Regulation of Transforming Growth Factor-β-dependent Cyclooxygenase-2 Expression in Fibroblasts

Abstract

Abnormal transforming growth factor-β (TGF-β) signaling is a critical contributor to the pathogenesis of various human diseases ranging from tissue fibrosis to tumor formation. Excessive TGF-β signaling stimulates fibrotic responses. Recent research has focused in the main on the antiproliferative effects of TGF-β in fibroblasts, and it is presently understood that TGF-β-stimulated cyclooxygenase-2 (COX-2) induction in fibroblasts is essential for antifibroproliferative effects of TGF-β. Both TGF-β and COX-2 have been implicated in tumor growth, invasion, and metastasis, and therefore tumor-associated fibroblasts are a recent topic of interest. Here we report the identification of positive and negative regulatory factors of COX-2 expression induced by TGF-β as determined using proteomic approaches. We show that TGF-β coordinately up-regulates three factors, heterogeneous nuclear ribonucleoprotein A/B (HNRPAB), nucleotide diphosphate kinase A (NDPK A), and nucleotide diphosphate kinase A (NDPK B). Functional pathway analysis showed that HNRPAB augments mRNA and protein levels of COX-2 and subsequent prostaglandin E₂ (PGE₂) production by suppressing degradation of COX-2 mRNA. In contrast, NDPK A and NDPK B attenuated mRNA and protein levels of COX-2 by affecting TGF-β-Smad2/3/4 signaling at the receptor level. Collectively, we report on a new regulatory pathway of TGF-β in controlling expression of COX-2 in fibroblasts, which advances our understanding of pathophysiological mechanisms of TGF-β.

Introduction

Transforming growth factor-β (TGF-β)³ is a multifunctional cytokine that regulates a multitude of physiological and pathological processes. TGF-β controls various cell activities ranging from cell differentiation, proliferation, and migration to extracellular matrix production (1–5). Therefore, abnormal regulation of TGF-β-dependent signaling pathways often leads to diseases, such as fibrosis-related diseases (6, 7), cancer (8–11), cardiovascular disease (12, 13), and autoimmune diseases (7). Among the wide range of biological functions regulated by TGF-β, its effect on fibrosis remains one of the most extensively studied research areas because excessive TGF-β signaling is a critical contributor to the pathogenesis of various fibrotic diseases (2, 7).

TGF-β directly promotes expression of type I collagen, a major component of the extracellular matrix, during fibrosis. TGF-β further induces connective tissue growth factor expression in fibroblasts, which synergizes with TGF-β to induce fibrosis. Epithelial to mesenchymal transition by TGF-β is another contributor to the fibrotic response. However, in addition to its well established profibrotic effects, several recent reports have demonstrated a key role of TGF-β as an antifibrotic regulator through regulation of cyclooxygenase-2 (COX-2).

Cyclooxygenase is a rate-limiting enzyme in the synthesis of prostaglandins, which are autocrine mediators of multiple cellular processes. Although cyclooxygenase-1 (COX-1) is constitutively expressed in most tissues and involved mainly in homeostasis, COX-2 is usually absent under basal conditions and induced by stimuli, such as growth factors and cytokines. Many lines of evidence have shown that the COX-2-prostanoid pathway is involved in many physiological and pathogenetic pathways, including those that regulate fibrosis, cancer, inflammation, angiogenesis, hemodynamics, and renal function (14–17). Although COX-2 and its major product, prostaglandin E₂ (PGE₂), are generally considered potent proinflammatory mediators, they also possess antifibrotic effects (14). Increased secretion of PGE₂ in response to TGF-β has been documented in fibroblasts and shown to be responsible for its antiproliferative effects. Suppression of COX-2 up-regulation and subsequent PGE₂ production, which acts in an autocrine fashion to inhibit proliferation and production of collagen, results in loss of the antiproliferative effects of TGF-β (18). Further, COX-2-deficient mice show enhanced susceptibility to pulmonary fibrosis (19) and cardiac fibrosis (20). Thus, TGF-β-stimulated COX-2 induction in fibroblasts is thought to be essential for the antifibroproliferative effects of TGF-β.

Moreover, recent mounting evidence has confirmed that tumor-associated stromal fibroblasts play critical roles in tumor development and progression (9, 10, 21). Stromal fibroblasts have been shown to be the predominant source of COX-2 in colon adenomas (22), and increased expression of COX-2 has been demonstrated in not only cancer cells per se but also the surrounding fibroblasts in invasive carcinomas (23). Considering that up-regulation of COX-2 and the subsequent prostaglandin cascade play crucial roles in tumorigenesis, tumor invasion, and metastasis (15), TGF-β-induced COX-2 expression in fibroblasts is envisioned to be a critical contributory factor in the tumor microenvironment. However, the involved molecular mechanisms still remain a subject of intense investigation.

Thus, how TGF-β-mediated COX-2 induction is controlled, especially in fibroblasts, is an important question to be addressed for elucidating the biology of these diseases. Here we show a new regulatory pathway of TGF-β-dependent COX-2 expression as centered on coordinated regulation of three newly identified factors using a differential proteomic study. Our findings advance our understanding of the underlying mechanisms of TGF-β pathophysiology in various diseases, including fibrotic and cancer diseases.

EXPERIMENTAL PROCEDURES

Cell Culture

Murine 10T1/2 fibroblasts were maintained in Dulbecco's modified Eagle's medium (Sigma) supplemented with 10% fetal bovine serum with 100 μg/ml streptomycin and 100 units/ml penicillin G. After 24 h of growth arrest with serum-free medium, subconfluent 10T1/2 cells were stimulated with recombinant human TGF-β1 (R&D Systems) (1 ng/ml) or lipopolysaccharide (LPS) (Sigma) (1 μg/ml) and harvested at the indicated times.

