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1 Departments of Biomolecular Science and 2 Pulmonary Medicine, Fukushima Medical University School of Medicine, Fukushima, 960-1295 Japan
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Introduction |
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The pathogenesis of IPF is complex, and the precise molecular mechanisms underlying the development of fibrosis are not fully elucidated (White et al. 2003). It has been believed that IPF represents a disease induced by persistent lung inflammation, resulting in the fibrotic response. However, accumulating observations suggest that foci of proliferating fibroblasts/myofibroblasts are more prominent than evidence of inflammation, as well as the poor efficacy of anti-inflammatory agents in patients with IPF, has led to a focus of the fibrotic pathway itself (Selman & Pardo 2003). Currently, it is considered that IPF results from cycles of epithelial injury, and the activation and proliferation of mesenchymal cells with the formation of fibroblastic foci, leading to the accumulation of extracellular matrix and abnormal wound repair (Selman & Pardo 2003). The extent of fibroblastic foci in lung biopsies is strongly associated with disease progression and a worsening prognosis, suggesting that myofibroblasts are the key effector in the pathogenesis of IPF (King et al. 2001). Thus, understanding the pathophysiological role of myofibroblasts, as well as mechanisms for their proliferation and persistence in lung tissue, should shed deeper insight into the basis of disease progression.
Recently, a new possibility for the mechanism of IPF was presented based on the analysis of oxidant and antioxidant expression in IPF (Kinnula et al. 2005). Reported observations include the up-regulation of biomarkers of oxidative stress (hydrogen peroxide, 8-isoprostane and NO) in the expired breath condensate (EBC) and bronchoalveolar lavage fluid (BALF) from patients with IPF (Montuschi et al. 1998; Kharitonov & Barnes 2001; Psathakis et al. 2006). Other studies reported the down-regulation of anitioxidants such as reduced glutathione in the lungs of affected patients (Behr et al. 1995). In addition, a number of antioxidant enzymes have been reported to be up-regulated in IPF (Lakari et al. 2000, 2001; Tiitto et al. 2003; Peltoniemi et al. 2004). These results suggest that oxidant–antioxidant imbalances may play a key role in the pathogenesis of IPF. This is further supported by the results of randomized clinical trials of treatments for IPF patients with N-acetylcysteine (NAC), a precursor of the major antioxidant glutathione. Although additional studies are needed for routine therapy, the data presented suggest that NAC, either alone or in combination with immunosuppressive therapy, may have beneficial effects in patients with IPF (Behr et al. 1997; Demedts et al. 2005; Walter et al. 2006). Thus, it is clear that the cellular redox state is dysregulated, and the resultant generation of reactive oxygen species (ROS) plays a significant role in the pathogenesis of IPF. However, the precise mechanisms of ROS metabolism, and their functions in the progression of IPF remain largely elusive.
In this study, we examined the gene expression profiles of myofibroblasts established from patients with IPF. This analysis reveals the selective up-regulation of enzymes responsible for the metabolism of lipid hydroperoxides. We further demonstrate that selenoprotein P, one of the most highly up-regulated proteins, regulates the cellular lipid redox state and maintains cell viability. Thus, selenoprotein P represents a novel effector in the pathogenesis of IPF, which may contribute to the persistence of myofibroblasts in an oxidative environment.
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Results |
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Two primary myofibroblast cell cultures (LF1, LF2) were isolated from individual patients with IPF (Supplementary Fig. S1). Microarray analysis of these cells was carried out to explore the pathophysiological function of myofibroblasts in IPF. We used Affymetrix HG Focus oligonucleotide microarrays comprising approximately 8700 human mRNAs. The data sets were compared with the profile of TIG7 cells, a human embryonic lung fibroblast cell line. Unsupervised hierarchical clustering revealed that the gene expression profiles of LF1 and LF2 cells are more similar to each other than to TIG7 cells (data not shown), implying that genes consistently up- or down-regulated in LF1 and LF2 cells may represent pathophysiological features of myofibroblasts in IPF. As expected, myofibroblast marker proteins were up-regulated in LF1 and LF2 cells (Table 1). Gene ontology analysis of the top 300 genes that were significantly up-regulated in both LF1 and LF2 cells (P < 0.005) reveals active features of myofibroblasts. Analysis of molecular function profiles shows an enrichment of genes associated with extracellular matrix structural constituents and proteoglycans, potentially reflecting active remodeling of the extracellular matrix in IPF (Fig. 1A). The profiling of biological processes showed enrichment of genes involved in stress responses (Fig. 1B). This result was further enhanced by the enrichment of genes related to the biology of ROS (Fig. 1A, oxidoreductase activity, selenium binding, electron transporter activity). Thus, the gene expression profiles reveal that redox regulation is one of the key features of myofibroblasts in IPF.
