wound closure by primary HCEC (Fig. 4A, p < 0.05) and HCE-T (Fig. 4B, p < 0.05) was induced by the addition of exogenous CTGF (in the absence of exogenous TGFβ1). This suggests that TGFβ1 stimulated re-epithelialisation occurs via the up-regulation and response to CTGF by epithelial cells rather than by a direct response to TGFβ1. This increased scratch wound closure was due to stimulated migration (Fig. 4D, p < 0.05) and inhibition of proliferation (Fig. 4C, p < 0.05).
To identify at least one potential signalling pathway involved in CTGF regulation of epithelial cells, immunoblotting was performed on cell lysates. Phosphorylation of ERK 2 was evident throughout the time course, whilst phosphorylation of ERK 1 did not occur until 10 min (Fig. 6A). Phosphorylation of ERK 1 after CTGF stimulation occurred up to 7 min later than that detected following TGFβ1 stimulation (Fig. 2A). By 30 min, phosphorylation of ERK 1/2 in response to CTGF was still evident (Fig. 6A) whereas dephosphorylation had begun to occur by this time in TGFβ1 treated cells (Fig. 2A). To confirm a regulatory role for the Ras/MEK/ERK pathway in epithelial cell response to CTGF, the scratch wound assay was repeated in the presence of CTGF with or without inhibitors of Ras, MEK and ERK phosphorylation. All three of the inhibitors prevented CTGF mediated scratch wound closure (Fig. 6B, p < 0.05).
CTGF is a downstream mediator of TGFβ1 stimulated re-epithelialisation It has been previously demonstrated that the effects of CTGF can be blocked using antisense oligonucleotides (AS) targeting mRNA [17]. The requirement of CTGF as a downstream mediator of TGFβ1 regulation of epithelial cells was confirmed by the dose dependent inhibition of TGFβ1 stimulated scratch wound closure by the addition of AS [17] to both primary HCEC (Fig. 5A, p < 0.05) and HCE-T (Fig. 5B, p < 0.05). CTGF protein production was inhibited in a dose dependant manner by the addition of antisense oligonucleotides in the presence of serum for 24 h in HCE-T cells. The corresponding scrambled control had no effect on CTGF production (Fig. 5C).
CTGF stimulates epithelial cell migration via the ERK MAPK signalling pathway CTGF stimulated epithelial cell migration was prevented by the addition of inhibitors of Ras, MEK and ERK in the colony dispersion assay (Fig. 6C, p < 0.05).
Fig. 6 – CTGF stimulates ERK 1/2 phosphorylation and inhibition of Ras/MEK/ERK prevents CTGF stimulated re-epithelialisation and migration. (A) Phosphorylated ERK 1/2 expression over time by immunoblotting of whole cell lysates stimulated with CTGF (10 ng/ml) using specific antibodies to phosphorylated ERK1/2 and GAPDH. (B) A wound was introduced into HCE-T serum starved monolayers with a pipette tip. Scratch wound closure was determined in cultures incubated in SFM with (+) or without (−) CTGF (10 ng/ml) in the presence and absence (Control) of Ras/MEK/ERK inhibitors. (C) Confluent HCE-T cells cultured in cloning rings were growth arrested prior to cell migration area determination in cultures incubated in SFM with (+) or without (−) CTGF (10 ng/ml) in the presence and absence (control) of Ras/MEK/ERK inhibitors. Error bars represents S.E.M. Asterisks indicate values that are significantly different from CTGF (+) control (p < 0.05).
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Fig. 7 – TGFβ and CTGF also regulate epithelial cells of non-corneal origin. (A,B) A wound was introduced into HCE-T serum starved monolayers with a pipette tip. Scratch wound closure was determined in cultures incubated in SFM with (0.1, 1, 10) or without (0) TGFβ1 (ng/ml) or CTGF (ng/ml). Both TGFβ and CTGF stimulated HaCaT scratch wound closure. (B), Cell proliferation was determined using WST-1 assay. Subconfluent serum starved HCE-T cells were incubated in SFM with (0.1, 1, 10) or without (0) TGFβ1 (ng/ml) or CTGF (ng/ml) for 24 h. Both TGFβ1 and CTGF inhibited HCE-T cell proliferation. (C) Confluent HCE-T cells cultured in cloning rings were growth arrested prior to ring removal and addition of treatment groups. Cells were incubated in SFM with (0.1, 1, 10) or without (0) TGFβ1 (ng/ml) or CTGF (ng/ml) for 72 h. Both TGFβ1 and CTGF stimulated migration, with TGFβ1 producing the stronger effect. Error bars represents S.E.M. Asterisks indicate values that are significantly different from TGFβ1 (0) or CTGF (0) (p < 0.05).
CTGF regulates the activity of more than one type of epithelial cell In order to rule out the possibility that responsiveness to TGFβ1 and CTGF may be peculiar to epithelial cells of corneal origin,
the scratch wound, proliferation and migration assays were repeated using the epidermal keratinocyte cell line HaCaT. TGFβ1 stimulated HaCaT scratch wound closure (Fig. 7A, p < 0.05), inhibited proliferation (Fig. 7B, p < 0.05) and stimulated
Fig. 8 – Proposed mechanism of TGFβ/CTGF signalling in epithelial cells during re-epithelialisation and migration. TGFβ1 binds to its receptor activating the Ras/MEK/ERK pathway which can be blocked with a MEK inhibitor (PD98059). This stimulates production of CTGF which can be inhibited using CTGF antisense oligonucleotides (AS). p38 is also activated which may play a role in enhancing CTGF production as the p38 inhibitor (SB203580) only blocks CTGF production by 50%. CTGF is released from the cell, upon binding to its receptor the Ras/MEK/ERK pathway is activated and migration occurs. This can be demonstrated by the blocking of migration using inhibitors against Ras (FTS), MEK (PD98059) and ERK (ERK 1/2 (I)) inhibitor.
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migration (Fig. 7C, p < 0.05). CTGF stimulated similar response trends in HaCaT cells (Figs. 7A–C, p < 0.05).
Proposed mechanism of action The diagram in Fig. 8 describes a proposed mechanism of action for CTGF as a downstream mediator of TGFβ in the migration element of re-epithelialisation and also indicates a role for the MAPK family in both TGFβ and CTGF signalling.
