Retraction
Following publication of this work [1], concerns were raised about the following:
- Fig 2E Actin panel appears similar to lanes 2–4 of the GAPDH panel in Fig 7D. These panels also appear similar to images in Fig 2A (Actin), Fig 2B (Actin), Fig 4A (NS5A), 4B (NS5A), 7D (Actin), and 9A (Actin panel) of a 2003 article published in the Journal of Biological Chemistry [2] by the corresponding author of [1]. The 2003 publication has been withdrawn [3], and the withdrawal notice for that article notes similarity of the image in question to figures in several other articles.
- Fig 7B Albumin panel lanes 2,3 flipped horizontally appear similar to Fig 1A ASC panel lanes 3,4 in a 2016 article published in the Journal of Biological Chemistry [4] by two of the authors of [1]. The 2016 article has been withdrawn [5], and the withdrawal notice indicates similarity between this image and figures published in several other articles, including two published in 2010 and 2012 [6, 7].
While the authors stand by the validity of the experimental data and the conclusions reached in the study, they were unable to provide original images or underlying data for Figs 2E, 7D and 7E.
In light of the image concerns and the unavailability of the underlying data, the PLOS ONE Editors retract this article.
GW did not agree with retraction. LDP and SM did not respond.
Owing to the concerns about similarities with previously published content [2–7] that is not licensed for reproduction and distribution under the terms of the Creative Commons Attribution License, Fig 2E Actin panel, Fig 7B Albumin panel, and Fig 7D GAPDH panel have been removed from the retracted article [1].
30 Apr 2019: The PLOS ONE Editors (2019) Retraction: Activation of TGF-β1 Promoter by Hepatitis C Virus-Induced AP-1 and Sp1: Role of TGF-β1 in Hepatic Stellate Cell Activation and Invasion. PLOS ONE 14(4): e0216025. https://doi.org/10.1371/journal.pone.0216025 View retraction
Figures
Abstract
Our previous studies have shown the induction and maturation of transforming growth factor-beta 1 (TGF-β1) in HCV-infected human hepatoma cells. In this study, we have investigated the molecular mechanism of TGF-β1 gene expression in response to HCV infection. We demonstrate that HCV-induced transcription factors AP-1, Sp1, NF-κB and STAT-3 are involved in TGF-β1 gene expression. Using chromatin immunoprecipitation (ChIP) assay, we further show that AP-1 and Sp1 interact with TGF-b1 promoter in vivo in HCV-infected cells. In addition, we demonstrate that HCV-induced TGF-β1 gene expression is mediated by the activation of cellular kinases such as p38 MAPK, Src, JNK, and MEK1/2. Next, we determined the role of secreted bioactive TGF-β1 in human hepatic stellate cells (HSCs) activation and invasion. Using siRNA approach, we show that HCV-induced bioactive TGF-β1 is critical for the induction of alpha smooth muscle actin (α-SMA) and type 1 collagen, the markers of HSCs activation and proliferation. We further demonstrate the potential role of HCV-induced bioactive TGF-β1 in HSCs invasion/cell migration using a transwell Boyden chamber. Our results also suggest the role of HCV-induced TGF-β1 in HCV replication and release. Collectively, these observations provide insight into the mechanism of TGF-β1 promoter activation, as well as HSCs activation and invasion, which likely manifests in liver fibrosis associated with HCV infection.
Citation: Presser LD, McRae S, Waris G (2013) Activation of TGF-β1 Promoter by Hepatitis C Virus-Induced AP-1 and Sp1: Role of TGF-β1 in Hepatic Stellate Cell Activation and Invasion. PLoS ONE 8(2): e56367. https://doi.org/10.1371/journal.pone.0056367
Editor: Ching-Ping Tseng, Chang Gung University, Taiwan
Received: July 16, 2012; Accepted: January 14, 2013; Published: February 21, 2013
Copyright: © 2013 Presser et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: This work was supported in part by the American Cancer Society, Illinois Division, Inc., 09-14, National Institutes of Health/National Institute of Allergy and Infectious Diseases grant R21:A1078532, and by the Rosalind Franklin University of Medicine and Science-H.M. Bligh Cancer Research Fund to G.W. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: Gulam Waris, Ph.D., currently serves as an academic editor of PLOS ONE. This does not alter the authors’ adherence to all the PLOS ONE policies on sharing data and materials.
Introduction
HCV infection causes chronic hepatitis in a significant number of infected individuals, which may gradually progress to liver fibrosis, cirrhosis and subsequently to hepatocellular carcinoma (HCC) [1]. HCV is an enveloped, single-stranded, positive-sense RNA virus which is approximately 9.6 kb in length, contains both 5′ and 3′ untranslated regions (UTRs), and encodes a single polyprotein of about 3000 amino acids [2]. The 5′ UTR contains the internal ribosome entry site (IRES) which is required for cap-independent translation of the polyprotein. The polyprotein is cleaved by host and viral proteases into structural proteins (core, E1, and E2) and nonstructural proteins (p7, NS2, NS3, NS4A, NS4B, NS5A, and NS5B) [2], [3]. Until 2005, the studies of molecular mechanisms of HCV replication and pathogenesis had been hampered by the lack of an efficient cell culture system or a suitable small-animal model. The development of a productive HCV (genotype 2a) infection system provided a major breakthrough which allows the production of infectious virions in cell culture [4], [5], [6].
The molecular mechanisms underlying liver injury and fibrosis in chronic HCV remain unclear. TGF-β1 is the major profibrogenic cytokine which regulates the production and deposition of the major extracellular matrix molecules (ECM) [7]. It has been reported that HCV infection is associated with a significant increase in TGF-β1 expression and secretion in liver and serum respectively [8], [9]. Previously, we and others have demonstrated an increased secretion of bioactive TGF-β1 from HCV-infected cells [10], [11], [12], [13], [14]. In addition, several other viruses have been shown to activate TGF-β1, and in some cases, TGF-β1 has a positive effect on the replication of the virus [15]. For instance it has been previously reported that TGF-β1 enhances replication of respiratory syncytial virus in lung epithelial cells [16]. Human cytomegalovirus induces TGF-β1 activation in renal tubular epithelial cells after epithelial-to-mesenchymal transition [13]. TGF-β1 has also been shown to play an important role in HIV/HCV co-infection as HIV increases HCV replication in a TGF-β1 dependent manner [11].
TGF-β1 has been shown to be regulated by transcription factors such as AP-1, Sp1, NF-κB, EGR-1, USF, ZF9/core promoter binding protein, and STAT-3 in various experimental systems [10], [17], [18], [19], [20], [21], [22], [23], [24], [25], [26], [27]. It has been well documented that cellular kinases play key roles in HCV-mediated pathogenesis by activating downstream transcription factors. We and others have shown the activation of various cellular kinases in response to HCV-infection such as JNK, p38 MAPK, ERK, Src, PI3K and JAK, and these kinases induce transcription factors Nrf2, NF-κB, AP-1, Sp1, HIF-1α, ATF6, SREBPs, and STAT-3 [10], [12], [17], [26], [28], [29], [30], [31], [32], [33], [34], [35].
Human hepatic stellate cells (HSCs) comprise approximately 15% of all liver cells and are the major cell type involved in liver fibrogenesis [36], [37], [38]. HSCs are normally in a “quiescent” or quiet state but can become activated by the binding of bioactive TGF-β1 to TGF-β1 receptors on HSCs [37], [39]. Upon activation, HSCs up regulate the production of ECM proteins and become invasive [37], [39].
In the present study, we first demonstrate the mechanisms of TGF-β1 promoter activation and then the effect of secreted bioactive TGF-β1 on HSC activation and invasion. We show that transcription factors AP-1, Sp1, NF-κB, and STAT-3 play critical role in TGF-β1 gene expression. Furthermore, we demonstrate increased HSCs activation and invasion when HSCs were incubated with conditioned medium (CM) from HCV-infected cells which contain bioactive TGF-β1. We also determined the role of TGF-β1 in HCV production and release. These data collectively demonstrate the mechanisms for TGF-β1 gene expression by HCV infection, and the role of TGF-β1 on HSC activation and invasion which leads to liver fibrosis.