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A closer look on the list of the genes highly up-regulated in LF1 and LF2 cells provides further information on redox regulation in myofibroblasts. As shown in Table 1, significant up-regulation was observed in genes associated with the metabolism of lipid hydroperoxides. These include phospholipid hydroperoxide reductase (selenoprotein P), the scavenger receptor of phosphatidylcholine hydroperoxide (CD36), oxidoreductases of reactive aldehydes formed from the breakdown of lipid peroxidation (aldehyde dehydrogenase 1, aldo-keto reductase 1C1, 1C2, 1C3) and phospholipase A2. These results suggest that proteins responsible for the metabolism of lipid hydroperoxides are selectively up-regulated in myofibroblasts. Subsequent analysis using real time PCR confirmed this observation (Supplementary Fig. S2). Expression levels of phospholipid hydroperoxide reductases (selenoprotein P, glutathione peroxidase 4) were significantly up-regulated in LF1 and LF2 cells. In contrast, differences in the expressions of hydrogen peroxide reductases and protein disulfide reductases were not evident or consistent. Thus, these results reveal a previously unrecognized physiological function of myofibroblasts characterized by a highly selective response to lipid hydroperoxides.
Enhanced generation of lipid hydroperoxides in myofibroblasts
We found this myofibroblast-specific response in isolated cultures without any stimulation. Thus, the up-regulation of reductases might reflect gene settings imprinted in response to exogenous stresses during disease progression or, more likely, a direct consequence of enhanced lipid peroxidation within the myofibroblasts. To address this question, we measured the levels of lipid hydroperoxides using diphenylpyrenylphosphin (DPPP), a molecular probe that becomes fluorescent upon oxidation by lipid hydroperoxides (Takahashi et al. 2001). It is well known that oxidized DPPP is stably retained in the cell membrane; thus this probe is useful for the estimation of long-term and cumulative lipid peroxidation within live cells. After labeling with DPPP for 10 h, we measured the fluorescence intensities of cells and culture medium. As shown in Fig. 2A, fluorescence intensities were significantly higher in LF1 and LF2 cells compared with TIG7 cells, indicating that the level of lipid hydroperoxides is significantly increased in myofibroblasts. Similar increases were detected in the culture medium from LF1 and LF2 cells (Fig. 2B), suggesting that these cells actively secrete lipid hydroperoxides into the culture medium, or that oxidized DPPP is actively secreted with other cellular lipids. Oxidized DPPP in the culture medium may not be detected as a consequence of passive equilibrium between the cells and medium. In a model experiment in which an ROS generator was over-expressed in retina pigment epithelial (RPE) cells, we could detect an increase in the fluorescence intensity in the cell fraction but not in the culture medium (data not shown). Together, these results reveal a novel physiological feature of myofibroblasts characterized by an enhanced generation of lipid hydroperoxides, which may play a significant role in the pathogenesis of IPF.
