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Multiple mechanisms of apoptosis induced by 20(S)-protopanaxadiol (aPPD) in glioma cells Liu, Guo Yu 2004

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Multiple Mechanisms of Apoptosis Induced by 20(S)-Protopanaxadiol (aPPD) in Glioma Cells By Guo Yu Liu B. Med., China Medical University, 1987 M. Med., China Medical University, 1992 A THESIS SUBMITTED IN PARTIAL F U L F I L L M E N T OF THE REQUIREMENTS FOR THE DEGREE OF M A S T E R OF SCIENCE IN THE FACULTY OF GRADUATE STUDIES (Department of Surgery) We accept this thesis as conforming to the required standard The University of British Columbia August 2004 © Guo Y u Liu, 2004 T H E U N I V E R S I T Y O F BRITISH C O L U M B I A F A C U L T Y OF G R A D U A T E S T U D I E S Library Authorization In presenting this thesis in partial fulfillment of the requirements for an advanced degree at the University of British Columbia, I agree that the Library shall make it freely available for reference and study. I further agree that permission for extensive copying of this thesis for scholarly purposes may be granted by the head of my department or by his or her representatives. It is understood that copying or publication of this thesis for financial gain shall not be allowed without my written permission. Name of Author (please print) Date (dd/mm/yyyy) Title of Thesis: f^^-hipl^ He-(À(MiîStY\S rrf ApûPJV^'S XhduCejj 3.0 c s ; - Pr&fovfrhct^qAîa.i LÀj>/>Ty ) )/\ Glioma Ce-tls Degree: Department of The University of British Columbia Vancouver, BC Canada HWfe / P r ê t i e z Year: grad.ubc.ca/forms/?formlD=THS page 1 of 1 last updated: 20-Jul-04 ABSTRACT Ginsenosides are major pharmaceutical components in ginseng, cytotoxicity of ginsenoside Rh2, which exists in ginseng extract with trace concentrations, on cancer cells has been known. In our previous study, we identified that Careseng®, a specially formulated ginseng product, had the equal cytotoxicity on tumor cells as Rh2. In this study I investigated the mechanism of apoptosis in glioma cells with different PTEN and P53 status induced by 20(S) protopanaxadiol (aPPD), which is a derivative of Rh2 and the major constituent of Careseng®. aPPD can be obtained in lower cost from ginseng. Firstly, based on the viability analysis and D N A staining of U87MG (PTEN negative and wild type p53) and SF188 (PTEN positive and p53 mutant) cells treated by aPPD, it was found that aPPD, as effectively as Rh2, rapidly induced apoptosis in two glioma cell lines. The cell apoptosis of U87MG and SF188 cells induced by aPPD was confirmed by D N A fragmentation, T U N E L DNA-staining, typical apoptotic nuclear changes and cell membrane blebbing. By Western Blot analysis, it was demonstrated that caspase-3, -7, -8 and -9 were activated in the SF188 cells with dose- and time-response relationship to aPPD treatment but not in the U87MG cells, and found that cells with caspase activation revealed much more sensitive to aPPD treatment than those without caspase activation. Phosphorylated Akt on Ser473 was significantly decreased in U87MG but not SF188 cells. APPD-caused increase in intensity of superoxide anion and number of positive cells in both cell lines. Antioxidant gene activator, tBHQ or general caspase inhibitor partially blocked the aPPD-induced cell when applied alone but demonstrated a nearly complete protection when used together in SF188 cells. Although, I N K phosphorylation was significantly increased in aPPD treated glioma cells, their activation was not related to the apoptosis as I N K inhibitors had effect on the cell viability. The above results suggest that aPPD is a promising candidate for anti-cancer agent that induces apoptosis in a wide spectrum of cancer types. The bioactivity of aPPD is related to multiple signalling pathways and apoptosis induced by aPPD may be mediated by different mechanisms in different cells. T A B L E OF CONTENTS ABSTRACT .• ii TABLE OF CONTENTS iv LIST OF FIGURES '. I vii LIST OF TABLES ix ABBREVIATIONS x ACKNOWLEDGEMENTS xiii CHAPTER 1 GENERAL INTRODUCTION 1 1. General understanding of ginseng 1 2. History of usage and study of ginseng and ginsenosides 2 3. Relationship between chemical structures and function of ginsenosides 4 4. Metabolism of ginseng saponin 7 5. Ginsenosides effect on differentiation, invasion, growth arrest and apoptosis of tumor cells 9 6. Aglycone of ginsenosides 12 CHAPTER 2 MATERIALS AND METHODS , 19 1. Equipments 19 2. Reagents 19 3. Cell cultures 20 4. Preparation of ginsenoside solutions and treatment 21 5. MTT assay 21 6. Hoechst 33342 staining and morphological analysis 22 7. D N A ladder 22 8. D N A Staining with ApopTag® 23 9. Apoalert™ mitochondrial membrane permeability assay 24 r 10. Extraction of subcellular fractions and mitochondria 24 11. Cytochrome c-depleted mitochondria and cytoc 25 12. Protein extraction and western blotting analysis 26 13. Cell-free system for caspase-3 activation assay 27 14. Caspase and JNK inhibition assay 27 15. Superoxide anion staining assay 28 16. Statistical analysis 28 CHAPTER 3 EFFECTS OF CASPASES ON aPPD-INDUCED CELL DEATH IN CELLS WITH DIFFERENT p53 AND PTEN STATUS 29 Introduction 29 Result 34 1. Cytotoxicity induced by aPPD on different tumor cells 34 2. The apoptotic cell-death induced by aPPD 35 3. aPPD-caused cytochrome c release from mitochondria 36 4. Multiple caspase activation in glioma cells treated with aPPD 37 5. Activation of caspase-3 by aPPD in SF188 cells requires a membrane component 37 6. Caspase-8 but not -3 or -9 partially responsible for aPPD-induced apoptosis in SF188 cells 38 7. Caspase activation is not essential for aPPD-induced cell-death 39 Discussion 55 CHAPTER 4 aPPD ACTIVATES STRESS SIGNALING PROTEIN KINASE WHILE INHIBITING PRO-GROWTH SIGNALING PROTEIN KINASE IN GLIOMA CELLS 60 Introduction 60 Result ..67 1. aPPD induced JNK phosphorylation in SF188 and U87MG cells in dose- and time-dependent fashions 67 2. JNK inhibitor did not block apoptosis induced by aPPD ; 67 3. Effect of aPPD on phosphorylation level of Akt Ser473 68 Discussion 72 CHAPTER 5 SUPEROXIDE ANION AND APOPTOSIS OF GLIOMA CELLS INDUCED BY aPPD 76 Introduction 76 Results 80 1. Dose-response and time-course of superoxide anion production induced by aPPD in glioma cell lines 80 2. Stimulation of antioxidant gene expression partially inhibited aPPD-induced apoptosis 80 Discussion 85 CHAPTER 6 GENERAL DISCUSSION AND CONCLUSION '. 88 REFERENCES 93 Figure. 1 -1. Chemical structure of ginsenosides and their related compounds 5 Figure 1-2. Schematic metabolism routes of 20(S)-protopanaxadiol type ginsenosides.. 15 Figure 1-3. Structural relation among 20(S)-protopanaxadiol type ginsenosides and aglycone 16 Figure 1 -4 Pathways of related to apoptosis induced by aPPD 18 Figure 3-1. Different cell-death induced by aPPD in dose- and time-dependent manner 41 Figure 3-2. Apoptosis induced by aPPD in SF188 cells at different time points 42 Figure 3-3. D N A ladder in SF188 cells induced by aPPD 43 Figure 3-4. Apoptosis induced by aPPD in U87MG cells at different time Points : 44 Figure 3-5. Detection of D N A strand breaks as biochemical markers of apoptosis by ApopTag® staining 45 Figure 3-6. Change of mitochondrial membrane permeability induced by aPPD treatment in SF188 cells stained with BD ApoAlert™ Mitochondrial Membrane Sensor Kit 45 Figure 3-7. Release of cytochrome c induced by aPPD from intact cell or separated mitochondria 46 Figure 3-8. Time-and dose- dependent activation of multiple caspases induced by Appd in SF188 cells... : 47 Figure 3-9. aPPD-induced caspase-3 activation in cell-free system 47 Figure 3-10. Effects of caspase inhibitors on apoptosis induced by aPPD in SF188 Cells 48 Figure 3-11. aPPD-induced apoptosis in SF188 cells pre- treated with general caspase inhibitor (Z-VAD-FMK) 49 Figure 3-12. General caspase inhibitor (Z-VAD-FMK) entirely blocked caspase activity 50 Figure 3-13. aPPD induced caspase-8 activation and apoptosis is independent of caspase-3 in MCF-7 cells 51 Figure 3-14. Caspase inhibitors did not affect viability of aPPD-treated MCF-7 cells 52 Figure 3-15. Caspase-3 and-8 activity in aPPD-treated U87MG cells 53 Figure 3-16. Detection of caspase-8 and -3 in SF210 and U138 cells treated with aPPD 54 Figure 4-1. aPPD induced dose- and time- dependent JNK activation in SF188 and U87MG glioma cells by Western blotting analysis 69 Figure 4-2. JNK inhibitors did not block apoptosis induced by aPPD 70 Figure 4-3. aPPD had no effect on phosphorylation levels of Akt Ser473 in SF188 cells 71 Figure 4-4. aPPD induced reduction of P-Akt in U87 M G cells 71 Figure 5-1. Superoxide anion generation in glioma cells induced by aPPD 82 Figure 5-2-1 Effect of lOuM tBHQ and 50uM Z - V A D - F M K (general caspase inhibitor, GI) on 86.8uM aPPD induced cell-death in SF188 cells 83 Figure 5-2-2. tBHQ had no effect on aPPD-induced cell death in U87MG cells 83 Figure 5-3. Vitamin C or E did not affect the aPPD-induced cell death in SF188 cells 84 Figure 6-1. Schematic representation of proposed intracellular signaling leading to apoptosis by aPPD in glioma cells 91 LIST OF TABLES Table 6-1. p53 and PTEN status in related brain tumors •OH hydroxyl radicals A A P H 2,2'-azobis(2-amidinopropane hydrochloride) AIF apoptosis inducing factor aPPD 20(S)-protopanaxadiol aPPT 20(S)-protopanaxatriol A R E antioxidant response element ATF activating transcription factor B H Bcl-2 homology caspase cyteine aspartate-specific proteases CDKs cyclin-dependent kinases CED-3 cell death gene-3 DISK death inducing signaling complex DR death receptor ERK extracellular signal-related kinases FADD Fas-associated death domain FasL Fas-ligand H 2 0 2 hydrogen peroxide HE Hydroethdine/Dihydroethidium HO-1 heme oxygenase-1 HPLC High-performance liquid chromatography Hsp heat shock protein IAP inhibitor of apoptosis proteins ICE mammalian interleukin-1/3 converting enzyme ILK Integrin-linked kinase JNK/SAPK c-Jun NH2-terminal kinases/ stress-activated protein kinases M A P K mitogen-activated protein kinase MTT 3-(4,5-dimethythinzol -2yl) -2,5-diphenyl tetrazolium bromide NAD(P)H nicotinamide adenin dinucleotide phosphate reduced NF- K B nuclear factor- K B NO nitric oxide N Q O l NAD(P)H: quinone oxidoreductase Nrf2 nuclear factor E2-related factor 2 02 - - superoxide anion ONOO- peroxynitrite anions PAK2 p21-activated protein kinase PARP poly (ADP-ribose) polymerase PCD programmed cell death PDK1 3 -phosphoinositidedependent protein kinase-1 PH pleckstrin homology PI3K phosphatidylinositol 3-kinase PIP2 phosphatidylinositol 4,5-bisphosphate PIP3 phosphatidylinositol- 3,4,5-trisphosphate PKB protein kinase B PTEN phosphatase and tensin homolog deleted from chromosome 10 ROS Reactive oxygen species SAPK stress-activated protein kinase SOD superoxide dismutase tBHQ tert-butyl-hydroquinone TLC thin layer chromatography TNF tumor necrosis factor TNFR tumor necrosis factor receptor TUNEL terminal deoxynucleotidyl transferase mediated dUTP nick end labeling X l l A C K N O W L E D G E M E N T S I would like to express my special and heartfelt gratitude to my supervisor, Dr. William W. Jia, for his intellectual guidance, inspiration, and financial support. I would like to thank my supervisory and oral defense committee members, Dr. Mercel B Bally, Dr. Shoukat Dedhar, and Brian Thiessen, for their helpful discussion, constructive instruction. I also like to express my sincerest appreciation to PanaGin Pharmaceuticals Inc and Dong Huang for their kind gifts of ginsenoside products. I will remember the help from virtually all people in Brain Research Center. I would like to express my special thankfulness to Xue-xian Bu, Hang Yan, Zhong Jun Ma, Coquinco Ainsley, Dong Qiang, Yan-huan Wen, Jian Wang, Ying Ru liu, L i Pang, Yuping L i , Jie Lu, L i Dong Liu, Yitao Liu and Yushan Wang for the one time or another help. I also want to thank some people who work outside of Brain Research Center, but they give me many kindly help. I am indebted to Dr. Andy Shih for providing some regents and Andy Johnston for his instruction to use flow cytometry equipment. Many thanks also go to the administrative and secretarial staff in Surgery Department of U B C and Brain Research Center. They are Joanne Clifton, Dr. York Hsiang, Chris Crossfield, Faye Pederson. Finally, I would like to express my special appreciation and gratitude to my family: my parents, my husband and my son for their love, understanding and support. This program is supported by BC Cancer Agency. CHAPTER 1 GENERAL INTRODUCTION 1. General understanding of ginseng Ginseng (Panax) is a member of the plant family Araliaceae, phylogenetically one of the oldest plant families. There are three tribes in this family differentiated by their petal shape and aestivation. Ginseng has petals which are broad based and slightly imbricate in the bud (i.e. slightly overlapping at the tip of side) (Court, 2000). There are only about a dozen species of Panax in the world (Duke, 1989). They are: Panax ginseng C. A . Meyer, Panax quinquefolius L, Panax japonicus C. A . Meyer, Panax trifolius L, Panax pseudoginseng Wallich, Panax notoginseng F. H . Chen, Panax elegantior, panax zingiberensis C. Y . Wu & K . M.Feng, Panax bipinnatifidus, and Eleutheroxoccus sentocosus (Duke, 1989). They belong to the genus Panax. Most studies of ginseng have utilized constituents from three common species: Panax ginseng (Asian mainly in China), Panax quinqeufolium (American ginseng) and Panax japonicus (Japanese ginseng) (Yun et al., 2001). With the exception of their biological types, ginsengs can be divided to three types based on the differences in preparation: fresh ginseng, white ginseng (dried after peeling off) and red ginseng (steamed and dried). Ginseng has been known as a non-toxic herb. Chronic treatment of ginseng on rats, mice, dogs, and rabbits has shown very few observable signs of toxicity (Chen et al., 1980; Hess, 1983; Nakata et al., 1998). There were no treatment-related changes in body weight, hematology, or clinical chemistry after 90 days of daily administration of as much as 15 mg/kg to both male and female dogs (Hess, 1983; Nakata et al., 1998). 2. History of usage and study of ginseng and ginsenosides Ginseng is a traditional Chinese medicine and has been used for thousands of years (Yun, 2001). The earliest written record about its pharmacological function comes from Shen Nung's materia Medica (Shen Nung Ben Cao Jing). In this book, ginseng was described as a non-toxic herb. In 196A.D., almost 2000 years ago, prescription of ginseng appeared in the Shanghan Lun (Treatise on Fever). In other Asian countries such as Japan and Korea, the history of application of ginseng is very obscure. Although the efficacy of ginseng was known in the West by the 18 th century (Huang K C , 1999), it was recorded that Panax quinqeufolium (American ginseng) has been exported to Asia since 1700s (Bahrke and Morgan, 2000). One of the earliest recorded chemical investigations of a panax species was published in 1854, when Rafinesque commented on a camphor-like substance named panacene which he had isolated from an extract of the root of the American ginseng (Court, 2000). After that, the active compounds of ginseng began to be extracted and studied widely. Forty years ago, Dr. Shibata and his colleagues classified the saponin glycosides-ginsenosides as protopanaxadiol type and protopanaxatriol type (Shibata et al., 1963; Shibata et al., 1966). The protopanaxadiol and protopanaxatriol are the genuine sapogenins(Shibata et al., 1966; Shibata, 2001). In 1966, his research group isolated and described ginseng saponins which they named "ginsenosides" (Shibata et al., 1966) and this terminology has now been universally accepted. Based on Dr. Shibata (Shibata et al., 1966), the ginsenosides (genuine sapogenins) are named ginsenosides Rx(x=0, a-1, a-2, b-1, b-2, b-3, c, d, e, f, g-1, -2, -3... h-1, -2, ...), x corresponds to the sequence of Rf value (retention factor =distance traveled by the target compound divided by the distance traveled by the solvent) of the spots on the thin layer chromatography (TLC) from the bottom to the top (Shibata et al., 1966). Ginsenosides are triterpenoid glycosides of the dammran series. In general, the ginseng saponins are divided into three groups according to the non-sugar (aglycone) part of the molecule: Protopanaxadiol type (PPD), Protopanaxatriol type (PPT), and Oleanolic acid (Odashima et al., 1985; Liu and Xiao, 1992; Tachikawa et a l , 1999; Yun, 2001). Thirty-five kinds of ginsenosides have been isolated so far from fresh, white or red ginseng and novel structures continue to be extracted and reported. Among them 22 of these ginsenosides are protopanaxadiol type, such as Ra, Rb, R b l , Rb2 Rb3, Rc, Rd, Rg3, Rh2 and aglycone PPD (aPPD); 11 are protopanaxatriol type, such as ginsenosides Re, Rf, R g l , Rg2, R h l , and aglycone PPT (aPPT); and only one ginsenoside Ro is the oleanane type (Odashima et al., 1985; Liu and Xiao, 1992; Yun, 2001). The biological active constituents of ginseng are considered to be a series of saponin glycosides (Shin et al., 2000) even though there are some other compounds in ginseng such as carbohydrates, nitrogenous compounds, fat-soluble compounds, vitamins and minerals. Mounting evidences have indicated therapeutic potential of ginseng and ginsenosides on the central nervous system (CNS) (memory, learning and behavior), neuroendocrine function, carbohydrate and lipid metabolism, immune function, and the cardiovascular system (Chen, 1996; Gillis, 1997; Attele et al., 1999; Takei et al., 2004; Tohda et al., 2004), although their pharmacological mechanisms are still unclear. 