Liquid Chromatography

Cell pellets before and 12 h after TGF-β stimulation were suspended in lysis buffer containing 6 m urea, 2 m thiourea, 10% glycerol, 50 mm Tris- HCl, 2% n-octyl glucoside, 5 mm Tris(2-carboxyethyl)phosphine hydrochloride, and 1 mm protease inhibitor (Sigma) and then centrifuged for 1 h at 18 °C. The supernatants were recovered and stored at −80 °C for further use. Cell extracts were analyzed using a two-dimensional liquid chromatography system (ProteomeLab PF2D, Beckman Coulter), according to the manufacturer's instructions and as previously described (24). Briefly, as the first dimension, chromatofocusing was performed on a high performance chromatofocusing column with a linear pH gradient generated with two buffers, the start buffer (pH 8.5 ± 0.1) and the eluent buffer (pH 4.0 ± 0.1). Fractions were collected, and then residual proteins were washed with 1 m NaCl. The collected fractions were automatically and sequentially applied to a high performance reversed phase column as the second dimension separation, which consists of a linear gradient of 0–100% B against A for 30 min, where A is 0.1% trifluoroacetic acid in water and B is 0.08% trifluoroacetic acid in acetonitrile. Collected fractions were stored at −20 °C for further analysis.

Matrix-assisted Laser Desorption Ionization Time-of-flight (MALDI-TOF) Mass Spectrometry

Fractions were reduced to a volume of 16 μl using a vacuum concentrator, and 2 μl of 1 m NH₄HCO₃, 1 μl of 10 mm dithiothreitol, and 50–150 ng of modified porcine trypsin (Promega) were added. After overnight incubation at 37 °C, 1 μl of 1% trifluoroacetic acid was added to stop digestion. Digests were desalted with C18 ZipTips (Millipore) and then eluted with 70% acetonitrile and 0.1% trifluoroacetic acid. The eluted peptide samples were mixed with a saturated matrix of α-cyanohydroxycinnamic acid, spotted onto a MALDI plate, and analyzed with a MALDI-TOF mass spectrometer (Voyager-DE STR; Applied Biosystems) in reflectron mode. All data sets were analyzed using the mass spectrometer software (Data Explorer, version 4.0, Applied Biosystems). Each spectrum was base line-corrected with noise reduction and smoothing before it was deisotoped, leaving only the monoisotopic masses. The spectra were internally calibrated using trypsin autolysis peaks (generated at 842.5099 Da and 2211.1046 Da). The peak detection threshold was manually adjusted over the background. Known contaminant peaks (e.g. trypsin and keratin) and matrix peaks were excluded manually from all mass lists.

Protein Identification

The mass list of each sample was entered into the Mascot public search engine (available at the Matrix Science web site) and used to query the Swiss-Prot data base 55.4 (385,721 sequences; 138,434,015 residues) using the following parameters: taxonomy, Mus musculus; enzyme, trypsin; one possible missed cleavage; monoisotopic masses; and peptide tolerance of 0.1 Da. Possible modifications included oxidation of Met and modification of Cys by carbamidomethylation. Results were scored using the probability-based Mowse score. Protein scores greater than 54 are considered significant (p < 0.05).

Preparation of Plasmid Constructs

The expression vector pCAG was previously described (25, 26). Full-length Ndpk a (nucleotide diphosphate kinase A), Ndpk b (nucleotide diphosphate kinase B), and Hnrpab (heterogeneous nuclear ribonucleoprotein A/B) were PCR-amplified from aortic cDNA and subcloned into pCAG vector with an N-terminal FLAG tag (Sigma) to construct a mammalian expression vector for FLAG-tagged proteins.

Quantitative RT-PCR Assay

Total RNA was obtained by the RNeasy preparation kit (Qiagen) and reverse transcribed, and then quantitative PCR was performed with a gene-specific primer set and a QuantumRNA 18 S internal standard primer set (Ambion). For real-time quantitative RT-PCR, reactions were performed in duplicate using the QuantiTect SYBR Green PCR kit (Qiagen) and a Light Cycler instrument (Roche Applied Science) according to the manufacturer's instructions. The sequences of gene-specific primer sets were as follows: HNRPAB, 5′-GGGAGGTCTAAACCCTGAAG-3′ and 5′-GGGCAACCTTGATTTCACAC-3′; HNRPAB (for discrimination of two isoforms), 5′-GGGAGGTCTAAACCCTGAAG-3′ and 5′-ATTCTGATGACCACCACGTC-3′; NDPK A, 5′-GGACCTTCTCAAGGAGCACTAC-3′ and 5′-ACCACAAGCTGATCTCCTTCTC-3′; NDPK B, 5′-TCTGAAGAACACCTGAAGCAGC-3′ and 5′-TAGTCGATCAGTTCTTCGGG-3′; COX-2, 5′-CATTCTTTGCCCAGCACTTC-3′ and 5′-CCTGAGTGTCTTTGACTGTG-3′. Error bars indicate S.E.

Western Blot Analysis

Cell extracts 24 h after TGF-β stimulation were subjected to SDS-PAGE analysis and then immunoblotted with anti-COX-2 rabbit polyclonal antibody (Cayman Chemical) and anti-COX-1 mouse monoclonal antibody (Cayman Chemical). For analysis of Smad proteins, anti-phospho-Smad2 rabbit polyclonal antibody (Cell Signaling) and anti-Smad2/3 mouse monoclonal antibody (BD Biosciences) were used.

Immunocytochemistry

Cells plated on glass coverslips were analyzed 24 h after TGF-β stimulation. Cells were washed in phosphate-buffered saline, fixed with 4% paraformaldehyde for 20 min, permeabilized with 0.1% Triton X-100 in phosphate-buffered saline for 10 min, and then blocked with 1% bovine serum albumin for 30 min. Subsequently, cells were incubated with the same primary antibodies as those for Western blot analysis, exposed to Alexa Fluor 488-conjugated goat anti-rabbit or anti-mouse IgG antibody (Invitrogen), and counterstained with propidium iodide. Fluorescent images were collected with a confocal laser microscope (MRC 1024; Bio-Rad). COX-2, COX-1, or Smad2/3 was visualized using green fluorescence, and nuclei were visualized using red fluorescence.