Discussion During homeostasis and particularly following injury, epithelial cells are exposed to a plethora of signals including soluble mediators and extracellular matrix molecules which direct their behaviour and fate. TGFβ is a very potent regulator of epithelial cells governing cell cycle progression, cell movement, differentiation and apoptosis [29,30]. Eukaryotic cells often utilise the mitogen-activated protein kinase (MAPK) signalling system to transduce extracellular signals into intracellular responses. One of the three major subgroups identified are the extracellular signalling regulated kinases 1 and 2 (also referred to as p44/p42 MAPKs) [31]. In our experiments TGFβ1 was found to inhibit epithelial cell proliferation and stimulate migration in agreement with previous findings [1,6]. The data presented in this study suggest wound closure was achieved via cell migration and that TGFβ1 stimulated re-epithelialisation was dependent upon both the ERK and p38 MAPK signalling pathways as inhibition of each pathway completely inhibited migration. This is in contrast to the response of rabbit corneal epithelial cells to HGF (hepatocyte growth factor) and KGF (keratinocyte growth factor) where inactivation of the ERK signalling pathway slightly increased migration via the activity of p38 in an organ culture wound model system. Inhibition of p38 almost completely blocked migration of the rabbit cells [32]. Little is known about the potential effects of CTGF on epithelial cell activity. TGFβ induction of CTGF expression has previously been thought to be restricted to post-natal and adult fibroblasts and not to induce CTGF in epithelial cells [13,20,33,34], however, it has been recently demonstrated that TGFβ1 induces CTGF expression in human proximal tubule epithelial cells [22]. This discrepancy may be explained due to the use of different cell lines and species. Our data clearly show the ability of primary corneal epithelial cells to produce CTGF. Furthermore, in response to TGFβ1 via activation of the ERK, MAPK pathway signalling, we have shown the absolute requirement of CTGF for TGFβ-stimulated re-epithelialisation via migration. Previous studies in corneal epithelial cells have indicated two-way cross talk between p38 and ERK 1/2 during hepatocyte growth factor induced migration [32]. Whilst inhibition of p38 activation only inhibited TGFβ1-induced CTGF production by 50% (indicating a supplementary rather than a primary role for p38 in regulation of CTGF production), the complete inhibition of TGFβ-stimulated epithelial cell migration in the presence of the p38 inhibitor confirmed a key role for the p38 pathway, warranting future investigation. In common with epithelial cells in this study, the Ras/MEK/ERK signalling pathway has previously been found to be required for
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TGFβ induction of CTGF promoter expression in fibroblasts while the JNK MAP kinase pathway acted as an inhibitor [34]. Furthermore, inhibition of RhoA genanylation by simvastatin in cultures of human lung fibroblasts blocked TGFβ-induction of CTGF expression, which demonstrates the importance of GTPase pathways in TGFβ signalling in other types of fibroblasts [35,36]. Upon exposure to 25 ng/ml TGFβ2, human foreskin fibroblasts expressed CTGF as determined by western blotting whereas the epithelial cell line MvLu did not. Further, a CTGF promoter fragment was found to respond to TGFβ2 only in the fibroblasts [34]. Inhibition of MEK 1/2 signalling blocked fibroblast CTGF induction by TGFβ2 and a dominant negative Ras inhibited TGFβ2 induced CTGF promoter activity. However, inhibition of the Ras/MEK/ERK cascade did not prevent TGFβ2mediated Smad activation of target genes [34]. It has been shown that minimum CTGF promoter is required for TGFβ dependant induction of CTGF expression in fibroblasts [9,21]. It will be interesting in the future to discover how the CTGF promoter is induced in epithelial cells and to discover further positive and negative regulators of CTGF in these cells. It has previously been hypothesised that changes in growth factor production during re-epithelialisation serve to direct stromal wound healing [37] and CTGF clearly plays a significant role in the regulation of fibroblast activity. However, the location of CTGF expression in the epithelium during wound healing in the rat cornea [26] and the production of CTGF in culture by HCEC in the absence of fibroblasts suggest that CTGF may also have an autocrine function in epithelial cells. A clear role for CTGF in the re-epithelialisation of stratified epithelium and specifically epithelial cell migration has now emerged from this study. Exogenous addition of CTGF to epithelial cells induced scratch wound closure and migration which could be inhibited by inactivation of Ras, MEK and ERK signalling. In MDCK cells, inhibition of ERK 1/2 activation inhibited sheet movement during scratch wound healing and disruption of the actin cytoskeleton suppressed ERK 1/2 activation. Therefore ERK 1/2 activation and cell movement form a positive feedback loop which facilitates sheet migration [38]. ERK and JNK activation has also been shown to be associated with TGFβ-induced breast cancer cell migration, independently of Smad-4 signalling [4]. CTGF stimulated mesangial cell migration was also found to be dependent upon p42/p44 (ERK 1/2) MAPK signalling possibly via ‘outside-in’ signalling via integrins [39]. Furthermore, CTGF was shown to induce actin disassembly and loss of focal adhesions to facilitate cell migration [40]. It is interesting to speculate that CTGF may influence corneal epithelial cells in a similar way. TGFβ has been shown to enhance tyrosine phosphorylation of α- and βcatenin, which results in the dissolution of the E-cadherin/ catenin complex from the actin cytoskeleton and reduced cell–cell adhesion in pancreatic tumour cells [41]. During development and tumourigenesis epithelial cells may undergo epithelial-to-mesenchymal transition (EMT) which essentially involves the conversion of the epithelial cell into a myofibroblast. In adult tissues such as the kidney and lung, epithelial cells undergo EMT leading to pathophysiological tissue changes resulting in fibrosis. This process has been shown to involve CTGF acting downstream of TGFβ [22,42–44]. However, the potential role of EMT during re-epithelialisation
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of stratified epithelium where the epithelial cells do not directly engage in fibrosis is unclear. Multilayered epithelium such as in skin and cornea is constantly regenerated by a population of stem cells which reside in discreet locations within the basal layer of the interfollicular epidermis and limbus, respectively [45–47]. Once these cells leave there niche they divide giving rise to daughter transient amplifying cells [48]. Interestingly during limbal explant culture, limbal epithelial cells can undergo intrastromal invasion (possibly via EMT) [49] whereas corneal epithelial cells seeded onto the same region of the limbus do not [50]. This suggests a fundamental change in the properties of limbal stem cell daughters once they have exited the niche. One study has suggested that epithelial cell migration during corneal wound re-epithelialisation does indeed shares features with EMT, specifically the up-regulation of Slug expression which was found to be associated with internalisation of E-cadherin and β-catenin from the cell membrane, enhanced expression of alpha smooth muscle actin, tropomyosin and matrix metalloproteinases at the epithelial wound margin [51]. The potential role of CTGF in the regulation of disassembly of E-cadherin/catenin complexes in cellular adherens junctions from the actin cytoskeleton to facilitate epithelial cell migration without stromal invasion, as occurs during re-epithelialisation, is unknown. It has previously been suggested that pharmacological inhibition of CTGF could become an alternative target for antifibrosis therapy via selective inhibition of elements of the TGFβ response in fibroblasts. Indeed, one such approach suggested in the context of skin fibrosis was targeting of the TGFβ induced Ras/MEK/ERK pathway to prevent expression of CTGF [34]. Whilst this approach may not interfere with keratinocyte function in the skin e.g. in the potential treatment of the connective tissue disease scleroderma, our data would suggest a detrimental effect upon re-epithelialisation of the cornea. Hence, testing of potential CTGF antiscarring strategies should include screening of the potential functional effects upon epithelial cells likely to come into contact with the reagent. In our hands the human skin keratinocyte HaCaT cell line was also capable of inducing wound healing type responses to both TGFβ and CTGF, indicating that CTGF regulation of epithelial cells from stratified epithelium is not restricted to cells of corneal origin. In summary, we propose a novel role for CTGF during tissue repair outside of its established involvement in fibrosis, which is in the regulation TGFβ-stimulated re-epithelialisation of stratified epithelium via epithelial cell migration. Furthermore, we present evidence to suggest that, TGFβ is unable to induce epithelial cell migration in the absence of CTGF. Upon binding of TGFβ to its receptor on epithelial cells, the Ras/MEK/ERK MAPK signalling pathway is activated followed by CTGF gene expression and protein production. We postulate that activation of p38 plays a cooperative role, possibly by enhancing CTGF protein translation since blocking of this signalling pathway only reduces, but does not eliminate, CTGF protein production. CTGF is then released from the cell whereupon it binds to its receptor, inducing epithelial cell migration in an autocrine manner, which occurs at least in part via the Ras/MEK/ERK signalling pathway. Our data demonstrates that CTGF can stimulate cells of stratified epithelia to engage in the non-
fibrotic process of re-epithelialisation and further highlights the complexity of TGFβ regulation of epithelial cell function.