3. Relationship between chemical structures and function of ginsenosides The principle active constituents in ginseng are ginsenosides, saponin glycosides, whose distribution appears to be restricted to the Panax genus (Duke, 1989). Ginsenosides share a similar basic structure, consisting of gonane steroid nuclei (dammarane skeleton), which includes 17 carbon atoms arranged in four rings with a modified side-chain at C20 (Byun et al., 1997). Although they have the common nuclear structure as illustrated in Fig.1-1 (Chen et al., 1980; Yoshikawa et al., 1998; Kim et al., 1999c; Hasegawa et a l , 2000; Bae et al., 2002b; Cai et al., 2002), it has been found that ginsenosides exhibit considerable structural variation. Their structure diversities come from three factors. First, they differ from one another by the type, number and attachment site of sugar moieties. Some sugar moieties present are glucose, maltose, fructose, and saccharose. They are attached to C-3, C-6, or C-20 (Attelé et al., 1999). Second, ginsenosides differ in hydroxyl groups for their number and site of attachment (Kim et al., 1998). Another factor is stereochemistry at C-20. Most ginsenosides that have been isolated are naturally present as enantiomeric mixtures (Soldati and Sticher, 1980). araf: a-L-arabinofuranosyl arap: oL-arabinopyranosyl glc: /3-D-glucopyranosyl rha: a-L-rhamnopyranosyl xyl: /?-D-xylopyranosyl Ac: acetyl Name of Ginsenosides R l R2 R3 Type of Ginsenosides aPPD H H OH PPD Rbl -Glc 2 - 'Glc H -O-Glc 6 - 1 Glc PPD Rb2 -Glc 2 - 1 Glc H -O-Glc 6 - 1 Arap PPD Rb3 -Glc 2 - 1 Glc H -O-Glc 6 - 1 X y l PPD Rc - G l c ^ G l c H -O-Glc^Ara f PPD Rd -Glc 2 - 1 Glc H -O-Glc PPD Rg3 -Glc 2 - 1 Glc H OH PPD Rg5 - G l c ^ G l c H H PPD Rh2 -Glc H OH PPD Rs l -Glc 2 - 1 AcGlc H -O-Glc 6 - 1 Arap PPD Rs2 -Glc 2 - ] AcGlc H -O-Glc 6- 1 Araf PPD Rs4 - G l c ^ A c G l c H OH PPD Compound K H H -O-Glc PPD aPPT H OH -OH PPT Rb3 H -O-Glc^Rha -OH PPT Re H -0-Glc 2 - ] Rha -O-Glc PPT R f H -0-Glc 2 - ] Glc OH PPT Rg l H -O-Glc -O-Glc PPT Rg2 H -O-Glc OH PPT Rhl H -O-Glc OH PPT Fig .1-1 Chemical structure of ginsenosides and their related compounds Position of sugar moieties has been shown to influence biological activity. For instance, ginsenosides R h l and Rh2 are structurally similar, except for the position of the B-D-glucopyranosyl group. In R h l the sugar is at C-6, and in Rh2, at C-3 as shown in Fig. 1-1. Rh l and Rh2 have different effects on tumor cells, for example, Rh2 induced differentiation of HL-60 cells into morphologically and functionally granulocytes by modulation of P K C isoform levels but Rh l did not (Kim et al., 1998). Rh2, which has the capacity to inhibit the growth and to stimulate melanogenesis in B16 melanoma cells (Odashima et al., 1985; Xia and Han, 1996), causes flattening of the cells cultured in a collagen gel, and leads to organized, non-overlapping monolayers. Cell-to-cell adhesiveness and cell-to-substrate adhesiveness are markedly increased in the B16 melanoma cells treated with Rh2. In contrast, R h l , which appears no effect on cell growth but does stimulate melanogenesis, does not cause morphological changes of the cells or exerts any effect on cell adhesiveness(Odashima et al., 1985; Ota et al., 1987). Rh2 is found to incorporate in the lipid fraction of the B16 melanoma cell membrane, but Rh l is not detected in the lipid fraction of B16 melanoma cells (Ota et al., 1987). It has also been reported that the presence of sugars in PPD and PPT aglycone structures reduces the potency to induce apoptosis and alternately alters membrane integrity (Popovich and Kitts, 2002). Certainly, the same ginsenoside can produce different activities for different tissues and organs. The 20(S)-protopanaxadiol group, represented by R b l , shows effective anti-inflammatory action, obvious vasodilating effect and tranquilizing function to central nervous system; the 20 (S)-protopanaxatriol group, represented by R g l , possesses the properties of exciting the central nervous system, anti-fatigue and hemolysis (Chen, 1996; Jiang et al., 1996; K im et al., 1999b; Shen, 2000). 4. Metabolism of ginseng saponins Biological functions of ginsenosides may be dependent on the way they are administrated. For example, intravenous administration of ginsenoside-Rb2 in syngeneic mice on day 1, 3 or 7 after B16-BL6 melanoma cell inoculation, achieves a remarkable reduction in the number of vessels, but does not cause a significant inhibition of tumor growth. In contrast, intra-tumoral or oral administration of ginsenoside-Rb2 causes a marked inhibition of both neovascularization and tumor growth (Sato et al., 1994). In vitro Rb2 does not affect the growth of rat lung endothelial (RLE) cells, B16-BL6 melanoma cells or various types of murine normal cells (Sato et al., 1994). This suggests that the metabolism of ginseng saponins in organs or tissues might affect their functions. Ginseng is usually taken by oral administration, therefore, the metabolism of ginsenosides in the digestive organs can affect the function of a ginseng product. The time course change of ginsenosides in the blood of healthy men after oral administration of red ginseng extracts, which mainly include R b l , Rb2, Rc, Re, and R g l , is examined based on an enzyme immunoassay (EIA) method combined with high-performance liquid chromatograph (HPLC) analysis developed by Kanaokaet al. (Kanaoka et al., 1992). The result shows that ginsenoside Rb l is not detected in the blood. It can partially be transferred to 20-O-/3-D-glucopyranosyl-20 (S)-protopanaxdiol (20(S)- compound K or IH-901) by Fusobacterium K-60, a ginsenoside Rbl-metabolizing bacterium, which is isolated from human intestinal feces (Hasegawa et al., 1996; Park et al., 2001), or to aglycone protopanaxadiol via ginsenoside Rd by bacterium Eubacterium sp. A-44 (Akao et al., 1998). After ginsenosides R b l and Rb2 are orally administered, they also can be partially transformed to ginsenoside Rg3 in the stomach. When Rg3 is anaerobically incubated with human fecal microflora, all specimens are quickly metabolized to ginsenoside Rh2 and aPPD by different bacteria such as Bacteroides sp., Eubacterium sp., and Bifidobacterium sp. in human fecal microflora as shown by T L C and HPLC analysis (Bae et al., 2000). The main metabolite is ginsenoside Rh2 (Bae et al., 2002a). 20(S)-protopanaxadiol (aPPD) and 20(S)-ginsenoside Rh2 exhibit the most potent cytotoxicity against tumor cell lines (Bae, 2002). When 20(S)-protopanaxadiol ginsenoside Rc is anaerobically incubated with human fecal microflora, it can be metabolized to compound K via Rd and aPPD by different metabolic pathways depending on the intestinal bacteria populations (Bae et al., 2002b). The main metabolite of ginsenoside Rc is compound K (Bae et al., 2002b). In the same experimental process, ginsenoside R g l is converted into 20(S)-protopanaxatriol via ginsenoside Rh l (Hasegawa et al., 1996). In addition, concentration of 20(S)-protopanaxadiol (aPPD), which is an aglycone of Rh2, was found increased in Rh2 treated B16 melanoma cells as a result of deglycosylation of Rh2 (Ota et al., 1991). From these experiments, 20(S)-protopanaxadiol type of ginsenosides: R b l , Rb2, Rg3, Rc and Rd are mainly metabolized to Rh2 and finally transformed to aPPD in intestine or the cells directly or indirectly. 5. Ginsenosides effect on differentiation, invasion, growth arrest and apoptosis of tumor cells Ideally, chemotherapeutic drugs should specifically target only tumor cells and decrease tumor burden by inducing cytotoxic and/or cytostatic effects with minimal collateral damage to normal cells. In reality, chemotherapy has suffered from a range of confounding factors including systemic toxicity due to a lack of specificity, rapid drug metabolism, and hard to avoid drug resistance(Johnstone et al., 2002). Ginsengs have been used for thousands of years, recent studies on individual ginsenoside separated from ginseng identify that some of them have obvious antitumor activity in vivo and in vitro studies (Nakata et al., 1998; Yun TK, 1998; Kim et al., 2003; K im et al., 2004). These ginsenosides may be potentially effective chemotherapeutics for cancer treatment. A prospective cohort study in Korea shows a decreased risk of cancers in various organs with a rise in the frequency of ginseng intake in a dose-dependent manner (Yun TK, 1998). Ginsenoside Rh2 has non- cell specific anticancer potential in vitro or in vivo. Rh2 has been demonstrated to have a cell-growth suppressive effect on various cancer cells, including murine B16 melanoma cells, human ovarian tumor (HRA cell), human hepatoma SK-HEP-1 cells, NTH 3T3 fibroblasts, rat C6 gliomal cells, et al. (Odashima et al., 1985; Ota et al., 1987; Ota et al., 1991; Lee et a l , 1996a; Lee et al., 1996b; Byun et al., 1997; Ota et al., 1997; K i m et a l , 1999a). In vivo oral administration of Rh2 showed potent antitumor effect without adverse side effects in nude mice with H R A human ovary cystadenocarcinoma cells (Nakata et al., 1998). Ginsenosides can induce differentiation of various tumor cells. They may interact with the polar heads of membrane phospholipids and the /3-OH of the cholesterol through their OH groups. And their hydrophobic steroid backbone could intercalate into the hydrophobic interior of the bilayer. Both of these effects may contribute to altering the lipid environment around membrane proteins (Park, 1996). When B16 cells are treated with Rh2, its lipophilic structure makes Rh2 incorporate into the cell membranes of B16 cells. This subsequently changes the cell surface properties and lipid organization of which the functions are related to differentiation (Ota et al., 1987) . Other authors reported that Rh2 induced SMMC-7721 cell (hepatocarcinoma cell line) differentiation and growth arrest activation were related to reduced telomerase activity by affecting transcription levels of human telomerase reverse transcriptase (Zeng and Tu, 2003). However, ginsenosides R h l and Rh2 cause the differentiation of F9 teratocarcinoma stem cells seems to be exerted via binding with a glucocorticoid receptor (GR) or its analogous nuclear receptor (Lee et al., 1996b). Ginseng and ginsenosides inhibit tumor angiogenesis and metastasis (Sato et al., 1994). Except for inhibition of neovascularization, tumor growth, and lung metastasis in syngeneic mice after B16-BL6 melanoma cell inoculation (Sato et al., 1994), ginsenoside-Rb2 also inhibits the invasiveness of human uterine endometrial cancer cells of H H U A and H E C - l - A cell via matrix metalloproteinase (MMP)-2 suppression (Fujimoto et al., 2001), therefore it may be used for inhibition of secondary spreading of uterine endometrial cancers (Fujimoto et al., 2001). The 20(R)- and 20(S)-ginsenoside-Rg3 from red ginseng also shows a significant inhibition on lung metastasis produced by two highly metastatic tumor cells, B16-BL6 melanoma and colon 26-M3.1 carcinoma, in syngeneic mice by consecutive intravenous administrations or oral administrations (Mochizuki et al., 1995). The mechanism of their antimetastatic effect is related to inhibition of the adhesion and invasion of tumor cells, and also anti-angiogenesis activity (Mochizuki et al., 1995). Using a cell monolayer invasion model, shows Rg3 significantly inhibited adhesion and invasion of B16-BL6 melanoma (Mochizuki et al., 1995; Shinkai et al., 1996). In addition, inhibition of tumor invasion of Rg3 has been verified in different tumor cell lines including rat ascites hepatoma cells (MM1), B16FE7 melanoma cells, human small cell lung carcinoma (OC10), and human pancreatic adenocarcinoma (PSN-1) cells, when examined in a cell monolayer invasion model (Mochizuki et al., 1995; Shinkai et al., 1996). Although more than 35 kinds of ginsenosides have been isolated from ginseng, only afew of them appear to inhibit tumor growth. It is believed that ginsenosides induced tumor growth inhibition and /or apoptosis may involve different signaling pathways: a. Ginsenosides inhibit cell growth by activating caspase-3 In vitro studies suggest that ginsenosides and their metabolites including Rh2, Rg3, and DH-901 (compound K) suppress tumor cell growth by apoptosis and they can activate caspases-3 and -8 by different pathways (Park et al., 1996; K im et al., 1999a; Liu et al., 2000; Fei et al., 2002). Rh2-induced tumor cell-death in human malignant melanoma, A375-S2 cells, C6 cell, A375-S2 cells and SK-HEP-cells is partially dependent on caspase-8 activated caspase-3 pathway in a Bcl-XL-independent manner (Park et al., 1996; K i m et al., 1999a; Fei et al., 2002). However, IH-901 (compound K) inhibits proliferation and induces apoptosis of Human HL-60 cells and human hepatoblastoma HepG2 cell death through activation of caspase-3 protease, which occurs via mitochondrial cytochrome c release independently of Bcl-2 modulation (Lee et al., 2000; Oh SH, 2004). Caspase activation was a necessary requirement for apoptosis induced by TH-901 because caspase inhibitors significantly inhibited IH-901-induced apoptosis (Oh SH, 2004). On the other hand, cell-cycle progression is regulated by two protein classes, the cyclins and their kinase partners, the cyclin-dependent kinases (CDKs) (Swanton, 2004). CDKs are serine/threonine kinases that become active only when associated with a regulatory partner (e.g. cyclins or other proteins). CDK/cyclin holoenzymes are activated through phosphorylation catalyzed by CDK-activator kinase (CAK). The activity of CDKs is negatively regulated by direct interactions with proteins referred to as C D K inhibitors (CDKIs) such as p21 c i p l / w a f l and p27 K i p l(Dai and Grant, 2003). p27 K i p l leads to cell-cycle arrest at G l /S phase (Polyak K et al., 1994). Ginsenoside Rh2 arrests the SK-HEP-1 cell cycle at the G l /S transition phase by selectively inducing expression of p27 K i p l and, as a consequence, down-regulating C D K (Lee et al., 1996a; Ota et al., 1997). It was reported that Rh2 induced apoptosis in human hepatoma cells was due to activation of cyclin A-CDK2 by caspase 3-mediated cleavage of p 2 1 W A F 1 / C I P 1 (Jin et al., 2000). CDK2 has been viewed as a key cell cycle regulator that is essential for S phase progression. A target molecule that may account for the chemoprevention effects of ginsenoside Rh 2 is CDK2 (Ota et al., 1997). Similarly, ginsenoside Rg3, Rs4 and Rs3 up-regulate the expression of Cyclin-kinase inhibitors, p21 and p27, arrested LNCap and cells SK-HEP-1 cells at GI phase, and subsequently inhibited cell growth through a caspase-3-mediated apoptosis mechanism (Attele et al., 1999; Kim et al., 1999c, d; Liu et al., 2000). b. Rh2 induces persistent JNK activation resulting in apoptosis The c-Jun N-terminal kinase (INK), also known as stress-activated protein kinase (SAPK), belongs to the mitogen-activated protein kinase (MAPK) family. In general, the JNKs mediate intracellular signals that control a variety of cellular functions, including cell differentiation and apoptosis (Xia et a l , 1995; Herdegen et al., 1997; Schaeffer and Weber, 1999). Transient activation of I N K promotes cell survival, whereas persistent INK activation induces apoptosis (Chang, 1999; Kobayashi and Tsukamoto, 2001). I N K activation has been found to be an essential component of the apoptotic process in a number of systems (Jarpe et al., 1998; Huang et al., 1999; Wang et al., 1999a). Rh2 induced early and later phase of JNK1 activation in the human hepatoma SK-HEP-1 cells. The early activation of JNK1 maybe associated with a protective role for cell survival until the induced cells commit to apoptosis, whereas the later activation is linked to execution of apoptotic progression (Ham et al., 2003b) c. Other death signaling pathways induced by ginsenosides Other apoptotic pathways involved include inhibiting phospholipase C by ginsenoside Rh2, which blocks generation of second messengers necessary for the activation of protein kinase C (Byun et al., 1997) . The apoptosis induced by this pathway does not seem to involve Bcl-2, Bcl-xL or Bax in Rat glioma C6Bu-l cells (Kim et al., 1999e). Additionally, the 20(S)-protopanaxatriol (aPPT), ginsenoside Rh2, and compound K greatly enhanced the cytotoxicity of the anti-cancer drugs in multi-drug resistant P388/ADM cells (Hasegawa et a l , 1995). Rg3 increased the life span in mice implanted with adriamycin (DOX)-resistant murine leukemia P388 cells in vivo and inhibited body weight loss significantly (Kim et al., 2003). 6. Aglycones of Ginsenosides A number of studies demonstrated that ginsenosides Rh2 perturbed normal cell cycle events and induced apoptosis on different cancer cells (Lee et a l , 1996a; Park et al., 1997; Liu et al., 2000), however, the trace amounts of Rh2 in regular ginseng extracts and the high cost of extraction of Rh2 are prohibitive for its application. As shown in the Fig. 1-2, ginsenosides R b l , Rb2, and Rc can be converted to different 20(S)-protopanaxadiols via hydrolysis, epimerization and hydration in the side-chain of the aglycone moiety of them under an improved alkaline cleavage procedure (Cui et al., 1993), mild acidic treatment (pH 3.0-3.5) (Cui et al., 1996; Bae, 2002), destabilization in tumor cells (Ota et al., 1991) or digestive system (Hasegawa et al., 1996; Park et al., 2001; Hasegawa, 2004). aPPD are the final products of all ginsenosides in the above reactions (Ota et al., 1991; Bae, 2002), which is also the focus of my study. The amount of aPPD in ginseng is undetectable, but it can be produced by different destabilization methods from all sources of protopanaxadiol type ginsenosides (Fig. 1-2 and 1-3) (Hasegawa et al., 1996; Park et al., 2001; Bae, 2002). I Rg3 VI. Rd À • i f Rh2 •v* Compound K ' an 901) Fig. 1-2 Schematic metabolism routes of 20(S)-protopanaxadiol type ginsenosides ® , (2), and (3) represent different reaction conditions. ® : mild acidic treatment or in human stomach (2): bacteria isolated from human fecal microflora © : human intestine Glc-Glc-GioGlc-< Glc-Glc-O i 1 3 -0-[/?-D-glucopyranosyl-( 1 -*£)-/?-D-gluco pyranosyl] -20-O-[/?-D-glucopyranosyl ( 1 — • 6) -/?-D-glucopyranosyl]-20(iS)-protopanaxadiol 3-0-[/?-D-glucopyranosyl-(l—•2)-/?-D-gluco pyranosyl]-20(5)-protopanaxadiol G l c -(3-O-/?