Adenoviral Infection

Recombinant adenoviruses harboring FLAG-tagged Ndpk a, Ndpk b, Hnrpab p37, and Hnrpab p42 were prepared with the AdEasy system using homologous recombination in bacteria as previously described (27). Forty-eight hours after adenoviral infection at 9 MOI, cells were harvested for RT-PCR or Western blot analysis. For determination of mRNA stability, 48 h after adenoviral infection at 3 MOI, transcription was stopped by the addition of 100 μm 5,6-dichlorobenzimidazole riboside (DRB) (Sigma). RNA samples were isolated at 0, 40, 80, and 120 min following DRB treatment. For assays of NDPKs, cells were growth-arrested with serum-free medium for 24 h at 24 h after infection at 30 MOI and then stimulated with TGF-β (1 ng/ml).

Measurement of PGE₂ Synthesis

Subconfluent cells maintained in Dulbecco's modified Eagle's medium (Sigma) with 10% fetal bovine serum were infected at 10 MOI with control empty adenovirus or adenovirus harboring HNRPAB. Culture medium was collected at 48 h after adenoviral infection. After the addition of indomethacin, a prostaglandin synthesis inhibitor, at a final concentration of 10 μg/ml, the medium was centrifuged and stored at −80 °C for further analysis. The concentration of PGE₂ in cell culture supernatant was estimated by using an enzyme-linked immunosorbent assay kit following the manufacturer's protocol (Cayman Chemical). The background concentration of PGE₂ in medium containing 10% fetal bovine serum was negligible (i.e. below the sensitivity of the assay). The enzyme-linked immunosorbent assay was performed in duplicate, and results are shown as means and S.E. Shown are representative data from one of three independent experiments.

RNAi Transfection

Silencer predesigned small interfering RNA targeted to NDPK A, NDPK B, and HNRPAB and Silencer Negative Control small interfering RNA was purchased from Ambion. Cells were harvested for RT-PCR or Western blot analysis at 24 or 48 h after transfection of double-stranded small interfering RNA using Lipofectamine 2000 (Invitrogen) according to the manufacturer's instructions.

Chromatin Immunoprecipitation Assay

Approximately 1 × 10⁶ cells were fixed by adding formaldehyde directly to media at 37 °C for 10 min to a final concentration of 1%, washed twice with ice-cold phosphate-buffered saline, and then harvested. After resuspending in 200 μl of SDS lysis buffer (50 mm Tris-HCl, pH 8.1, 10 mm EDTA, 1% SDS, 0.5 mm phenylmethylsulfonyl fluoride, 0.5 μg/ml leupeptin, and 1 μg/ml pepstatin), chromatin was sheared with a sonicator (Bioruptor, Cosmo Bio), centrifuged to remove debris, and then diluted to 2 ml in dilution buffer (16.7 mm Tris-HCl, pH 8.1, 167 mm NaCl, 1.2 mm EDTA, 0.01% SDS, 1.1% Triton X-100, 0.5 mm phenylmethylsulfonyl fluoride, 0.5 μg/ml leupeptin, and 1 μg/ml pepstatin). Ten microliters of Dynabeads (Invitrogen) preincubated with 1 μg of anti-Smad2/3 antibody (BD Biosciences) or normal control IgG (Santa Cruz Biotechnology, Inc., Santa Cruz, CA) were added to 900 μl of the diluted supernatant, rotated overnight at 4 °C, and then washed twice with low salt wash buffer (20 mm Tris-HCl, pH 8.1, 150 mm NaCl, 2 mm EDTA, 0.1% SDS, and 1% Triton X-100), once with LiCl wash buffer (10 mm Tris-HCl, pH 8.1, 250 mm LiCl, 1 mm EDTA, 0.5% Nonidet P-40, and 0.5% SDS), and once with TE (10 mm Tris-HCl, pH 8.1, and 1 mm EDTA). Immunoprecipitated DNA was eluted in 200 μl of elution buffer (10 mm Tris-HCl, pH 8.1, 300 mm NaCl, 5 mm EDTA, and 0.5% SDS), incubated at 65 °C for 4 h to reverse cross-links, and then purified using a PCR purification kit (Qiagen). The region of the Cox-2 promoter from −366 to −151 bp was amplified by PCR using the following primer pair: 5′-GAAGCTGTGACACTCTTGAG-3′ and 5′-CAGGCTTTTACCCACGCAAA-3′.

RESULTS

Differential Protein Expression Profile for TGF-β-treated and -untreated Cells Generated Using the Two-dimensional Liquid Chromatographic Fractionation System with Subsequent Protein Identification Using MALDI-TOF Mass Spectrometry

Extracts of control and TGF-β-treated cells were first applied to liquid chromatography, which separated proteins with chromatofocusing based on pI in the first dimension. Subsequently, selected fractions separated with chromatofocusing were sequentially and automatically applied to reversed phase chromatography as the second dimension, thus making a two-dimensional protein map based on pI and hydrophobicity (Fig. 1A). The center panel in Fig. 1A is a differential display between the treated and untreated cells, calculated by point-to-point subtraction of corresponding data. As shown in this panel, comparison before and after TGF-β stimulation revealed dozens of protein peaks, whose intensity increased or decreased after stimulation. These peaks were identified with MALDI-TOF mass spectrometry, based on the second dimensional elution profiles, whose representative data are presented in Fig. 1B. The identified proteins are listed in Table 1 along with the corresponding band label in Fig. 1, protein name, accession number in the Swiss-Prot data base, Mascot score, and percentage of coverage.

FIGURE 1.

A differential display protein expression map for TGF-β-treated and -untreated cells. After 24 h of growth arrest with serum-free medium, subconfluent cells were stimulated with recombinant human TGF-β1 (1 ng/ml) for 12 h and then harvested. A, cellular protein separation of control cells (left) and TGF-β-stimulated cells (right) by two-dimensional liquid chromatographic fractionation. The x axis shows decreasing pH from 6.2 to 4.2, and the y axis displays increasing hydrophobicity (%B). The intensity of blue bands represents the relative protein amount as detected by UV absorption at 214 nm. The center panel is a two-dimensional differential map of the protein expression of the treated versus untreated cells. The lanes in the map are obtained by point-to-point subtraction of corresponding data. The green bands indicate up-regulated proteins, and the red bands represent down-regulated proteins in TGF-β treated cells. Protein bands identified by subsequent MALDI-TOF mass spectrometry analysis are labeled in a corresponding manner with those in Table 1 and B. B, representative elution profiles of selected pI fractions (fractions 4 and 9). The blue line depicts cell extract of untreated cells, and the red line shows that of TGF-β-treated cells. Three peaks higher in the TGF-β-stimulated cells (C, M, and L′) in these graphs were identified as NDPK B, HNRPAB, and vimentin, respectively.