Acknowledgments We would like to thank Priscilla Philips for the ELISA work. This work was supported by The Eranda Foundation (GAS), The Special Trustees of Moorfields Eye Hospital (JTD) and NEI grant EY05587 (GSS).
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a v a i l a b l e a t w w w. s c i e n c e d i r e c t . c o m
w w w. e l s e v i e r. c o m / l o c a t e / y e x c r
Research Article
TGFβ stimulated re-epithelialisation is regulated by CTGF and Ras/MEK/ERK signalling Genevieve A. Secker a , Alex J. Shortt a , Edith Sampson b , Quenten P. Schwarz c , Gregory S. Schultz b , Julie T. Daniels a,⁎ a
Department of Pathology, Cells for Sight Transplantation and Research Programme, UCL Institute of Ophthalmology, 11-43 Bath Street, London, EC1V 9EL, UK b Department of ObGyn, The University of Florida, Gainsville, FL 32610, USA c Department of Cellular Therapy, UCL Institute of Ophthalmology, London, EC1V 9EL, UK
ARTICLE INFORMATION
ABS T R AC T
Article Chronology:
The complex mechanisms by which transforming growth factor β (TGFβ) regulate re-
Received 18 May 2007
epithelialisation following injury of stratified epithelia are not fully understood. TGFβ
Revised version received
signals via binding to distinct receptors activating downstream effectors, including Smads
31 August 2007
which initiate transcription of target genes. However, studies have shown that TGFβ can
Accepted 4 September 2007
also signal independently of Smads through MAPK pathways, demonstrating the diversity
Available online 7 September 2007
of TGFβ signalling. Connective tissue growth factor (CTGF) is strongly induced by and acts downstream of TGFβ causing pathophysiology in tissues by inducing matrix deposition,
Keywords:
conversion of fibroblasts into contractile myofibroblasts (e.g. dermis and corneal stroma)
TGFβ
and stimulation of epithelial-to-mesenchymal transition (e.g. kidney and lung) all of which
Re-epithelialisation
are known to cause fibrosis. However, a role for CTGF in epithelial cell function which does
Migration
not involve direct contribution to fibrosis has not been demonstrated. We show for the first
CTGF
time that synthesis of CTGF in cultures of human corneal epithelial cells is induced by TGFβ
Ras/MEK/ERK
through the Ras/MEK/ERK MAPK signalling pathway and that this is required for reepithelialisation to occur through cell migration. These data reveal a novel function for CTGF in the regulation of epithelial tissue repair beyond its established role in fibrosis, and further highlight the complexity of TGFβ regulation of epithelial cell function. © 2007 Elsevier Inc. All rights reserved.
Introduction The pathways involved in TGFβ regulation of re-epithelialisation of stratified epithelia, specifically inhibition of proliferation and promotion of migration of epithelial cells to facilitate wound closure are unclear [1]. Members of the TGFβ family can initiate cellular responses by binding to distinct receptors with
intrinsic serine/threonine kinase activity and activation of specific downstream intracellular effectors termed Smad proteins. The Smads relay the signal from the cell membrane to the nucleus, where they affect transcription of target genes [2]. For example Smad 4 is necessary for TGFβ-induced cell cycle arrest and migration in immortalised keratinocytes (HaCaT) and human pancreatic tumour cells (Colo-357) [3].
