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Malignant cells are subjected to high levels of oxidative stress that arise from the increased production of reactive oxygen species (ROS) due to their altered metabolism. They activate antioxidant mechanisms to relieve the oxidative stress, and thereby acquire resistance to chemotherapeutic agents. In the present study, we found that PGC1α, a key molecule that both increases mitochondrial biogenesis and activates antioxidant enzymes, enhances chemoresistance in response to ROS generated by exposure of cells to ovarian sphere-forming culture conditions. Cells in the cultured spheres exhibited stem cell-like characteristics, and maintained higher ROS levels than their parent cells. Intriguingly, scavenging ROS diminished the aldehyde dehydrogenase (ALDH)-positive cell population, and inhibited proliferation of the spheres. ROS production triggered PGC1α expression, which in turn caused changes to mitochondrial biogenesis and activity within the spheres. The drug-resistant phenotype was observed in both spheres and PGC1α-overexpressing parent cells, and conversely, PGC1α knockdown sensitized the spheres to cisplatin treatment. Similarly, floating malignant cells derived from patient ascitic fluid included an ALDH-positive population and exhibited the tendency of a positive correlation between expressions of multidrug resistance protein 1 (MDR1) and PGC1α. The present study suggests that ROS-induced PGC1α mediates chemoresistance, and represents a novel therapeutic target to overcome chemoresistance in ovarian cancer.
Tumors undergo metabolic reprogramming to meet the increased energetic and anabolic demands . A number of previous reports have shown that according to the Warburg effect, cancer cells typically rely on aerobic glycolysis due to exhibited defects in mitochondrial function [2, 3]. Tumor cells, however, still possess a capacity to synthesize significant amounts of triphosphate (ATP) if challenged by glucose depletion, via the mitochondrial oxidation of fatty acids and amino acids. This cellular production of ATP and the corresponding intermediate metabolites is a major function of mitochondria. The TCA intermediates and the reducing coenzymes, such as NADH and FADH2, contribute to the production of macromolecules as building blocks for cancer cells . In fact, cancer cells exhibit highly functional mitochondria that efficiently produce both energy and anabolic materials.
Reactive oxygen species (ROS) are byproducts of mitochondrial metabolism, and function as second messengers in the transduction of extracellular signals that control cellular proliferation and cell cycle progression. The altered metabolism exhibited by tumor cells causes ROS production and sustains ROS at an aberrantly high level , thereby driving cancer cells to develop an adaptive system against oxidative stress . In one such detoxifying mechanism, PGC1α, as a major regulator of mitochondrial biogenesis and respiratory function, is required for induction of ROS-detoxifying enzymes, and also contributes to the reduction of ROS generation by mitochondrial metabolism [7–9]. Under the normal conditions, PGC1α is sustained at low levels, but its expression is elevated in response to increasing bioenergetic demands and metabolic alterations . Recently, PGC1α was reported to exhibit oncogenic properties in some cancer cells [10–13]. However, the effect of PGC1α on chemoresistance in ovarian cancer has not yet been elucidated.
Ovarian cancer is usually diagnosed at advanced stages due to difficulty in early detection . More than one third of patients with ovarian cancer suffer from malignant ascites, which is a condition considered to mark the development of chemoresistance and metastasis, resulting in poor prognosis [15–18]. Ascites is composed of acellular fluid containing soluble factors and heterogenic cellular components as a form of single cells or more commonly aggregates and spheres . Floating condition, either as aggregates or spheres, in the ascitic fluid helps cells to survive in an anchorage-independent environment and protects them against penetration of chemotherapeutic agents . In addition, a proportion of malignant cells forming aggregates or spheres have features of cancer stem cell (CSC)-like phenotypes contributing to drug resistance and metastasis [20, 21]. In vitro models in which a non-adherent culture condition mimics the ascitic environment can be used to effectively investigate advanced ovarian cancer, and identify novel therapeutic targets. In the present study, we attempted to identify biological changes associated with chemoresistance using a non-adherent culture consisting of multicellular spheres with CSC-like phenotypes. Using this sphere cell culture, we demonstrated that PGC1α induced by ROS generation facilitates mitochondrial biogenesis and attenuates mitochondrial activity to confer chemoresistance to ovarian cancer cells.