TABLE 1.

A list of proteins increased in TGF-β-treated cells as identified by MALDI-TOF mass spectrometry

Band	Protein name	Swiss-Prot accession no.	Mascot score	No. of peptides	Percentage of coverage	Increasea
					%	-fold
A	Peroxiredoxin-2	Q61171	71	5	24	9.2
B	T-complex protein 1 subunit β	P80314	71	6	19	1.9
C	Nucleoside diphosphate kinase B	Q01768	110	6	51	2.0
C'	Nucleoside diphosphate kinase B	Q01768	82	5	45
D	Phosphoenolpyruvate carboxykinase (GTP), mitochondrial precursor	Q8BH04	61	5	9	4.2
E	Ornithine aminotransferase, mitochondrial precursor	P29758	76	8	22	1.4
F	Stress-70 protein, mitochondrial precursor	P38647	80	7	14	3.5
G	Actin, cytoplasmic 1	Q6ZWM3	67	5	20	2.1
	Actin, cytoplasmic 2	P63260	67	5	20
G'	Actin, cytoplasmic 1	Q6ZWM3	67	5	20
	Actin, cytoplasmic 2	P63260	67	5	20
H	60-kDa heat shock protein, mitochondrial precursor	P63038	122	9	24	2.7
I	Protein-disulfide isomerase A3 precursor	P27773	166	12	25	+
J	Heat shock cognate 71-kDa protein	Q3U9G0	134	13	28	+
K	ATP synthase subunit β, mitochondrial precursor	Q3U774	186	14	38	3.5
L	Vimentin	P20152	211	14	36	1.8
L'	Vimentin	P20152	170	13	39
M	Heterogeneous nuclear ribonucleoprotein A/B	Q99020	105	7	26	+
N	78-kDa glucose-regulated protein precursor	P20029	92	9	18	3.8

^a-Fold increase was estimated by calculating change in peak areas. +, proteins whose peaks could not be detected in the control cell lysate.

Up-regulation of NDPK B, HNRPAB, and Their Related Factors as Confirmed by RT-PCR Analysis and Increased COX-2 by Western Blot and Immunocytochemistry

Heterogeneous nuclear ribonucleoprotein A/B (HNRPAB) and nucleotide diphosphate kinase B (NDPK B) were among the proteins identified as being up-regulated by TGF-β treatment. HNRPAB has been previously reported to bind to the COX-2 3′-untranslated region and stabilize an exogenous COX-2 3′-untranslated region mRNA reporter (28). On the other hand, NDPK B belongs to the NDPK/NM23 gene family, whose member protein NM23-H1, a human orthologue of NDPK A, up-regulates the COX-2 promoter activity in reporter assays (29). Because the effect of these factors on endogenous COX-2 expression level is hitherto unknown, we focused our subsequent studies on understanding the mechanisms underlying the actions of these factors on COX-2 regulation.

First, to validate the proteomic screening analysis, mRNA levels of HNRPAB and NDPK B were analyzed by conventional RT-PCR and real-time PCR assays. As shown in Fig. 2A, the transcription levels of both factors were augmented after TGF-β stimulation in a time-dependent manner. mRNA levels of their related factors, NDPK A and COX-2, were also increased in a similar time-dependent manner. Next, because HNRPAB exists as two isoforms, heterogeneous nuclear ribonucleoprotein A/B (Q99020) and S1 protein C2 (Q20BD0) that differ by a 47-amino acid insertion close to the C terminus (28), we examined which isoform of HNRPAB is induced by TGF-β, using a PCR primer pair that discriminates the two isoforms. As shown in the lowest left panel in Fig. 2A, TGF-β led to the expression of both isoforms of HNRPAB. We refer to the smaller isoform as HNRPAB p37 and to the larger isoform as HNRPAB p42 hereafter.

FIGURE 2.

Effect of TGF-β stimulation on expression levels of HNRPAB, NDPK A, NDPK B, and COX-2. After 24 h of growth arrest with serum-free medium, cells were stimulated with recombinant human TGF-β1 (1 ng/ml) and then harvested. A, induction of NDPK B, HNRPAB, their putative downstream gene, COX-2, and a related gene, NDPK A, by TGF-β as shown by RT-PCR studies. Further, the lowest left panel shows that both isoforms of HNRPAB were up-regulated after TGF-β stimulation. Real-time PCR assay results are shown in the graph on the right. In the upper graph, the solid line and squares indicate NDPK A, the dotted line and circles show NDPK B, and the dotted line and triangles show HNRPAB. B, Western blot analysis confirming the increased protein level of COX-2 24 h after TGF-β stimulation. Note that the expression level of COX-1, which is known to be a relatively constitutive protein, was not affected by TGF-β as confirmed in the lower panel. C, immunofluorescence staining of TGF-β-stimulated cells (a and c) and control cells (b and d) for COX-2 (a and b) and COX-1 (c and d), indicating the induction of COX-2 protein 24 h after TGF-β stimulation, in contrast to the constitutive expression of COX-1, as shown by green fluorescence. Red fluorescence represents nuclei stained with propidium iodide. The subcellular location pattern of COX-2 is consistent with previous reports showing expression in the nuclear envelope and endoplasmic reticulum.

Further, effects on COX-2 expression were examined by Western blot, which showed a resultant increase in COX-2 protein in contrast to the constitutive expression of COX-1 (Fig. 2B) and immunocytochemistry (Fig. 2C) with further documentation of characteristic prominent staining of the nuclear envelope (30).