⁎ Corresponding author. Fax: +44 2076086887. E-mail address: [email protected] (J.T. Daniels). Abbreviations: TGFβ, transforming growth factor β; SFM, serum free medium; DMEM, Dulbecco's modified Eagle's medium; GAPDH, glyseraldehyde-3-phosphate dehydrogenase; BSA, bovine serum albumin; PBS, phosphate buffered saline; KSFM, keratinocyte serum free medium; HCEC, human corneal epithelial cells; S.E.M., standard error 0014-4827/$ – see front matter © 2007 Elsevier Inc. All rights reserved. doi:10.1016/j.yexcr.2007.09.001
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However, there is also evidence that TGFβ-stimulated epithelial cell migration can occur independently of Smad 4 with signalling via the ERK and JNK MAPK pathways [4,5]. During re-epithelialisation of mouse corneas TGFβ was found to enhance re-epithelialisation by suppressing cell division and enhancing cell migration via the p38 MAPK pathway, independent of Smad signalling [6]. These findings illustrate the complexities of TGFβ signalling and regulation [7]. During early development and tissue repair, TGFβ gene expression in fibroblasts is co-ordinately regulated with that of connective tissue growth factor (CTGF) [8,9]. CTGF is a member of the CCN (CTGF, Cyr61 and Nov) protein family [10] and was first identified in conditioned medium collected from cultured human vascular endothelial cells [11]. CTGF is a cysteine-rich, heparin binding protein,[11] whose gene expression is strongly induced by TGFβ in fibroblasts [8,12]. A 1-h exposure to TGFβ is sufficient to induce CTGF gene transcription for up to 36 h in fibroblasts [13]. A novel TGFβ response element controls transcription in both human and murine CTGF promoters
[13]. These observations suggest that CTGF is a downstream mediator of TGFβ activity in fibroblasts [13]. Indeed, CTGF is known to mediate key cellular events in response to TGFβ including migration [13], proliferation [14,15], matrix production [16] and contraction and differentiation of fibroblast into myofibroblasts [14,17,18], which are all elements of a fibrotic response. Previous reports have indicated that CTGF induction by TGFβ is specific for connective tissue cells and does not occur in epithelial cells [19,9,13,20,21,22]. Nevertheless, a growing body of evidence supports the notion that a variety of epithelial cell types can produce CTGF during development and in adult tissues [23–27]. However, the functional significance of CTGF production during homeostasis and following epithelial injury of stratified epithelia such as those of the cornea and skin remain unclear. In the present study, we report that CTGF is produced by differentiating human corneal epithelial cells (HCEC) and that CTGF has an additional role for mediating the activity of TGFβ
Fig. 1 – TGFβ stimulates scratch wound closure in primary HCEC and HCE-T through migration. (A,B) A wound was introduced into primary human corneal epithelial cells and HCE-T serum starved monolayers with a pipette tip. Scratch wound closure was determined in cultures incubated in SFM with (0.1, 1, 10) or without (0) TGFβ1 (ng/ml). All three concentrations significantly stimulated primary corneal epithelial cell (A), and HCE-T cell (B), re-epithelialisation after 20 and 8 h respectively. (C) Cell proliferation was determined using WST-1 assay. Subconfluent serum starved HCE-T cells were incubated in SFM with (0.1, 1, 10) or without (0) TGFβ1 (ng/ml) for 24 h. TGFβ1 significantly inhibited HCE-T cell proliferation at concentrations of 0.1 and 10 ng/ml. (D) Confluent HCE-T cells cultured in cloning rings were growth arrested prior to ring removal and addition of treatment groups. Cells were incubated in SFM with (0.1, 1, 10) or without (0) TGFβ1 (ng/ml) for 72 h, with all concentrations stimulating migration. Error bars represents S.E.M. Asterisks indicate values that are significantly different from TGFβ1 (0) (p < 0.05).
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during tissue repair beyond the regulation of fibrosis; namely regulation of re-epithelialisation. Specifically, CTGF is required for TGFβ-stimulated epithelial cell migration. Furthermore we have demonstrated a role for ERK and p38 MAPK signalling in both TGFβ-stimulated CTGF production and migration in epithelial cells derived from stratified epithelium.
Materials and methods
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described previously [19], were used at concentrations 0– 10 ng/ml in serum free media (SFM) consisting of basal media supplemented with 0.1% bovine serum albumin (BSA). ERK 1/2, p38 and Ras pathways were inhibited using PD98059 (100 μM; Cell Signalling Technology, ERK 1/2 inhibitor, SB203580 (10 μM; Promega, Madison, WI) and Farnesylthiosalicylic acid (5 μM; FTS; Biomol International, Exeter, UK). Antisense oligodeoxynucleotides (AS), synthesised as described previously [17], were used at concentration of 1, 10 and 100 μM and the scrambled control was used at 100 μM.
Cells, cell culture and reagents Scratch wound model Human corneas supplied by Moorfields Eye Hospital National Health Service (NHS) Foundation Trust Eye Bank were used in the study, in accordance with the Declaration of Helsinki for the use of human tissue. Human corneal epithelial cells (HCEC) were isolated and cultured in keratinocyte serum free medium (KSFM) with supplements (Invitrogen, Scotland, UK). Corneal segments were dissected into quadrants and placed in 1x Dispase II (Roche Diagnostics Ltd, East Sussex, UK) overnight at 4 °C before epithelial cells were gently scraped off using forceps. Experiments were performed with cells between passages 1 and 3. HCE-T cells (Riken, Tsukuba Institute) were maintained in corneal epithelial cell medium (CECM) consisting of DMEM:F12 (1:1) (Invitrogen) supplemented with 5% FCS (Invitrogen), 10 ng/ml Epidermal Growth Factor (Invitrogen), 0.1 ng/ml Cholera Toxin (Sigma-Aldrich Chemical Co. Ltd., Poole, UK) and 5 ug/ml Insulin (Sigma-Aldrich). HaCaT cells (a kind gift from F. Watt, Cancer Research UK, London, UK) were cultured in DMEM (Invitrogen) supplemented with 10% FCS. All cells were cultured in the presence of 5% CO2 in air at 37 °C and were grown in the presence of antibiotics. Growth factors, TGFβ1 (R&D Systems Europe Ltd., Oxon, UK) and CTGF produced by baculovirus expression in insect cells, as
HCE-T or HCEC cultured to confluence in marked 96-well plates were serum starved in SFM for 24 h. A 200-μl plastic pipette tip was used to scratch the monolayers, the wounded cells were digitally photographed under an inverted phase contrast microscope prior to addition of treatment groups and 8 (HCE-T) or 18 h (HCEC) post wounding. The distance between the scratch wound edges were measured using Image Tool software (UTHSCSA, San Antonio, TX), with average wound areas calculated to determine percentage wound closure (re-epithelialisation) between time points.
Cell proliferation assay HCE-T proliferation was monitored using the water soluble tetrazolium-1 (WST-1) reagent (Roche Diagnostics Ltd) according to the manufacturer's instructions. Briefly, HCE-T cells were seeded into 96-well tissue culture plates, at a density of 5 × 103 cells per well in CECM and cultured for 18 h. CECM was removed, and the cells were washed three times prior to the addition of SFM for a 24-h serum starvation period. Treatment groups and serum free baseline control
Fig. 2 – TGFβ utilises ERK 1/2 and p38 pathway signalling for re-epithelialisation. (A) Phosphorylated ERK 1/2 and p38 expression over time was determined by immunoblotting whole cell lysates stimulated with TGFβ1 (10 ng/ml) using specific antibodies to phosphorylated ERK 1/2, p38 and GAPDH. ERK 1/2 phosphorylation occurred at 1 min and was most prominent at 10 min with p38 phosphorylation occurring at 3 min and then becoming most prominent at 10 min. GAPDH was used as a loading control. (B) A wound was introduced into HCE-T serum starved monolayers with a pipette tip. Scratch wound closure was determined in cultures incubated with (+) or without (−) TGFβ1 (10 ng/ml) in presence and absence (control) of MEK1 (PD98059) and p38 (SB203580) inhibitors. Inhibition of both pathways prevented TGFβ1 stimulated wound closure. Error bars represents S.E.M. Asterisks indicate values that are significantly different from TGFβ1 (+) control (p < 0.05).