Although PA1 is an ovarian cancer cell line derived from the ascites of a patient with teratocarcinoma, we selected PA1 sensitive to a platinum-based chemotherapy to examine whether sphere-forming culture conditions induce chemoresistance. As expected, sphere-culture conditions resulted in an enriched CSC population with a high ALDH activity. Compared to the parent cells (attached/two dimensional-cultured), the ALDH activity exhibited by the CSCs in ovarian tumor spheres was significantly increased (Figure (Figure1A).1A). Serial subculturing of the spheres (passage 1 and 5) enriched the ALDH-positive population (Figure (Figure1A),1A), and mRNA expression for two subtypes of ALDH and stemness-related genes including Nanog, Sox2, and Bmi1 was also increased in spheres relative to parent cells (Figure 1B and 1C). To confirm the resistance of spheres to a platinum-based chemotherapeutic agent, cisplatin (CDDP), we assessed the effect of treating both parent cells and spheres with the serial concentrations of CDDP or paclitaxel, and found that the spheres exhibited a higher IC50 than their parent cells (Figure (Figure1D,1D, Supplementary Figure 2A). The number of apoptotic cells was found to be significantly decreased (Figure (Figure1E),1E), while conversely, the expression of drug-resistance-related MDR1 and ABCG2 proteins (Figure (Figure1F)1F) was significantly increased in spheres. Taken together, these results suggest that sphere formation enriches the population of stem-like cells in the PA1, and thereby confers drug-resistance.
Sphere formation has been previously shown to stimulate ROS generation . In the present study, hydrogen peroxide (H2O2) and superoxide (O2−) were increased and decreased, respectively, in the spheres compared to their parent cells (Figure (Figure2A).2A). Furthermore, the spheres exhibited relatively high antioxidant gene expression levels in response to endogenous ROS (Figure (Figure2B),2B), while N-acetyl-cisteine (NAC, ROS scavenger) treatment decreased ROS level produced in spheres, and reduced the ALDH activity increased in spheres (Figure 2C and 2D). NAC treatment also decreased the size of the spheres, but did not affect the morphology nor viability of parent cells (Figure (Figure2E).2E). These findings indicate that the intracellular ROS generation caused by sphere formation induces phenotypical changes in CSCs.
To confirm that the ROS-induced PGC1α expression promotes cell detoxification, we analyzed the expression of genes related to mitochondrial biogenesis (PGC1α, −1β, and NRF1) and oxidative phosphorylation (OXPHOS; SDHA, SDHD, and COX4I; Figure Figure3A).3A). Among the genes considered, we focused on PGC1α involved in mitochondrial biogenesis and metabolism . The expressions of OXPHOS complex II, III, and IV, and PGC1α (Figure 3B and 3C) was enhanced by sphere formation, and PGC1α expression was observed to be localized near the center of the spheres (Figure (Figure3D).3D). In addition, sphere formation elevated the mitochondrial mass, and reduced mitochondrial activity (Figure 3E and 3F), which is influenced by mitochondrial dynamics (i.e. fusion and fission) . As predicted, sphere formation altered mitochondrial structure. In contrast to parent cells exhibited elongated mitochondria, the spheres were observed to be markedly fragmented at the perinuclear region (Figure (Figure3G).3G). These findings suggest that sphere-induced ROS generation increases PGC1α expression and mitochondrial biogenesis, but reduces mitochondrial activity via induced mitochondrial fission.