Materials and Methods
Cell Lines
The human hepatoma cell line, Huh-7.5, was obtained from Dr. C. Rice (Rockefeller University, NY) (Blight et al., 2002). Huh-7.5 cells were cultured at 37°C in a humidified atmosphere containing 5% CO2 with Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal calf serum, 100 U of penicillin/ml, and 100 µg of streptomycin sulfate/ml. The human hepatic stellate cell (HSC) line, LX-2 was obtained from Dr. S. Friedman (Mount Sinai Hospital, NY) [40]. LX-2 cells were cultured in DMEM as described above. For HSCs activation experiments LX-2 cells were serum starved in serum free DMEM for 48 h before use.
Plasmids, Reagents and Antibodies
The infectious J6/JFH-1 cDNA (HCV genotype 2a) was obtained from Dr. C. Rice (Rockefeller University, NY). The TGF-β1 promoter luciferase-reporter plasmids (phTG1 −1362/+11, phTG5 −453/+11, phTG6 −323/+11, phTG7 −175/+11, phTG7-4 −60/+11) were provided by Dr. S. J. Kim (National Cancer Institute, MD) [19], [41]. The HCV nonstructural (NS) protein NS3 expression plasmids pFlag-NS3, pFlag-NS3/4A, were provided by Dr. M. Gale (University of Washington, WA) [41]. The wild-type HCV NS5A expression vector was obtained from Dr. A. Siddiqui (University of California San Diego, CA). The dominant negative c-Jun plasmid (pcDNA3.1/His-TAM67) as well as (AP-1)4-luciferase-reporter construct was obtained from Dr. Nancy Colburn (National Cancer Institute, MD). The dominant negative mutants of STAT-3 (pSG5hSTAT3-β; a mutation at Tyr 705 of STAT-3, and STAT3-S727A, a mutation at Ser 727 of STAT-3), and the STAT-3 reporter plasmid pLucTKS3 (driven by the thymidine kinase promoter with seven copies of upstream STAT-3 binding sites from the human C-reactive protein gene) was obtained from Dr. Richard Jove (Moffitt Cancer Center, FL). The plasmid bearing the IκBα S32A/S36A (Ser to Ala) mutated gene under control of the CMV promoter was obtained from Dr. Robert Scheinman (University of Colorado Health Sciences Center, CO). The plasmid p3x-κB-Luc (luciferase reporter driven by the minimal fos promoter with three upstream NF-κB binding sites from MHC class I) was a generous gift from Dr. J. Martin (University of Colorado at Boulder, CO). All the primary antibodies were used according to the manufacturer’s protocol: HCV NS3 (Virogen, MA), α-SMA (Abcam, MA), GAPDH, STAT-3, IκBα, c-jun, c-fos, Sp1 and TGF-β1 (Cell Signaling, MA), furin and TSP-1 (Santa Cruz Biotechnology, CA).
HCV Cell Culture Infection System
The plasmid pFL-J6/JFH1 encoding the HCV J6/JFH-1 genome was linearized with XbaI for in vitro transcription using the Ampliscribe T7 transcription kit (Epicentre Technologies, WI). Fifteen micrograms of J6/JFH-1 RNA was delivered into Huh-7.5 cells by electroporation as described previously [6], . Cells were passaged every 3–5 days; the presence of HCV in these cells and the corresponding supernatants were determined as described previously [6], [29]. The cell-free virus was propagated in Huh-7.5 cell cultures, as described previously [4]. The expression of HCV protein in HCV-infected cells was analyzed using western blot assays. The viral titer in cell culture supernatant was expressed as focus forming unit (ffu) ml−1, which was determined by the average number of HCV-NS5A-postitive foci detected at the highest dilutions as described previously [6]. The HCV positive cell culture supernatant was used to infect naive Huh-7.5 cells at appropriate dilutions (moi of 1) for 5–6 h at 37°C and 5% CO2 [12], [29]. Cells were then plated in complete DMEM for 5–7 days. In most of the experiments, HCV-infected cells at day 6–7 were used. The cell culture supernatant collected from Huh-7.5 cells expressing JFH-1/GND (replication defective virus) was used as a negative control.
Western Blotting
Cells were harvested and cellular lysates were prepared by incubating in radioimmune precipitation (RIPA) buffer (50 mM Tris-HCl, pH 7.5, 150 mM NaCl, 1% NP-40, 0.5% sodium deoxycholate, 0.1% SDS, 1 mM sodium orthovanadate, 1 mM sodium formate, and 10 µl/ml protease inhibitor cocktail (Thermo Scientific, IL) for 30 min on ice. Cellular lysates were subjected to SDS-PAGE. Gels were electroblotted on to nitrocellulose membrane (Thermo Scientific, IL) in 25 mM Tris, 192 mM glycine and 20% methanol. Membranes were incubated for 1 h in blocking buffer [20 mM Tris-HCl, pH 7.5, 150 mM NaCl, 0.5% Tween-20, 5% dry milk], probed with primary antibody for 1 h at room temperature (RT) and washed twice for 10 min with blocking buffer without milk followed by incubation with secondary antibody for 1 h at RT. After an additional washing step with blocking buffer, immunoblots were visualized using the Odyssey Infrared Imaging System (LI-COR Biosciences, NE).
Preparation of Nuclear Lysates
Mock and HCV-infected cells were washed with ice-cold PBS and lysed in hypotonic buffer (20 mM HEPES, pH 7.9, 10 mM KCl, 0.1 mM Na3VO4, 1 mM EDTA, 10% glycerol, 1 mM PMSF, 3 mg/ml aprotinin, 1 mg/ml pepstatin, 20 mM NaF and 1 mM DTT with 0.2% NP-40) on ice for 15 min followed by centrifugation at 4°C for 1 min. The nuclear pellet was washed one time with ice-cold PBS and resuspended in high salt buffer (hypotonic buffer with 20% glycerol and 420 mM NaCl) at 4°C by rocking for 30 min. After centrifugation for 5 min the clarified nuclear lysates were used for electrophoretic mobility shift assay.
Electrophoretic Mobility Shift Assay (EMSA)
EMSA were performed using the Odyssey infrared EMSA kit according to the manufacturer’s protocols (LI-COR Biosciences). The duplex oligonucleotides containing the putative AP-1 and Sp1 binding sites on human TGF-β1 promoter (AP-1; 5′-CTTGTTTCCCAGCCTGACTCTCCTTCCGTTCT-3′; Sp1; 5′-AGCCGGGGAGCCCGCCCCCTTTCCCCCAGGG-3′ the AP-1 and Sp1 binding sites are italicized) were labeled at 5′-end with IRDye 700 infrared dye (Integrated DNA Technologies). For mobility shift assay, equal amounts of nuclear lysates (10 µg) were incubated with 50 nM of IRDye 700 labeled probes. The competition reactions were performed with a 200-fold molar excess of unlabeled consensus probe prior to addition of labeled probe. The supershift was performed by incubation of nuclear lysate and probe complex with antibody for 20 min. The DNA-protein complexes were resolved by 5% polyacrylamide gel electrophoresis in 0.5 X TBE buffer. The gels were visualized using a LI-COR Odyssey imaging system.