-D-glucopyranosyl-20(,S)-protopanaxadiol) (20(5)-protopanaxadiol) aPPD Fig. 1-3 Structural relation among 20(S)-protopanaxadiol type ginsenosides and aglycone Since ginsenosides R b l , Rb2, Rc, Rd, and Re are the major constituents of ginseng especially in American ginseng extracts (Wang et al., 1999b; Assinewe et al., 2002) and total ginsenosides in ginseng can be as high as 20%(W/W) (Huang K C , 1999; Assinewe et al., 2003), so high yield of aPPD can be obtained in relatively lower costs. A specially formulated ginseng product, Careseng®, mainly contains aglycone ginsenoside such as aPPD and aPPT, has been tested on different cultured tumor cell lines and tumor animal models. The results showed that in vitro, it caused apoptosis in dose- and time- dependent manner and related to caspase activation in different tumor cell lines (Jia et al., 2003). Most importantly, cytotoxicity of aPPD seems identical to Rh2 or stronger on cancer cells (Popovich and Kitts, 2002). This may be explained by the observations that Rh2 was rapidly metabolized into aPPD in cells (Ota et al., 1991; Bae, 2002). Relatively low costs of production of aPPD but equally potent tumor-inhibiting effects as Rh2 renders further investigation on this compound for potential development as an anti-cancer drug. In the present study, mechanisms of aPPD-induced apoptosis were investigated in glioma cell lines from different aspects. They include verification of apoptotic nature of aPPD-induced cell death, activation of caspase cascade in apoptosis of aPPD treated glioma cells, involvement of JNK activation, superoxide anion production and Akt survival pathway in aPPD-induced apoptosis as shown in Fig. 1-4. Growth factor Death receptor M A P K Integrin Growth factor receptor-(P^ T R A I L — • T R A D D F A D D aspase-8 Hiid ROS (O 2 ,H 2 O 2 ) J N K C-Jun<P) AP-1 p53 Proteins Lipids DNA Cyto-C ' se-9 Caspase-3, -6 and -7 PTEN NFkB BAD p53 Apoptosis Fig. 1-4 Pathways of related to apoptosis induced by aPPD CHAPTER 2 MATERIALS AND METHODS 1. Equipments The C 0 2 water-jacketed incubator was manufactured by NuAir (Plymouth, MN). The inverted microscope with phase contrast and fluorescent filters (Axiovert 200 microscope) equipped with software Northern Eclipse was made by Carl Zeiss light microscope (Germany). The Mini-Protein II System for protein electrophoresis and the Semi-Dry /Wet Transfer Cell for protein blotting were purchased from Bio-Rad Laboratories (Mississauga, ON), p-Quant OD Reader with software KC4 2.6 Rev 3 was from Bio-TEK Instruments, INC. (Winooski, USA). The Eppendorf centrifuge 5415D was made by Brinkmann Instruments, Inc (N.Y., USA), the ultracentrifuge was an Optima™ TLX Ultracentrifuge made by Beckman (Germany), the 4°C Microtube centrifuge was Biofuga Fresco made by Kendro Laboratory Products (Germany). The Image Station 440 CF and the software Kodak ID 3.5 for western blotting visulization and analysis were purchased from Eastman Kodak Company (USA), the BD FACS Calibur (Becton-Dickinson, USA) equipped with the software CellQuest for flow cytometry analysis was made by Becton Dickinson (Mountain View, CA.). The tissue culture dishes: 10cm dishes, 6 well plates and 96 well plates were manufactured by Corning (Corning, N . Y., USA). 2. Reagents 20(S)-protopanaxadiol (aPPD) was kindly provided by Pegasus Pharmaceuticals Inc. (Vancouver, BC). ApopTag® Plus Peroxidase In Situ Apoptosis Detection Kit was purchased from Chemicon International, Inc.(Temecula, CA). Antibodies for anti-caspase-8 polyclonal antibody, anti-caspase-3 (CPP32) antibody, purified mouse anti-human caspase-7 monoclonal antibody, and purified mouse anti-cytochrome c monoclonal antibody were purchased from BD Biosciences (Mississauga, ON). Z-DEVD-FMK (caspase-3 inhibitor), Z-IETD-FMK (caspase-8 inhibitor), Z-LEHD-FMK (Caspase-9 Inhibitor), Z-VAD-FMK (General caspase inhibitor), and BD ApoAlert™ Mitochondrial Membrane Senser Kit were also purchased from BD Biosciences (Mississauga, ON). Purified mouse anti-human caspas-9 antibody was purchased from Calbiochem (San Diego, CA). Phosho-SAPK/JNK (Thrl83/Tyrl85) antibody, SPRK/JNK antibody, Phospho-Akt (Ser473), and Akt antibody were purchased from Cell Signalling Technology, Inc. (Mississauga, ON). INK inhibitor I (L)-form (cell permeable), INK inhibitor I (L)-form (cell permeable), Negative control I, and INK inhibitor TJ (SP60025) were purchased from Calbiochem (San Diego, CA). Dihydroethidium (hydroethidine) was purchased from Molecular Probes (Leiden, The Netherlands). Protease inhibitor cocktails were obtained from Roche Molecular Biochemicals (Mannheim, Germany). Modified DC Protein Assay Kit and nitrocellulose membrane were purchased from BIO-RAD (USA). TNF-a, cycloheximide and most of the chemicals were purchased from Sigma. 3. Cell cultures SF188 cells (Brain Tumor Research Center, UCSF, San Francisco, CA), U87 cells (American Type Culture Collection (ATCC), Manassas, USA), SF210 and U138 glioma cells, which were gracious gifts of Dr. Shoukat Dedhar (Cancer Research Center, Vancouver BC), and MCF-7, MCF-7 c3 and MCF-7 vec that were from Dr. Nancy L. Oleonick (Case Western Reserve University School of Medicine, Cleveland, Ohio) were cultured in D M E M (Dulbecco's modified Eagle's medium, GIBCO BRL) supplemented with 10% fetal bovine serum (FBS) (GIBCO BRL), lOOunits /ml penicillin and 100ug/ml streptomycin at 37°C in a humidified atmosphere with 5% CO2 and were fed every 3-4 days. 4. Preparation of ginsenoside solutions and treatment Stocking solutions of 50mg/ml of aPPD (108mM) was prepared in 100% ethanol. aPPD was diluted in D M E M supplemented with 2% fetal bovine serum (final ethanol concentration 0.25%(v/v)) immediately before each experiment. For time-course studies, the cells were treated with 108.4uM of aPPD (50ug/ml). The treatments were ended at selected time-points by adding MTT (3-(4,5-dimethythinzol -2yl) -2,5-diphenyl tetrazolium bromide (Sigma) for viability experiments or adding lx sample buffer (62.5mMTris-HCl. pH6.8 at 25°C, 2%SDS, 10%glycerol, 50mM DTT, 0,01%(W7V) bromo-phenol blue) for western blotting assay. For dose-response experiments, cell cultures were treated with different concentrations of aPPD for 5 to 9 hours followed by adding lx sample buffer for western blot assay. If viability was to be measured, cells were typically treated with the agents for 24 hours or specially indicated before adding 0.5mg/ml MTT. 5. MTT assay Viability of cultured cells was determined with MTT assay, based on MTT reduction by viable cells (Korting et al., 1994). Cells were seeded in 96 well plates in a concentration of 2.5X104/well one day prior to experiments. Cell cultures were incubated with 2% FBS culture medium containing aPPD of indicated concentrations for various periods. The cell culture medium was then removed and lOOul of 0.5mg/ml of MTT in serum-free medium was added in each well. Following incubating the cells at 37°C and 5% CO2 incubator (NuAir, Plymouth, MN) for 4 hours, lOOul of the lyses buffer (50 V/V N , N -dimethylformamide (Sigma, USA), 20% SDS (BIO-RAD, USA) and 0.4% V/V glacial acetic acid in distill water, pH 4.8) was added and the plates were incubated at 37°C in a humidified atmosphere with 5% CO2 incubator overnight. The optical density of each well was read at 595nm absorbance in a microplate reader (BIO-TEK, USA). 6. Hoechst 33342 staining and morphological analysis Cells were seeded in 6-well plates or 35mm plates 24 hours prior to experiment to allow 80% confluency on the day of treatment of aPPD or other agents. The Hoechst 33342 (Sigma) was dissolved in distilled water to make a 2mM stocking solution and stored at 4°C in the dark. After treatment, the DMEM medium was removed and 2ml of medium containing 2uM Hoechst 33342 dye was applied to each well followed by incubation at RT for 1-2 min. The dye was then removed and fresh medium was added to the cultures. The cells were visualized under fluorescent microscope (Zeiss, Oberkochen, Germany) at UV illumination (380nm) and the micrographs were taken with Northern Eclipse software (Germany). 7. DNA ladder SF188 cells were plated in 10cm dishes to reach 90% confluency before adding aPPD of various concentrations. The cells were scraped 5 hours after the treatment with plastic scrapers, and collected by centrifugation at 1500 rpm for 5min at RT. After washing the cells with PBS once, the cells were lyzed in 1ml of fresh-prepared cell lyses buffer (lOmM Tris.Cl, lOmM EDTA, and 5% Triton-100). The cell lysates were rotated at 4°C for 15min and centrifuged at 10,000rpm for 25min at RT. After the supernatant was treated with RNase A at 37°C for 80min, 140 ul of 10% SDS, 75ul of 20 mg/ml of proteinase K were added and the mixture was incubated at 50°C for 2 hours. After DNA was extracted with phenol and chloroform once each, lOOul of 3M sodium acetate and about 2.5 ml of ethanol dehydrate were added in each tube, and the DNA was precipitated at -20°C overnight. After centrifuging at 13.000rpm 4°C for 40min, equal amount of DNA in each well was analyzed with 2% gel electrophoresis fractionation at 50 vol. and 4°C. The gel was then examined under U V light with ethidium bromide staining for visualization and photographed with software Kodak ID 3.5 (Eastman Kodak Company, USA). 8. DNA Staining with ApopTag® The SF188 and U87MG cells were seed on coverlips 24 hours followed by 3 or 7 hours treatment with aPPD of various concentrations. The ApopTag® Peroxidase in situ Apoptosis Detection Kit (CHEMICON International, Inc. Temecula, CA) was used according to the instruction provided by the manufacturer to detect DNA strand breaks as biochemical markers of apoptosis. The slides were fixed in 1 % paraformaldehyde in PBS, pH 7.4 for 10 min. at RT. After two 5-minute washes in PBS, the cells were permeabilized in precooled ethanol: acetic acid 2:1 for 5 min. at -20°C and quenched in 3.0% hydrogen peroxide in PBS for 5 min at RT to inactivate endogenous peroxidase. Following incubating in equilibration buffer for 5min at RT, incubate the slides with working strength TdT enzyme in a humidified chamber at 37°C for 1 hour and the reaction was stopped with working strength stop/wash buffer for 10 min. at RT. After three 1-minute washes in PBS, the slides were incubated with anti-digoxigenin peroxidase conjugate in a humidified chamber for 30 min. Following 4x2-minute washes in PBS at RT, the reaction products were developed in DAB peroxidase substrate for 15 minutes at RT. In order to determine the optimal staining time, color development was monitored under the microscope. The reaction was stopped by rinsing the slides in distilled water and the slides were counterstained with 0.5% (w:v) methyl green for 5minutes at RT. After rinsing in distilled water and 100% 1-butanol for three times separately and dehydrating three 2-minutes through xylene, the counterstained slides were mounted with Permount and viewed under microscope (Zeiss, Oberkochen, Germany). Cells were considered apoptotic if they exhibited brown staining and characteristic apoptotic morphology (cell shrinkage, pyknotic nuclei, cell blebbing/apoptotic bodies). 9. Apoalert™ mitochondrial membrane permeability assay MitoSensor reagent (manufacturer) is a cationic dye with green fluorescence in its monomelic form. When it is taken up by the mitochondria, it forms aggregates with intense red fluorescence. MitoSensor reagent cannot aggregate in the mitochondria of apoptotic cells because of altered mitochondria membrane potentials. As a result, the dye remains in monomeric form in the cytoplasm, where it fluoresces only green. Thus, normal cells show red and green fluorescence while cells with permeabilized mitochondrial membrane only exhibit green fluorescence. The mitochondrial membrane permeability assay protocol was as described in the instruction provided by the manufacture. Cells grown in 6-well plates were rinsed with serum-free medium. Immediately prior to use, lui of BD MitoSensor Reagent was added to 1 ml of Incubation Buffer (provided by BD Biosciences), vortexed and centrifuged to remove possible aggregates. One milliliter of above reagent was applied in each well followed by incubation at 37°C in a 5% CO2 incubator for 20 min. Then 3 ml of fresh serum-free medium were added to the cells, and the cells were photographed with a fluoresce microscope using a fluorescein (488 nm) or rhodamine (568 nm) filter. 10. Extraction of subcellular fractions and mitochondria Subcellular fractions were obtained by a series of centrifugation as described by Gross et al (Gross et a l , 1998; Gross et a l , 1999) with some modifications. aPPD-treated or -untreated SF188 cells were collected by scraping with a plastic scraper and washed in ice-cold PBS once, resuspended in three volumes of isotonic HIM buffer (HIM buffer: 200mM mannitol, 70mM sucrose, ImM EGTA, and lOmM HEPES PH 7.5) with a mixture of protease inhibitors (Complete Protease Cocktail, Roche, added at 1:500 dilution) and kept on ice for 20min. Cells were homogenized with a tight glass homogenizer to keep more than 85% of cell broken and the nuclei intact under the microscope monitoring and spun at 1000 x g for 10 min at 4°C. Pellets containing large cell debris, unbroken cells and nuclei were discarded. The supernatant was spun at 10,000x g for 20 min at 4°C to obtain heavy membrane pellet mainly including mitochondria. The pellet was washed with HIM buffer once and resuspended in lOOul HIM buffer. The post-mitochondria supernatant was spun at 100,000x g for 1 hour at 4°C to collect cytosol supernatant and light membrane pellet including the endoplasmic reticulum and plasma membrane (light membrane system). Protein concentration was determined by Modified DC Protein Assay (Bio-Rad, USA). The mitochondria were used for cytochrome c release experiments. The cytosol and light membrane used for cell-free assay to detect caspase activation was incubated with aPPD, cytochrome c or dATP as described previously. The cytosol samples also were separated on 12% SDS-polyacrylamide gels, blotted on nitrocellulose membrane, and probed with an antibody against cytochrome c (BD Biosciences). 11. Cytochrome c -depleted mitochondria and cytochrome c release assay Cytochrome c depleted mitochondria were prepared as described by M . Marita et al (Narita et a l , 1998). Briefly, 2mg/ml of isolated mitochondria suspended in a hypotonic buffer consisting of 2mM HEPES with lOmM of KC1 or in HIM buffer as a control were incubated on ice for 20min and centrifuged to collect the supernatant. To deplete the mitochondria, 150mM KC1 in 2 mM HEPES was used to wash the mitochondria twice and then resuspended in HIM Buffer. For aPPD-induced cytochrome c release, aPPD or vehicle was added to mitochondria in HIM buffer and incubated on ice for 20 min to collect treated supernatant and mitochondria followed by Western blotting to measure the levels of cytochrome c in both fractions. 12. Protein extraction and western blotting analysis Cells were treated with aPPD of various concentrations or control vehicle in 2% FBS D M E M for different time as designed. Both detached and attached cells were collected by scraping with scrapers followed by centrifugation. After wash with ice-cold PBS, the cells were lyzed by adding lx sample buffer (62.5mMTris-HCl. pH6.8 at 25°C, 2%SDS, 10%glycerol, 50mM DTT, 0,01%(W/V) bromo-phenol blue) and heated to 95-100°C for 5 minutes. Protein concentration was measured using modified DC Protein Assay (BIO-RAD) similar to Lowry assay (Lowry et al., 1951). Equal weight of protein (50-100ug/lane in different experiments) from each sample was taken for immunoblot analysis. The protein was size-fractionated with 10% SDS-polyaccarylamide gel electrophoresis for caspase-8, INK, Phospho-JNK, Akt or Phospho-Akt, and 12% gel for Caspase-3, -7, -9, and cytochrome c followed by electroblotting onto nitrocellulose membranes using semidry transfer system (BIO-RAD) for caspases and cytochrome c or using wet transfer system for INK, Phospho-JNK, Akt and Phospho-Akt. The membranes were blocked in 5% non-fat milk (BIO-RAD, USA) for 1 hour at room temperature (RT), and probed with primary antibodies at 4°C overnight. Antibodies for caspase-3, -8, -9, cytochrome c, INK and Akt were 1:1000 diluted in PBS buffer with 1% bovine serum albumin (BSA) and 0.1% sodium azide; antibodies for caspase-7, Phospho-JNK and Phospho-Akt were 1:500 diluted in PBS buffer with 1% BSA and 0.1% sodium azide. After washed with PBST (phosphate buffered saline with 0.05% Tween 20) for 3 times, the membrane was incubated with 1:5000 of HRP-conjugated anti-mouse or anti-rabbit IgG antibody (horseradish peroxidase, PerkinElmer Life Sciences, Boston MA) for 2 hours at room temperature, then the reaction products were detected with enhanced chemiluminescence substrate (PerkinElmer Life Sciences, Boston MA) and visualized under Kodak image system (Kodak, USA). Data were analyzed with Kodak ID 3.5 software. 13. Cell-free system for caspase-3 activation assay Based on Liu et al (Liu et a l , 1996), cell-free system was used to detect caspase-3 activation in cytosol fraction and light membrane system (including endoplasmic reticulum and plasma membrane) of SF188 which were extracted from untreated SF188 cells as described in section 10 of this chapter. 50ug cytosol protein for each experiment was applied and incubated with ImM dATP, lOuM cytochrome c, lOug of cell membrane protein in the presence or absence of 108uM aPPD. The mixtures were incubated at 30°C for 2 hours, and then western blotting assay was used to detect caspase-3 activation. 14. Caspase and JNK inhibition assays Cells were grown in 96-well plates overnight before each experiment. Caspase inhibitors (BD Biosciences, Mississauga, ON) were dissolved in DMSO to make a lOmM stocking solution. JNK inhibitor I and its negative control were dissolved in distill water for stock solutions (lOmM) and stored at -20°C. 4mg/ml of JNK inhibitor II was dissolved in DMSO (based on manufacturer) to make a stocking solution. Before use, the stock solutions were diluted with D M E M with 2% FBS to designed concentrations. Cells were pretreated with lOOul of inhibitor for 2 hours at 37°C, in 5% C 0 2 incubator, followed by adding aPPD of various concentrations. Cell viability was determined with MTT assay as described earlier in this chapter. 