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LF1 and LF2 cells show relatively good proliferation rates (doubling times of approximately 2 and 3 days for LF1 and LF2 cells, respectively). This raises the question of how LF1 and LF2 cells deal with their own self-generated oxidative stresses, leading us to analyze selenoprotein P, one of the most highly up-regulated proteins in LF1 and LF2 cells. Although selenoprotein P has never been linked to the pathogenesis of IPF, a potential function of selenoprotein P in myofibroblasts was suggested by its well-known activity as an antioxidant. Biochemical characterization of selenoprotein P showed that it selectively reduces lipid hydroperoxides (Takebe et al. 2002). Thus, it seemed possible that myofibroblasts express selenoprotein P to suppress lipid hydroperoxides within the cells. To test this possibility, we first verified the gene expression data at the level of protein expression. Western blot analysis showed that the expression levels of selenoprotein P in LF1 and LF2 cells are higher than those in TIG7 cells (Fig. 3A). Analysis of lung tissue extracts showed abundant expression of selenoprotein P in patients with IPF (Fig. 3B). Immunohistochemical analysis showed expression of selenoprotein P in the fibrotic regions and epithelial cells (Fig. 3C). Positive immunostaining was detected in samples from all patients analyzed (n = 4, data not shown). In contrast, selenoprotein P was not detected in normal lung parenchyma (Fig. 3C). These results show clearly the association of selenoprotein P expression with the pathogenesis of IPF. We next examined the function of selenoprotein P in the metabolism of lipid hydroperoxides by carrying out RNAi-mediated knockdown of selenoprotein P in myofibroblasts. Treatment with RNAi oligonucleotide significantly suppressed selenoprotein P expression (Fig. 4A, inset). Next, we measured lipid hydroperoxides by DPPP. Interestingly, a significant increase in fluorescence intensity was detected in RNAi-treated cells (Fig. 4A). Furthermore, RNAi treatment led to a more remarkable increase in fluorescence intensity in the culture medium (Fig. 4B). These results strongly suggest that selenoprotein P suppresses lipid hydroperoxides in myofibroblasts. Finally, we examined the effect of selenoprotein P knockdown on the viability of myofibroblasts. As expected, the suppression of selenoprotein P resulted in a partial but significant decrease in the number of viable cells (Fig. 5A). LF1 and LF2 cells showed similar responses to selenoprotein P knockdown (representative data from the analysis on LF2 cells are shown in Figs 4 and 5A). Together, these results signify that myofibroblast viability depends on the expression of selenoprotein P to suppress self-generated oxidative stresses.
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Importantly, the suppression of selenoprotein P had a partial effect on cell viability (Fig. 5A), suggesting that the signaling pathway for maintaining cell viability is activated in myofibroblasts. Thus, we assessed the activity status of MAP kinase pathway proteins, which are known to be activated in response to various types of stress (Lewis et al. 1998). Western blot analysis using active form-specific antibodies showed that JNK and p38 MAP kinase pathway proteins are activated in response to selenoprotein P suppression (Fig. 5B), suggesting that these pathways may be involved in the regulation of myofibroblast viability. To test this possibility, we examined the effects of specific inhibitors on cell viability. Treatment with SP600125, a specific inhibitor of JNK, resulted in considerable loss of viability in cells treated with selenoprotein P RNAi oligonucleotide (Fig. 5A), suggesting that JNK pathway plays a critical role in the regulation of myofibroblast survival. Importantly, JNK inhibitor alone caused a significant loss of cell viability without selenoprotein P RNAi. Thus, these results suggest that endogenous JNK activity, which is detectable in Fig. 5B, may also have a critical function in maintaining viability of myofibroblasts. Our preliminary results show that, although selenoprotein P is highly up-regulated, myofibroblast cells still suffer significant amount of endogenous oxidative stress, resulting in the spontaneous apoptosis (data not shown). Together, endogenous activity of JNK may function for maintaining cell viability in response to this background (leaked) oxidative stress. In contrast to JNK inhibitor, treatment cells with an inhibitor of p38 MAP kinase (SB20385) had no significant effect on viability, although SB20385 effectively blocked p38 MAP kinase activity, which was confirmed by the suppression of the phosphorylation of MAPKAP2, a substrate of p38 MAP kinase (data not shown). LF1 and LF2 cells showed similar responses to inhibitor treatment (representative data from the analysis of LF2 cells are shown in Fig. 5). Together, these results suggest that JNK, but not P38, have a critical role in the regulation of myofibroblast viability.
Selenoprotein P functions as an anti-apoptotic factor against oxidative stress
Loss of viability in cells treated with selenoprotein P RNAi may be due to increased apoptosis caused by self-generated oxidative stress. To determine whether selenoprotein P has a role in apoptosis, we measured cell apoptosis in RNAi-treated cells. As shown in Fig. 6A, the number of apoptotic cells was significantly increased in response to RNAi treatment, suggesting that selenoprotein P functions as an anti-apoptotic factor in myofibroblasts. We next examined the effect of selenoprotein P over-expression on oxidative stress-induced apoptosis and MAP kinase activation. As expected, over-expression of selenoprotein P significantly suppressed hydrogen peroxide-induced apoptosis in RPE cells (Fig. 6B). Selenoprotein P over-expression also significantly suppressed hydrogen peroxide-induced activation of JNK and P38 (Fig. 6C). Together, these results reveal the function of selenoprotein P as an antioxidant and anti-apoptotic protein.