Chromatin Immunoprecipitation (ChIP) Assay
The ChIP assay was performed using SimpleChIP Enzymatic Chromatin IP Kit (Cell Signaling, Cat#9003). Briefly, mock- and HCV-infected cells (5×107 cells) were fixed in 1% formaldehyde for 10 min to crosslink the DNA and the DNA-associated-proteins. The reaction was quenched using 125 nM glycine for 5 min. The cell pellet was washed two times with ice-cold PBS and suspended in ice-cold buffer A containing DTT, PMSF and protease inhibitor cocktail. The nuclei were pelleted by centrifugation at 3,000 rpm for 5 min at 4°C. The supernants were removed and the pellet was suspended in ice-cold buffer B+ DTT. The lysate was incubated with 5 µl micrococcal nuclease for 20 min at 37°C to digest DNA to length approximately to 150–900 bp. The samples were suspended in 1X ChIP buffer and sonicated for 30 seconds using Qsonica Q700. The sonicated lysate was centrifuged at 10,000 rpm for 10 min at 4°C to remove debris. The supernatant (cross-linked chromatin preparation) were incubated with anti-c-Jun (1∶50), c-Fos (1∶50) and Sp1 (1∶100) antibody or a normal rabbit IgG followed by an isolation procedure using Protein G magnetic beads. The DNA-protein interaction was reversed by heating to 65°C for 12 h. The AP-1 and Sp1 binding sites on the immunoprecipitated DNA was determined by quantitative RT-PCR using SYBR green dye and AP-1 and Sp1 primers (Table 1). The PCR products were visualized on 1% agarose gel stained with 0.5 µg/ml ethidium bromide.
Lipid Droplet Staining
Mock- and HCV-infected cells on glass cover slips were fixed with 4% paraformaldehyde and permeabilized as described above. The cells were incubated with a fluorescent dye; BODIPY 493/503 (4,4-difluoro-3a,4a-diaza-s-indacene) (Invitrogen, CA) for lipid droplet staining. After washing with PBS, cells were mounted with antifade reagent containing DAPI (4, 6-diamidino-2 phenylindole) (Invitrogen, CA) and observed under a laser scanning confocal microscope (Zeiss LSM 510).
RNA Interference
Mock- and HCV-infected cells were transfected with TGF-β1 siRNA (siTGF-β1), sifurin, siTSP-1 and siGFP according to the manufacturer’s protocol (Santa Cruz Biotechnology, CA). Each siRNA consists of pools of three to five target-specific 19–25 nt siRNA designed to knockdown the target gene expression. For each transfection, two solutions were prepared. Solution A: 60 pmol of siRNA duplex was mixed with 100 µl siRNA transfection medium. Solution B: 6 µl of transfection reagent was added to 100 µl siRNA transfection medium. Solutions A and B were allowed to incubate at RT for 20 min. Solutions A and B were combined, and allowed to incubate another 20 min at RT. The combined solutions were then added to the cells in six well plates, and then incubated for 5 h at 37°C and 5% CO2, and the transfection solution was replaced with complete DMEM growth media.
Luciferase Assay
Mock- and HCV-infected Huh-7.5 cells were transfected with wild-type and mutant TGF-β1 promoter-luciferase constructs. At 36 h post transfection, cells were serum starved for 4–5 h. Cells were harvested and cellular lysates were analyzed for luciferase activity using the dual-luciferase reporter assay kit (Promega, WI). All transfections included a renilla expression vector to serve as an internal control. In all the experiments the data represent luciferase activity relative to mock cells.
Inhibitor Treatments
The cells were serum starved for 4 h and treated with inhibitors against p38 MAPK (SB203580, 10 µM), JNK (SP600125, 30 µM), PI3K (LY294002, 50 µM), Src (SU6656, 10 µM), JAK 2/3 (AG490, 100 µM), and MEK1/2 (UO126, 20 µM) (Calbiochem, MA) at indicated concentrations for 12 h. The cells were also treated with inhibitors against transcription factors AP-1 (Tanshinone IIA, 80 µM) (Enzo, NY), phosphorylation of IkBα/NF-κB pathway (Bay 11-7085, 80 µM) (Calbiochem, MA), NF-κB (80 µM) (Cat# 481406; Calbiochem, MA) and SP1 (Mithramycin A, 100 µM) (Sigma, MO).
TGF-β1 ELISA
The cell culture supernatant from mock- and HCV-infected cells were harvested, and centrifuged at 1000 rpm for 10 min to remove cell debris. After centrifugation, the conditioned medium (CM) were collected. The secreted TGF-β1 protein in CM was determined by ELISA according to the manufacturer’s protocol (Promega, WI). A standard curve was constructed by serial dilutions of human recombinant TGF-β1. TGF-β1 levels were measured in triplicate determinations.
Detection of Bioactive TGF-β1 Using Mink Lung Epithelial Cell Luciferase Assay
Mink lung epithelial cells (MLECs) containing bioactive TGF-β1 sensitive plasminogen activator inhibitor promoter-luciferase construct (PAI/L) was a kind gift from Dr. D. B. Rifkin and were assayed as previously described [12], [42], [43]. The assay is based on the ability of bioactive TGF-β1 to bind to MLEC receptors. This results in a dose-dependent increase in luciferase activity. Briefly, MLEC were plated in 96 well plates at a concentration of 2.5×105 cells per well in regular DMEM and incubated for 24 h at 37°C. Next, cells were incubated with CM from HCV-infected cells for 24–48 h. Cells were then washed twice with PBS, and lysed with 50 µl of reporter lysis buffer (Promega, WI). Twenty microliter of cell extract and 90 µl of luciferase assay reagent were added to 96 well white opaque flat bottom plate and light emission is measured for 10 s in a Bio-TEK Synergy HT Multi-Detection microplate reader. TGF-β1 standards were prepared by adding 2 µl human recombinant TGF-β1 to 500 µl of 0.2% FBS DMEM into a polypropylene tube. The standard stock solution is then serially diluted to obtain standards from 1000–125 pg/ml.
Quantitative Real-time RT-PCR
Total RNA was extracted from mock- and HCV-infected cells using TRIzol (Invitrogen, CA). HCV RNA was quantified by real-time RT-PCR using an ABI PRISM 7500 Sequence Detector (Applied Biosystems, CA). Amplifications were conducted in triplicate using HCV specific primers and 6-carboxyfluorescein (6FAM)- and tetrachloro-6-carboxyfluorescein (TAMRA)-labeled probes (Applied Biosystems, CA). The sequences for the primers and probes were designed using Primer Express software (Applied Biosystems, CA) (Table 1). Amplification reactions were performed in a 25 µl mix using RT-PCR core reagents kit and the template RNA. Reactions were performed in a 96-well spectrofluorometric thermal cycler under the following conditions: 2 min at 50°C, 30 min at 60°C, 10 min at 95°C, 44 cycles of 20 s at 95°C and 1 min at 62°C. Fluorescence was monitored during every PCR cycle at the annealing step. At the termination of each PCR run, the data was analyzed by the automated system and amplification plots were generated. To determine the HCV RNA copy number, standards ranging from 101 to 108 copies/µg were used for comparison.
SYBR Green RT-PCR
The expression of cellular genes in mock- and HCV-infected cells were quantified by real-time RT-PCR using their respective primers (Table 1). Total cellular RNA was extracted using TRIzol and treated with DNase using RQ1 RNase-free DNase prior to cDNA production. The cDNA was reverse-transcribed from 1 µg of total RNA using oligo(dT) primers according to the manufacturer’s protocol (Applied Biosystems, CA). Quantitative RT-PCR was carried out using SYBR green master mix and specific primer sets. Amplification reactions were performed under the following conditions: 2 min at 50°C, 10 min at 95°C, 40 cycles for 15 s at 95°C, and 1 min at 60°C. Relative transcript levels were calculated using ΔΔCt method as specified by the manufacturer.