15. Superoxide anion staining assay Hydroethdine (HE) is a dye specifically staining for superoxide anion (Ye et al., 1999). HE was dissolved in DMSO to make a 20mM stock solution and stored at -20°C in the dark. Cells were seeded in 6-well plates 24 hours before experiments. The cells were then treated with aPPD for 2 hours. It was followed by adding 2ml 5uM HE in PBS in each well. The cells were incubated for another 20min at 37°C and harvested with a plastic scraper followed by centrifugation at 3400 RPM and washed twice with PBS. The cells were resuspended in 500ul of PBS for analysis with flow cytometry (excitation, 480nM; emission, 567nM). 16. Statistical analysis Data were obtained from at least two independent experiments. Triplicates or more were performed in each experiment. The data were statistically analyzed for significance by one-tailed Student's r-test. Difference with P-values <0.01 was regarded as statistically significant and indicated by a * in figures. CHAPTER 3 EFFECTS OF CASPASES ON aPPD-INDUCED C E L L DEATH IN CELLS WITH DIFFERENT p53 AND PTEN STATUS INTRODUCTION Tumorigenesis is a multistep process (Soria et a l , 2003) in which mutations in key cellular genes produce a series of acquired capabilities. The most important among them is to evade apoptosis for mutated cells, which is also an essential "hallmark of cancer" (Hanahan and Weinberg, 2000). Apoptosis was first described by Kerr et al in 1972 (Kerr et al , 1972), however, after 20 years, in the 1990s due to molecular identification of the key components including caspases (cyteine aspartate-specific proteases) (Yuan et a l , 1993) and the Bcl-2 family proteins (Tsujimoto et a l , 1984; Bakhshi et a l , 1985; Cleary et a l , 1986), the mechanism of apoptosis began to be understood at the molecular level. Apoptosis or programmed cell death (PCD) is a gene-directed mechanism activated as a suicidal event to eliminate excess, damaged, or infected cells. Characteristic features of apoptosis include chromatin condensation, membrane blebbing, and DNA fragmentation, which are features that generally distinguish cells committed to apoptosis from other types of death such as necrosis (Kerr, 1971; Kerr et a l , 1972; Wyllie et al , 1980). Major advances have been made toward understanding the molecular mechanism of programmed cell death (Chinnaiyan A M , 1996; Hengartner, 2000; Leist and Jaattela, 2001). The faithful and central executors of apoptosis are a family of cyteine aspartate-specific proteases, termed caspases (Salvesen and Abrams, 2004). Caspases are implicated in apoptosis with the discovery that Caenorhabditis elegans (C. elegans) cell death gene-3 (CED-3), the product of a gene required for cell death in the nematode Caenorhabditis elegans, is related to mammalian interleukin-lP-converting enzyme (ICE or caspase-1) (Thornberry et a l , 1992; Yuan et a l , 1993). Caspase-1 itself has been identified to have no effect on the apoptosis mechanism (Martinon and Tschopp, 2004). To date, the caspase gene family contains 14 members in mammalian species, Human have 11 of these enzymes (Thornberry and Lazebnik, 1998). Members of caspase family share similarities in amino acid sequence, structure, and substrate specificity. They are all initially expressed as proenzymes (30 to 50 kD) that contain three domains: an NH2-terminal prodomain, a large subunit (-20 kD), and a small subunit (-10 kD) (Nicholson and Thornberry, 1997; Fulda et al., 2001). Based on phylogenetic analysis, this family is composed of two major subfamilies that are related to either ICE (caspase-1; inflammation group) or the mammalian counterparts of CED-3 (apoptosis group) (Nicholson, 1999; Martinon and Tschopp, 2004). Caspases (apoptosis group) maybe subclassified as initiators (e.g., caspase-8, -10, -2, or -9) or effectors, also known as executioners (e.g., caspase-3, -6, or -7), based on whether they have a large or small prodomain (Thornberry and Lazebnik, 1998; Nicholson, 1999). Caspases are among the most specific proteases, with an unusual and absolute requirement for cleavage after aspartic acid (Asp) residues (Stennicke and Salvesen, 1998). Procaspases are activated to fully functional caspases by two cleavage events. The first proteolytic cleavage divides the chain into large and small caspase subunits, and a second cleavage removes the N-terminal prodomain (Ramage et al., 1995; L i et al., 1997a; Zimmermann et al., 2001). The active caspase is a tetramer of two large and two small subunits (Wolf and Green, 1999). There are two well-characterized caspase-activation pathways which involve apical caspases. One is the extrinsic pathway initiated from the cell surface death receptor. Here, the death signal is transmitted through binding of an extracellular death ligand such as tumor necrosis factor (TNF) to its cognate receptor, the TNF receptor (TNFR). The death receptors transmit signals to the interior of the cells, where the apical proteases of the extrinsic pathway, caspases-2, -8 and -10 are recruited (Ashkenazi and Dixit, 1998) and activated. The other is triggered by changes in mitochondrial integrity as a consequence of cellular stress, and is mediated by cytochrome c in the mitochondrion, which leads to caspase-9 activation (Green and Reed, 1998) and then cleave downstream effector caspases-3, -6, and -7 (Hengartner, 2000). Active caspase-3 is capable of cleaving and activating pro-caspase-8, which engages the extrinsic cascade, amplifying and speeding up the apoptotic process. Apoptosis is initiated either by recruitment of caspase-8 (Walczak and Sprick, 2001) into the death inducing signaling complex (DISC) (Martin et a l , 1998; Muzio et a l , 1998), which consists of the receptor, FADD (Fas-associated death domain) as an adaptor and caspase-8 as an initiator caspase (Muzio et a l , 1998), to activate death receptors or by activation of caspase-9 to form apoptosome in the case of mitochondrion dependent events (Zimmermann et a l , 2001). It is thought that the initiator caspases can undergo self-activation to start the caspase cascade. Although the precise mechanism for the activation of initiator caspases are still not clear, abundant evidence suggests that oligomerization of caspase-8 and caspase-9 leads to their processing to form active caspases containing two large and two small subunits of the protease domain (MacCorkle et a l , 1998; Srinivasula et a l , 1998; Yang et a l , 1998; Salvesen and Dixit, 1999). This could be achieved by recruitment of two or more procaspases into the DISC or apoptosome (Annunziato et a l , 2003; Aouad et a l , 2004). After caspase activation, they are capable of cleaving a vast array of proteins at specific consensus sites (Nicholson and Thornberry, 1997; Green, 1998; Hengartner, 2000). The execution phase of apoptosis is produced by the caspase-dependent cleavage of hundreds of cellular proteins (Fischer et a l , 2003) . To date, more than 280 caspase substrates including themselves have been identified, these substrates are generally categorized as proteins involved in the scaffolding of the cytoplasm and nucleus, cell cycle progression and DNA repair-related proteins (Stroh and Schulze-Osthoff, 1998; Fischer et al., 2003). Some of identified caspase substrates are protein kinases or other proteins involved in signal transduction. For instance, proteolytic activation of the PAK2 (p21-activated protein kinase) (Bagrodia, 1999) has been reported during CD95 and TNF-mediated apoptosis (Rudel and Bokoch, 1997). As PAK2 is able to trigger stress-activated kinases of the INK pathway (c-Jun NH2-terminal kinases/ stress-activated protein kinases) (Rudel et al., 1998), this may provide a link between caspases and INK activation during apoptosis signaling. The p53 protein is a major target for mutational inactivation in human cancer, and represents a major difference between normal cells and cancer cells. The p53 tumor suppressor protein is a nuclear phosphoprotein that can bind to DNA in a sequence-specific manner to activate gene expression in response to DNA damage, elicits either cell cycle arrest to allow DNA be repaired or apoptosis if the damage is excessive (Vogelstein et al., 2000). Indeed, it is now clear that the functioning of the p53 DNA damage signaling pathway is lost in most, if not all, human cancer (Hollstein et al., 1991; Levine, 1997). For example, p53 is mutated in nearly one-third to half of glioblastoma patients (Chen et al., 1995; Kleihues and Ohgaki, 1999; Kleihues et al., 2002), which is the most common malignant brain tumor of adults, and is among the most lethal of all cancers (Kleihues et al., 2002; Mischel and Cloughesy, 2003). Mutation and consequent loss of p53 function has been shown to play a crucial role in tumorigenesis and development of resistance to therapeutic regimens in gliomas and other human malignancies (Iwadate Y et al., 1996). Transient or irreversible p53-mediated cell cycle arrest in the transition from GI to S phase is mediated by the transcriptional activation of the cyclin-dependent kinase (CDK) inhibitor p21WAFl/CTPl (El-Deiry et al., 1993; Harper et al., 1993; Vogelstein et al., 2000). Additionally, the extrinsic and intrinsic pathways appear to cooperate in p53-induced apoptosis. p53 promotes the expression of a number of genes that are involved in apoptosis, including those encoding death receptors (Muller et a l , 1998) and proapoptotic members of the Bcl-2 family (Miyashita and Reed, 1995; Oda et a l , 2000). Two targets of p53, which engage the extrinsic pathway, are FAS (CD95 or APO-1) (Owen-Schaub et a l , 1995) and KILLER/DR5 (death receptor 5) (Takimoto and El-Deiry, 2000; Wu et a l , 2000). It is thought that the p53-induced up-regulation in the levels of these death receptors results in their oligomerization and activation in the absence of their respective ligands. Both FAS and KILLER/DR5 are capable of inducing apoptosis when overexpressed in cells in the absence of additional ligand. In most cases, p53-induced apoptosis proceeds through mitochondrial release of cytochrome c, which leads to caspase activation (Schuler et a l , 2000). Additionally, p53 directly activates the proapoptotic Bcl-2 protein Bax in the absence of other proteins to permeabilize mitochondria and results in DNA damage, the mitochondrial release of cytochrome c, caspase activation, and apoptosis (Chipuk et a l , 2004). The most commonly used cancer therapeutics eliminates tumors by inducing apoptosis and inhibiting proliferation in sensitive tumor cells (Fisher, 1994). The mechanism for chemotherapy-induced apoptosis might be damage to DNA, lipid components of cell membranes, and cellular proteins causing an imbalance of the cellular homeostasis (Herr and Debatin, 2001). Caspases play an important role as effector molecules in apoptosis including cytotoxic therapy-induced cell death. Although the cellular targets for different cytotoxic agents are diverse, most signaling pathways activated by anticancer drugs ultimately result in activation of caspases (Kaufmann, 1989; Fisher, 1994; Herr and Debatin, 2001). Caspase-mediated apoptosis is the principal program of cell death in many developmental and physiological settings (Kerr et a l , 1972; Strasser and Dixit, 2000; Kaufmann and Hengartner, 2001), however, it is not the single executive effector of cell-death . Recent data indicate that programmed cell death can occur in the complete absence of caspases (Leist and Jaattela, 2001). To date, the cellular components of caspase-independent apoptosis are not identified. Although activation of caspases is ultimately produced in extrinsic and intrinsic apoptotic systems, z-VAD-fmk (general caspase inhibitor) does not block apoptosis from mitochondrial changes, such as loss of the membrane potential, production of ROS, or the release of apoptogenic factors such as cytochrome C and apoptosis inducing factor (AD?) (Green and Reed, 1998). As mentioned earlier in the chapter 1, recent studies have found that ginseng extraction and some of ginsenosides are effective on inhibition of tumor angiogenesis and metastasis (Sato et al., 1994; Mochizuki et al., 1995), induction of cell cycle arrest at GI and/or S phases (Odashima et al., 1985; Ota et al., 1987) and tumor cell apoptosis (Kikuchi et al., 1991; Liu et àl., 2000). Our previous results showed that in vitro, Careseng®, the mixture with 51% 20(S) protopanaxadiol (aPPD) caused apoptosis in tumor cells related to caspase activation independent on p53 in different tumor cell lines (Jia et al., 2003). Therefore, my study on aPPD was started from caspase pathways. In this chapter, the apoptotic effect of aPPD on glioma cells and MCF-7 cells were tested and the involvement of caspase pathways was described. RESULTS 1. Cytotoxicity induced by aPPD on different tumor cells Different tumor cell lines were treated with aPPD and cytotoxicity was measured with MTT assay (see Chapter 2 for methods). aPPD-induced cell-death on glioma cell lines, SF188, U87MG, SF210, and U138 was shown in Fig.3-la and b. It was apparent that aPPD had more potently cytotoxic effect on SF188 than other glioma cells. When these cells were treated for 24 hours, 43.4uM aPPD resulted in near 100% of cell killing on SF188 cells but 108.4uM was required to achieve 90% cell death on U87 cells (Fig. 3-la). The difference of the two cell lines in sensitivity to aPPD-induced cytotoxicity was also evident by the time course experiment as shown in Fig.3-lb. At the concentration of 108.4uM aPPD, near 90% of SF188 cells died within 10 hours but it required 24 hours to reach the similar level of cell killing on U87 cells. Similar results were also obtained in other aPPD-treated cell lines including prostate cancer cells LNCaP, PC3, lung cancer cells H460, H838, pancreatic cancer cells BXPC-3, MIA PaCa-2, and melanoma cells B16, etc. (data not shown). 2. The apoptotic cell death induced by aPPD To verify the apoptotic nature of aPPD-induced cell death, morphological changes of the glioma cells treated with aPPD were first observed under light microscope. SF188 cells were treated with 108.4uM of aPPD and micrographs were taken at 1, 4, 8 hours and control cells. In Fig. 3-2, typical apoptotic changes in SF188 cells: chromatin concentration and nuclei fragmentation appeared first in early hours in Hoechst 33342-stained cells (Fig.3-2 d, f, h, j) followed by cell blebbing (Fig3-2 c, e and i). At the same time, DNA fragmentation induced by aPPD in SF188 cells (Fig. 3-3) gave another evidence to indicate the apoptotic nature of aPPD-induced cell death. Interestingly, apoptosis in U87MG cells induced by aPPD appeared slightly different (Fig. 3-4). With 108uM aPPD treatment up to 24 hours, no nuclei fragmentation was found and only few cell blebbing were seen (open arrow in Fig. 3-4g). The dying cells were rounded (white arrows in Fig. 3-4c and e) with apparent chromatin condensation (white arrows in Fig. 3-4d, f and h). To further confirm that the cell death was apoptosis in both cell lines treated with aPPD, DNA fragmentation was determined using TUNEL staining with ApopTag® Plus Peroxidase In Situ Apoptosis Detection Kit (Temecula, CA) (Fig. 3-5). As shown in Figure 3-5, numerous TUNEL positive cells were seen in both SF188 and U87MG cells treated with aPPD at 3 hours and 7 hours, respectively (Fig. 3-5 b and d). In the third experiment, changes in mitochondria membrane permeability following treatment with aPPD were studied. It has been shown that changes in mitochondrial transmembrane potential is associated with apoptosis (Green and Reed, 1998). These changes result in altered membrane permeability, which can be tested using BD ApoAlert™ Mitochondrial Membrane Sensor Kit (BD Biosciences, USA). In normal mitochondria, the dye is polymerized when maintained inside of mitochondria, showing red-colored grains (Fig.3-6, b and e, white arrows). In the cytosol, the dye stays in monomer status and appears diffused green florescence (Fig.3-6 c, stealth). Fig.3-6 shows SF188 cells treated with aPPD at an early (a to c) and late (d to f) time points. It was clear that at later time point when cells underwent apoptosis as shown by their morphology in Fig. 3-6e, much less grainy red florescence could been seen, indicating an altered permeability of mitochondrial membrane in the dying cells. 3. aPPD-caused cytochrome c release from mitochondria Since the altered permeability of mitochondrial membrane causes release of cytochrome c as one step of the apoptotic pathways, levels of cytochrome c were measured in the cytosol of aPPD-treated SF188 cells. In Fig. 3-7 a and c, following treatment with aPPD, amount of cytochrome c in the cytosol of SF188 was increased with time in SF188 cells. However, when mitochondria was isolated from the cells through centrifugation and treated with aPPD, no cytochrome c was released (Fig. 3-14d). To verify that the cytochrome c was still kept in the mitochondria after the procedure, a cytochrome c depletion experiment (Fig. 3-14e) was conducted. It was confirmed that while aPPD did not directly stimulate cytochrome c release from the isolated mitochondria, a large amount of cytochrome c could still be released to supernatant with K+ depletion, indicating the mitochondria membrane was intact. Therefore, these results clearly suggest that aPPD did not directly act on mitochondrial membrane to cause cytochrome c release but it may act through other cellular component upstream to the signal transduction to alter the mitochondria membrane permeability resulting release of the cytochrome c. 4. Multiple caspase activation in glioma cells treated with aPPD Previous study indicated that Rh2 caused caspase-3 cleavage and activation on human hepatoma SK-HEP-1 cells to induce apoptosis (Park et a l , 1997). To investigate whether caspases were involved in aPPD-induced apoptosis, the activity of caspases was determined by Western blot analysis. Fig. 3-8 a, b and c show that caspase-3, -8, and -9 in SF188 cells were cleaved within 2 or 3 hours after treatment with 108.4uM aPPD and the amounts of activated caspases increased with the time of treatment. In Fig. 3-8 d, e and f, activated caspase-3, -8 and -7 were seen in cells treated with aPPD at concentration of 65uM for 4.5 hours. These results indicated that multiple caspases were activated by aPPD in SF188 cells. 