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Discussion |
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Recently, oxidant–antioxidant imbalance and the mechanisms of redox regulation in human IPF have received much attention mainly due to the beneficial effects of antioxidant agents in patients with IPF (Behr et al. 1997; Demedts et al. 2005; Walter et al. 2006). Although a number of studies have suggested a significant role for redox regulation in the progression of human IPF, little is known about the mechanistic details of this regulation (Kinnula et al. 2005). Some major questions remaining to be answered include: What kinds of ROS contribute to the progression of IPF? What kinds of cells generate ROS? What is the function of the generated ROS? How do myofibroblasts and type II alveolar epithelial cells (dominant cells in fibrotic lesions) survive under oxidative conditions? Recent studies have shed some light on these questions, especially in relation to myofibroblasts. Waghray et al. (2005) reported that fibroblasts isolated from the lungs of patients with IPF generate extracellular hydrogen peroxide in response to transforming growth factor-ß1 (TGF-ß1). They also reported that secreted (extracellular) hydrogen peroxide is toxic to pulmonary epithelial cells, which may cause injury in the alveolar epithelium. Larios et al. (2001) stimulated lung fibroblasts with TGF-ß1, and found that extracellular matrix proteins are cross-linked in an oxidant-dependent manner, presenting a potential mechanism for the deposition of extracellular matrix. In contrast, the results of several immunohistochemical studies suggest that the levels of antioxidant and detoxification enzymes are elevated in areas of epithelial regeneration, but not in the fibrotic lesions of interstitial lung diseases. These enzymes include superoxide dismutase (Lakari et al. 2000), catalase (Lakari et al. 2000), thioredoxin (Tiitto et al. 2003), glutaredoxin (Peltoniemi et al. 2004) and heme-oxygenase 1 (Lakari et al. 2001). Together, these results suggest that the species, distributions and functions of ROS may vary in distinct tissue microenvironments in IPF. Thus, the redox balance in fibrotic lesions, which consist mainly of fibroblasts and myofibroblasts, appears to be regulated by an unknown mechanism. Our study provides novel evidence that myofibroblasts function as a generator of lipid hydroperoxides, whose function in the pathogenesis in IPF remains virtually unidentified. Lipid hydroperoxides and reactive aldehydes are known to function as inducers of apoptosis, regulators of transcription factors, substrates for protein carbonylation and protein cross-linkers (Niki et al. 2005). The identification and biochemical characterization of lipid hydroperoxides generated in myofibroblasts should provide novel information about the pathophysiological function of myofibroblasts in IPF. A recent observation suggest that vitamin E supplementation may suppress the progression of experimental lung fibrosis, supports our results (Deger et al. 2006). Immunohistochemical analysis to identify the species and localizations of lipid hydroperoxides in fibrotic lung tissues will be one of the next challenges.
Our study identified selenoprotein P as one of the key effectors of redox regulation and cell viability in myofibroblasts. Selenoprotein P is an abundant extracellular protein that functions as a phospholipid hydroperoxide reductase and as a selenium supply protein. Our results are consistent with the reported function of selenoprotein P as an antioxidant. In our study, we detected the expression of the selenoprotein P protein in cell cultures and tissue sections from patients with IPF. Further analyses should be carried out on serum, EBC and BALF in order to understand whether the dysregulation in selenoprotein P expression is limited to the lung alveoli. Recently, there have been several reports of the association of selenoprotein P with human diseases including cirrhosis (Burk et al. 1998) and Crohn's disease (Andoh et al. 2005). In these reports, a suppression of selenoprotein P expression, which may cause oxidative stress in patients, was associated with disease severity. Our results revealed a paradoxical feature of selenoprotein P, whose expression protects disease-promoting cells from oxidative damage. In addition, our immunohistochemical analysis revealed the expression of selenoprotein P in the alveolar epithelium, as well as in interstitial fibrotic lesions. Thus, the expression of selenoprotein P may also play a significant role in the viability of epithelial cells in an oxidative environment. However, the precise regulatory mechanisms for the expression of selenoprotein P remain to be determined.