Site-directed Mutagenesis
The base substitution mutations of AP-1 and Sp1 binding sites on TGF-β1 promoter-luciferase reporter constructs were carried out by oligonucleotide-mediated mutagenesis as described previously [44]. Site-directed mutagenesis was performed using AP-1 and Sp1 primers (Table 1). The PCR reactions were performed with wild-type TGF-β1 promoter-luciferase construct and AP-1 and Sp1 mutagenesis primers according to the manufacturer’s protocol (Stratagene, CA). Briefly, reaction buffer (5 µL), dsDNA template (30 ng), oligonucleotide primers (125 ng each), dNTP mix (1 µL), 1 µL PfuTurbo DNA polymerase (2.5 U/µL), and 35 µL ddH20 were added to a final volume of 50 µL. PCR amplification reactions were performed under the following conditions: Segment 1, 1 cycle at 95°C for 30 s; Segment 2, 16 cycles at 95°C for 30 s; 55°C for 1 minute, 68°C for 8 min. At the end of reaction, samples were digested with DpnI and transformed into DH5α competent cells (Invitrogen, CA). Clones were tested by restriction digestion and the base substitution mutations were confirmed by DNA sequencing.
CytoTox-ONE™ Homogeneous Membrane Integrity Assay
Mock- and HCV-infected cells in 96-well plates were treated with various inhibitors. The assay plate was equilibrated to RT for 20–30 min. CytoTox-ONE reagent (Promega, WI) was added into the wells and incubated at RT while shaking for 30 seconds. Plate was next incubated at RT for 10 min without shaking. 50 µl stop solution was added to each well in the same order as CytoTox-ONE reagent. Plate was again incubated at RT while shaking for 10 seconds. Fluorescence was recorded with an excitation wavelength of 560 nm and emission wavelength of 590 nm. Cytotoxicity was calculated by subtracting the average fluorescence values of the culture medium background from all fluorescence values of the experimental wells. The average fluorescence values from kinase inhibitor treated cells were used to calculate the percent cytotoxicity for a given experimental treatment. Percent cytotoxicity = 100×(experimental - culture medium background)/(maximum LDH release – culture medium background). The cytotoxicity levels were measured in triplicate.
CytoSelect™ 24-well Cell Invasion Assay
Conditioned medium from mock- and HCV-infected cells transfected with siTGF-β1, and siGFP were collected. LX-2 cells were plated at a concentration of 1×106 cells/ml in serum free DMEM for 48 h in the upper chamber. Five hundred micro liters CM from mock- and HCV-infected cells transfected with siTGF-β1, and siGFP were placed into the lower chamber to stimulate cell invasion. Following incubation for 48 h at 37°C in 5% CO2 atmosphere, media was carefully aspirated from the insert. Cotton-tipped swabs were used to gently swab the interior of the inserts to remove non-invasive cells. Next, inserts were transferred to a clean well containing 400 µL of cell stain solution and incubated for 10 min at RT. Inserts were then washed several times in ddH20 and allowed to dry. Next, inserts were transferred to an empty well and 200 µL of extraction solution was added to the lower chamber of each well and incubated at RT for 10 min on an orbital shaker. One hundred micro-molar of each sample was transferred to a 96-well microtiter plate and the absorbance was recorded at 560 nm.
Results
HCV Activates TGF-β1 Promoter
In our previous studies we have shown the induction and maturation of TGF-β1 by HCV infection [12]. In this study, we sought to investigate the molecular mechanism(s) of TGF-β1 promoter activation leading to the secretion of bioactive TGF-β1, activation and invasion of human HSCs. To initiate this study, we have incubated human hepatoma cell line Huh-7.5 cells with HCV cell culture supernatant as described previously [12], [28], [29]. Mock- and HCV-infected cells were transiently-transfected with wild-type (phTG1 −1362/+11) and various deletion mutants of TGF-β1 promoter-luciferase reporter constructs (phTG5 −453/+11, phTG6 −323/+11, phTG7 −175/+11, phTG7-4 −60/+11) (Fig. 1A). We observed approximately 6 fold and 3 fold increase in luciferase activity by phTG5 and phTG1 respectively in HCV-infected cells compared to mock-infected cells (Fig. 1B). However, we did not observe TGF-β1 promoter-luciferase activation in cells transfected with deletion mutants (phTG7, and phTG7-4). These results suggest that the region between −1362 to −323 is responsible for the TGF-β1 promoter-luciferase activation in HCV-infected cells.
A) Wild-type TGF-β1 promoter luciferase reporter (phTG1) and various deletion constructs (phTG5, phTG6, phTG7, and phTG7-4) are shown. Two negative regulatory sequences (NRS) are shown in the region between −453 and −1362 as well as an USF1/2 binding region. Two AP-1 binding sites are located between −453 and −323. Two Sp1 binding sites are located between −323 and −175. Three Sp1 sites and two Egr-1 sites are located between −175 and −60. B) Mock- and HCV-infected cells were transfected with TGF-β1 promoter-luciferase reporter constructs (phTG1, TG5, TG6, TG7, TG7-4) or vector control (500 ng) using Lipofectamine 2000 according to the manufacturer’s instructions (Invitrogen, CA). At 36 h post transfection cellular lysates were subjected to dual-luciferase reporter assay. The values represent the means standard deviations of three independent experiments performed in triplicate. * denotes p<0.05 compared to mock cells. C) Mock cells were transfected with TGF-β1 promoter-luciferase reporter constructs (phTG1, TG5, TG6, TG7, TG7-4) or vector control (500 ng) along with plasmids expressing HCV NS3, NS3/4A, or NS5A (500 ng), as described above. At 36 h post transfection cellular lysates were subjected to dual-luciferase reporter assay. The values represent the mean standard deviations of three independent experiments performed in triplicate. * denotes p<0.05 compared to mock cells. The data represent luciferase activity relative to mock cells.
Previously, we have shown that HCV nonstructural (NS) proteins (NS3, NS3/4A, and NS5A) were able to induce TGF-β1 activation and secretion [12]. To demonstrate the effect of HCV NS3, NS3/4A, and NS5A on TGF-β1 promoter activation, Huh-7.5 cells were cotransfected with TGF-β1 promoter-luciferase reporter constructs along with HCV NS3, NS3/4A, and NS5A expression vectors. The results show increased luciferase activity of phTG5 and phTG1 by NS3, NS3/4A, and NS5A (Fig. 1C). However, deletion mutant phTG6 showed modest activity. In contrast, deletion mutants, phTG7 and phTG7-4 did not show any activity by HCV NS proteins (Fig. 1C).
Role of HCV-induced Transcription Factors on TGF-β1 Promoter Activation
Previously, several transcription factors such as EGR-1, USF, ZF9/core promoter biding protein, AP-1, Sp1, NF-kB, and STAT-3 have been shown to bind to TGF-β1 promoter [10], [17], [18], [20], [21], [22], [23], [25], [27]. Since we observed increased luciferase activity of wild-type (phTG1) and deletion mutant (phTG5) in HCV-infected cells, we have used these luciferase constructs in our further studies. To determine if HCV-induced transcription factors activate TGF-β1 promoter, mock- and HCV-infected cells were transfected with phTG1 and phTG5 promoter-luciferase reporters followed by treatment with the inhibitors of AP-1 (Tanshinone IIA), Sp1 (Mithramycin), IkBα phosphorylation (Bay 11-7085), NF-κB, and cotransfected with the dominant negative (dn) mutants of NF-kB and STAT-3. The results show increased luciferase activity of phTG1 and phTG5, which was reduced upon treatment with AP-1 and Sp1 inhibitors (Fig. 2A). In contrast, we did not observe any effect of the inhibitors of IkBα and NF-kB as well as dn mutants of IkBα and STAT-3 (Fig. 2A). This is not unexpected as phTG1 and phTG5 do not contain the binding sites for NF-kB and STAT-3 (Fig. 1A).