HNRPAB Increases the mRNA and Protein Levels of COX-2 and Leads to the Production of PGE₂

To understand the effects of HNRPAB on COX-2, we overexpressed HNRPAB by adenoviral infection. As indicated in Fig. 3A, both isoforms of HNRPAB augmented the mRNA level of COX-2. HNRPAB p42 was more potent in inducing transcription levels of COX-2 than HNRPAB p37. These effects were accompanied by similar increases in the expression level of COX-2 protein as shown by Western blot analysis (Fig. 3B) and immunofluorescence (Fig. 3C). Also, the secretion of PGE₂, a downstream product of COX-2 metabolism, in cell culture supernatant was measured by enzyme immunoassay. Although the prominent induction of COX-2 protein by HNRPAB p42 did not lead to proportional accumulation of PGE₂, the significant induction of PGE₂ in culture medium of cells overexpressing HNRPAB was seen (Fig. 3D). Thus, overexpression of HNRPAB augmented COX-2 transcript, protein, and enzymatic activity.

FIGURE 3.

Effect of HNRPAB on expression levels of COX-2. Cells were harvested at 48 h after adenoviral infection. A, augmented mRNA expression levels of COX-2 by adenoviral overexpression of HNRPAB as shown by RT-PCR. The right panel shows quantification by real-time PCR assay. B, Western blot analysis confirming up-regulated protein level of COX-2 in the upper panel in contrast to constitutive expression of COX-1 in the lower panel. C, induction of COX-2 by HNRPAB as demonstrated by immunocytochemistry. Adenovirus-mediated forced expression of HNRPAB increased COX-2 protein (b and c) compared with cells infected with control empty adenovirus (a) as shown by green fluorescence. Red fluorescence represents nuclei stained with propidium iodide. D, increased secretion of PGE₂ in culture medium by HNRPAB as measured by enzyme immunoassay. Culture medium was collected at 48 h after adenoviral infection at 10 MOI. E, the synergistic effect of TGF-β and HNRPAB on COX-2 mRNA induction. Cells were growth-arrested with serum-free medium for 24 h at 24 h after adenoviral infection, stimulated with TGF-β (1 ng/ml) for 8 h, and then harvested. Under the condition in which TGF-β increased COX-2 mRNA (lane 1 versus lane 4), HNRPABs augmented the stimulatory effect of TGF-β on COX-2 (lane 4 versus lanes 5 and 6). The right graph shows the result of a real-time PCR assay. Overexpression of HNRPAB (gray bars for HNRPAB p37 and black bars for HNRPAB p42) resulted in synergistic increase in mRNA levels of COX-2, as compared with infection of control empty adenovirus (white bars). F, Western blot analysis confirming increased protein levels of TGF-β-induced COX-2 by HNRPABs (lane 4 versus lanes 5 and 6) under the condition in which TGF-β increased COX-2 protein (lane 1 versus lane 4). The treatment condition was the same as that of E, but cells were harvested 24 h after TGF-β stimulation. Note that neither TGF-β nor HNRPABs affected the expression level of COX-1 as demonstrated by the lower panel.

In addition, we examined the effect of HNRPAB on TGF-β-induced COX-2 mRNA and protein. As shown in Fig. 3, E and F, TGF-β and HNRPAB showed a synergistic effect on COX-2 expression.

Adenoviral Transfer of HNRPAB Stabilizes COX-2 mRNA Levels

HNRPAB has been previously reported to stabilize an exogenous COX-2 3′-untranslated region mRNA reporter in HeLa cells (28). However, it is unknown whether overexpression of HNRPAB can stabilize endogenous mRNA of COX-2. To evaluate this potential post-transcription regulation, HNRPAB was overexpressed in cells using adenoviral transfer with further inhibition of transcription by the addition of DRB at 48 h. Quantitative RT-PCR assay (Fig. 4A) and real-time PCR (Fig. 4B) showed rapid degradation of COX-2 mRNA after DRB treatment in cells infected with control empty adenovirus (lanes 1–4 in Fig. 4A and solid line in Fig. 4B) and prolonged stabilization of COX-2 mRNA in cells overexpressing HNRPAB, especially p42 (lanes 5–12 in Fig. 4A and dotted lines in Fig. 4B). On the other hand, knockdown of HNRPAB using RNAi led to enhanced degradation of COX-2 mRNA (Fig. 4C). In addition, HNRPAB did not show any transactivation effects on a murine Cox-2 promoter in cotransfection reporter assays (data not shown). Collectively, the mechanism of HNRPAB-dependent up-regulation of COX-2 is mainly due to post-transcriptional stabilization of COX-2 mRNA.

FIGURE 4.

Regulation of endogenous COX-2 mRNA stability by HNRPAB. A, quantitative RT-PCR assay demonstrating suppressed degradation of COX-2 mRNA in cells overexpressing HNRPAB. Forty-eight hours after adenoviral infection, transcription was stopped by the addition of 100 mm DRB. RNA samples were isolated at 0, 40, 80, and 120 min following DRB treatment. COX-2 mRNA was rapidly diminished after DRB treatment in cells infected with control empty adenovirus (lower panel, lanes 1–4). Note that, in contrast, adenoviral transfer of HNRPAB contributed to prolonged stabilization of COX-2 mRNA over 120 min after DRB treatment (upper and middle panels, lanes 5–12). The three panels show the results of PCR with identical samples and increasing cycle numbers (24, 26, and 29 cycles). B, degradation curves of COX-2 mRNA as shown by real-time PCR assay. Although cells with empty adenovirus showed rapid decrease in COX-2 mRNA (solid line and squares), overexpression of HNRPAB resulted in no significant change in mRNA levels of COX-2 for 120 min (dotted lines and circles for HNRPAB p37 and triangles for HNRPAB p42). The expression level of COX-2 mRNA was normalized to that of 18 S rRNA and subsequently that at 0 min. C, augmented degradation of COX-2 mRNA by RNAi silencing of HNRPAB as revealed by real-time PCR assay. Twenty-four hours after RNAi transfection, transcription was stopped by the addition of 100 mm DRB. RNA samples were isolated at 0 and 60 min following DRB treatment. Cells transfected with RNAi targeting HNRPAB (dotted line and circles) showed enhanced degradation of COX-2 mRNA, as compared with cells transfected with negative control RNAi (solid line and squares). The expression level of COX-2 mRNA was normalized to that of 18 S rRNA and subsequently that at 0 min.