5. Activation of caspase-3 by aPPD in SF188 cells requires a membrane component As an attempt of identifying the upstream element responsible for aPPD-induced apoptosis, a cell free system was utilized. Cell-free system has been well developed to study many aspects of apoptotic cell death in vitro, such as morphological transformation characteristic of apoptosis in isolated nuclei (Lazebnik et a l , 1993), DNA fragmentation into nucleosomal fragments and dATP or cytochrome c induced activation of caspases (Liu et a l , 1996), and function of pro-apoptotic Bcl-2 family members (Cosulich et a l , 1997). As shown in Fig. 3-9 (lane 1), aPPD in the cytosol did not induce caspase-3 activation. However, the caspase-3 activity was significantly elevated in the cytosol treated with aPPD in the presence of cell membrane (Fig. 3-9 lane 5). Thus, it appeared that some components in the cytomembrane (without mitochondrial membrane) were required in aPPD-induced caspase-3 activation. 6. Caspase-8 but not -3 or -9 partially responsible for aPPD-induced apoptosis in SF188 cells. Park et al. and Fei et al have reported that caspase-3 inhibitor effectively prevented Rh2 induced DNA fragmentation and partially inhibited Rh2-induced apoptosis(Park et al., 1997; Fei et al., 2002). Since aPPD also induced caspase activation and apoptosis, caspase inhibitors were used to test whether they could block the cleavage of caspases and prevent apoptosis caused by aPPD in SF188 cells. The cells were pre-treated with a caspase-8 inhibitor (Z-IETD-FMK) for 2 hours followed by adding different concentrations of aPPD (Fig. 3-10a). Viabilities of cells at 24 hours showed that those pre-treated with Z-IETD-FMK had significantly higher viability than the untreated cultures at every concentrations of aPPD (PO.001). However, the inhibitor only prevented aPPD-induced cell death by approximately 20-30%. Similar results were obtained using a general caspase inhibitor (Z-VAD-FMK) that blocks activity of all caspases (Fig.3-10d). In the latter, SF188 cells were pre-treated with different concentrations of Z-VAD-FMK for 2 hour followed by treatment with 43.4uM aPPD for 24 hours. Z-VAD-FMK prevented aPPD-induced cell death dose-dependently. The maximum protection was approximately 40% (PO.001) at 40uM of Z-VAD-FMK. Interestingly, inhibitors for caspase-3 (Z-DEVD-FMK) and caspase-9 (Z-LEHD-FMK) had no effect on the cell death induced by aPPD (Fig.3-10 b and c). Therefore, although all the major caspases were activated in aPPD-treated SF188 cells, it seemed that caspase-8 was only partially involved in the initiation of apoptosis while caspase-3 and -9 might not be responsible at all. As illustrated in Fig. 3-11 aPPD-induced cell death in the absence of caspase activation was still apoptotic, as all the apoptotic morphology such as membrane blebbing (Fig.3-11 a and c), nuclei fragmentation (Fig. 3-11 b and d), chromatin condensation (Fig.3-1 If) and apoptotic body formation (Fig. 3-11 c) were present as indicated with arrows in figures. To verify that the concentration of Z-VAD-FMK utilized in the above study was sufficient, activity of caspases was measured by Western blotting in SF188 cells treated with aPPD in the presence of Z-VAD-FMK. As shown in Fig. 3-12 a-d, Z-VAD-FMK entirely blocked caspase-8 and -3 at all the time points. Furthermore, inactivation of caspases was also confirmed by cell viability assay on TNF-a and cycloheximide treated SF188 cells. TNF-a (25ng/ml) together with cycloheximide (25ng/ml) has been shown to induce apoptosis through activation of caspases (Pober, 1998; Sawada et a l , advance online publication 7 May 2004). As shown in Fig.3-12e, Z-VAD-FMK blocked cell death induced by TNF-a and cycloheximide in a dose-dependent fashion. At concentration of 30-50uM, the inhibitor completely blocked the TNF-a and cycloheximide induced cell death. As a comparison, 50uM of Z-VAD-FMK only partially prevented the cell death induced by aPPD on the same cells (Fig 3-12f). Results in Fig 3-12 a-e together indicated that concentrations of Z-VAD-FMK utilized in the experiments were sufficient in completely blocking caspase activity. Thus, the partial blockage by the inhibitor on aPPD-induced apoptosis on SF188 cells was not due to residual activity of caspases but rather suggesting the presence of additional mechanisms responsible for the apoptosis caused by aPPD. 7. Caspase activation is not essential for aPPD-induced cell-death To further demonstrate that caspase activation is only partially responsible for aPPD induced apoptosis in tumor cells, the human breast cancer MCF-7 cells that were caspase-3 null due to a mutation were used (Janicke et a l , 1998). As shown in Fig.3-13, although caspase-8 was activated by aPPD (Fig. 3-13d), no downstream effector caspases were activated in MCF-7 cells'(Fig.3-13c). In addition, MCF-7 c3 and MCF-7 vec cells that were tranfected with caspase-3 cDNA and vector only were used, respectively. The MCF-7 vec cells served as a negative control. As shown in Fig.3-13a, all the three cell lines had similar dose-response viability curves when they treated with aPPD, suggesting that caspase-3 activation was not essential in aPPD induced apoptosis in MCF-7 cells. At the same time, caspase inhibitors were used to pre-treat the MCF-7 cells, as shown in Fig. 14, inhibition of caspase-8 by Z-IETD-FMK or Z-VAD-FMK did not have any effect on cell death induced by aPPD. These results suggest that caspase-3 and -8 are not essential in aPPD-induced apoptosis. Furthermore, the fact that caspase-8 was activated in a caspase 3-null MCF-7 cells indicated that activation of caspase-8 by aPPD was not through activation of caspase- 3 as seen in some other systems (Ferguson et al., 2003; Rabi et al., 2003). Another example of caspase-independent cell death was seen on aPPD treated U87MG cells. Fig. 3-15, a and b show Western blots of caspase-3 and -8 in aPPD- treated U87MG cells. On U87MG cells, neither caspase-8 nor caspase-3 was activated in the presence of aPPD although normal levels of pro-caspase-8 and -3 were expressed in the cells. Pre-treatment with caspase inhibitors (general caspase inhibitor, Z-VAD-FMK) did not affect aPPD-induced cell death on U87 MG cells (Fig. 15f). Similarly, aPPD also failed to activate caspases in SF210 cells as shown in Fig. 3-16 a and b. 120 r 100 -=_ GO TJ 80 -O 6 g 5 60 -<— 0 — 40 -> 20 -0 -a Ci • SF188 • U87MG • SF210 • U138 D. Dose of aPPD(uM) 140 120 c - 100 fl& ~& 0 80 -o .£ 60 Viab 40 -20 -0 • 4 6 Time (hours) 10 24 Fig.3-1 Different cell-death induced by aPPD in dose- and time-dependent manner. MTT assay was used to determine the viability of cells, a, dose-dependent viabilities in glioma cells: SF188, U87MG, SF210 and U138 treated with different concentrations of aPPD as indicated for 24 hour. b, time-dependent viability change of SF188 and U87MG cells treated with 108.4uM aPPD at different time-points. Membrane blebbing Nucleus fragmentation Fig. 3-2 Apoptosis induced by aPPD in SF188 cells at different time points! Left panels show the micrographs of SF188 cells taken under phase-contrast light microscopy; the right panels show Hoechst 33342 fluorescence microphotographs in the same field under 200X (a to h) or 320X(i and j). Arrows point to membrane blebbing, nuclei fragmentation and chromatin condensation. 1 2 3 4 5 Fig. 3-3 DNA ladders in SF188 cells induced by aPPD. SF188 cells were treated with different concentrations of aPPD for 5 hours. 1. DNA marker (lkb, Invitrogen, Frederick, MD). 2. SF188 negative control. 3. aPPD 21.7uM. 4. aPPD 65.1uM. 5. aPPD 108uM. Control 4h 7h 24h O Fig. 3-4 Apoptosis induced by aPPD in U87MG cells at different time points. A l l of the photographs were taken under 320X power, the left panels show the phase-contrast light micrographs of U87MG cells; the right panels show the Hoechst 33342 fluorescence microphotographs of U87MG cells in the same field, a and b were U87MG control, c to h were U87MG cells treated with 108.4uM aPPD at the time points as indicated. The left panels show rounded cells at earlier time point (white arrows), the stealth arrows point to cell blebbing and the open arrows indicate apoptotic body. In right panels, the arrows are condensed nuclei. SF188 U87MG _ b ! • ' ^ A+ Fig. 3-5 Detection of D N A strand breaks as biochemical markers of apoptosis by ApopTag® staining (CHEMICON, Temecula, CA) . a, control SF188 cells; b, SF188 cells treated with 108uM of aPPD for 3 hours; c, control U87MG cells and d, U87MG cells treated with 108uM aPPD for 7 hours. A l l of the photographs were taken under 400X. The representative nuclei of apoptotic cells were stained in dark-brown as indicated by arrows in the b and d. 2hour Fig. 3-6 Change of mitochondrial membrane permeability induced by aPPD treatment in SF188 cells stained with B D ApoAlert™ Mitochondrial Membrane Sensor Kit. a to f show morphological changes and mitochondrial permeabilization of SF188 cells induced by 108.4uM of aPPD treatment for early (2 hours) (a, b and c) or late (6 hours) stages (d, e and f). a and d, phase-contrast micrographs; b and e, fluorescence micrographs under rhodamine filter to show mitochondrial staining; c and f, merged micrographs of fluorescein filter micrographs and rhodamine filter micrographs. 1 2 3 4 5 6 1 2 3 4 1. Control supernatant 2. Ethanol-treated supernatant 3. aPPD-treated supernatant 4. Control mitochondria 5. Ethanol-treated mitochondria 6. aPPD-treated mitochondria 1. Control 2. Depleted mitochondria 3. Control supernatant 4. Depleted mitochondrial supernatant Fig.3-7 Release of cytochrome c induced by aPPD from intact cell or separated mitochondria, a and c, the level of cytochrome c in the cytosol induced by aPPD at different time points(a) and the data from western blotting result was analyzed by Kodak ID 3.5 software(c).Cytosol of SF188 cells was prepared as illustrated in the Material and Method. The amount of loaded protein in graph a was controlled by actin imminoblot in b. d, release of cytochrome c from mitochondria separated from SF188 cells induced by aPPD. e, cytochrome c depleted from mitochondria using hypotonic buffer as outlined in the Materials and Methods. Fig. 3-8 Time-and dose- dependent activation of multiple caspases induced by aPPD in SF188 cells. Western blotting assay was used to test the cleaved fragments of caspase-3, -7, -8, and -9 in SF188 treated with aPPD. The left panels (a, b, and c) are time course of caspase-3, -8 and -9 in SF188 cell treated with 108.4uM aPPD at 1, 2, 3, 4, and 5 hours. The right panels (d, e and f) are levels of caspase-3, -8 and -7 in SF188 cells treated with aPPD of indicated concentrations for 5 hours. For each graph, the up arrow shows the pro-caspase and the low arrows point to the activated caspases. dATP _ + _ _ _ _ aPPD + - - - + + C e l l membrane - - + - + -Cyto-c mmm. + - + w m am mm mm mm i 2 3 4 5 6 Fig. 3-9 aPPD-induced caspase-3 activation in cell-free system. Caspase-3 activation in SF188 cytosol was detected with Western blotting assay. SF188 cytosol fraction treated with: 1, 108uM aPPD only, 2, lOuM dATP only, 3, incubating with cell membrane, 4, lOnM cytochrome c , 5, 108uM aPPD and cell membrane, 6, 108uM aPPD and lOnM cytochrome c. con 21.7 32.5 43.4 54.2 65.1 86.8 0 5 10 20 30 40 50 Con Dose of aPPD (uM) Dose of General Caspase Inhibitor(uM) Fig. 3-10 Effects of caspase inhibitors on apoptosis induced by aPPD in SF188 cells, a, caspase-8 inhibitor b, caspase-9 inhibitor c, caspase-3 inhibitor and d, general caspase inhibitor, * PO.001). Ç LI - _ j : j } ^ ! 0 & j I y M i I D ""b d \ i f * *«,* • 0 ' ^ # 1 • * * • n ft j l h 4h 8h Fig. 3-11 aPPD-induced apoptosis in SF188 cells pre- treated with general caspase inhibitor (Z-VAD-FMK). a, c and e: phase-contrast micrographs; b, d and f, Hoechst 33342 fluorescence micrographs of the same field (200X). Fig. 3-12 General caspase inhibitor (Z-VAD-FMK) entirely blocked caspase activity, a and b, caspase-3 and -8 of SF188 cells treated with aPPD (108.4uM). c and d densitormetry results of a and b. e, Effect of GI on TNF-a (25ng/ml) and cycloheximide (25ng/ml) treated SF188cells; f, The same concentration of GI only partially protected SF188 cells treated with aPPD (108.4uM).(*P<0.01) GI = general caspase inhibitor. Dose of aPPD (uM) Time of treatment(hour) Con 21.7 43.4 65.1 86.8 108.4 Con+(uM) Con 21.7 43.4 65.1 86.8 108.4(uM) Fig. 3-13 aPPD induced caspase-8 activation and apoptosis is independent of caspase-3 in MCF-7 cells, a and b, dose-response and time-response study for viabilities of MCF-7 cells treated with aPPD. MCF-7 c3, stable expression of transfected caspase-3, MCF-7vec, vector control, c, aPPD did not induced expression or activation of caspase-3 in MCF-7 cells, d, caspase-8 activation in MCF-7 cells treated with aPPD. ISO 140 120 100 u "S 80 1 60 > 40 20 0 • aPPD only • Caspase-8 inhibitor • General caspase inhibitor 21.7 43.4 54.2 65.1 74.9 86.8 108.4 Dose of aPPD(uM) Fig. 3-14 Caspase inhibitors did not affect viability of aPPD-treated MCF-7cells (P>0.1 except for 108uM aPPD group). a Casp-8 0 21.7 43.4 65.1 86.8 108.4 Con + (uM) Actin Casp-8 Con- 1 3 5 7 10 Con+(hours) e -Casp-3 0 21.7 43.4 65.1 86.8 108 130 43.4* Con+(uM) laPPD jfZlGeneral caspase inhibitor and a P P D JS . " 5 Dose of a P P D (uM ) Fig. 3-15 Caspase-3 and-8 activity in aPPD-treated U87MG cells, a, caspase-8 in U87MG treated with various concentrations of aPPD; c, caspase-8 treated with 108uM aPPD at various time points. The first lane was negative control of U87MG cells, the second to sixth lanes were aPPD-treated groups at different time points, the seventh was positive control. The amount of loaded protein in each lane of a and c was controlled by the actin immunoblot in b and d. e, caspase- 3 in U87 MG cells treated with different concentrations of aPPD for 9 hours and the lane labeled with * was U87MG cells treated with 43.4uM aPPD for 24 hours. The upper arrows in each graph show the pro-caspase-8 or -3; the lower arrows in each graph point to the activated caspase- 8 or -3. f, the dose-response viabilities of U87MG cell induced by different concentrations of aPPD for 24 hours with or without pre-treated with general caspase inhibitor (Z-VAD-FMK) for 2 hours by MTT assay analysis. Casp-8 Casp-3 d Con- 21.7 43.4 65.1 108.4 Con+(uM) Fig.3-16 Detection of caspase-8 and -3 in SF210 and U138 cells treated with aPPD. SF210 and U138 were treated with different concentrations of aPPD for 7 hours and western blotting assay was applied to determine activation of caspase-8 and -3. a and b, caspas-8 and -3 in SF210 cells separately; c and d, caspas-8 and -3 in U138 cells separately. In each of graphs the upper arrows showed pro-caspases, the lower arrows showed activated caspases. DISCUSSION Many cancer therapeutics eliminate tumors by inducing apoptosis (Herr and Debatin, 2001; Johnstone and Astrid A. Ruefli 2002). Apoptosis was initially characterized by the activation of a specific family of cysteine proteases, the caspases, followed by a series of caspase-mediated typically morphological changes (Kerr et al., 1972; Strasser and Dixit, 2000; Kaufmann and Hengartner, 2001). This study showed that aPPD effectively induced cell death in different human tumor cell lines in a dose- and time- dependent fashion. Like Rh2, the aglycone derivative aPPD also exhibited effective anti-cancer activity by inducing apoptosis in various cancer cells with different genetic background and tissue origins. Our results were in consistent with previous studies in Rh2 (Kim et al., 1999a; Fei et al., 2002) that aPPD caused activation of caspases in many types of tumor cells. The most striking finding from this study is that caspases are not required for aPPD-induced apoptosis. This conclusion is supported by the following facts: a) Caspase-8 inhibitor could only partial block aPPD-induced apoptosis in SF188 cells by 20-30% and inhibition on caspase-3 and other caspases had no protective effect at all; b) Consistent to the latter, the caspase-3-null cells MCF-7 were equally effectively killed by aPPD and restoring the caspase-3 did not result in more cell death, indicating that caspase-3 was indeed not required. Furthermore, the general caspase inhibitor also had no effect on aPPD-induced apoptosis in this cell line; c) More directly, no caspases were activated in U87MG cells while the cells underwent apoptosis in the presence of aPPD; d) The caspase-independent cell death observed above was not due to incomplete inhibition of caspases as the same concentrations of inhibitors completely blocked apoptosis induced by TNF-O! together with cycloheximide and Western blotting also showed complete absence of activated fragments of caspases while the non-activated pro-caspases were still present. The next question is that in the absence of caspase activation, whether or not the cell death induced by aPPD was still apoptosis. To answer this question, the apoptotic nature of aPPD induced cell death in the presence and absence of caspase activation was extensively studied. Evidence supporting the apoptotic death in the absence of caspase activation were: 1) similar apoptotic morphology in aPPD-treated SF188 cells with or without caspase inhibitors; 2) numerous TUNEL-positive cells induced by aPPD in both SF188 cells with activated caspases and U87MG that no caspases were activated. According to the nuclear morphology, PCD can be divided into three subclasses: classic apoptosis, apoptosis-like PCD and necrosis-like PCD (Leist and Jaattela, 