One of the key features of the pathogenesis of IPF is the persistence of myofibroblasts in the fibrotic lesions. The elimination of myofibroblasts by apoptosis is a critical process during normal cutaneous wound healing, a process that is suppressed in fibrotic diseases such as IPF (Thannickal & Horowitz 2006). In addition, the alveolar microenvironment in IPF is characterized by high concentrations of various cytokines, ROS and reactive biomolecules, which induce the apoptosis of type 1 alveolar epithelial cells (Thannickal & Horowitz 2006). Thus, the acquisition of an anti-apoptotic phenotype in this microenvironment is a critical process for the persistence of myofibroblasts in IPF. Several studies have shown that the anti-apoptotic phenotype may be regulated by specific signaling molecules in myofibroblasts. These include TGF-ß1 (Horowitz et al. 2004), FAK (Vittal et al. 2005), PI-3K (Horowitz et al. 2004), AKT (Horowitz et al. 2004) and XIAP (Tanaka et al. 2002). Our study reveals that JNK plays an important role in cell viability, providing evidence that a specific cell survival signaling pathway is activated in response to oxidative stress. Although function of JNK and selenoprotein P in the regulation of cell viability is evident, mechanism of myofibroblast survival remains to be analyzed. Therefore, assessment of mechanism of JNK activation, as well as the identification of downstream targets of JNK, will be important as the next challenge. Recently, strategies to induce selective myofibroblast apoptosis are expected to yield an effective approach to the treatment of fibrotic diseases (Tan et al. 1999). Thus, a detailed analysis of this previously unidentified signaling pathway, as well as other apoptosis-regulating pathways, may provide basic information for future targeted therapies.
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Experimental procedures |
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Total RNA was extracted from LF1 and LF2 myofibroblast cell cultures using Isogen (Nippon gene, Tokyo, Japan). The preparation of cDNA from RNA derived from the cell cultures, sample hybridization and scanning of the GeneChipR Human Genome Focus Arrays (Affymetrix) were performed at the Bio Matrix Research (Kashiwa, Chiba, Japan) in accordance with the procedures established by Affymetrix. The Affymetrix Genome Focus chip contains 11 replicate probe sets per gene that are used to address the reliability of the generated data and to determine a P-value for probe set binding using GCOS software (Affymetrix). Hybridization experimental data for each myofibroblast sample were normalized to the data from TIG7 cells. Differentially expressed genes were selected at a threshold of 1.5-fold difference between groups with a significance level of P < 0.005. Clustering analysis of genes was carried out using a TIGR MultiExperiment Viewer <http://www.tm4.org/mev.html>. Gene ontology analysis of the significantly up-regulated genes was carried out using DAVID (Database for Annotation, Visualization and Integrated Discovery) and EASE (Expression Analysis Systematic Explorer), web-based applications <http://david.abcc.ncifcrf.gov/>, which allow access to a relational database of functional annotations. The EASE score, a variant of the Fisher exact probability that weights significance in favor of themes supported by more genes, was used to judge the over-presentation of specific functional annotations. Data are expressed as enrichment score, the geometric mean (in -log scale) of P-values in a corresponding annotation. The original microarray data was submitted in Gene Expression Omnibus (GEO) at National Center for Biotechnology Information <http://www.ncbi.nlm.gov/geo>. Data accession number is GSE6804.
Immunoblotting analysis
Cell lysates were prepared from subconfluent cultures of myofibroblasts. The cells were washed with ice-cold phosphate-buffered saline and lysed in 50 mM Tris–HCl, pH 7.2, 2% SDS, 10 mM dithiothreitol, 10 µg/mL leupeptin, 10 µg/mL aprotinin, 1 mM sodium orthovanadate. Protein amount was determined by the DC assay (BioRad, Hercules, CA). Extracts (20 µg) were separated by SDS-PAGE and transferred to PVDF membranes. The membranes were blocked with 5% non-fat dry milk in TBS containing 0.1% Tween 20, and incubated with primary antibodies diluted with 5% bovine serum albumin in TBS. Primary antibodies included mouse anti-
SMA (1 : 200, Dako, Glostrup, Denmark) and goat anti-selenoprotein P (1 : 500, sc-22639, Santa Cruz, CA), mouse anti-GAPDH (1 : 1000, 1D4, Covance, Berkeley, CA), mouse anti-ERK1 (1 : 5000, BD Bioscience, San Jose, CA), mouse anti-ERK1/2-pThr202/pTyr204 (1 : 1000, BD Bioscience), rabbit anti-JNK/SAPK1 (1 : 1000, BD Bioscience), mouse anti-JNK- pThr183/pTyr185 (1 : 100, BD Bioscience), mouse anti-p38/SAPK2a (1 : 5000, BD Bioscience), mouse anti-p38 MAPK (1 : 2500, BD Bioscience) and rabbit anti-MAPKAP2-pThr334 (1 : 1000, Cell Signaling, Danvers, MA), rabbit anti-SEK1-pThr261 (1 : 200, Cell Signaling), rabbit anti-MKK3-pSer189 (1 : 200, Cell Signaling). Blots were probed with goat anti-mouse (1 : 10 000, BioRad), goat anti-rabbit (1 : 10 000, BioRad) or donkey anti-goat (1 : 10 000, Santacruz, Santacruz, CA) secondary antibody coupled to horseradish peroxidase and visualized by enhanced chemiluminescence (GE Healthcare, Piscataway, NJ).