NDPK A and NDPK B Suppress TGF-β-dependent Augmentation of COX-2

Next, we assessed whether NDPK A and NDPK B increase COX-2 expression as is consistent with a previous report using reporter assays (29). Unexpectedly, however, ectopic overexpression of these factors diminished TGF-β-dependent induction of COX-2 mRNA (Fig. 5A). Suppressed COX-2 protein levels were also confirmed by Western blot analysis (Fig. 5B) and immunofluorescence (Fig. 5C). RNAi analysis was further used to confirm the effect of NDPK A and NDPK B on COX-2 by depleting these factors. As shown in Fig. 5, D and E, knockdown of either NDPK A or NDPK B led to increased COX-2 expression at both the mRNA and protein levels. Thus, NDPK A and NDPK B repress COX-2.

FIGURE 5.

Effect of NDPK A and NDPK B on the expression level of COX-2. A, suppression of mRNA expression level of COX-2 by adenoviral overexpression of NDPK A or NDPK B as shown by RT-PCR. At 24 h after adenoviral infection, cells were starved for an additional 24 h, treated with TGF-β (1 ng/ml) for 8 h, and then harvested. Under the condition in which TGF-β increased COX-2 mRNA (lane 1 versus lane 4), either NDPK A or NDPK B diminished the stimulatory effect of TGF-β on COX-2 (lane 4 versus lanes 5 and 6). The right graph shows the result of the real-time PCR assay. Overexpression of NDPKs (gray bars for NDPK A and black bars for NDPK B) led to an attenuated increase in mRNA levels of COX-2, compared with infection of control empty adenovirus (white bars). B, Western blot analysis confirming reduced protein levels of TGF-β-induced COX-2 by NDPK A or NDPK B (lane 4 versus lanes 5 and 6) under the condition in which TGF-β increased COX-2 protein (lane 1 versus lane 4). The treatment condition was the same as that of A, but cells were harvested 24 h after TGF-β stimulation. Note that neither TGF-β nor NDPKs affected the expression level of COX-1 as demonstrated by the lower panel. C, attenuation of TGF-β-stimulated COX-2 expression by NDPK A or NDPK B as demonstrated by immunocytochemistry. The treatment condition was the same as that of B. Adenoviral transfer of either NDPK A or NDPK B impaired the expression level of COX-2 protein (b and c) compared with cells infected with control empty adenovirus (a) as shown by green fluorescence. Red fluorescence represents nuclei stained with propidium iodide. D, augmentation of mRNA expression level of COX-2 by RNAi silencing of NDPK A or NDPK B, as indicated by RT-PCR. Cells were harvested 48 h after RNAi transfection. The middle and lower panels on the left demonstrate the specificity of RNAi constructs. The right graph shows the result of a real-time PCR assay. Note that RNAi targeting either NDPK A or NDPK B increased COX-2 mRNA (black bars) under the condition in which each RNAi construct decreased mRNA of each target specifically (gray bars for NDPK A and white bars for NDPK B). E, increased COX-2 protein by RNAi silencing of NDPK A or NDPK B as shown by Western blot analysis. Cells were harvested 48 h after RNAi transfection. The difference of effects between NDPK A and NDPK B might attribute to the different efficiency of RNAi suppression as demonstrated in D.

NDPK A and NDPK B Impair TGF-β-dependent Induction of COX-2 Not at the Transcriptional Level but at the TGF-β Receptor Level

With these apparently conflicting results, we conducted reporter assays similar to that used in a previous report to rule out cell type- dependent effects, which showed that NDPK A and NDPK B can transactivate the Cox-2 promoter (supplemental Fig. S1). It was therefore thought unlikely that NDPK A and NDPK B act on the Cox-2 promoter to impair transcription. We thus speculated that an alternative mechanism might be that NDPK A and NDPK B attenuate COX-2 expression not at the transcription level but at the receptor level, because a recent report has shown that NM23-H1, a human orthologue of NDPK A, interacts with serine-threonine kinase receptor-associated protein, a TGF-β receptor-interacting protein, and negatively regulates a number of TGF-β target genes in other cell lines (31).

TGF-β receptors can activate both Smad-dependent pathways and Smad-independent pathways (32, 33). First, to address whether TGF-β-induced COX-2 expression is mediated by Smad proteins, a chromatin immunoprecipitation assay using anti-Smad2/3 antibody was done. PCR analysis was performed to amplify the region of the Cox-2 promoter that contains possible Smad binding sites (34), including multiple Smad binding elements (AGAC), one AP-1-like site (TGCGTGG), and one Sp1 site (GGGCGG) (35). In vivo binding of Smad2/3 to the Cox-2 promoter was clearly induced in response to TGF-β stimulation (Fig. 6A). In addition, real-time PCR analysis showed that recruitment of Smad2/3 on the promoter is increased by the addition of TGF-β in a time-dependent manner (Fig. 6B). These findings suggest that TGF-β-dependent COX-2 expression is at least in part regulated by Smad2/3/4 pathways.

FIGURE 6.