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medium (100 μl) were added to the cells (eight replicates) and incubated for 22 h. Ten microlitres of WST-1 reagent was added to each well and the plate was incubated at 37 °C for 2 h. The absorbance, related to the number of viable cells converting the reagent to coloured formazan crystals, was read at 450 nm.
Collection of conditioned medium
Colony dispersion
Immunoblot analysis
Cell migration was assessed using a colony dispersion assay as described by Pilcher et al. [28] Briefly, HCE-T or HaCaT cells were seeded and cultured to confluence within cloning rings. The cells were cultured for a further 24 h in the presence of 1 mM hydroxyurea (Sigma-Aldrich) to induce growth arrest. After removal of the rings, the cells were thoroughly washed with PBS prior to the addition of treatment groups. Cells were allowed to migrate for 72 h, then washed with PBS three times, fixed with 100% chilled methanol for 10 min and stained with Harris hematoxylin (Shandon Life Sciences International, Europe, Ltd., UK) for 30 min prior to washing with water. Dispersion areas were digitally photographed and measured in pixels with Image Tool analysis software.
Sub confluent HCE-T were serum starved in SFM for 48 h prior to addition of treatment groups. Cell monolayers were collected at time points 0 to 1 h using radioimmune precipitation assay (RIPA) buffer (1% Nonidet P-40/0.5% deoxycholic acid/ 0.1% SDS/158 mM NaCl/50 mM Tris/pH 7.2) and centrifugated at 13,000 rpm for 5 min prior to supernatant collection. Proteins were resolved by SDS/10% PAGE (NuPAGE; Invitrogen, Groningen, The Netherlands) for 60 min at 150 V in Tris-acetate running buffer (Invitrogen) and transferred to nitrocellulose for 2 h at 30 V with a NOVEX transfer apparatus and Transfer Buffer (Invitrogen). Nitrocellulose was blocked with TBSTT (TBS/0.1% Tween-20/0.2% Triton-X) with 1% BSA for 1 hr, probed with antiACTIVE® MAPK (Promega, Southampton, UK; 1:2000) and anti-
Cells were seeded at 60% confluence and allowed to adhere overnight prior to serum starvation for 24 h. Conditioned media was collected following addition of treatment groups for 24 h or directly following serum starvation period.
Fig. 3 – CTGF is produced during HCEC differentiation and TGFβ stimulates CTGF production in epithelial cells via the ERK 1/2 and p38 pathways. (A) Conditioned media collected from serum starved primary human corneal epithelial cells analysed by CTGF ELISA demonstrated a temporal production of CTGF during differentiation. (B) CTGF ELISA was performed on conditioned media collected from serum starved HCE-T incubated with (0.1, 1, 10) or without (0) TGFβ1 (ng/ml) for 24 h. All concentration of TGFβ1 stimulate CTGF protein production, (C) HCE-T cells were incubated with (+) or without (−) TGFβ1 (10 ng/ml) in the presence or absence (control) of MEK1 (PD98059) and p38 (SB203580) inhibitors demonstrated reduced CTGF protein production. Representative images of differentiating HCEC (D) Day 4, (E) Day 7 and (F) Day 14. Error bars represent S.E.M. Asterisks indicate values that are significantly different from TGFβ1 (0) (p < 0.05).
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ACTIVE® p38 (Promega; 1:5000) primary antibodies in block for 1 h, then washed 3 ×30 min prior to addition of Donkey antiRabbit-HRP (Jackson Laboratories) secondary antibody in TBSTT+ 0.1% BSA for 1 h followed by 3×30 min washes. Immunocomplexes were detected by enhanced chemiluminescence (ECL) (Amersham, Amersham, UK) and images analyzed and processed using an image reader (LAS-1000 Pro, ver.2.1; Fuji, Bedford, UK).
CTGF enzyme linked immunosorbant assay CTGF was measured using a capture sandwich ELISA with biotinylated and non-biotinylated affinity purified goat polyclonal antibodies to human CTGF as described [26]. Briefly, a flat-bottom ELISA plate was coated with 50 μl of goat antihuman CTGF antibody at a concentration of 10 μg/ml in PBS/ 0.02% sodium azide for 18 h at 4 °C. Wells were washed three times and incubated with 300 μl of blocking buffer (PBS/0·02%
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sodium azide/1% BSA) for 1 h at room temperature. The wells were washed three times and 50 μl of recombinant human CTGF protein (from 0.156 to 100 ng/ml) or sample was added and incubated at room temperature for 2 h. After washing three times, 50 μl of biotinylated goat anti-human CTGF (2 μg/ml) was added and incubated at room temperature in the dark for 1 h, then washed three times, and 50 μl of alkaline phosphatase-conjugated streptavidin (1·5 μg/ml, Zymed, South San Francisco, CA, USA) was added and incubated at room temperature for 1 h. The wells were washed again three times and incubated with 100 μl of alkaline phosphatase substrate solution (1 mg/ml p-nitrophenyl phosphate, Sigma Chemicals, St. Louis, MO, USA) in sodium carbonate/bicarbonate buffer, pH = 9.6. Absorbance at 405 nm was measured using a microplate reader (Molecular Devices, Sunnyvale, CA). CTGF levels were expressed as ng/ml for three replicate samples for each condition.
Fig. 4 – CTGF stimulates scratch wound closure of HCEC and HCE-T through migration. (A, B) A wound was introduced into primary human corneal epithelial cells and HCE-T serum starved monolayers with a pipette tip. Scratch wound closure was determined in cultures incubated in SFM with (0.1, 1, 10) or without (0) CTGF (ng/ml). All three concentrations significantly stimulated re-epithelialisation of primary corneal epithelial cell (A) with (1 and 10 ng/ml) stimulating HCE-T cells (B) after 20 and 8 h respectively. (C) Cell proliferation was determined using WST-1 assay. Subconfluent serum starved HCE-T cells were incubated in SFM with (0.1, 1, 10) or without (0) CTGF (ng/ml) for 24 h. CTGF significantly inhibited HCE-T cell proliferation at concentrations of 0.1 and 1 ng/ml. (D) Confluent HCE-T cells cultured in cloning rings were growth arrested prior to ring removal and addition of treatment groups. Cells were incubated in SFM with (0.1, 1, 10) or without (0) CTGF (ng/ml) for 72 h, with all concentrations stimulating migration. Error bars represent S.E.M. Asterisks indicates values that are significantly different from CTGF (0) (p < 0.05).