2001). Caspase-mediated cleavage of specific substrates explains several of the characteristic features of apoptosis such as chromatin condensation, fragmentation, and cell membrane blebbing (Earnshaw, 1995; Rudel and Bokoch, 1997; Hengartner, 2000; Nagata, 2000; Leist and Jaattela, 2001; Petit et a l , 2002). However, in the present study, both SF188 and U87MG cells showed chromatin condensation and DNA fragmentation after treated with aPPD regardless the status of caspase activity. In the case of SF188, cell membrane blebbing was the most remarkable morphology change when the cells underwent aPPD-induced apoptosis, and it was not affected by caspase inhibitors. Similar phenomena were analyzed in cell membrane dynamics in different death models. The studies demonstrated that the plasma membrane blebbing on a dying cells could occur independently of caspase activation (McCarthy, 1997; Foghsgaard et a l , 2001; Leist and Jaattela, 2001). Thus, although the other possible forms of cell death that does not require activation of caspases in aPPD-treated cells cannot be ruled out, the above results strongly suggest that aPPD is able to induce apoptosis without activating caspases. In a recent study, it has been demonstrated that the expression and activation of caspase-3 was not related to the apoptosis in some cells, while it was required for other cellular processes such as neuronal differentiation and migration (Rohn et a l , 2000; Yan et a l , 2001). Some studies also indicate that the apoptosis or programmed cell death can occur in the complete absence of caspase or in a caspase-independent manner where non-caspase proteases and other death effectors can be executioners of apoptosis (Susin et a l , 1999; Leist and Jaattela, 2001; Lockshin and Zakeri, 2002), such as JNK activation (Yang et a l , 1997), production of ROS (Green and Reed, 1998). In the present study, aPPD induced activation of caspases in SF188 cells but not U87MG. It was noticed that SF188 is P53- / PTEN+ while U87MG is P53+/PTEN-(Chen et a l , 1995; Furnari et a l , 1997; Li et a l , 1997b; Maier et a l , 1999; Zundel et a l , 2000; Pore et a l , 2003; Kano et a l , 2004), leading to the suspician that the caspase-8 activation might be related to the PTEN status. However, when the cells were expanded to other glioma cells with different PTEN status, the results became less conclusive. Although SF188 cells with positive PTEN clearly showed activation of caspase-8 when treated with aPPD, the protease was also activated in U138 that have been thought PTEN negative (Li et a l , 1997b) as showen in Fig. 3-16c. Since PTEN status was not measured in this study and the so-called PTEN negative cells were all determined only by western blotting, which may not detect low-level expression of the protein; nevertheless, it is worthwhile further investigating the mechanism for activation of caspase-8 in SF188 but not U87MG in aPPD-treated cells. Another well-studied gene that involves apoptosis is p53. In most cell systems, chemotherapy and radiation induce apoptosis through functionally wild type p53 because it acts to negatively regulate cell division by controlling a set of gene expression (Buckbinder et a l , 1994). Normal p53 status is associated with a positive reaction to anti-tumor therapy (Lotem J and L , 1993; Fujiwara T et a l , 1994) and dysfunction of p53 causes uncontrolled cell growth to enhance tumorigenesis (Greenblatt et a l , 1994). Therefore, it is largely believed that the restoration of p53 function is therapeutically useful to enhance the beneficial response to chemotherapeutic regimens (Lotem J and L., 1993; Fujiwara T et al., 1994). Thus, replacement of the altered gene that related to tumorigenesis has been used to develop basic gene therapy for cancer (Hamada et al., 1996). Interestingly, a recent study found that U373MG glioma cells, which had a mutant p53 gene, were more sensitive to cisplatin as compared to U87MG glioma cells with normal p53. Further, U87MG cells blocked the p53 response with antisense oligonucleotides became more sensitive to cisplatin and shifted the cellular response from cell circle arrest to caspase 3-mediated apoptosis (Datta et al., online publication 28 May 2004). Another experiment showed that U87MG cells with endogenous functional p53 were relatively resistant to 7-radiation and that the loss of functional p53 rendered them susceptible to 7-radiation-induced apoptosis (Hara et al., advance online publication 16 April 2004). In the present study, three cell lines were used: U87 MG and MCF-7 express wild-type p53 while SF188 expresses mutated p53 (Chen et al., 1995; Abramovitch and Werner, 2003; Kano et al., 2004). It appeared that cells with wild type p53 needed higher concentrations of aPPD to induced cell death. While those results seem supporting the notion that wild-type p53 may render the cells more resistant to chemotherapy agents, there are two factors must also be considered. Firstly, while both MCF-7 and U87MG have wild-type p53, they both also lack caspase activation induced by aPPD. Secondly, p53 mutation in many cancer cells does not mean p53-null. Instead, mutant forms of p53 are expressed, which may show some additional functions different from the wild-type p53 (He et al., 2002). Thus, cautions must be taken to interpret our results and further investigation is needed to clarify the relationship between p53 status and the resistance to aPPD-induced apoptosis. Taking the above together, results from the present study provided another example of caspase-independent apoptosis, suggesting aPPD is not only a potential chemotherapy drugs for cancer treatment but also an interesting compound that may provide a valuable model for investigating the complex mechanisms of apoptosis. But for the current study, the most important conclusion revealed from this series of experiments was that there should be additional mechanisms mediating aPPD induced apoptosis in those glioma cells, which will be described in the rest of this thesis. CHAPTER 4 aPPD ACTIVATES STRESS SIGNALING PROTEIN KINASE WHILE INHIBITS PRO-GROWTH SIGNALING PROTEIN KINASE IN GLIOMA CELLS INTRODUCTION Kinases are enzymes that add phosphates to small molecules or other enzymes, creating active signaling molecules or inactivate signaling pathways (Downward, 1998a). In this part of my study, effects of aPPD on activity of two kinases: JNKs (c-Jun NH2-terminal kinases) and PKB/Akt (protein kinase B) were tested in glioma cells. Mitogen-activated protein kinase (MAPK) superfamily, which was first reported in 1986 (Sturgill and LB, 1986), has been implicated in the growth factor-mediated regulation of diverse cellular events such as proliferation, senescence, differentiation and apoptosis (Dent et a l , 2003a). Mammals express at least four distinct groups of MAPKs including extracellular signal-related kinases (ERK)-l/2, p38 proteins (p38 a IBIj 18), extracellular signal regulated kinase 5 (ERK5), and c-Jun-N-terminal kinases (JNK1-3/SAPK) (Seger and Krebs, 1995; Zhou et a l , 1995; Chang and Karin, 2001; Cavanaugh, 2004; Raviv et a l , 2004), which are initially described to be stress-induced protein kinases (SAPK) that phosphorylate the NH2-terminus of the transcription factor c-Jun (Karin, 2001). Among the MAP kinases, ERK is mainly activated by mitogens and growth factors, while p38 and JNKs are activated by many environmental stress stimuli including UV- and y -irradiation, cytotoxic drugs and reactive oxygen species (e.g., H2O2) (Chen et a l , 1996; Dreskin et a l , 2001; Dent et a l , 2003b). The JNKs are initially identified and purified by Kyriakis et al. as a kind of protein kinase that are activated in the liver of rodents exposed to cycloheximide (Kyriakis and Avruch, 1990). As a typical serine/threonine kinases, JNKs comprise 11 protein kinase subdomains (I-XI subdomains) (Gupta et a l , 1996; Manning and Davis, 2003). Their protein kinase activation loop is located between domains VII and VIII, which contains the threonine and tyrosine residues that are phosphorylated for full kinase activation (Gupta et al., 1996; Manning and Davis, 2003). JNKs are encoded by three genes: JNK1, JNK2 and JNK3, each of which can produce 46- and 54-kDa isoforms (Ip and Davis, 1998). Ten JNK isoforms are created by alternative splicing of messenger RNA transcripts derived from these three genes (Gupta et al., 1996; Davis, 2000). There are two key alternative splicing sites: the first is between subdomain IX and X of the C-terminal lobe of the protein, which results in splicing forms with altered substrate specificity; the second alternative splicing site occurs at the C terminus of the protein, and results in proteins in differ length by either 42 or 43 amino acids (Gupta et al., 1996; Dreskin et al., 2001 ; Manning and Davis, 2003). Each M A P K pathway consists of a set of three kinases: a M A P K kinase kinase (MAPKKK), which phosphorylates and activates a M A P K kinase (MAPKK), which in turn, phosphorylates and activates a MAPK (English et al., 1999; Chang and Karin, 2001; Mizuno et al., 2004). Al l MAPKs recognize similar phosphoacceptor sites composed of serine or threonine followed by a proline, and the amino acids that surround these sites further increase the specificity of recognition by the catalytic pocket of the enzyme (Karin, 2001). The INK pathway was discovered and described in the early to mid-1990 (Derijard et al., 1994; Kallunki et al., 1994; Kannan and Jain, 2000; Curtin et al., 2002). It has been identified that activated JNK has double effects on cell survival and apoptosis. The influence of JNK on cell death not only depends on the death-inducing stimulus and cell type, but also appears to be determined by the duration of JNK activation. Early activation or transient activation of JNKs are essential for protecting cells against TNF-mediated apoptosis and promotes cell survival, whereas persistent JNK activation induces apoptosis (Chang, 1999; Kobayashi and Tsukamoto, 2001; Chang et al., 2003). The transcription factor complex AP-1 has been identified as a target of MAP kinase signaling pathways (Karin, 1995). AP-1 is not a single protein, but a homo- or heterodimer composed of members of the c-Jun, c-Fos, activating transcription factor (ATF), and other protein families (Shaulian and Karin, 2002), which can bind to specific control elements present in the promoters of genes to regulate cell differentiation and proliferation (Angel and Karin, 1991). c-Jun is one of the best characterized components of AP-1 (Mechta-Grigoriou et al., 2001; Shaulian and Karin, 2001). It is an immediate early gene whose level can increase within an hour of exposure to a variety of extracellular stimuli and the same stimuli that induce c-Jun expression also trigger its phosphorylation at Ser63 and Ser73 in the N-terminal domain, which is required for it to become transcriptionally active (Smeal et a l , 1991; Davis, 2000). The phosphorylation of these residues is thought to be mediated by the isoforms of JNK (Derijard et al., 1994; Davies et al., 2000). Genetic evidence suggests that c-Jun is essential for development and proliferation (Mechta-Grigoriou et al., 2001; Shaulian and Karin, 2001, 2002). c-Jun has also been implicated in the regulation of apoptosis, which can promote or counteract, depending on the tissue, the developmental stage and the nature of the death stimulus (Leppa and Bohmann, 1999; Herdegen and V., 2001; Shaulian and Karin, 2002; Cuadrado et al., 2004). Besides, it has been found that JNKs phosphorylate several proteins involved in the apoptotic process including c-Myc, p53, Bcl-2 and Bcl-x L (Fuchs et al., 1998b; Fuchs et al., 1998a; Noguchi et al., 1999; Fan et al., 2000). Therefore, JNK activation has been believed to be an essential component of the apoptotic process in a number of systems (Jarpe et al., 1998; Huang et al., 1999; Wang et al., 1999a; Phelan et al., 2001). Some studies found that the nuclear factor- K B ( N F - K B ) -dependent inhibition of the JNK pathway (Tang G et al., 2001) was central to the N F - K B anti-apoptotic function (Smaele et al., 2001) and abrogation of N F - K B activation resulted in prolonged JNK activation which was required for TNF-induced apoptosis (Smaele et al., 2001; Tang G et a l , 2001), which further verified that JNKs involve in apoptosis pathway. Similarly, different JNK inhibitors such as small peptide inhibitor (JNK inhibitor I) and pan JNK inhibitor SP6000125 (JNK inhibitor II) are used to suppress JNK activity to investigate the role of JNK in cell proliferation, cell cycle progression and ultimately the sensitivity of cells to apoptosis (Bonny et a l , 2001; Holzberg et a l , 2003). PI3K-PKB/Akt kinase (Phosphatidylinositol 3-kinases-protein kinase B/Akt) pathway has emerged as a critical pathway for cell survival in cancers. PI3Ks function to generate intracellular second messengers by catalyzing the transfer of phosphate from ATP to the D-3 position of the inositol ring of membrane-localized phosphoinositides (Toker and Cantley, 1997). There are three kinds of PI3K (Vanhaesebroeck et a l , 1997). Class I PI3K consists of a family of heterodimeric complexes composed of a pi 10 catalytic subunit and a regulatory subunit that exists predominantly in a p85 form (Toker and Cantley, 1997; Vanhaesebroeck et a l , 1997). Class I P13Ks phosphorylate phosphatidylinositol 4-phosphate and phosphatidylinositol 4,5-bisphosphate (PIP2) in vivo, however, their preferred substrate is likely to be PIP2. Class II PI3Ks are larger (> 200 kDa) enzymes that phosphorylate in vitro phosphatidylinositol and phosphatidylinositol 4-phosphate, but not PIP2. Class III P13Ks have a substrate specificity restricted to phosphatidylinositol (Toker and Cantley, 1997; Vanhaesebroeck et a l , 1997). The products of PI3-kinase activation, phosphatidylinositol- 3,4,5-trisphosphate (PIP3) and its immediate breakdown product phosphatidylinositol- 3,4-trisphosphate (PI(3,4)P2), trigger physiological processes, by interacting with proteins possessing pleckstrin homology (PH) domains (Isakoff et a l , 1998; Lemmon and Ferguson, 2000). The N-terminal PH domain is common to numerous signaling proteins and provides a lipid binding module to direct Akt to PI3K-generated PD?3 and PI(3,4)P2. One of the best characterized PTP3/PI(3,4)P2 effector proteins is PKB, also known as Akt (Musacchio et a l , 1993; Gibson et a l , 1994; Blomberg et a l , 1999), which is a human homologues of the v-akt oncogene of AKT8 provirus cloned at 1987. AKT8 provirus genome contains both viral and nonviral sequences, v-akt is the nonviral sequence and viral oncogene of the AKT8 virus (Staal, 1987; Bellacosa et a l , 1991). Akt is a serine/threonine kinase belonging to the ' A G C superfamily of protein kinases (named after family members protein kinase A, protein kinase G and protein kinase C). In mammals, there are three isoforms of Akt (PKBa, /3, 7 or Akt l , 2, 3), while different Akt isoforms are activated similarly and phosphorylate downstream substrates with equal specificity and efficiency (Franke, 2000). Al l of the substrates identified by an oriented peptide library approach or peptide library screening contained Arg-Xaa-Arg-Xaa-Xaa-(Ser/Thr) motifs (Obata et a l , 2000). Al l three isoforms share a high degree of amino acid identity and are composed of three functionally distinct regions: an N-terminal PH domain, a central catalytic domain, and a C-terminal hydrophobic motif (HM). Together, these regions encompass a phosphoprotein of approximately 56 kDa. PH domain was first identified in 1993 as a 100-120-residue stretch of amino-acid-sequence similarity that occurs twice in pleckstrin and is found in numerous proteins involved in cellular signaling (Haslam et a l , 1993; Mayer et a l , 1993). It was originally proposed that PH domains might be involved in protein-protein interactions in cellular signaling (Haslam et a l , 1993; Mayer etal, 1993). Growth factors that act to increase PI3K activity and lead to the subsequent generation of PIP3 and PI(3,4)P2 provide a plasma membrane recruitment for Akt. Membrane recruitment of Akt is a hallmark of activation (Vanhaesebroeck et a l , 1997; Downward, 1998a; Scheid and Woodgett, 2003). Based on a large number of studies, the PI3K7 Akt pathway clearly has a key role in cellular survival and transformation. Activated Akt phosphorylates several pro- and anti-apoptotic proteins, including the Bcl-2 family member BAD (Downward, 1999), caspase-9 (Cardone et al., 1998; Cheung et al., 2004), cyclic AMP response element-binding protein, the inhibitor of NF-/cB kinase IKKa, forkhead transcription factor-1 (Datta et al., 1999; Kops et al., 1999), which are known to induce the expression of genes that are critical for apoptosis, and preventing the release of cytochrome c from mitochondria (Datta et al., 1999; Di Cristofano and PP., 2000). Inhibition of Akt has been shown to induce apoptosis (Asselin et al., 2001b; Asselin et al., 2001a; Franke et al., 2003). Phosphorylation of Thr308 partially activates Akt, while phosphorylation of both Thr308 and Ser-473 sites is required for full activation of Akt (Alessi et al., 1996; Downward, 1998b; Datta et al., 1999). Akt Thr-308 lies within the activation loop of the kinase domain, upstream kinases may induce Akt activity by phosphorylation of the Thr-308 (Datta et al., 1999). The protein kinase termed 3-phosphoinositidedependent protein kinase-1 (PDK1) plays a central role in activating AGC subfamily members (Peterson and SL., 1999) . PDK1 phosphorylates Akt at Thr308 (Alessi et al., 1997; Stokoe et al., 1997). Integrin-linked kinase (ILK) is a PI-3 kinase-dependent serine/threonine kinase that interacts with the cytoplasmic tail of integrin |3 subunits (Hannigan et al., 1996; Delcommenne et al., 1998), and acts as an effector of integrin-mediated cell adhesion as well as growth factors (Delcommenne et al., 1998; Persad et al., 2001). ILK is also an upstream regulator of Akt and has also been suggested to regulate Akt by