Effect of NDPK A and NDPK B on the Smad2/3/4 signaling pathway and resultant Smad2/3 recruitment on the promoter region of Cox-2. A, in vivo recruitment of Smad2/3 on the promoter of Cox-2 upon TGF-β stimulation. After 24 h of growth arrest with serum-free medium, cells were stimulated with TGF-β (1 ng/ml). Chromatin was harvested 8 h after TGF-β treatment and then immunoprecipitated (IP) with anti-Smad2/3 antibody (lanes 5 and 6) or normal mouse IgG as a negative control (lanes 3 and 4). Note that direct recruitment of Smad2/3 to the Cox-2 promoter was only seen with TGF-β treatment (lane 5 versus lane 6). Lanes 1 and 2 are input (1%), confirming that the applied chromatin amounts were the same. B, time course of Smad2/3 binding to the Cox-2 promoter. After 24 h of growth arrest with serum-free medium, cells were stimulated with TGF-β (1 ng/ml) and then harvested at the indicated times. Chromatin precipitated by anti-Smad2/3 antibody (solid line and squares) or normal mouse IgG (dotted line and circles) was quantified by real-time PCR. Data were normalized by input DNA. Error bars, S.E. C, suppression of phosphorylation levels of Smad2 by adenoviral transfer of NDPK A or NDPK B as shown by Western blot analysis. Cells were growth-arrested with serum-free medium for 24 h at 24 h after adenoviral infection, stimulated with TGF-β (1 ng/ml) for 12 h, and then harvested. Under the condition in which TGF-β increased phosphorylated Smad2 (lane 1 versus lane 4), either NDPK A or NDPK B reduced the phosphorylation level of Smad2 (lane 4 versus lanes 5 and 6). D, immunofluorescence staining of Smad2/3, indicating that NDPK A or NDPK B suppressed TGF-β-stimulated nuclear translocation of Smad2/3. Cells were growth-arrested with serum-free medium for 24 h at 24 h after adenoviral infection, stimulated with TGF-β (1 ng/ml) for 8 h, and then harvested. Although TGF-β induced nuclear translocation of Smad2/3 as shown by prominent green fluorescence staining of nuclei of TGF-β-treated cells (a versus b), overexpression of NDPK A or NDPK B attenuated the nuclear translocation of Smad2/3 proteins (c and d), compared with cells infected with control empty adenovirus (b). Red fluorescence represents nuclei stained with propidium iodide. E, chromatin immunoprecipitation assay demonstrating the attenuated recruitment of Smad2/3 on the promoter of Cox-2 by NDPK A and NDPK B. At 24 h after adenoviral infection, cells were starved for an additional 24 h, treated with TGF-β (1 ng/ml) for 8 h, and then harvested. Chromatin was immunoprecipitated with anti-Smad2/3 antibody (lanes 9–12) or normal mouse IgG as a negative control (lanes 5–8). Under the condition in which TGF-β recruited Smad2/3 onto the promoter of Cox-2 (lane 9 versus lane 10), adenoviral forced expression of either NDPK A or NDPK B attenuated the binding of Smad2/3 to the promoter, as compared with infection with empty virus (lane 10 versus lanes 11 and 12). Lanes 1–4 are input (1%), confirming that the applied chromatin amounts were the same. The graph on the right shows results of real-time PCR assay. Data were normalized by input DNA.

In the Smad-dependent pathways, the TGF-β receptor phosphorylates Smad2 and Smad3. Phosphorylated Smad2 and Smad3 then form oligomers with or without Smad4 that translocate to the nucleus, where they regulate the transcription of downstream genes (32–34). Thus, we assessed whether NDPK A and NDPK B can suppress each step of these pathways. First, adenoviral overexpression of these factors decreased TGF-β-mediated phosphorylation of Smad2 as shown by Western blot (Fig. 6C). Next, subsequent nuclear translocation of Smad2/3 was impaired as demonstrated by immunofluorescence (Fig. 6D). Moreover, chromatin immunoprecipitation showed that resultant TGF-β-induced recruitment of Smad2/3 onto the Cox-2 promoter was clearly attenuated by forced expression of NDPK A or NDPK B (Fig. 6E). Collectively, these findings confirmed that NDPK A and NDPK B can suppress TGF-β-stimulated induction of COX-2 by down-regulating the Smad pathways at the receptor level.

LPS Does Not Up-regulate HNRPAB and NDPKs, and Their Effects on LPS-induced COX-2 Are Limited

To examine the generality and specificity of the involved mechanisms, we assessed whether LPS can up-regulate HNRPAB and NDPKs (Fig. 7A). Although LPS induced transient increase of COX-2, LPS did not show significant effects on the expression levels of HNRPAB, NDPK A, or NDPK B.

FIGURE 7.

Effects of HNRPAB, NDPK A, and NDPK B on the expression level of LPS-induced COX-2. A, effect of LPS stimulation on expression levels of COX-2, NDPK A, NDPK B, and HNRPAB. After 24 h of growth arrest with serum-free medium, cells were stimulated with LPS (1 μg/ml) and then harvested at the indicated times. The right graph shows the results of real-time PCR assay. Although LPS induced transient increase of COX-2 (solid heavy line and large squares), LPS did not show significant effects on the expression levels of NDPK A (solid line and squares), NDPK B (dotted line and circles), or HNRPAB (dotted line and triangles). B, effect of HNRPAB on COX-2 mRNA induction stimulated by LPS. Cells were growth-arrested with serum-free medium for 24 h at 24 h after adenoviral infection at 10 MOI, stimulated with LPS (1 μg/ml), and then harvested at the indicated times. Under the condition in which LPS increased COX-2 mRNA (lane 1 versus lane 4), HNRPABs augmented the stimulatory effect of LPS on COX-2 at 6 h (lane 7 versus lanes 8 and 9). However, little effect was seen on the peak induction level of COX-2 at 2 h (lane 4 versus lanes 5 and 6). The right graph shows results of real-time PCR assay. Overexpression of HNRPAB (dotted lines and circles for HNRPAB p37 and triangles for HNRPAB p42) up-regulated mRNA levels of COX-2 at 6 h, as compared with infection of control empty adenovirus (solid line and squares). C, effect of NDPKs on LPS-dependent COX-2 mRNA induction. Cells were growth-arrested with serum-free medium for 24 h at 24 h after adenoviral infection at 30 MOI, stimulated with LPS (1 μg/ml), and then harvested at the indicated times. Under the condition in which LPS increased COX-2 mRNA (lane 1 versus lane 4), NDPKs did not show significant inhibitory effect on LPS-stimulated COX-2 expression (lanes 4–9). The right graph shows the results of a real-time PCR assay. Overexpression of NDPKs (dotted lines and circles for NDPK A and triangles for NDPK B) resulted in no decrease or only a slight decrease in mRNA levels of COX-2, as compared with infection of control empty adenovirus (solid line and squares).

Further, we overexpressed HNRPAB and NDPKs in the presence of LPS. As demonstrated in Fig. 7B, although HNRPAB showed no significant effect on the peak induction level of COX-2 (lanes 4–6), HNRPAB slightly increased COX-2 mRNA at later phases (lanes 7–9), suggesting its mRNA- stabilizing effect. However, as a whole, its regulatory ability on LPS-induced COX-2 seems to be limited in this setting. Moreover, overexpression of NDPKs resulted in no suppression or only mild suppression of COX-2 induction as shown in Fig. 7C. This observation that NDPKs did not efficiently attenuate the expression level of LPS-induced COX-2 is consistent with the proposed mechanism that they impair COX-2 expression mainly at the TGF-β receptor level.