A) Mock- and HCV-infected cells were transfected with 500 ng of wild-type (phTG1 −1362/+11), deletion mutant (phTG5 −453/+11) TGF-β1 promoter-luciferase reporter constructs and vector control. At 36 h posttransfection, cells were incubated with inhibitors of AP-1 (Bar#1; Tanshinone IIA, 80 µM), IkB kinase (Bar#4 and 5; Bay 11-7085, 80 µM and NF-κB activation inhibitor 80 µM), and Sp1 (Bar# 6; Mithramycin A, 100 µM) as described in Materials and Methods. Mock- and HCV-infected cells were also transfected with 500 ng of TGF-β1 promoter-luciferase reporter and 500 ng of the dominant negative mutants of STAT-3 (Bar#7; pSG5hSTAT3-β; mutation of STAT-3 tyrosine phosphorylation site 705 into phenylalanine; Bar#8; STAT3-S727A; mutation of STAT-3 serine phosphorylation site 727 into alanine) and the inhibitory subunit of NF-κB (Bar#9; IκBα S32A/S36A; mutation of IkBα serine 32,36 into alanine). Cellular lysates were subjected to dual-luciferase reporter assay. The values represent the means standard deviations of three independent experiments performed in triplicate. * denotes p<0.05 compared to mock cells. ** denotes p<0.05 compared to untreated promoter. B) Mock- and HCV-infected cells were treated with the inhibitors and dn mutants as described above. Total cellular RNA was extracted and TGF-β1 mRNA was analyzed using TGF-β1 specific primers and SYBR green probe. The values represent the means standard deviations of three independent experiments performed in triplicate. * denotes p<0.05 compared to mock cells. ** denotes p<0.05 compared to untreated promoter. # denotes p<0.05 compared to vector control. C) Mock- and HCV-infected cells were treated with inhibitors or transfected with dominant negative mutants as described above. Samples were tested for cytotoxicity using CytoToxONE homogenous membrane integrity assay as described in Materials and Methods. The data represent luciferase activity relative to mock cells. D, E, and F) HCV-infected cells were transfected with vectors expressing dominant negative proteins of IkBα (IkBα S32A/S36A), STAT-3 (pSG5hSTAT3-β and STAT3-S727A), and AP-1 (pcDNA3.1/His-TAM67). At 48 h posttransfection, cellular lysates were prepared and subjected to western blot analysis using anti-IkBα, STAT-3, and c-Jun antibodies. Actin was used as a protein loading control.
To examine if HCV-induced NF-kB and STAT-3 have any effects on endogenous TGF-β1 promoter activation, mock- and HCV-infected cells were incubated with the inhibitors and dn mutants as described in figure 2A. Total cellular RNA was harvested and subjected to quantitative RT-PCR. We observed a 20 fold increased TGF-β1 mRNA expression, which was reduced in cells treated with the inhibitors of AP-1, Sp1, IkBα, NF-κB, and dn mutants of IkBα and STAT-3 (Fig. 2B). These results suggest that endogenous TGF-β1 promoter is regulated by HCV-induced AP-1, Sp1, NF-kB, and STAT-3. The cellular toxicity assay was performed by CytoTox-One cytotoxicity assay. Untreated cells showed approximately 2.5–3.5% cytotoxicity, whereas the positive lysis control cells showed approximately 7–9% cytotoxicity (Fig. 2C). Mock- or HCV-infected cells treated with the inhibitors did not cause significant cytotoxicity (Fig. 2C). The expression of IkBα, STAT-3 and AP-1 dominant negative proteins in HCV-infected cells are shown by western blot assay (Fig. 2D–2F).
Role of AP-1 and Sp1 Binding Sites on HCV-induced TGF-β1 Promoter Activation
To demonstrate the role of AP-1 binding site (–432/−398) and Sp1 binding site (–229/−198) on HCV-induced TGF-β1 promoter activation, we mutated the binding sites of AP-1 and Sp1 in phTG1 and phTG5 using site-directed mutagenesis (Fig. 3A). The mutations were confirmed by DNA sequence analysis from the Genomics Core Facility at Northwestern University, IL. Mock- and HCV-infected cells were transfected with wild-type and mutated reporter constructs. The results showed increased luciferase activity of wild-type phTG1 and phTG5, however, a mutation in AP-1 and Sp1 binding sites individually resulted in a decrease in HCV-mediated TGF-β1 promoter-luciferase activity (Fig. 3B). To determine if these effects were synergistic, we introduced double mutations in TGF-β1 promoter constructs. Our results showed a significant decrease in TGF-β1 promoter activity in HCV infected cells that were transfected with TGF-β1 promoter-luciferase constructs containing mutations in AP-1 and Sp1 binding sites (Fig. 3B), suggesting the synergistic effect of AP-1 and Sp1 on TGF-β1 promoter-luciferase reprter.
A) AP-1 and Sp1 binding sites on TGF-β1 promoter sequences were deduced from sequence data as described previously [19]. The pGL3 vector includes the wild-type TGF-β1 fragment –1362/+11 fused to the luciferase gene and was used as a template for the site-directed mutagenesis as described in Materials and Methods. The AP-1 and Sp1 binding site were mutated by a change of two bases in the consensus sequence as indicated. Substituted bases are italicized and highlighted with a box. The wild-type and mutated plasmids used for the transfection experiments are shown. B) Mock- and HCV-infected cells were transfected with 500 ng of the wild-type phTG1 and phTG5 and mutant TGF-β1 luciferase constructs (Mut-1, Mut-2, Mut-3). At 36 h posttransfection cellular lysates were harvested and subjected to dual-luciferase reporter assay. The values represent the means standard deviations of three independent experiments performed in triplicate. * denotes p<0.05 compared to mock cells. ** denotes p<0.05 compared to wild type phTG1 and phTG5 constructs. C) Cross-linked chromatin preparation from mock and HCV-infected cells were immunoprecipitated with anti-c-Jun, c-Fos and Sp1 antibody or a normal rabbit IgG. The AP-1 and Sp1 binding sites on the immunoprecipitated DNA was determined by quantitative RT-PCR using the primers and SYBR green probe. Amplification of input chromatin (input) prior to immunoprecipitation were served as positive controls for chromatin extraction and PCR amplification. Chromatin immunoprecipitation using a non-specific antibody (normal human IgG) served as negative controls. The data represent luciferase activity relative to mock cells. The values represent the means standard deviations of three independent experiments performed in triplicate. * denotes p<0.05 compared to mock cells. D) Five microliters of the PCR products were visualized on 1% agarose gel stained with 0.5 µg/ml ethidium bromide.
To determine whether AP-1 and Sp1 interact with the TGF-β1 promoter in vivo in HCV-infected cells, ChIP assay was performed using c-Jun, c-Fos, and Sp1 antibody. The DNA qRT-PCR analysis showed that c-Jun, c-Fos and Sp1 specific antibodies immunoprecipitated chromatin from HCV-infected cells (Fig. 3C). However, immunoprecipitation with non-specific antibody (normal human IgG) did not amplify the DNA fragments. The PCR amplification of input chromatin before immunoprecipitation was served as positive control. The amplified DNA fragments were further confirmed by agarose gel electrophoresis (Fig. 3D). These results indicate that AP-1 and Sp1 form a protein-DNA transcriptional regulatory complex by binding to the TGF-β1 promoter in HCV-infected cells.
Effect of HCV-induced Signaling Pathways on Transcription Factor Activation
To determine the role of HCV-induced Ca2+ signaling and induction of reactive oxygen species (ROS) on the activation of HCV-induced transcription factors, mock- and HCV-infected cells were transfected with STAT-3, NF-κB, and AP-1 responsive luciferase reporter plasmids. Our data show increased activity of STAT-3, NF-κB, and AP-1 responsive luciferase reporters which were decreased when treated with intracellular Ca2+ chelator (BAPTA-AM) or antioxidant (PDTC; pyrrolidine dithiocarbamate) (Fig. 4A).