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Statistical analysis One-way analysis of variance (ANOVA) was used to compare data. A difference between means of p < 0.05 was considered to be statistically significant.
Results TGFβ1 stimulates re-epithelialisation through the up-regulation of migration and inhibition of proliferation An in vitro scratch wound assay was used to determine the effect of TGFβ1 on re-epithelialisation. All concentrations of TGFβ1 tested (0.1 to 10 ng/ml) stimulated scratch wound closure of both primary HCEC (Fig. 1A, p < 0.05) and the HCE-T cell line (Fig. 1B, p < 0.05). HCE-T were able to respond more quickly than HCEC, however the response trends were comparable indicating the validity of the cell line in the analysis of corneal epithelial cell behaviour. The process of re-epithelialisation is driven by cell migration and proliferation. Using a colony dispersion assay, in which the cells are growth arrested to eliminate the proliferation component [28], TGFβ1 was found to stimulate HCE-T migration (Fig. 1D, p < 0.05) with an inverse dose response of cellular migratory activity to TGFβ1 concentration. Conversely, all concentrations of TGFβ1 tested, with the exception of 1 μg/ml, significantly inhibited HCE-T proliferation (Fig. 1C, p < 0.05).
TGFβ1 utilises the ERK and p38 MAPK signalling pathways during TGFβ1 stimulated re-epithelialisation To identify intracellular signalling pathways involved in TGFβ1 regulation of epithelial cells, western blotting was performed on cell lysates to detect phosphorylated proteins
of the MAPK pathways known to be involved in cell proliferation and migration. Following stimulation of serumstarved HCE-T with TGFβ1, phosphorylation of ERK 1/2 were detected at 1 min (Fig. 2A). By 3 min phosphorylation of ERK 2 was more pronounced than ERK 1. By 10 min maximal phosphorylation of ERK 1/2 had occurred. De-phosphorylation was evident by 30 min indicating a specific growth factor response. Unlike the ERKs, phosphorylation of p38 was delayed until 3 min post-stimulation with TGFβ1. Subsequent to this time point phosphorylation of p38 followed the same pattern as ERK 1/2 phosphorylation. The functional significance of ERK and p38 MAPK signalling pathway activation was demonstrated by the inhibition of TGFβ1 stimulated scratch wound closure in the presence of the MEK1 inhibitor PD98059 and the p38 inhibitor SB203580 respectively (Fig. 2B, p < 0.05).
Epithelial cells produce CTGF and TGFβ stimulates CTGF production via the ERK 1/2 and p38 MAPK signalling pathways We have previously found CTGF to be an autocrine downstream mediator of TGFβ1 activity in fibroblasts [17,18,15]. To determine if epithelial cells are capable of producing CTGF, conditioned media collected from serum starved primary HCECs exhibiting characteristic morphology of differentiating cells (Figs. 3C, D and F) were analysed by CTGF ELISA [26]. Over a 21-day period increasing levels of CTGF were released from the cells (Fig. 3A) confirming the ability of corneal epithelial cells to produce CTGF. Furthermore, TGFβ1 stimulated the production of CTGF in HCE-T (Fig. 3B) via the MEK1 and p38 (Fig. 3C) MAPK pathways. Inhibition of the MEK1 and p38 MAPK pathways reduced TGFβ1 stimulated CTGF production by 100% and approximately 50% respectively, potentially indicating a secondary cooperative rather than a primary role for p38 signalling in CTGF production.
Fig. 5 – CTGF antisense inhibits TGFβ mediated scratch wound closure in HCEC and HCE-T. (A, B) Antisense (AS) and Scrambled control (Scr) oligodeoxynucleotides targeting CTGF mRNA were used to block CTGF protein production. A wound was introduced into primary human corneal epithelial cells and HCE-T serum starved monolayers with a pipette tip. Scratch wound closure was determined in cultures incubated in SFM with (10 ng/ml) or without (Control) TGFβ1 in the presence (1, 10, 100 μM) and absence (−) of AS along with a control (scr 100μM). (A) 10 μM and 100 μM AS inhibited TGFβ1-stimulated primary human corneal epithelial cell scratch wound closure at 18 h. (B) 100 μM AS inhibited TGFβ1-stimulated HCE-T scratch wound closure at 8 h. (C) Demonstration of antisense oligonucleotides ability to knockdown CTGF expression in HCE-T cells. Error bars represents S.E.M. Asterisks indicate values that are significantly different from AS (−) control (p < 0.05).
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CTGF stimulates re-epithelialisation through the up-regulation of migration and inhibition of proliferation
CTGF activation of ERK MAPK signalling in epithelial cells is required for re-epithelialisation
Scratch wound closure by primary HCEC (Fig. 4A, p < 0.05) and HCE-T (Fig. 4B, p < 0.05) was induced by the addition of exogenous CTGF (in the absence of exogenous TGFβ1). This suggests that TGFβ1 stimulated re-epithelialisation occurs via the up-regulation and response to CTGF by epithelial cells rather than by a direct response to TGFβ1. This increased scratch wound closure was due to stimulated migration (Fig. 4D, p < 0.05) and inhibition of proliferation (Fig. 4C, p < 0.05).
To identify at least one potential signalling pathway involved in CTGF regulation of epithelial cells, immunoblotting was performed on cell lysates. Phosphorylation of ERK 2 was evident throughout the time course, whilst phosphorylation of ERK 1 did not occur until 10 min (Fig. 6A). Phosphorylation of ERK 1 after CTGF stimulation occurred up to 7 min later than that detected following TGFβ1 stimulation (Fig. 2A). By 30 min, phosphorylation of ERK 1/2 in response to CTGF was still evident (Fig. 6A) whereas dephosphorylation had begun to occur by this time in TGFβ1 treated cells (Fig. 2A). To confirm a regulatory role for the Ras/MEK/ERK pathway in epithelial cell response to CTGF, the scratch wound assay was repeated in the presence of CTGF with or without inhibitors of Ras, MEK and ERK phosphorylation. All three of the inhibitors prevented CTGF mediated scratch wound closure (Fig. 6B, p < 0.05).
CTGF is a downstream mediator of TGFβ1 stimulated re-epithelialisation It has been previously demonstrated that the effects of CTGF can be blocked using antisense oligonucleotides (AS) targeting mRNA [17]. The requirement of CTGF as a downstream mediator of TGFβ1 regulation of epithelial cells was confirmed by the dose dependent inhibition of TGFβ1 stimulated scratch wound closure by the addition of AS [17] to both primary HCEC (Fig. 5A, p < 0.05) and HCE-T (Fig. 5B, p < 0.05). CTGF protein production was inhibited in a dose dependant manner by the addition of antisense oligonucleotides in the presence of serum for 24 h in HCE-T cells. The corresponding scrambled control had no effect on CTGF production (Fig. 5C).