PGC1α as a signaling molecule regulates the expression of antioxidant enzymes . To confirm that PGC1α expression within the spheres is induced by ROS generation, including particularly H2O2, we assessed the effect of treating parent cells with various concentrations of H2O2. The results of the analysis showed that increased exogenous H2O2 levels correlated with increased PGC1α expression (Figure (Figure4A).4A). However, exogenous H2O2 addition to the spheres did not increase PGC1α, MDR1, and OXPHOS protein levels (Supplementary Figure 1). Conversely, ROS elimination via NAC treatment caused a reduction in PGC1α and OXPHOS complex expression in spheres, as well as a corresponding decrease in mitochondrial mass (Figure (Figure4B,4B, and and4D).4D). Interestingly, inhibiting ROS generation induced down-regulation of MDR1 and ABCG2 in spheres (Figure (Figure4E).4E). Taken together, these results suggest that ROS-mediated PGC1α induction results in mitochondrial biogenesis and structural changes, and scavenging ROS sensitizes the spheres to CDDP treatment via down-regulation of PGC1α and associated drug resistance-related proteins.
To determine whether expression of PGC1α is correlated with drug-resistance, we evaluated the effect of overexpressing PGC1α in parent cells. We conducted an immunoblot analysis to confirm that PGC1α was overexpressed following pcDNA-PGC1α transfection in parent cells (Figure (Figure5A).5A). Expectedly, PGC1α-overexpression in parent cells caused an increase in mitochondrial mass to a level comparable to that exhibited by the spheres (Figure (Figure5B).5B). We next further investigated the effect of PGC1α overexpression on the acquisition of CSC-like phenotypes by conducting an ALDEFLUOR assay, and verified conferred drug-resistance via an MTT assay and annexin V-PI staining. The results of these analyses showed that compared to parent cells, PGC1α overexpression increased both the ALDH activity and expression of drug-resistant proteins in spheres (Figure 5C and 5D). To evaluate drug resistance, we treated parent cells and PGC1α-overexpressing cells with various concentrations of CDDP or paclitaxel for 48 h. The PGC1α-overexpressing cells exhibited a higher IC50 value of CDDP or paclitaxel than parent cells (Figure (Figure5E,5E, Supplementary Figure 2C), and the number of apoptotic cells was also significantly lower in the PGC1α-overexpressing cells than in parent cells following CDDP treatment (Figure (Figure5F).5F). In contrast, PGC1α silencing sensitized spheres to CDDP or paclitaxel treatment (Figure (Figure5G,5G, Supplementary Figure 2B). Thus, these results indicate that PGC1α mediates chemoresistance in ovarian cancer cells.
Malignant cells are present in ascites of patients with ovarian cancer, either as single cells, or aggregates (Supplementary Figure 3) . Recent studies have demonstrated that a small proportion of the total ascites cells expresses pluripotent genes and putative CSC markers such as ALDH enzymatic activity [25, 26]. Therefore, we hypothesized that putative CSCs in ascites may be associated with PGC1α-mediated chemoresistance. To test the relationship between PGC1α and chemoresistance in patient-derived cancer cells, we collected and isolated the cellular portion of 14 ascites from patients with serous, clear cell, and endometrioid ovarian cancer at advanced stage III – IV (Table (Table1).1). Of 12 collected patient samples, the majority of ascites-derived cells expressing PGC1α displayed MDR1 expression (Figure (Figure6A),6A), such that we identified a positive correlation between PGC1α and MDR1 expression that was independent of histologic subtypes (p = 0.0027; Figure Figure6B).6B). To enrich cancer cells from heterogenic ascites-derived cells, we performed serial subcultures of ascites 13 and 14 which were classified as being serous and clear cell carcinoma subtype, respectively. Two enriched cancer cells (A13 at passage 14 and A14 at passage 11) had a large ALDH-positive population as much as PA1 spheres (Figures (Figures1A1A and and6C).6C). We found that ascites-derived cancer cells with ALDH-positive population expressed high levels of PGC1α and MDR1 (Figure (Figure6D),6D), and were relatively resistant to CDDP treatment compared to PA1 parent cells (Figures (Figures1D1D and and6E).6E). When we assessed the relationship between chemoresistance and ROS production, we observed that superoxide anion and hydrogen peroxide generation was significantly increased in the ascites-derived cancer cells compared to immortalized normal ovarian surface epithelial cells (IOSE; Figure Figure6F).6F). Reduction of ROS levels via NAC treatment decreased the viability of the ascites-derived cancer cells (Figure (Figure6G),6G), and inhibited their expression of PGC1α and MDR1 (Figure (Figure6H).6H). These findings suggest that, similar to the results generated during analysis of the spheres, ROS-induced PGC1α mediates the chemoresistance of ascites-derived cancer cells with ALDH-positive population.