A) Mock- and HCV-infected cells were transfected with 500 ng of pLucTKS3, p3x-κB-Luc, or (AP-1)4-Luc reporter constructs. At 36 h posttransfection cells were treated with intracellular calcium chelator (BAPTA-AM, 50 µM) or antioxidant (PDTC, 100 µM). Cellular lysates were harvested and subjected to dual-luciferase reporter assay. The values represent the means standard deviations of three independent experiments performed in triplicate. *denotes p<0.05 compared to mock cells. **denotes p<0.05 compared to HCV-infected mock-treated with inhibitors. The data represent luciferase activity relative to mock cells. B and C) EMSA was carried out in the presence of IRDye 700-labeled oligonucleotides derived from TGF-β1 promoter (contains AP-1 and Sp1 binding sites) and nuclear lysates from mock (Huh7.5) and HCV-infected cells. Lane 1, probe alone; lanes 2 and 3, equal amounts of nuclear lysates from mock and HCV-infected cells; lane 4, same as lane 3 but incubated with 200-fold excess of unlabeled oligonucleotides; lane 5, DNA-protein complex incubated with anti-c-Jun or anti-Sp1 for 20 min at RT.
To determine the binding of HCV-induced AP-1 and Sp1 with oligonucleotide derived from TGF-β1 promoter, we performed the EMSA of c-Jun and Sp1 with labeled probe. Our results showed the increased DNA-protein complex formation in HCV-infected nuclear lysates (Fig. 4B and 4C). The specificity of DNA-protein complexes were confirmed by competition with 200-fold molar excess of unlabeled consensus probe and a supershift of DNA-protein complex in the presence of anti-c-Jun and anti-Sp1 antibodies.
Role of HCV-induced Cellular Kinases on TGF-β1 Promoter Activation
Previously, we and others have shown that the activation of transcription factors are regulated by cellular kinases in HCV expressing cells [10], [29], [31], [45]. To demonstrate the role of HCV-induced cellular kinases in TGF-β1 promoter activation, mock- and HCV-infected cells were transiently-transfected with phTG1 and phTG5 promoter-luciferase reporters followed by treatment with the inhibitors of p38 MAPK (SB203580), JNK (SP600125), Src (SU6656), PI3K (LY294002), JAK (AG490), and MEK1/2 (U0126). The results show increased activity of phTG1 and phTG5 in HCV-infected cells, which was abrogated in HCV-infected cells treated with inhibitors of p38 MAPK, JNK, Src, and MEK1/2, but not with PI3K and JAK (Fig. 5A).
A) Mock- and HCV-infected cells were transfected with 500 ng of phTG1 and phTG5 TGF-β1 promoter-luciferase reporter. At 36 h posttransfection cells were serum starved for 4 h and treated with inhibitors against p38 MAPK (SB203580, 10 µM), JNK (SP600125, 30 µM), PI3K (LY294002, 50 µM), Src (SU6656, 10 µM), JAK 2/3 (AG490, 100 µM), and MEK1/2 (UO126, 20 µM) for 12 h. Cellular lysates were subjected to dual-luciferase reporter assay. The values represent the means standard deviations of three independent experiments performed in triplicate. *denotes p<0.05 compared to mock cells. **denotes p<0.05 compared to HCV-infected mock-treated cells. B) Mock- and HCV-infected cells were treated with kinase inhibitors as described in Materials and Methods. Total cellular RNA was extracted and TGF-β1 mRNA was analyzed using TGF-β1 specific primers and SYBR green fluorescent dye. The values represent the means standard deviations of three independent experiments performed in triplicate. *denotes p<0.05 compared to mock cells. **denotes p<0.05 compared to HCV-infected mock-treated cells. C) Mock- and HCV-infected cells were treated with kinase inhibitors as described in Materials and Methods. Cells were subjected to growth inhibition assay using CytoToxONE homogenous membrane integrity assay as described in Materials and Methods.
To demonstrate the effect of these kinases on endogenous TGF-β1 gene expression, mock- and HCV-infected cells were incubated the kinase inhibitors as described above. Total cellular RNA was subjected to quantitative RT-PCR. We observed increased expression of TGF-β1 mRNA in HCV-infected cells, which was abrogated in the presence of the inhibitors of p38 MAPK, JNK, Src, and MEK1/2 (Fig. 5B). To determine the level of toxicity caused by the kinase inhibitors in the HCV-infected cells, CytoTox-One cytotoxicity assay was performed. We did not observe any cytotoxicity in cells treated with above kinase inhibitors (Fig. 5C).
Effect of HCV-induced Transcription Factors on TGF-β1 Secretion
To determine the role of HCV-induced AP-1, Sp1, NF-κB, and STAT-3 on TGF-β1 secretion, mock- and HCV-infected cells were incubated with the inhibitors of AP-1 (Tanshinone IIA), Sp1 (Mithramycin), IκBα (Bay 11-7085), and NF-κB, or transfected with the dn mutants of AP-1 (TAM67), STAT-3 (STAT-3β, STAT-3 S727A), and IkBα (IkBα S32, 36 to alanine 32, 36) as described in figure 2. Conditioned media (CM) were collected from these cells and subjected to TGF-β1 ELISA. We observed approximately 1250 pg/ml of TGF-β1 in CM collected from HCV-infected cells, which was significantly reduced by treatment with above inhibitors or transfected with dn mutants (Fig. 6A).
A) Mock- and HCV-infected cells were treated with inhibitors of AP-1, Sp1, IkB kinase or transfected with dn mutants of AP-1, STAT-3, or IκBα. At 36 h posttransfection, total secreted TGF-β1 in CM were determined by TGF-β1 specific ELISA. The values represent the means standard deviations of three independent experiments performed in triplicate. *denotes p<0.05 compared to mock cells. **denotes p<0.05 compared to HCV-infected mock-treated cells. B) CM as described above, were subjected to growth inhibition assay using mink lung epithelial cells as described in Materials and Methods. The data shown here represent the means standard deviations of three independent experiments performed in triplicate. *denotes p<0.05 compared to mock cells. **denotes p<0.05 compared to HCV-infected mock-treated cells.
The bioactive TGF-β1 in CM was quantified by a standard growth inhibition assay using mink lung epithelial cells (MLEC) as described previously [12], [42], [43]. In this assay, MLEC stably transfected with the PAI/L demonstrate a dose-dependent increase in luciferase activity which indirectly corresponds to growth inhibition. MLEC were incubated with CM from mock- and HCV-infected cells treated with above inhibitors or transfected with above dn mutants. MLEC cells were then lysed, and subsequent luciferase assay was performed. HCV-infected cells secreted approximately 2.6 fold more bioactive TGF-β1 compared to mock-infected cells. Increased secretion of bioactive TGF-β1 by HCV-infection was significantly reduced by treatment with above inhibitors or dn mutants (Fig. 6B).
Effect of HCV-induced TGF-β1, Furin, and TSP-1 on Hepatic Stellate Cells Activation
Hepatic stellate cells (HSCs) are the primary cell type involved in liver fibrosis [7]. To demonstrate the effect of secreted TGF-β1 from HCV-infected cells on HSCs, LX-2 cells were incubated with CM from mock- and HCV-infected cells as well as HCV-infected cells transfected with siGFP, siTGF-β1, siTSP-1, and sifurin. In our previous studies we have shown that furin and TSP-1 are involved in the proteolytic processing (maturation) of TGF-β1 [12]. To determine the knock down of TGF-β1, TSP-1, and furin by their siRNA, quantitative RT-PCR and western blot assay were performed. We observed reduced expression of TGF-β1, TSP-1, and furin mRNA and protein at 72 h posttransfection (Fig. 7A and 7B).