phosphorylation of Akt at Ser-473 (Delcommenne et al., 1998; Persad et al., 2001). The PTEN (phosphatase and tensin homolog deleted from chromosome 10) tumor-suppressor gene, also known as MMAC1 (mutated in multiple advanced cancers), is located on chromosome 10q23, a genomic region that suffers loss of heterozygosity (LOH) in many human cancers (Cantley and Neel, 1999). In early embryos, loss of Akt activity after overexpression of PTEN results in ectopic apoptosis in Drosophila (Scanga et al., 2000). The PTEN, a phosphatase for the lipid products of PI3K, specifically cleaves, in vitro and in vivo, the D3 phosphate of this second messenger lipid PIP3 produced by the activity of PI3-kinase (Maehama and Dixon, 1998). The actions of PTEN reduce the entire pool of lipids capable of binding with Akt and make PIP3 level be very low in quiescent cells, but it rapidly increases upon stimulation by growth factors, through activation of PI3K. However, somatic deletions or mutations of PTEN gene have been identified in a large fraction of tumors including glioblastomas (Di Cristofano and PP., 2000), accumulation of PD?3 at the membrane allows recruitment of proteins containing a PH domain such as Akt to bind PIP3, which leads to constitutively activated Akt. As result, Akt exerts antiapoptotic activity as a survival factor (Datta et al., 1999). Akt pathway is considered a key determinant of biologic aggressiveness of solid tumors, and a major potential target for novel anti-cancer therapies. It has become an attractive target for antitumor drug development (Vara et al., 2004). Like many antitumor drugs, ginsenosides as antitumor drugs, their mechanisms of antitumor have been studied in different fields. It has been identified that ginsenosides, especially Rh2, induced tumor cell apoptosis in different ways. Some studies find that in SK-HEP-1 (human liver adenocarcinoma cells), later phase of JNK1 activation, which is linked to a caspase-dependent mechanism, is associated with the induction of apoptosis; while the early JNK1 activation at the first 10-30 minutes of Rh2 treatment had been found which is associated with a SEK1 -mediated (where SEK-1 stands for SAPK/ERK kinase 1 (Sanchez I et al., 1994) mechanism and acts to prolong cell survival in response to apoptosis-inducing agents (Ham et al., 2003a). Like Rh2, aPPD induced apoptotic cell-death in different tumor cell lines as shown in the previous chapter. In this chapter, activation of JNK and its relation to aPPD induced apoptosis were described. The effect of aPPD on survival pathway were tested in glioma cells with different PTEN: PTEN null tumor cell line U87 M G and PTEN positive cell line SF188 (Freeman et a l , 2003; Pore et a l , 2003). Result 1. aPPD induced J N K phosphorylation in SF188 and U87MG cells in dose- and time-dependent fashions Western blotting analysis showed that aPPD increased phophorylated -JNK (P-JNK) in dose- and time-dependent manners as illustrated in Fig. 4-1. Comparing to the control group, the levels of P-JNK in aPPD treated groups were 5-10 times higher. In Fig. 4-1, 1 toi .5 hour-treatment with 108.4uM aPPD was enough to induce JNK phosphorylation in both U87 M G and SF188 cells. Interestingly, 5 hours treatment with 21.7uM of aPPD, the concentration did not cause apoptosis in U87MG, but induced increase in JNK phosphorylation. 2. J N K inhibitors did not block apoptosis induced by aPPD To study the relationship between JNK and apoptosis induced by aPPD, different concentration (0-20uM) of the JNK inhibitor I (cell-permeable peptide inhibitor) and a negative control were used to pre-treat SF188 cells for 2 hours and then the cells were treated with 34uM aPPD for overnight. As shown in the Fig. 4-2, neither JNK inhibitor nor the negative control affected the apoptosis induced by aPPD. In the groups treated with different concentrations of JNK inhibitor I or JNK inhibitor I negative control only, the cell viabilities were the same as the control groups treated with vehicle (0 concentration of JNK inhibitor I or JNK inhibitor I negative control), but almost all the cells died when treated with JNK inhibitor I or the negative control in the presence of 34uM aPPD. Similarly, JNK inhibitor II (SP600125) was also not effective to block cell-death induced by aPPD (Fig. 4-2 c and d). These results suggested that JNK activation was not related to aPPD-induced cell death in the glioma cells. 3. Effect of aPPD on phosphorylation level of Akt Ser473 As a crucial step in the survival pathway, phosphorylation on Akt was investigated in aPPD-treated SF188 and U87MG glioma cells. Based on the results from the Western blot analysis with an antibody specific for Ser473 phosphorylated Akt, SF188 cells revealed very low level of phosphorylation in the control group (Fig.4-3a). The levels of phosphorylated Akt remained unchanged in SF188 cells treated with aPPD up to 108.4uM for 5 hours. But in U87MG cell treated with different concentrations of aPPD for 7 hours, the levels of P-Akt were declined in a dose-dependent fashion (Fig.4-4a). P-JNK JNK Actin a •aaas ssee m 1-12 g 10 < 2 8 ta! 5 6 S 2 05 „ P-JNK (46kD) Con 21.7 65.1 86.8 108.4 (uM) I a con 21.7 65.1 108.4 Dose of aPPD(uM) SF188 P-JNK .INK Con 1.5 2.5 3.5 4.5 5.5 (hour) o 1 as 0 |P-JNK(46kD) DP-JNK(54kD) ^ 5 S 4 _ J i t SF188 con 1.5 2.5 3.5 4.5 5.5 Time of treatment(hour) P-JNK JNK Actin |P-JNK(54kD) • P-JNK(46kD) Con 1 3 5 (hour) _ U87MG Con 1 3 5 Time of aPPD treatment(hour) Fig. 4-1 aPPD induced dose- and time- dependent J N K activation in SF188 and U87 M G glioma cells by Western blotting analysis, a to c were SF188 cells treated with different concentrations of aPPD for 5 hours, a, phosphorylated JNK(P-JNK), b, JNK and c, actin. a, b, and c were from the same experiment, d, was the quantitative analysis of P-JNK using Kodak ID 3.5 softwere. e and f were time course results in SF188 cells treated with aPPD 108.4uM for different time, e, P-JNK; f, JNK; g was the quantitative analysis of P-JNK. e, f, and g came from the same experiment, h, i , and j were the results of U87 cells treated with 108uM aPPD at different time-points, k was the quantitative analysis of P-JNK in h comparing with actin using Kodak ID 3.5 softwere. The upper arrow shows 54kD band, and the lower arrow shows 46kD band in J N K or P-JNK results. 140 120 100 JS. ~-~> o 80 o 60 40 > 20 0 SF188 • JNK. inhitior I • JNK inhibitor I and aPPD I 1 1.5 2.5 5 10 20 Dose of JNK inhibitor I (uM) 140 120 ? 100 « o & 60 > 40 20 0 SF188 • Negative control • Negative control andaPPD X x. je. X T I 1 1.5 2.5 5 10 Dose of negative control (uM) 20 SF188 HaPPD llOOnM JNK inhibitor II 0 ^ ^ f ^ Dose of aPPD (ug/ml) 160 140 120 100 SO 60 40 20 0 U87MG BaPPDonly • 40nM JNK inhibiotrll V fob-Dose of aPPD(uM) Fig.4-2 JNK inhibitors did not block apoptosis induced by aPPD. SF188 cells were pretreated with different concentration of JNK inhibitors or JNK inhibitor negative control before 1 hour of adding aPPD treatment or vehicle, a was the result of SF188 cells treated with different concentrations of JNK inhibitor I with or without aPPD as indicated in graph; b was the result of SF188 cells treated with different concentrations of JNK inhibitor I negative control with or without aPPD. c, SF188 cells pre-treated with or without different concentrations of JNK inhibitor II with different concentrations of aPPD as indicated in the graph, and d U87MG cells pre-treated with or without different concentrations of JNK inhibitor II with different concentrations of aPPD as indicated in the graph. For each group in a and b, PO.01; in c and d, P>0.01 P-Akt Akt Con 21.7 65.1 108.4 (aPPD uM) Fig. 4-3 aPPD had no effect on phosphorylation levels of Akt Ser473 in SF188 cells, a, P-Akt, b, total Akt P-Akt Akt Actin Con 21.7 65.1 86.8 108.4 (uM) P- Akt Akt Actin Con 5 (hours) Con 21.7 43.4 65.1 86.8 108.4 Dose of aPPD (uM) Con 1 3 5 Treatment time (hour) Fig. 4-4 aPPD induced reduction of P-Akt in U87 M G cells. U87 cells were treated with aPPD at indicated concentrations for 7 hours or at 108.4uM for indicated time points, a to c and e to g were Western blotting results for P-Akt, Akt or actin, d and h were densitometry analysis using Kodak 1D3.5 software. DISCUSSION Although the results showed that aPPD upregulated levels of P-JNKs in both SF188 and U87MG cells, two specific JNK inhibitors failed to block apoptosis induced by aPPD in the glioma cells. Thus, it appears that JNK does not involve in the apoptotic mechanisms of aPPD and the role of JNK activation in aPPD-treated cells remains unknown. Much attention has been focused on the opposing roles of JNK in the regulation of both cell death and, under certain conditions, cell survival (Potapova et a l , 2000). JNK activation has controversial functions depending on types of stimuli and cell lines. For instance, thymocyte apoptosis induced by U V irradiation or anti-Fas antibody is not affected by the absence of either JNK1 or JNK2, although the JNKs could be activated by both stimuli; JNK2 is required for apoptosis of immature thymocytes induced by anti-CD3 antibody but not for apoptosis induced by anti-Fas antibody, UVC or dexamethasone (Sabapathy et a l , 1999; Sabapathy et a l , 2001). Additionally, apoptosis in mouse hepatocytes, induced by human TNF-a, was blocked by SP600125, indicating that TNF- receptor (TNF-R) 1-mediated JNK activation is important for TNF-a-induced cell death (Schwabe et a l , 2004). The apoptotic effect of Eiger, TNF-a homolog of Drosophila does not require the activity of the caspase-8 homolog DREDD, but it completely depends on its ability to activate the JNK pathway (Moreno et a l , 2002). Another study reported that JNK1 and JNK2 were protective in TNF-a -mediated apoptosis (Hochedlinger et a l , 2002). It suggests that JNK activation coming from the same stimulus may give rise to different reactions in different cells and it also appears that the same cell may have different JNK activations depending on the stimuli. There are at lest three kinds of JNK commercial inhibitors can regulate the activation of JNK. In the present study, two types of the inhibitors have been used. JNK inhibitor I, bioactive cell-permeable peptide inhibitors of JNK, is engineered by linking the minimal 20-amino acid inhibitory domains of the islet-brain (IB) proteins to the 10-amino acid HIV-TAT sequence that can rapidly transfer the inhibitor peptide into cells (Bonny et al., 2001; Barr et al., 2002). Kinase assays indicate that the inhibitor I blocks activation of JNK by blocking activation of transcription factor c-Jun. JNK inhibitor II (SP600125), a small molecular compound, is a reversible inhibitor that competes with ATP for binding to kinases and has the highest selectivity toward JNK (20-fold more than other kinases) (Bennett et al., 2001). As a pan blocker of JNK activation, SP600125 is used to protect dopaminergic neurons from l-methyl-4-phenyl-l,2,3,6-tetrahydropyridine (MPTP) neurotoxicity on PD mice (Human Parkinson's disease mice model) (Wang et al., 2004). The last commercial inhibitor, JNK inhibitor III, is a cell-permeable peptide containing the JNK-binding (ô) domain of human c-Jun. It strongly and specifically induces apoptosis in HeLa tumor cells, which is paralleled by inhibition of serum-induced c-JUN phosphorylation and up-regulation of the cell cycle inhibitor p21 w a f 7 c , p (Holzberg et al., 2003). In the present study, JNK inhibitors I and II were applied to block the activation of JNK, but neither had effect on aPPD induced apoptosis in both SF188 and U87MG cells. While both inhibitors failed to protect two glioma cell lines from apoptosis induced by aPPD is a strong indication that JNK is not involved in the apoptotic mechanism, the conclusion may still need further verification. For instance, the levels of phosphorylated JNK in the presence of inhibitors will be measured to confirm that the inhibitors indeed block the activation of JNK. In addition, it seems that JNK inhibition by different inhibitors may only affect a specific pathway. Liedtke et al. found that inhibition of JNK by JNK dominant negative TAK1 accelerated TNF-a -induced apoptosis in hepatoma cells but the same effect did not seen when SP600125 was used as an JNK inhibitor (Liedtke et al., 2002). Further, JNK exert its proapoptotic effect in hepatocytes independently of transcription and c-Jun (Schwabe et al., 2004), which means using JNK inhibitor I may not block the apoptosis in this type of cells. Thus, differently approaches for inhibiting JNK should be applied to test this possibility in the glioma cells. The phosphorylation of Akt on Ser-473 was also tested, which is activated by integrin-linked kinase (ILK), a PI-3 kinase-dependent serine/threonine kinase (Hannigan et a l , 1996; Delcommenne et a l , 1998). PTEN inhibits PI3-kinase-dependent activation of ILK (Obara et a l , 2004) and Akt (Stambolic et a l , 1998). Further, aPPD significantly decreased the level of P-Akt in PTEN negative U87 M G cell but not for PTEN positive SF188 cells. The activation of Akt is regulated in a complicated manner via phosphorylation of Akt on Thr-308 or Ser-473 (Downward, 1998b). Besides PTEN, ILK also affects the P3K-dependent regulation of Akt (Delcommenne et a l , 1998). Overexpression of ILK in epithelial cells results in an increased activity of Akt (Dedhar, 2000) due to phosphorylation of Akt on serine-473 by ILK (Delcommenne et a l , 1998). ILK and Akt are constitutively activated in cells lack PTEN expression (Obara et a l , 2004). ILK is thought critical for the PTEN-sensitive regulation of Akt-dependent cell cycle progression and cell survival (Persad et a l , 2000). ILK activity is PI3K and phosphoinositide-dependent (Delcommenne et a l , 1998; Lynch et a l , 1999; Persad et a l , 2000) and the kinase activity of ILK can be modulated by the interaction of cells with components of the extracellular matrix or by integrin clustering (Wu et a l , 1998) and growth factors (Delcommenne et a l , 1998; Wu et a l , 1998). The preliminary results show that aPPD does not affect levels of activated integrin in SF188 cells (data not shown), which is in consistent to the unchanged phosphorylation on Akt in these cells. I will further test the activity of integrin in U87 cells that showed significant reduction in levels of P-Akt after treatment with aPPD. Besides serine-473, phosphorylation on Akt at Thr308 also contribute to cell survival (Alessi et a l , 1997; Stokoe et a l , 1997; Datta et a l , 1999). I will also investigate whether aPPD alters Akt pathway in different aspects in both SF188 and U87MG cells. Nevertheless, decreased P-Akt in aPPD treated U87MG cells indicates that aPPD reduced activity of survival pathway in the cells. As inhibition in survival pathway has been proposed as a mechanism that causes apoptosis in cancer cells (Jiang et al., 2000; Larribere et al., 2004), it may play a role in the cell death observed in U87MG in which caspases pathway is not activated by aPPD. Further study will be conducted to test this possibility. CHAPTER 5 SUPEROXIDE ANION AND APOPTOSIS OF GLIOMA CELLS INDUCED BY aPPD INTRODUCTION One of the upstream signals activating JNK (c-Jun-N-terminal kinases) is reactive oxygen species (ROS) (Wilmer et a l , 1997; Du et a l , 2001; Filomeni et a l , 2003). ROS has also been known to induce apoptosis through both caspase-dependent and -independent mechanisms (Higuchi and Matsukawa, 1997; Devadas et a l , 2003; Kaul et a l , 2003; Sato et al , 2004). These characters make ROS production as a potential mechanism for aPPD-induced apoptosis in glioma cells. ROS include free radicals, which are any atom or molecule containing one or more unpaired electrons (Halliwell, 1994) and non-radical oxidants, such as the superoxide anion (02-~), hydroxyl radicals ("OH) and the non-radical hydrogen peroxide (H2O2) and hypochlorous acid (HOC1), et al (Lee and Choi, 2003). ROS are constantly generated under normal conditions as a consequence of aerobic metabolism. 02*~ represents a primary source of free radicals in biological systems (Fridovich, 1995). Under normal physiological conditions, 02'~ is generated by NAD(P)H oxidases (nicotinamide adenin dinucleotide phosphate reduced oxidases), xanthine oxidase, or as a side product of the mitochondrial respiratory chain (Boveris et a l , 1972; Boveris and Chance, 1973; Boveris and Cadenas, 1975; Boveris et a l , 1976; Cadenas and Davies, 2000) in intracellular sites including mitochondria, endoplasmic reticula, and nuclear membranes (Hockenbery et a l , 1993). Approximately 1% - 4% of all electrons channeled through the mitochondrial respiratory chain form 02*~ (Fridovich, 1995; Klein and Ackerman, 2003) or about 1-3% of the oxygen that we breathe in is used to make 02*~ (Halliwell, 1994). Since human beings consume a lot of oxygen, we may produce over 2 kg of superoxide in the our body every year (Halliwell, 1994). The generation of ROS in normal cells is under tightly homeostatic control (Curtin et al., 2002). The ( V - is converted to H2O2 by Mn- SOD (Mn-superoxide dismutases) in mitochondria or Cu, Zn-SOD in the cytosol. Further, there are two major defense systems, which are glutathione (GSH) peroxidase by the GSH redox cycle presenting in both the cytosol and mitochondria or catalase presenting only in the peroxisome fraction, transform H202to H 2 0 (Curtin et al., 2002). In addition, biological antioxidants, including glutathione, a-tocopherol (Vitamin E), carotenoids, and ascorbic acid (Vitamin C) can react directly with most oxidants, such as oxygen, superoxide anion radical and hydroxyl radical non-catalytically (Chan, 1993; Halliwell, 1994; Kamal-Eldin and Appelqvist, 1996; Kannan and Jain, 2000; Curtin et al., 2002). ROS generation has been reported to occur in TNF- and Fas-mediated apoptosis (Wolfe et al., 1994; Suzuki et al., 1998) as well as following treatment of cells with various agents including ultraviolet irradiation and chemotherapeutic drugs (Zamzami et al., 1995; Gorman et al., 1997), when ROS overcome the defense systems of the cell and redox homeostasis is altered, the result is oxidative stress. Oxidative stresses provoke physiological responses that induce the expression of various cytoprotective genes (Nguyen T, 2003) such as nuclear factor E2-related factor 2 (Nrf2). The transcription factor Nrf2 (Moi P et al., 1994), which ubiquitously expressed in a wide range of tissues and cell types (Chan et al., 1993; Moi P et al., 1994; McMahon et al., 2001), is identified as the major regulator of the cytoprotective genes encoding phase 2 detoxication enzymes such as glutathione S-transferase (GST) and NAD(P)H: quinone oxidoreductase (NQOl) (Itoh et al., 1997) and antioxidants through the antioxidant response element (ARE) (Itoh et al., 1997; Itoh K et al., 1999). ARE is a cw-acting enhancer sequence that mediates transcriptional activation of genes in cells exposed to oxidative stress (Rushmore et a l , 1991). Now, ARE sequences have been characterized within the proximal regulatory sequences of genes encoding the antioxidant enzymes GST (Rushmore et a l , 1990), NQOl (Favreau and Pickett, 1991), heme oxygenase-1 (HO-1) (Prestera T, 1995), and y-glutamylcysteine synthetase (Chan and Kwong, 2000). Besides, ARE induction by small-molecule activators such as /ert-butylhydroquinone (tBHQ) has been shown to protect cell lines against the oxidative insult initiated by dopamine, hydrogen peroxide (H 20 2), and glutamate (Duffy et a l , 1998; Li et a l , 2002). ROS have been shown to resulting in DNA degradation, protein denaturation, and lipid peroxidation (Higuchi, 2003). Although 02"~ is mild oxidant with a short free diffusion pathway of a few micrometers (Saran and Bors, 1994) and short half-lives, it is reasonably recognized as a pivot of intracellular oxidative chain events due to its more actively chemical reactivity. It is well known that the unpaired electrons alter the chemical reactivity of an atom or molecule, usually making it more reactive than the corresponding non-radical (Halliwell, 1994). 