CTGF stimulates epithelial cell migration via the ERK MAPK signalling pathway CTGF stimulated epithelial cell migration was prevented by the addition of inhibitors of Ras, MEK and ERK in the colony dispersion assay (Fig. 6C, p < 0.05).
Fig. 6 – CTGF stimulates ERK 1/2 phosphorylation and inhibition of Ras/MEK/ERK prevents CTGF stimulated re-epithelialisation and migration. (A) Phosphorylated ERK 1/2 expression over time by immunoblotting of whole cell lysates stimulated with CTGF (10 ng/ml) using specific antibodies to phosphorylated ERK1/2 and GAPDH. (B) A wound was introduced into HCE-T serum starved monolayers with a pipette tip. Scratch wound closure was determined in cultures incubated in SFM with (+) or without (−) CTGF (10 ng/ml) in the presence and absence (Control) of Ras/MEK/ERK inhibitors. (C) Confluent HCE-T cells cultured in cloning rings were growth arrested prior to cell migration area determination in cultures incubated in SFM with (+) or without (−) CTGF (10 ng/ml) in the presence and absence (control) of Ras/MEK/ERK inhibitors. Error bars represents S.E.M. Asterisks indicate values that are significantly different from CTGF (+) control (p < 0.05).
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Fig. 7 – TGFβ and CTGF also regulate epithelial cells of non-corneal origin. (A,B) A wound was introduced into HCE-T serum starved monolayers with a pipette tip. Scratch wound closure was determined in cultures incubated in SFM with (0.1, 1, 10) or without (0) TGFβ1 (ng/ml) or CTGF (ng/ml). Both TGFβ and CTGF stimulated HaCaT scratch wound closure. (B), Cell proliferation was determined using WST-1 assay. Subconfluent serum starved HCE-T cells were incubated in SFM with (0.1, 1, 10) or without (0) TGFβ1 (ng/ml) or CTGF (ng/ml) for 24 h. Both TGFβ1 and CTGF inhibited HCE-T cell proliferation. (C) Confluent HCE-T cells cultured in cloning rings were growth arrested prior to ring removal and addition of treatment groups. Cells were incubated in SFM with (0.1, 1, 10) or without (0) TGFβ1 (ng/ml) or CTGF (ng/ml) for 72 h. Both TGFβ1 and CTGF stimulated migration, with TGFβ1 producing the stronger effect. Error bars represents S.E.M. Asterisks indicate values that are significantly different from TGFβ1 (0) or CTGF (0) (p < 0.05).
CTGF regulates the activity of more than one type of epithelial cell In order to rule out the possibility that responsiveness to TGFβ1 and CTGF may be peculiar to epithelial cells of corneal origin,
the scratch wound, proliferation and migration assays were repeated using the epidermal keratinocyte cell line HaCaT. TGFβ1 stimulated HaCaT scratch wound closure (Fig. 7A, p < 0.05), inhibited proliferation (Fig. 7B, p < 0.05) and stimulated
Fig. 8 – Proposed mechanism of TGFβ/CTGF signalling in epithelial cells during re-epithelialisation and migration. TGFβ1 binds to its receptor activating the Ras/MEK/ERK pathway which can be blocked with a MEK inhibitor (PD98059). This stimulates production of CTGF which can be inhibited using CTGF antisense oligonucleotides (AS). p38 is also activated which may play a role in enhancing CTGF production as the p38 inhibitor (SB203580) only blocks CTGF production by 50%. CTGF is released from the cell, upon binding to its receptor the Ras/MEK/ERK pathway is activated and migration occurs. This can be demonstrated by the blocking of migration using inhibitors against Ras (FTS), MEK (PD98059) and ERK (ERK 1/2 (I)) inhibitor.
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migration (Fig. 7C, p < 0.05). CTGF stimulated similar response trends in HaCaT cells (Figs. 7A–C, p < 0.05).
Proposed mechanism of action The diagram in Fig. 8 describes a proposed mechanism of action for CTGF as a downstream mediator of TGFβ in the migration element of re-epithelialisation and also indicates a role for the MAPK family in both TGFβ and CTGF signalling.
Discussion During homeostasis and particularly following injury, epithelial cells are exposed to a plethora of signals including soluble mediators and extracellular matrix molecules which direct their behaviour and fate. TGFβ is a very potent regulator of epithelial cells governing cell cycle progression, cell movement, differentiation and apoptosis [29,30]. Eukaryotic cells often utilise the mitogen-activated protein kinase (MAPK) signalling system to transduce extracellular signals into intracellular responses. One of the three major subgroups identified are the extracellular signalling regulated kinases 1 and 2 (also referred to as p44/p42 MAPKs) [31]. In our experiments TGFβ1 was found to inhibit epithelial cell proliferation and stimulate migration in agreement with previous findings [1,6]. The data presented in this study suggest wound closure was achieved via cell migration and that TGFβ1 stimulated re-epithelialisation was dependent upon both the ERK and p38 MAPK signalling pathways as inhibition of each pathway completely inhibited migration. This is in contrast to the response of rabbit corneal epithelial cells to HGF (hepatocyte growth factor) and KGF (keratinocyte growth factor) where inactivation of the ERK signalling pathway slightly increased migration via the activity of p38 in an organ culture wound model system. Inhibition of p38 almost completely blocked migration of the rabbit cells [32]. Little is known about the potential effects of CTGF on epithelial cell activity. TGFβ induction of CTGF expression has previously been thought to be restricted to post-natal and adult fibroblasts and not to induce CTGF in epithelial cells [13,20,33,34], however, it has been recently demonstrated that TGFβ1 induces CTGF expression in human proximal tubule epithelial cells [22]. This discrepancy may be explained due to the use of different cell lines and species. Our data clearly show the ability of primary corneal epithelial cells to produce CTGF. Furthermore, in response to TGFβ1 via activation of the ERK, MAPK pathway signalling, we have shown the absolute requirement of CTGF for TGFβ-stimulated re-epithelialisation via migration. Previous studies in corneal epithelial cells have indicated two-way cross talk between p38 and ERK 1/2 during hepatocyte growth factor induced migration [32]. Whilst inhibition of p38 activation only inhibited TGFβ1-induced CTGF production by 50% (indicating a supplementary rather than a primary role for p38 in regulation of CTGF production), the complete inhibition of TGFβ-stimulated epithelial cell migration in the presence of the p38 inhibitor confirmed a key role for the p38 pathway, warranting future investigation. In common with epithelial cells in this study, the Ras/MEK/ERK signalling pathway has previously been found to be required for