To date, it has been generally thought that cancer cells dominantly utilize glycolysis to meet the metabolic demands, due to an increased incidence of mitochondrial defects . However, emerging evidence reveals that cancer cells have functional mitochondria that mediate tumorigenesis. In the present study, we provide the evidence that ROS-induced PGC1α mediates chemoresistance in ovarian cancer cells. Currently, platinum-based therapy is the standard treatment used for ovarian cancer. We observed that PA1 cells sensitive to CDDP acquired drug resistance when they were stimulated by exposure to specific culture conditions to form spheres, because this process stimulated ROS generation, and induced the expression of detoxifying enzymes and PGC1α. In turn, PGC1α induction in the spheres caused mitochondrial biogenesis and structural changes, and enhanced drug resistance by mitigating oxidative stress induced by ROS-inducing drugs. However, exogenous H2O2 addition to the spheres after 3D-structure formation did not increase PGC1α and OXPHOS protein levels in a H2O2 concentration-dependent manner compared to mock-treated spheres, because PGC1α already activated inside the spheres during 3D-structure formation.
Exposure to culture conditions conductive to sphere formation is considered as an efficient method for the enrichment and isolation of CSCs [27–30]. In addition, three-dimensional (3D) culture via sphere formation is a representative method used to mimic in vivo tumor microenvironment. Because of limitation of oxygen penetration, the inside of in vitro-cultured spheres (3D culture) is in both oxidative stress and hypoxic conditions, which activate antioxidant proteins and HIF contributing to resistance to chemotherapy [22, 31–33]. In advanced ovarian cancers, malignant cells floating in ascites as a form of single cells, aggregates and/or sphere have the ability to metastasis to distant sites and to protect themselves against chemotherapy [15, 19]. It has been previously proposed that malignant cells with cancer stem-like phenotypes are enriched in aggregates and/or spheres within ascites. In fact, ascites-derived spheres expressing Oct4, STAT3, and CA125 have been shown to exhibit a strong tumorigenic ability in nude mice compared to single cells . According to the results of the present study, the microenvironmental condition of spheres mimics that of an in vivo tumor mass by enabling ovarian cancer cells to acquire stem-cell properties, including high ALDH activity and drug resistance. Interestingly, 2D and 3D culture conditions facilitated opposite cell responses to NAC treatment. In the patient-derived samples, the expressions of MDR1 was positively correlated with that of PGC1α in ascites cells. This implies that the physical tumor microenvironment should be considered a factor that critically influences cancer-cell phenotypes.
While antioxidants are generally established to exert a protective effect against carcinogenesis by reducing DNA damage, recent research revealed that ROS scavengers, including NAC, inhibit the growth and proliferation of glioblastoma-initiating cells via the impairment of cell cycle progression . In addition, it was reported that elevated ROS produced by prostate tumor initiating cells was required for the activation of IL-6/STAT3, which was related with carcinogenesis of human prostate cells . Intracellular ROS as second messenger stimulate cell proliferation and regulate . The results of the present study demonstrated that the utilized 3D culture condition facilitates ROS generation, and thus enables ROS to stimulate proliferation of tumor cells. A reduction in the level of ROS decreases the size and the number of incident spheres, concomitantly with a reduction in ALDH activity. In addition, up-regulation of antioxidant gene expression is observed in spheres accompanying ROS elevation. We observed a decreased level of superoxide and an increased level of hydrogen peroxide, because superoxide dismutase 2 (SOD2) may act in the process to convert superoxide to hydrogen peroxide in mitochondria. Similarly, accumulating evidence suggests that SOD2 expression is correlated with chemoresistance in lymphoma , basal-like breast carcinoma , and lung adenocarcinoma . In line with previous data, the results of the present study suggest that acquisition of chemoresistance and stem-like phenotypes in spheres is likely associated with the ROS-induced expression of detoxifying enzymes. Thus, removal of ROS may be a promising strategy to target proliferative CSCs in ovarian cancer.