DISCUSSION

A carefully controlled balance of positive and negative signaling pathways regulates normal physiological cellular activities. Their inappropriate alteration often leads to human diseases. Thus, understanding regulatory signaling networks, especially those of growth factors, is a critical step in elucidating mechanisms that regulate development of pathophysiological conditions. Although the complexity of TGF-β signaling is very challenging to analyze, a differential proteomic approach allowed us to discover both positive and negative components of TGF-β-stimulated regulation of COX-2 expression. Although few previous reports have described that TGF-β induces COX-2 in several cell lines (18, 36, 37), surprisingly, almost nothing has been described about underlying molecular mechanisms. This is the first report that shows that HNRPAB is a TGF-β-induced factor that directly increases the expression level of COX-2 protein. Further, our findings on NDPKs are intriguing, considering their possible roles as negative regulatory components of TGF-β-dependent pathways. Moreover, this is the first to report on a direct link between TGF-β-Smad2/3/4 signaling and COX-2 expression (supplemental Fig. S2).

Implications in Fibroproliferative Diseases

Fibroproliferative diseases, including cardiac fibrosis, pulmonary fibrosis, systemic sclerosis, liver cirrhosis, progressive kidney disease, and macular degeneration, are one of the leading causes of morbidity and mortality and can affect all tissues and organ systems (6). TGF-β plays a pivotal role in regulating fibrosis and can exert both pro- and antiproliferative effects on fibroblasts. TGF-β induces a biphasic response in lung fibroblasts with stimulation of proliferation at low concentrations and inhibition at high concentrations, the latter of which is accounted for by autocrine synthesis of PGE₂ (18). Interestingly, although inappropriate activation of TGF-β signaling leads to excess fibrosis in the pathogenesis of various fibrotic diseases, genetic ablation of Cox-2, one of the downstream targets of TGF-β, also results in excessive fibrosis, as shown in Cox-2 null mice (19, 20). Although antifibrotic therapies are considered to be promising, an incomplete understanding of the molecular machinery of TGF-β-dependent pathways has hindered the development of such therapies.

Here we have demonstrated that TGF-β up-regulates HNRPAB. Of note, HNRPAB is a single protein with multiple functions. Although HNRPAB acts as an RNA-binding protein, it was also identified independently as CArG box-binding factor-A on the basis that it binds to single-stranded and double-stranded DNA in a sequence-specific manner as a transcriptional regulator (38–40). A recent report has demonstrated the key role of this factor in epithelial-mesenchymal transition, a main contributor to tissue fibrosis (40). Thus, HNRPAB is a unique protein that shows bimodal actions; as a profibrotic factor, it is a DNA-binding protein that promotes epithelial-mesenchymal transition, and as an antifibrotic factor, it is an RNA-binding protein that stabilizes mRNA of COX- 2. Therapeutic modification of its RNA-binding ability that does not affect its DNA-binding capacity or vice versa may be potentially exploitable for targeted therapeutic intervention against fibroproliferative diseases.

Possible Roles in Carcinogenesis and Metastasis

The significance of TGF-β signaling in carcinogenesis and metastasis has been widely investigated. In general, TGF-β is a tumor suppressor early in carcinogenesis, inhibiting growth and promoting differentiation, and then acts as a tumor promoter later in cancer development to promote growth, survival, invasion, and metastasis (2, 8, 9, 11). COX-2 is also involved in cancer progression. Clinical and experimental data have demonstrated that increased COX-2 expression correlates with poor survival in various cancers, and selective inhibitors of COX-2 inhibit tumor growth (15). Stromal fibroblasts as well as cancer cells per se have been shown to be the source of COX-2 (15, 22, 23). Given that tumor-associated fibroblasts are important contributors to the tumor microenvironment (9, 10, 21), TGF-β-mediated COX-2 expression in fibroblasts is a topic that deserves further attention.

An earlier proteomic report identified HNRPAB as one of eight up-regulated genes in the gene expression signature associated with various tumor metastases (41). Notably, the authors suggested that a considerable proportion of the gene expression signature seems to be derived from stromal components of tumors. Thus, not only TGF-β and COX-2 but also HNRPAB is probably up-regulated in the tumor microenvironment. Although nothing has been shown concerning their relationship in tumor-associated fibroblasts, our findings lead us to propose a straightforward pathway by which TGF-β stimulates HNRPAB, which in turn increases COX-2 in tumor-associated fibroblasts, thus resulting in tumor development and progression.

Bifunctional Effects of NDPKs

NDPK A and NDPK B are members of the NDPK/NM23 family, which have multiple activities ranging from DNA-binding in the nuclei to protein-protein interaction in the cytosol. These family proteins have been shown to be involved in many cellular processes, including cell growth, differentiation, metastasis, and development (42–45). Although our findings in the present study are based on experiments using fibroblasts, this novel pathway potentially impacts on the biology of tumor cells. NDPK was first characterized as a metastasis suppressor in melanoma (46). Its high expression was associated with less metastatic potential in various tumors, including melanoma, breast cancer, colon cancer, and liver cancer. In contrast, increased expression in prostate cancer, neuroblastoma, and lymphoma is related to tumor aggressiveness (42, 47). Despite the widely accepted notion of close involvement of NDPKs in tumor progression, no clear consensus has been made regarding its precise roles in tumor biology, or rather, NDPKs are considered to have multifaceted roles in a variety of tumors in a highly context-dependent manner. We note that an earlier study showed that NM23-H1, a human orthologue of NDPK A, up-regulates COX-2 promoter activity in reporter assays (29). This is in agreement with the tumor-progressive property of NDPKs, considering the effect of COX-2 on tumor progression. On the other hand, our finding that NDPKs attenuate COX-2 expression is consistent with classically proposed functions of NDPK as a metastasis suppressor and possible tumor suppressor. The molecular basis of multiple NPDK-dependent signaling pathways is, however, more complex, and further investigation is warranted to better understand the precise underlying mechanisms.

In conclusion, we have shown a new regulatory pathway of TGF-β-dependent COX-2 expression by HNRPAB and NDPKs, which provides a new perspective in understanding the biology of various diseases, including fibrotic and cancer diseases.