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TGFβ induction of CTGF promoter expression in fibroblasts while the JNK MAP kinase pathway acted as an inhibitor [34]. Furthermore, inhibition of RhoA genanylation by simvastatin in cultures of human lung fibroblasts blocked TGFβ-induction of CTGF expression, which demonstrates the importance of GTPase pathways in TGFβ signalling in other types of fibroblasts [35,36]. Upon exposure to 25 ng/ml TGFβ2, human foreskin fibroblasts expressed CTGF as determined by western blotting whereas the epithelial cell line MvLu did not. Further, a CTGF promoter fragment was found to respond to TGFβ2 only in the fibroblasts [34]. Inhibition of MEK 1/2 signalling blocked fibroblast CTGF induction by TGFβ2 and a dominant negative Ras inhibited TGFβ2 induced CTGF promoter activity. However, inhibition of the Ras/MEK/ERK cascade did not prevent TGFβ2mediated Smad activation of target genes [34]. It has been shown that minimum CTGF promoter is required for TGFβ dependant induction of CTGF expression in fibroblasts [9,21]. It will be interesting in the future to discover how the CTGF promoter is induced in epithelial cells and to discover further positive and negative regulators of CTGF in these cells. It has previously been hypothesised that changes in growth factor production during re-epithelialisation serve to direct stromal wound healing [37] and CTGF clearly plays a significant role in the regulation of fibroblast activity. However, the location of CTGF expression in the epithelium during wound healing in the rat cornea [26] and the production of CTGF in culture by HCEC in the absence of fibroblasts suggest that CTGF may also have an autocrine function in epithelial cells. A clear role for CTGF in the re-epithelialisation of stratified epithelium and specifically epithelial cell migration has now emerged from this study. Exogenous addition of CTGF to epithelial cells induced scratch wound closure and migration which could be inhibited by inactivation of Ras, MEK and ERK signalling. In MDCK cells, inhibition of ERK 1/2 activation inhibited sheet movement during scratch wound healing and disruption of the actin cytoskeleton suppressed ERK 1/2 activation. Therefore ERK 1/2 activation and cell movement form a positive feedback loop which facilitates sheet migration [38]. ERK and JNK activation has also been shown to be associated with TGFβ-induced breast cancer cell migration, independently of Smad-4 signalling [4]. CTGF stimulated mesangial cell migration was also found to be dependent upon p42/p44 (ERK 1/2) MAPK signalling possibly via ‘outside-in’ signalling via integrins [39]. Furthermore, CTGF was shown to induce actin disassembly and loss of focal adhesions to facilitate cell migration [40]. It is interesting to speculate that CTGF may influence corneal epithelial cells in a similar way. TGFβ has been shown to enhance tyrosine phosphorylation of α- and βcatenin, which results in the dissolution of the E-cadherin/ catenin complex from the actin cytoskeleton and reduced cell–cell adhesion in pancreatic tumour cells [41]. During development and tumourigenesis epithelial cells may undergo epithelial-to-mesenchymal transition (EMT) which essentially involves the conversion of the epithelial cell into a myofibroblast. In adult tissues such as the kidney and lung, epithelial cells undergo EMT leading to pathophysiological tissue changes resulting in fibrosis. This process has been shown to involve CTGF acting downstream of TGFβ [22,42–44]. However, the potential role of EMT during re-epithelialisation
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of stratified epithelium where the epithelial cells do not directly engage in fibrosis is unclear. Multilayered epithelium such as in skin and cornea is constantly regenerated by a population of stem cells which reside in discreet locations within the basal layer of the interfollicular epidermis and limbus, respectively [45–47]. Once these cells leave there niche they divide giving rise to daughter transient amplifying cells [48]. Interestingly during limbal explant culture, limbal epithelial cells can undergo intrastromal invasion (possibly via EMT) [49] whereas corneal epithelial cells seeded onto the same region of the limbus do not [50]. This suggests a fundamental change in the properties of limbal stem cell daughters once they have exited the niche. One study has suggested that epithelial cell migration during corneal wound re-epithelialisation does indeed shares features with EMT, specifically the up-regulation of Slug expression which was found to be associated with internalisation of E-cadherin and β-catenin from the cell membrane, enhanced expression of alpha smooth muscle actin, tropomyosin and matrix metalloproteinases at the epithelial wound margin [51]. The potential role of CTGF in the regulation of disassembly of E-cadherin/catenin complexes in cellular adherens junctions from the actin cytoskeleton to facilitate epithelial cell migration without stromal invasion, as occurs during re-epithelialisation, is unknown. It has previously been suggested that pharmacological inhibition of CTGF could become an alternative target for antifibrosis therapy via selective inhibition of elements of the TGFβ response in fibroblasts. Indeed, one such approach suggested in the context of skin fibrosis was targeting of the TGFβ induced Ras/MEK/ERK pathway to prevent expression of CTGF [34]. Whilst this approach may not interfere with keratinocyte function in the skin e.g. in the potential treatment of the connective tissue disease scleroderma, our data would suggest a detrimental effect upon re-epithelialisation of the cornea. Hence, testing of potential CTGF antiscarring strategies should include screening of the potential functional effects upon epithelial cells likely to come into contact with the reagent. In our hands the human skin keratinocyte HaCaT cell line was also capable of inducing wound healing type responses to both TGFβ and CTGF, indicating that CTGF regulation of epithelial cells from stratified epithelium is not restricted to cells of corneal origin. In summary, we propose a novel role for CTGF during tissue repair outside of its established involvement in fibrosis, which is in the regulation TGFβ-stimulated re-epithelialisation of stratified epithelium via epithelial cell migration. Furthermore, we present evidence to suggest that, TGFβ is unable to induce epithelial cell migration in the absence of CTGF. Upon binding of TGFβ to its receptor on epithelial cells, the Ras/MEK/ERK MAPK signalling pathway is activated followed by CTGF gene expression and protein production. We postulate that activation of p38 plays a cooperative role, possibly by enhancing CTGF protein translation since blocking of this signalling pathway only reduces, but does not eliminate, CTGF protein production. CTGF is then released from the cell whereupon it binds to its receptor, inducing epithelial cell migration in an autocrine manner, which occurs at least in part via the Ras/MEK/ERK signalling pathway. Our data demonstrates that CTGF can stimulate cells of stratified epithelia to engage in the non-
fibrotic process of re-epithelialisation and further highlights the complexity of TGFβ regulation of epithelial cell function.
Acknowledgments We would like to thank Priscilla Philips for the ELISA work. This work was supported by The Eranda Foundation (GAS), The Special Trustees of Moorfields Eye Hospital (JTD) and NEI grant EY05587 (GSS).
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