A) Mock- and HCV-infected cells were transfected with siGFP, siTGF-β1, sifurin, and siTSP-1 as described in Materials and Methods. To determine the levels of silencing, total cellular RNA was extracted, and subjected to quantitative RT-PCR using furin, TSP-1, and TGF-β1 specific primers. The data shown here represent the means standard deviations of three independent experiments performed in triplicate. *denotes p<0.05 compared to mock cells. **denotes p<0.05 compared to HCV-infected mock-transfected cells. B) Equal amounts of cellular lysates from above siRNA transfected cells were subjected to western blot analysis using anti-TGF-β1, furin, and TSP-1 antibodies. Lane 1, Lysates from mock (Huh7.5) cells; lane 2, HCV-infected cells; lanes 3 and 4, HCV-infected cells transfected with siGFP or siTGF-β1, sifurin and siTSP-1. Actin and albumin were used as protein loading controls. C) HSC line, LX-2 was grown and serum starved for 48 h prior to incubation with conditioned medium (CM) collected from HCV-infected Huh-7.5 cells transfected with siGFP, siTGF-β1, sifurin, or siTSP-1 for 24 h. Total cellular RNA was extracted from LX-2 cells and the levels of α-SMA and Col 1 alpha 1 mRNA were determined by quantitative RT-PCR. The data shown here represent the means standard deviations of three independent experiments performed in triplicate. *denotes p<0.05 compared to LX-2 cells incubated with CM from mock cells. **denotes p<0.05 compared to LX-2 cells incubated with CM from HCV-infected cells. D) Cellular lysates from cells as described above were subjected to immunoblot analysis using antibodies against α-SMA and GAPDH. Lanes 1 and 2, lysates from LX-2 cells incubated with CM from mock- and HCV-infected cells; Lanes 3–6, lysates from LX-2 cells incubated with CM from HCV-infected cells transfected with siGFP, siTGF-β1, sifurin, and siTSP-1. Bottom panel shows the protein loading control immunoblotted with anti-GAPDH primary antibody.
LX-2 cells were incubated with CM from HCV-infected cells. The results showed increased expression of LX-2 cells activation markers, α-smooth muscle actin (α-SMA) and collagen type 1 α 1 (Col1A1) mRNA, which was reduced in LX-2 cells incubated with CM collected from HCV-infected cells transfected with siTGF-β1, siTSP-1, or sifurin (Fig. 7C). To further demonstrate the activation of LX-2, western blot analysis of α-SMA was performed. The results show a significant increase in α-SMA expression following incubation with conditioned media from HCV-infected cells (Fig. 7D), which was reduced in LX-2 cells incubated with conditioned media from HCV-infected cells transfected with siTGF-β1, siTSP-1, or sifurin (Fig. 7D).
Effect of HCV-induced TGF-β1 on HSC Invasion
To evaluate the effect of TGF-β1 from HCV-infected cells on HSCs, LX-2 cells in serum-free DMEM were plated in the upper chamber of the CytoSelect Cell Invasion Assay. CM from HCV-infected cells transfected with siTGF-β1 or siGFP was used in the lower chamber to stimulate cell invasion (Fig. 8A). The results showed increased invasion of LX-2 cells when incubated with CM from HCV-infected cells, which was reduced in LX-2 cells incubated with CM collected from HCV-infected cells transfected with siTGF-β1 but not with siGFP (Fig. 8B). Using extraction solution, we also quantified the invading cells by recording the absorbance of the samples at 560 nm. The results show an increased invasion of LX-2 cells when incubated with CM from HCV-infected cells, which was reduced in LX-2 cells incubated with CM from HCV-infected cells transfected with siTGF-β1 but not with siGFP (Fig. 8C).
A) Schematic of invasion assay. B) LX-2 cells were plated in upper chamber in serum free DMEM. CM from mock- and HCV-infected cells as well as HCV-infected cells transfected with siTGF-β1, and siGFP were used in the lower chamber. Boyden chamber was incubated at 37°C for 48 h to examine HSCs invasion. Invasion was assessed by counting migrated cells in multiple microscopic fields per well at 35× magnification. C) The HSC invasion was also quantified using extraction solution and absorbance was recorded at 560 nm according to manufacturer’s protocol (Cell Biolabs, Inc., CA). The data shown here represent the means standard deviations of two independent experiments performed in duplicate. *denotes p<0.05 compared to LX-2s treated with CM from mock cells. **denotes p<0.05 compared to LX-2s treated with CM from HCV-infected cells.
Effect of TGF-β1, Furin, and TSP-1 on HCV Replication and Release
To evaluate the effect of TGF-β1, furin, and TSP-1 on HCV replication, and release, we used RNA interference approach as described in figure 7A. Total cellular RNA was extracted from cells as well as supernatant from mock and HCV-infected cells and subjected to quantitative RT-PCR analysis using HCV-specific primers and Taqman probe. We observed an increase in HCV replication in HCV-infected cells (Fig. 9A), which was significantly reduced in HCV-infected cells transfected with siTGF-β1, siTSP-1 or sifurin (Fig. 9A). However, transfection of siGFP (negative control) did not show any effect on HCV replication (Fig. 9A).
A) Total cellular RNA was collected at day 4 from mock, HCV-infected cells, and those transfected with siRNA, and the level of HCV RNA was determined by quantitative RT-PCR. The data shown here represent the means standard deviations of three independent experiments performed in triplicate. *denotes p<0.05 compared to mock cells. **denotes p<0.05 compared to HCV-infected mock-transfected cells. B) Two milliliters of CM were collected from above cells at day 7, and the level of HCV release was determined by quantitative RT-PCR as described in Materials and Methods. The data shown here represent the means standard deviations of three independent experiments performed in triplicate. *denotes p<0.05 compared to mock cells. **denotes p<0.05 compared to HCV-infected, mock-transfected cells. C) Mock, HCV-infected cells, as well as HCV-infected cells transfected with siTGF-β1, and siGFP were prepared for confocal laser-scanning microscopy as described in Materials and Methods. Briefly, cells were incubated with BODIPY for 1 h at RT, and washed with PBS. DAPI was used as a nuclear stain. Arrows indicate lipid droplets. Bar, 10 µM.
Similarly, we observed an increase in HCV RNA in the supernatant of HCV-infected cells, which was significantly reduced in HCV-infected cells transfected with siTGF-β1, siTSP-1, or sifurin (Fig. 9B) but not with siGFP (negative control) (Fig. 9B). These results suggest the role of HCV-induced TGF-β1, furin, and TSP-1 in HCV replication and release.
Previously, lipid droplets have been shown to play a critical role in HCV assembly and secretion [46], [47], [48], [49]. To demonstrate the effect of HCV-induced TGF-β1 on lipid droplet formation, cells were subjected to lipid droplet staining as described in Materials and Methods. The results showed no change in lipid droplet formation (Fig. 9C).
Discussion
Chronic HCV infection can lead to liver fibrosis, cirrhosis, and eventually hepatocellular carcinoma by various mechanisms. Although induction of profibrogenic molecules such as TGF-β1 has been shown to play an important role in the pathogenesis of HCV, little is understood about the mechanism of HCV-mediated liver fibrosis [10],[12].
Liver fibrosis is defined as the excessive accumulation of ECM proteins including multiple types of collagens, fibronectin, laminin, and other molecules that are associated with chronic liver diseases [50]. Accumulation of ECM proteins distorts the hepatic architecture by forming scar tissue and the subsequent development of nodules of regenerating hepatocytes defines the progression of fibrosis to cirrhosis [51]. HSCs are the primary source of ECM and activation of HSCs by various stimuli often leads to fibrosis [52]. The initial activation of HSCs is likely to be a result of stimuli produced by neighboring cells e.g. hepatocytes, or Kupffer cells, these stimuli include ROS, lipid peroxides, growth factors, and inflammatory cytokines [52].
TGF-β1 is the most potent fibrogenic stimulus to HSCs and elevated TGF-β1 expression has been implicated in the pathogenesis of various diseases including liver fibrosis, and HCC [53]. Previous studies related to HCV-mediated liver fibrosis have been conducted in HSCs. In the absence of inflammation, TGF-β1 is secreted from HSC and Kupffer cells, but not from hepatocytes. However, during liver injury and inflammation, hepatocytes can become a major source of TGF-β1 [10], [12], [54], [55], [56], [57]. Secreted bioactive TGF-β1 from hepatocytes can activate HSCs leading to the secretion of ECM proteins [43].