0 2 ' - is a precursor of more reactive species to promote the production of secondary derivatives such as H 2 0 2 and "OH in intracellular oxidative chain events (Ose and Fridovich, 1976; Fridovich, 1995). On one hand, 02"~ is rapidly converted within the cell to H 2 0 2 by the superoxide dismutases (SOD1, SOD2, and SOD3). On the other hand, H 20 2can react with reduced transition metals, via the Fenton reaction, to produce the highly reactive hydroxyl radical ('OH), a far more damaging molecule to the cell. In addition to forming H 2 0 2 and 'OH, 02"~ radicals can rapidly react with nitric oxide (NO) to generate cytotoxic peroxynitrite anions (ONOO"). Peroxynitrite can react with carbon dioxide, leading to protein damage via the formation of nitrotyrosine and lipid oxidation (Klein and Ackerman, 2003). The role of ROS, in particular, superoxide radical, in mediating apoptosis is well established. Superoxide anions do not exhibit direct apoptosis inducing potential (Ivanovas B et al., 2002), whereas 02*~, as a primary signaling molecule, promotes the production of ROS downstream, which act as signaling molecules further to cause apoptosis through a redox-sensitive pathway. ROS induce cell apoptosis by causing lipid peroxidation and DNA damage (Higuchi and Matsukawa, 1997). Furthermore, the redox-sensitive signaling may interact with more classical signaling pathways of apoptosis (Pahl and Baeuerle, 1994; Lander, 1997). It is verified that both exogenously and endogenously generated ROS served to promote activation of initiator caspases such as caspase-8 (Devadas et al., 2003), further to initialize classical apoptosis. Additionally, ROS generation has been reported to occur in TNF- and Fas-mediated apoptosis (Hoffman and Weinberg, 1987; Wolfe et al., 1994; Suzuki et al., 1998; Leong and Karsan, 2000). ROS is a key mediator in activating the members of the mitogen-activated protein kinase (MAPK) superfamily pathway such as ASK1 (MEKK5) and JNK, which is a key step for the signal transduction pathway that contributes to progression of cell apoptosis (Wilmer et al., 1997; Shan et al., 1999; Du et al., 2001). These effects suggest that such oxygen derivatives could be used for cancer chemotherapy in vitro and in vivo (Davies and Goldberg, 1987). It was proposed that ginsenosides that with no sugar moieties attached to the 20-position of the triterpene dammarane, such as ginsenoside Rg3, Rh2, and Rg2 act as a prooxidant (Liu et al., 2003). Some researcher found that Rh2-induced cell death is mediated by the generated ROS and the activation of caspase pathway in a Bcl-XL-independent manner (Kim et al., 1999a). Aglycones of ginsenosides, such as aPPD or aPPT, play a prooxidative role in 2,2'-azobis(2-amidinopropane hydrochloride)(AAPH) -induced hemolysis of erythrocytes (Liu et al., 2003). AAPH is a good inducer of free radical in experimental model to research the free radical-induced membrane damage (Miki et al., 1987). Here, effect of aPPD on production of ROS especially O2' in glioma cells and the relationship between C>2~ production and apoptosis in glioma cells were studied. RESULTS 1. Dose-response and time-course of superoxide anion production induced by aPPD in glioma cell lines SF188 cells or U87 M G cells were seeded (5xl05/well) in 6-well plates 24 hours before treatment. Levels of superoxide anion (02*~) was determined 2 hours following the aPPD treatment by dihydroethidine (HE) staining and measured with flow cytometry as described in Chapter 2. Fig.5-1 a and c showed that aPPD markedly increased intensity of 0 2 " i n both SF188 cells and U87 M G cells by 588 +/- 57.08 PO.001 and 651+/-82.11, P<0.001 in 86.8 and 130uM aPPD groups comparing with HE staining group respectively; In Fig.5-1 b and d, the percentages of (^'"positive cells significantly increased in 108 and 130uM aPPD-treated SF188 and U87MG cell cultures by 74.38+/- 0.79% or 65.45+/-9.37%, P<0.001 comparing with HE staining group. Statistical significance between control group and aPPD treated groups appeared when concentration of aPPD was above 86.8uM for SF188 cells and above 65.1uM for U87 cells. In addition, the results also showed (Fig. 5-1 f) that aPPD caused rapid increase in 02*" levels in SF188 cells that reached a plateau in first 2 hours. 2. Stimulation of antioxidant gene expression partially inhibited aPPD-induced apoptosis To determine whether ROS is responsible for aPPD-induced apoptosis, two experiments were performed. SF188 cells were treated with tBHQ 48 hours prior to the treatment with aPPD and cell viability was determined by MTT assay. As shown in Fig. 5-2, in the presence of tBHQ or together with the general caspase inhibitor (Z-VAC-FMK), SF188 cells were partially or completely protected from apoptosis induced by aPPD at the concentration as high as 86.8uM (Fig.5-2-1), although each of them alone only blocked cell death by 20%. However, tBHQ pre-treatment did not protect U87MG cells from aPPD-induced apoptosis (Fig. 5-2-2). Also, Vitamin C and E, both are known antioxidants failed to protect SF188 cells from aPPD caused cell death as shown in Fig.5-3. SF188 DoseofaPPD(uM) o a. SF188 Dose ofaPPD(uM) U 8 7 M G & & <£' sc? Dose of aPPD(uM) TJ87MG & ^ i > t*> * s * <vv %fa-Dose of aPPD(uM) Fig. 5 -1 Superoxide anion generation in glioma cells induced by aPPD. a, b, SF188 cells; c, d, U87 M G cells, both treated with aPPD for 2 hours, e and f, time course of SF188 cells treated with 108.4 uM aPPD. a, c and e show the mean fluorescence intensity of positive cells, b, d and f show the percentage of positive cells. The bars with * marker indicate P< 0.01 comparing to control group. 140 120 -LQQ-- • aPPD • GI • tHIIQ(lOuM) • tBHQ(10uM) + GI f Cell viability^ 80 60 40 20 * 86.8 Dose ofaPPD(uM) Fig. 5-2-1 Effect of lOuM tBHQ and 50uM Z-VAD-FMK (general caspase inhibitor, GI) on 86.8uM aPPD induced cell-death in SF188 cells (PO.01 in groups with star marker) 140 Dose of aPPD (ug/ml) Fig. 5-2-2 tBHQ had no effect on aPPD-induced cell death in U87MG cells. For each one in 30uMtBHQ-pretreated group compares with aPPD treatment P<0.01. Fig. 5-3 Vitamin C or E did not affect the aPPD-induced cell death in SF188 cells. The SF188 cells were pre-treated with Vitamin C or Vitamin E for 1 hour prior to aPPD treatment. Cell viability was determined with MTT assay at 24 hours or 16hous separately. The above results demonstrate that 1) aPPD causes production of superoxidase anion in both SF188 and U87 cells; 2) The effect of ROS can be partially prevented by pre-treatment with /ert-butylhydroquinone (tBHQ), a stimulator of antioxidant genes; 3) When caspases and ROS were both blocked, the cells were completely protected from aPPD induced apoptosis in SF188 cells. 4) Vitamin C or E had no effect on cell death induced by aPPD. Vitamin C and Vitamin E are known to constitute a strong line of defense in retarding free radical induced cellular damage, amongst the antioxidants Vitamin C plays a central role as it functions in vivo to repair the membrane-bound oxidized vitamin E as to the regeneration vitamin E (Chan, 1993). Why Vitamin C or E did not protect glioma cells from aPPD-induced apoptosis? One explanation could be the complicated mechanism of Vitamin C and E for their antioxidant effects (Abudu et al., 2004). Although Vitamin E acts as a free radical scavenger that can directly react with oxygen, superoxide anion radical and hydroxyl radical (Kamal-Eldin and Appelqvist, 1996), due to its lipid solubility, it is mainly a chain breaking antioxidant within the lipoprotein. When Vitamin E reacts with a lipid peroxyl radical in a single electron reaction, a Vitamin E radical will produce (Abudu et al., 2004). If Vitamin E radical reacts with a second peroxyl radical or with another Vitamin E radical, the reaction comes to termination (Kamal-Eldin and Appelqvist, 1996). When this reaction cannot be terminated, Vitamin E radical forms to act as a pro-oxidant which is mediated through the one-electron oxidation catalysed by peroxyl radicals or by the reductive activation of Cu(II) to Cu(I) with the concomitant formation of the a-tocopheroxyl radical (Witting et al., 1995; Bagnati et al., 1999). Similarly, Vitamin C at low concentrations or the presence of redox-active transition metal ions (S. Udenfriend et al., 1954) can also undergo a one electron reaction to form an ascorbyl radical (Ashidate et al., 2003; Abudu et al., 2004). In rat liver homogenates, higher concentrations (1 and 3 mM) of ascorbate suppressed lipid peroxide productions but lower concentrations (0.1 and 0.3 mM) did not (Song et al., 2003). Additionally, Vitamin E involves up-regulation of TGF-beta receptor II expression and TGF-beta-, Fas-and JNK-signaling pathways to induce tumor apoptosis. In some senses, these data provide a better understanding of the anticancer actions of a dietary form of vitamin E (Shun et al., 2004). Since the intracellular concentrations of vitamin C and E or the distribution of the two agents were not measured, it was not ruled out if the levels of the two agents were not high enough in a subcellular location to block the chain of oxidation in the aPPD treated cells. The above results showed that tBHQ partially prevented aPPD-induced cell death. tBHQ has antioxidation activity, but unlike Vitamins, it does not directly react with ROS. tBHQ induces expression of several phase II detoxifying enzymes such as NADPHrquinone oxidoreductase (NQOl) and glutathione S-transferase (GST) via the transcriptional activation of antioxidant-responsive element (ARE) (Rushmore and Pickett, 1990; Prestera and Talalay, 1995), which contributes to the protection of cells from oxidative stress (Johnson et al., 2002). Activation of ARE by tBHQ is dependent on Nrf2 but not oxidative stress itself (Lee et al., 2001). Oxidative stress, electrophiles or tBHQ induce the release of Nrf2, resulting in nuclear translocation of Nrf2 and subsequent activation of ARE (Itho et al., 1999; Kraft et al., 2004) by binding to the ARE and mediate ARE-regulated gene expression such as induction of NQOl and other detoxifying enzyme genes (Venugopal and Jaiswal, 1996; Alam et al., 1999; Nguyen et al., 2000) to destroy ROS and decrease oxidative stress. It is well known that high levels of Nrf2 in glial cells (Shih et al., 2003) or tBHQ activated Nrf2 (Kraft et al., 2004) can protect neural cells from a variety of oxidative stressors. The results showed that ROS is involved in glioma cell apoptosis induced by aPPD, which is in consistence with some early work using Rh2, a precursor of aPPD (Kim et a l , 1999a). Most importantly, tBHQ and general caspase inhibitor together (Z-VAD-FMK) could completely block the apoptosis induced by aPPD in SF188 cells while two agents alone had only partial protection. These results indicated that aPPD-induced apoptosis in SF188 cells is mediated by at least two mechanisms: activation of caspases and production of ROS. Since inhibitors for each mechanism did not block entire apoptosis but combination use of both did, and since production of ROS did not cause activation of caspases as seen in U87MG cells, it is clear that the two mechanisms are active simultaneously and independent to each other. It is not clear why tBHQ partially protected SF188 but not U87MG cells from aPPD induced apoptosis. One possibility is that U87MG cells rely on the survival pathway. Any weakening on this pathway such as inhibition in Akt phosphorylation is sufficient to kill the cells. Thus, for its apoptosis, ROS caused damage becomes redundant when the survival pathways are blocked. Further study is needed to test this hypothesis. CHAPTER 6 GENERAL DISCUSSION AND CONCLUSION As described in the previous chapters, I have investigated possible mechanisms of aPPD-induced apoptosis in SF188 and U87MG glioma cells. Those results can be summarized as follows: 1. aPPD induces tumor cell-death in a p53- and PTEN-independent manner. 2. aPPD-induced cell death is largely apoptotic. 3. In some cells such as SF188, aPPD activates multiple caspases, especially caspase 8, which contributes to apoptosis. 4. Activation of caspases requires a cellular membrane component. 5. In other cells, such as U87MG, aPPD induces apoptosis through a caspase-independent mechanism. 6. Other possible mechanisms involved in aPPD-induced apoptosis may include: inhibited cell survival pathway by inhibiting Akt phosphorylation and generation of ROS. Figure 6-1 is a diagram of hypothesized mechanism for aPPD-induced apoptosis based on my results. aPPD acts on a cellular membrane component to activate multiple intracellular signal pathways. In SF188 cells, it activates caspase 8 and consequently other caspases. It also causes over production of ROS that can induce apoptosis through both caspase and non-caspase pathways. In U87MG cells, the mechanism is not clear, but aPPD can inhibit phosphorylation of Akt to block the survival pathway, which can also cause apoptosis. While the above diagram can explain some phenomena observed in both cell lines treated with aPPD, there are many questions remain unanswered and the followings are just a few: First of all, although it is demonstrated that there is a component in the cellular membrane interacting with aPPD to cause caspase activation, it is not still known what it is exactly. Based on the chemical structure of aPPD and its hydrophobic nature, it is possible that the molecule will interact with a lipid or lipid proteins in the membrane. Identifying the target molecule for aPPD is very important, as it not only fill in the unknowns for its mechanism, it may also explain the specificity of this compound for cancer cells as animal toxicity tests have shown that aPPD is not toxic in vivo at all. Secondly, although it is found that ROS is overproduced in aPPD-treated cells, we still have not identified the exact species of the free radicals yet. It is further puzzling that tBHQ could protect cells from apoptosis but not Vitamin C or E. Since tBHQ acts on an indirect mechanism to protect cells from ROS damage, more direct evidences including the source of ROS must be sought. Thirdly, it is not known what make U87MG cells different from SF188 in their response to aPPD. It is particularly interesting tp find why aPPD can activate caspases in SF188 but not in U87MG. The relations between caspase activation and PTEN /P53 status should be further studied. I will apply some methods such as siRNA interruption of PTEN mRNA or dominant-negative PTEN to different glioma cell lines to dtermine the effection of PTEN on cell' response to aPPD treatment. This may provide a model system to study the mechanism of caspase activation in cancer cells. Nevertheless, the present study has clearly shown that aPPD possesses some important therapeutic characteristics that it activates multiple pro-apoptotic pathways and inhibits pro-growth signaling protein simultaneously. Results and conclusions obtained from the present study render further investigation for potential development of the compound into an anti-cancer drug. Fig. 6-1 Schematic representation of proposed intracellular signaling leading to apoptosis by aPPD in glioma cells. This overview is based on the present findings as well as those previously published by other investigators and does not exclude participation of other signaling pathways or complementary signals within the presented diagram. The activated signaling component related to apoptosis induced by aPPD is in black lines, and the signaling events inhibited are in green dash lines. 1. When aPPD acts on cells, it decreases phosphoralated Akt level and releases cell from inhibition of caspase activation and ROS stress damage from P-Akt, the result is that cell survival is weaken and apoptosis occurs; 2. aPPD via cell membrane system induces multiple caspase activation and induces cell apoptosis; and 3. aPPD quickly increases superoxide anion production , and it can cause cell death with or without activating caspases. 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