PGC1α critically regulates the transcriptional control of mitochondrial metabolism and biogenesis [9, 23]. Recent studies have suggested that expression of PGC1 family members is associated with oncogenic processes, including metastasis and chemoresistance of cancers. PGC1α and mitochondrial transcription factor A (TFAM) were found to be increased in high grade serous ovarian cancers that were highly chemoresistant . Similarly, circulating breast cancer cells have been found to exhibit enhanced mitochondrial biogenesis and respiration as a result of induced PGC1α expression, leading to an increased rate of metastasis . An increased reliance on OXPHOS with a marked shift away from aerobic glycolysis was observed in glioma stem cells . Similarly, high PGC1α and HIF1α levels have been suggested as effective and non-invasive plasma biomarkers of poor prognosis for patient with breast cancer . The findings of the present study support a role for PGC1α in the regulation of oncogenic properties. Sphere formation-induced ROS generation was shown to stimulate PGC1α expression, leading to the increased expression of OXPHOS-related genes and proteins. Although mitochondrial fission was observed in spheres, mitochondrial mass was also increased to compensate for the inefficient energy metabolism caused by mitochondrial fission under low oxygen conditions.
Thus, the present study provides evidence that PGC1α induced by oxidative stress mediates chemoresistance in ovarian cancer. Many previous studies have focused on the increase of mitochondria that occurs in relation to caspase-dependent death pathways in cancer; however, the findings of the present study demonstrate the contradictory functions PGC1α-regulated mitochondria in mediating cancer cell survival. Although further studies are required to elucidate the molecular mechanisms underlying our findings, we propose that PGC1α is a key regulator that controls chemoresistant phenotypes in ovarian cancers.
Ovarian cancer cell line, PA1 was purchased from American Type Culture Collection, and cultured in DMEM/F12 (Life Technologies, Gaithersburg, MD) supplemented with 10% fetal bovine serum (FBS; Life Technologies), 1% penicillin and streptomycin (Life Technologies). PA1 cells were maintained at 37°C in humidified atmosphere of 5% CO2.
PA1 cells (1,000 cells/cm2) were plated onto poly-2-hydroxyethyl methacrylate (10 mg/ml, poly-HEMA; Sigma Aldrich, St. Louis, MO) coated plate, and cultured in DMEM/F12 containing 20 ng/ml human recombinant epidermal growth factor (EGF; Life Technologies), 20 ng/ml basic fibroblast growth factor (bFGF; Life Technologies), 5 μg/ml human recombinant insulin (Life Technologies), 0.1x B27 (Life Technologies), and 1% penicillin and streptomycin (Life Technologies). Spheres were maintained at 37°C in humidified atmosphere of 5% CO2 for two weeks.
Cancer stem-like cells with high ALDH activity were identified in spheres and parent cells (1 × 106 cells) using ALDEFLUOR® assay kit (STEMCELL Technologies, Vancouver, Canada). The spheres were dissociated with 0.05% trypsin-EDTA at 37°C for 2 min. Trypsin-EDTA treated spheres were centrifuged at 4°C to avoid further enzymatic damage (700 × g, 10 min). Dissociated spheres and parent cells (each 1 × 106 cells) were suspended in ALDEFLUOR assay buffer containing ALDH substrate without/with 50 mM DEAB (as a negative control, specific inhibitor to ALDH), and incubated for 30 min at 37°C. ALDH-positivity was analyzed by BD FACSCanto II flow cytometer (BD Biosciences, NorthRyde, Australia).