In the present study, we investigated the molecular mechanisms of TGF-β1 promoter activation in response to HCV, as well as the effect of secreted TGF-β1 on human HSCs activation and invasion. Using a series of TGF-β1 promoter-luciferase constructs, we demonstrate that the region between –323 and –453 is responsible for TGF-β1 promoter activation in response to HCV-infection (Fig. 1B). Previous studies have demonstrated two AP-1 binding sites between –323 and –453 [44]. In addition, our results show modest level of activity by phTG6 which contains known Sp1 binding sites. phTG1 showed decreased activity compared phTG5 because phTG1 is known to contain negative regulatory regions [19].
One of the effects of HCV translation/replication activities in the ER is the activation of cellular transcription factors [35]. Previously, HCV proteins (core, NS3, and NS5A) have been shown to induce various transcription factors (STAT-3, Sp1, NF-κB, and AP-1) through multiple signaling pathways [30], [31], [43], [50]. Our results showed a significant decrease in TGF-β1 promoter activation in HCV-infected cells treated with inhibitors of AP-1 and Sp1. However, we did not observe a reduction of TGF-β1 promoter activation when cells were treated with inhibitors of NF-κB, or transfected with dominant negative forms of NF-κB or STAT-3, as the TGF-β1 promoter phTG1 (–1362/+11) does not contain binding sites for NF-κB and STAT-3.
However, endogenous TGF-β1 gene expression was significantly reduced by NF-κB inhibitors and dominant negative forms of NF-κB and STAT-3 as well as inhibitors of AP-1 and Sp1. Previous studies have demonstrated a far upstream TGF-β1 promoter region at positions −3155 and −2515 upstream of the transcription initiation site [21]. This region contains STAT-3 binding site which is far upstream from TGF-β1 promoter-luciferase and would explain the discrepancy between TGF-β1 promoter-luciferase and endogenous TGF-β1 mRNA results. Similarly, NF-κB has been shown to be activated by HCV-infection and plays an important role in TGF-β1 promoter activation; however TGF-β1 promoter region −1362 to +11 does not contain any NF-κB binding sites. Therefore, it could be possible that NF-κB is either binding directly to a secondary promoter region upstream, or is indirectly regulating the TGF-β1 promoter region via interactions with other cellular proteins [10], [11], [45].
Previously, AP-1 and Sp1 transcription factors have been shown to play an important role in the induction of TGF-β1 in various systems [19], [22], [44], [58]. Transcriptional regulation of TGF-β1 by v-src gene products has been shown to be mediated through the AP-1 complex [58]. AP-1 proteins have been shown to mediate hyperglycemia-induced activation of TGF-β1 promoter in mesangial cells [44]. Sp1 is known to play an important role in HPV E6- and E7-mediated activation of the TGF-β1 promoter [22]. Our results are consistent with these previous studies.
AP-1, STAT-3, Sp1, and NF-κB are activated by upstream cellular kinases and belong to a category of rapid acting transcription factors. AP-1 and NF-κB are both complexes that have been shown to be phosphorylated and activated in response to HCV gene expression [31], [45]. STAT-3, although not a complex like AP-1 and NF-κB, has also been shown to be activated by HCV gene expression [30]. Sp1 has been shown to be activated by p38 MAPK but the mechanism has not been defined [59]. We and others have shown the activation of cellular kinases JNK, p38 MAPK, JAK2, ERK1/2, Src and PI3K/Akt signaling in HCV-infected cells [10], [29], [31], [34], [45], [60]. In this study, we observed that the activation of TGF-β1 promoter is mediated through the activation of cellular kinases such as JNK, p38 MAPK, Src, and ERK.
Human hepatic stellate cells are the primary cell type responsible for liver fibrosis following their activation into fibrogenic myofibroblast-like cells [7], [37]. In this study, the fibrogenic effect of TGF-β1 secreted from HCV-infected Huh-7.5 cells was studied by examining the status of the well known markers of HSCs activation, α-SMA and Col1A1. Our results showed a significant decrease of α-SMA and ColIAI mRNA expression and α-SMA protein expression in HSCs incubated with CM from HCV-infected cells transfected with siTGF-β1, siFurin, or siTSP-1. These data suggest that secreted bioactive TGF-β1 is regulated by host proteins furin and TSP-1, and bioactive TGF-β1 plays an important role in the activation of HSCs. Our data is in agreement with previously published work on TGF-β1 stimulation of HSCs and elucidates the role of secreted TGF-β1 from HCV-infected cells [61], [62], [63], [64]. Another hallmark of HSC activation is an invasive phenotype [64]. We observed an increase in LX-2 invasion when incubated with CM from HCV-infected cells, and a significant decrease of invasive phenotype with CM from HCV-infected cells transfected with siTGF-β1. This data suggests that TGF-β1 secreted from HCV-infected cells plays a critical role in invasive potential of HSCs.
Previous studies have shown that TGF-β1 increased replication of respiratory syncytial virus and JC virus [15], [65]. Our previous studies have demonstrated that siTGF-β1 decreased replication of HCV [12]. However, the underlying mechanism by which TGF-β1 enhances HCV replication is unknown. Previously, the stimulation as well as suppression of HCV replication by exogenous addition of TGF-β1 has been demonstrated in HCV replicon system [11], [66]. Endogenous TGF-β1 has been shown to induce intracellular signaling pathways including activation of hypoxia inducible factor-1 (HIF-1) and direct interaction of TGF-β1 with STAT-5 leading to liver fibrosis [17], [67].
Lipid droplets (LDs) are mainly involved in lipid storage but can also be involved in vesicular transport and cellular signaling [68]. Several clinical studies have demonstrated that chronic HCV infection is associated with enhanced accumulation of LDs in the liver [69], [70], [71]. Previous studies have shown that LDs have a critical role in the production of infectious HCV particles [46], [47], [48], [49]. Our data suggests that TGF-β1 is required for the release of infectious HCV particles without affecting LD biogenesis (Fig. 9C), suggesting that TGF-β1 may be regulating HCV release through LD-independent mechanisms.
In summary, we show TGF-β1 promoter activation by HCV-infection is dependent on transcription factors AP-1, Sp1, STAT-3, and NF-κB. Our results also show the activation of these transcription factors is dependent on the activation of cellular kinases. These studies provide greater insight into the molecular mechanisms of TGF-β1 promoter activation by HCV infection. Our results also demonstrate the role of secreted TGF-β1 from HCV-infected cells in the activation and invasion of HSCs suggesting invasive potential of activated HSCs. In addition, our results demonstrate the role of TGF-β1 in HCV replication and release. The results of these studies provide ideas for new concepts and a framework to develop novel strategies of treatment of chronic HCV infection associated with liver fibrosis.
Acknowledgments
We thank Dr. Takaji Wakita (NIID, Tokyo, Japan), and Dr. Charles Rice (Rockefeller University, NY) for the generous gift of HCV genotype 2a (JFH-1) and J6/JFH-1 infectious cDNA and Huh-7.5 cell line; Dr. S.J. Kim (NIH/HCI, MD); for TGF-β1 reporter constructs’ Dr. Daniel Rifkin (NYU, NY) for the mink lung epithelial cell line (Mv1Lu TGF-β1 sensitive PAI-1/L); Dr. Scott Friedman (Mount Sinai Hospital, NY) for the LX-2 cell line; Dr. Nancy Colburn (NCI, MD) for the dominant negative c-Jun plasmid (pcDNA3.1/His-TAM67) as well as (AP-1)4-luciferase-reporter construct.
Author Contributions
Conceived and designed the experiments: LP SM GW. Performed the experiments: LP SM. Analyzed the data: LP SM GW. Contributed reagents/materials/analysis tools: LP SM GW. Wrote the paper: LP GW.
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