Immunohistochemistry
Tissue samples were embedded in OCT compound (Cryo Mount, Muto Pure Chemical, Tokyo, Japan), cut into 4 µm-thick sections, and blocked with 5% non-fat dry milk for 30 min. The sections were washed with PBS, and the endogenous peroxidase activity was blocked with 3% hydrogen peroxide. The samples were incubated with anti-selenoprotein P antibody (Santacruz) followed by the second antibody (Simple stain MAX-PO (multi) Nichirei, Tokyo, Japan). After washing thoroughly, positive signals were obtained following incubation with 3,3-diaminobenzidine tetrahydrochloride (Nichirei, Tokyo, Japan). Carrazzi's hematoxylin was used as a counterstain.
Measurement of lipid hydroperoxides
Lipid hydroperoxides were detected using the fluorescence probe diphenylpyrenylphosphin (DPPP, Takahashi et al. 2001). Subconfluent cells were incubated with 100 µM DPPP in DMEM for 10 h. Then the labeled cells were trypsinized, and resuspended in PBS. Fluorescence intensities of cell suspensions (5 x 104 cells/mL) were measured with a Spectrofluorophotometer RF-5300PC (Shimazu CO., Kyoto, Japan) at excitation and emission wavelengths of 351 and 380 nm, respectively. Cell suspensions without DPPP labeling were used as negative controls. The fluorescence intensity of the culture medium used for cell labeling, equivalent to 2.5 x 104 cells, was also measured.
RNA interference
dsRNAi oligonucleotides (100 pmol) were transfected into LF cells using Fugene HD (Roche). The cells were then incubated for 96 h with two oligonucleotide sequences of selenoprotein P and examined for the presence of lipid hydroperoxides. Following sequences were used as dsRNAi oligonucleotides:
#1: 5'CAAGATCCAATGCTAAACT3'
#2: 5'GAAGCCATTAAGATTGCTT3'
Control siRNA oligonucleotide (Silencer® negative control #1 siRNA, Applied Biosystems, Foster City, CA) was also transfected into LF cells using Fugene HD.
Cell viability and apoptosis
Cell viability was measured by the MTT assay as described previously (Kabuyama et al. 1998). Cell apoptosis was measured by an ssDNA ELISA assay (ApoStrandTM ELISA Apoptosis Detection Kit, Biomol, Plymouth Meeting, PA), which detects single-stranded DNA specifically generated in apoptotic cells. Measurement was carried out according to the manufacture's instruction.
Selenoprotein P over-expression
Selenoprotein P over-expression was carried out as described previously (Tujebajeva et al. 2000). Briefly, the coding and 3'-untranslated region of selenoprotein P gene was PCR-amplified and cloned into mammalian expression vector pcDNA3. RPE cells were pretreated with 50 nM sodium selenite for 12 h and transfected with DNA using Fugene 6 (Roche, Valencia, CA). Cells were analyzed at 48 h post-transfection. Expressed selenoprotein P was recovered from culture medium by Ni-NTA agarose (Qiagen, Mannheim, Germany) and detected by Western blotting. PCR primers used are:
5'-GGATCCCCAACGATGTGGAGAAGCCTGGGGCTTGCC-3' (Forward)
5'-GAATTCTGAATTTATTTGGACAAATCCGTACTGTA-3' (Reverse)
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Acknowledgements |
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Footnotes |
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* Correspondence: E-mail: yoshihom{at}fmu.ac.jp
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References |
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Received: 26 April 2007
Accepted: 31 July 2007
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