ROS level of spheres and parental cells was measured by dihydroethidium (DHE; Sigma Aldrich, St. Louis, MO) and 6-carboxy-2,7-dichlorodihydrofluorescein diacetate (DCFH-DA; Sigma Aldrich) to detect superoxide anion and hydrogen peroxide, respectively. The spheres and parental cells were dissociated with 0.05% trypsin-EDTA at 37°C for 2 min. Trypsin-EDTA treated spheres were centrifuged at 4°C to avoid further enzymatic damage (700 × g, 10 min). The dissociated parental cells were centrifuged at 500 × g, 4°C for 4 min. Both dissociated spheres and parental cells were incubated in serum-free medium with DHE (5 μM for 10 min) or DCFH-DA (20 μM for 30 min) at 37°C in the dark. Relative fluorescence intensity of ethidium (ETH, oxidized form of DHE by superoxide) or DCF (oxidized form of DCFH by hydrogen peroxide) was measured by BD FACSCanto II flow cytometer (BD Biosciences).
Total RNA was extracted from spheres and parent cells with TRIzol reagent (Invitrogen). Complementary DNA (cDNA) was synthesized from 1μg of total RNA using superscript III first-strand synthesis system with oligo(dT)20 primers (Invitrogen). The synthesized cDNA was diluted with nuclease free water (five-fold), and mixed with Master Mix SYBR Green I dye (Bio-Rad, Hercules, CA). Relative gene expression levels were measured by CFX96 Real-Time PCR Detection System (Bio-Rad) according to the manufacturer's protocol. Amplification conditions were followed for 35 cycles at 94°C for 30 s, 60°C for 30 s, and 72°C for 30 s. The relative gene expression levels were quantified using the 2−ΔΔCtmethod and normalized to the Ct value of the reference genes, β-actin and GAPDH.
Spheres and parent cells were harvested and lysed with protein extraction buffer (0.5 M NaCl, 0.5 M Tris-HCl, 50 mM EDTA, 50 mM EGTA, 10% triton X-100, 1 mg sodium deoxycholate, 1 mM Na3VO4, 1mM phenylmethylsulfonyl fluoride, EDTA-free protease inhibitor, and distilled water). Proteins (20 μg) were separated on 6 - 10% SDS-PAGE, and transferred onto a nitrocellular membrane. After blocked with 5% skim milk solution in Tris-buffered saline with 0.1% Tween 20 (0.1% TBS-T), the membranes were incubated with primary antibodies specific for MDR1 (Cell Signaling Technology, Beverly, MA), ABCG2 (Santa Cruz Biotechnology, Santa Cruz, CA), total OXPHOS complexes (Abcam, Cambridge, MA), and PGC1α (Calbiochem, Darmstadt, Germany). Signals were visualized using a chemiluminescence detection kit (AbFrontier, Seoul, Korea).
To measure cell viability after treated with cisplatin (CDDP), cultured cells (spheres and parent cells) were incubated with 2 mg/ml of 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) dissolved in phosphate buffered saline (PBS) solution at 37°C in the dark. After incubation for 3 h, MTT was removed, and dimethyl sulfoxide (DMSO) was added to dissolve formazan in the live cells. Aliquots (200 μl) of DMSO solution were transferred into 96-well plates, and absorbance was recorded at 540 nm using a spectrophotometer (Labsystem Multiskan, Helsinki, Finland)
Spheres were cultured for two weeks and treated with various concentrations of CDDP for 48 h. Drug-treated cells were dissociated with 0.05% Trypsin-EDTA, and centrifuged at 500 × g for 4 min. The cell pellet was suspended in PBS and stained using annexin V-PI apoptosis detection kit (BD Bioscience Pharmingen, San Jose, CA). Apoptotic cells were analyzed by BD FACSCanto II flow cytometer using the CellQuest analysis program (BD Biosciences, NorthRyde, Australia).
To measure the relative mitochondrial activity compared to parent cells, both parent cells and spheres were dissociated and incubated in serum-free medium containing 200 nM MitoTracker® Orange CMTMRos (Invitrogen) at 37°C for 30 min. After incubation, the stained cells were centrifuged at 500 × g for 4 min, and washed with PBS. The cells were analyzed by BD FACSCanto II flow cytometer (BD Bioscience).
After culture for two weeks, sphere samples were fixed with 4% formaldehyde at 4°C overnight. Fixed samples were washed with PBS for 30 min (three times every 10 min) followed by dehydration through a graded ethanol of 25, 50, 70, 90, and 100% for 10 min in each step. Paraffin-embedded preparations of sphere samples were sectioned at 7 μm thickness by a microtome (HM340E; Microm, Waldorf, Germany). Water drops were put on VWR® Micro Slides (VWR International, West Chester PA), and the sectioned paraffin-embedded samples were put on the water drops to help the samples to attach to the slides. The sections on the slides were dried at 40°C overnight to vaporize water drops, and dewaxed with CitriSolv (Fisher BioSciences, Pittsburgh, PA). After rehydrated with a graded ethanol of 100, 70, 50, and 25%, and PBS for 10 min in each step, samples were blocked in PBS containing 0.1% Tween-20 (0.1% PBS-T) and 3% normal goat serum (Vector Laboratories, Burlingame, CA) at room temperature for 1 h.
Sectioned samples were incubated in 0.1% PBS-T containing 3% normal goat serum and primary antibody (anti-PGC1α mouse antibody, 1:200; anti-ABCG2 mouse antibody, 1:500) at 4°C overnight. After washed in 0.1% PBS-T for 60 min (three times every 20 min), samples were incubated in 0.1% PBS-T containing secondary antibody (anti-mouse IgG conjugated with AlexaFluor488, 1:250) for 4°C overnight. Samples were washed with 0.1% PBS-T for 60 min (three times every 20 min), and mounted with mounting medium (Vectashield with DAPI, H-1200; Vector Laboratories). Fluorescent images were captured using LSM 700 confocal microscope (Zeiss, Germany).
PA1 parent cells were transfected with pcDNA4-myc-PGC1α using Lipofectamine® 2000 transfection reagent (Life Technologies). The pcDNA4-myc-PGC1α was a gift from Toren Finkel (Addgene plasmid # 10974) . Transfected cells were selected with 400 μg/ml Zeocin (Invitrogen) during two weeks.
Small interfering RNA (siRNA) was obtained from Bioneer (Daejeon, Korea). Spheres were dissociated and transfected with siRNA against PGC1α (Sense: CAAUAACUCCACCAAGAAA, Antisense: UUUCUUGGUGGAGUUAUUG) and scrambled siRNA (negative control) using Lipofectamine® 2000 transfection reagent (Life Technologies). The transfected cells were cultured onto a poly-HEMA coated plate for one week.
Ascites was aspirated from ovarian cancer patients with different histological types including serous, clear cell, and endometrioid type, at the stage III and IV. Ascites was diluted with the same volume of PBS, and centrifuged (4°C, 1,400 × g, 10 min). Cells were gently overlaid onto Ficoll-PaqueTM-PREMIUM, and centrifuged (4°C, 1,400 × g, 30 min). The cell layer was collected and cultured in the complete culture medium.
Data were presented as mean ± SEM of at least three independent experiments. Student's t-test and one-way ANOVA were used for statistical analyses. Significant difference among experimental groups was analyzed by Scheffe's post hoc test. All analyses were conducted using IBM SPSS statistics 23 (SPSS Inc., Chicago, IL).
James E. Womack provided language and writing assistance.
CONFLICTS OF INTEREST
The authors have declared no conflicts of interest.
This work was supported by the BK21 plus program (5256-20140100), the Pioneer Research Center Program (2012-0009555), the National Institutes of Health (CA116984, CA123233), SNUH research fund (07-2017-0530), and the Korea Health Technology R&D Project (07-2017-1041).