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Molecular functions of the tumour suppressor p33ING2 Chin, Mei Yieng 2004

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MOLECULAR FUNCTIONS OF THE TUMOUR SUPPRESSOR p33ING2  by MEI YIENG CHIN B . S c , University Putra Malaysia, 2000 A thesis submitted in partial fulfillment of the requirements for the degree of MASTER OF SCIENCE in T H E F A C U L T Y OF G R A D U A T E STUDIES (Department of Medicine; Experimental Medicine Program) W e accept this thesis as conforming to the required standard  T H E UNIVERSITY O F BRITISH C O L U M B I A August 2004 © Mei Yieng Chin, 2004  THE UNIVERSITY OF BRITISH  COLUMBIA  F A C U L T Y OF G R A D U A T E STUDIES  Library Authorization  air  1  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.  MEI YIENG CHIN  August 27, 2004  Name of Author (please print)  Date (dd/mm/yyyy)  Title of Thesis:  Molecular Functions of the tumour suppressor p33ING2  Degree:  Master of Science  Department of  Medicine  Year:  2004  The University of British Columbia Vancouver, BC Canada  grad.ubc.ca/forms/?formlD=THS  page 1 of 1  last updated: 27-Aug-04  ABSTRACT  Since p33/A/G2 was cloned in 1998, there have been several studies indicating that p33ING2 is a tumour suppressor candidate as it regulates gene transcription, cell cycle arrest and apoptosis in various human carcinoma cell lines. These functions have been shown to be dependent on the activity of wild-type p53. p33ING1 b, which shares 58.9% homology with p33ING2, has been shown to play a role in cellular stress response to UV irradiation.  Due to the structural similarities  between  p33ING1b and p33ING2, we hypothesized that p33ING2 enhances UVB-induced apoptosis. induced  To test this hypothesis, we investigated the role of p33ING2 in UV-  apoptosis  and the  repair  of  UV-damaged DNA.  We  report that  overexpression of p33ING2 enhances UVB-induced apoptosis in wild-type  p53  M M R U melanoma cells, but not in mutant p53 M e W o melanoma cells.  We  demonstrate that the enhancement of UVB-induced apoptosis by p33ING2 requires the presence of functional p53.  Furthermore, we found that overexpression of  p33ING2 upregulates the expression of endogenous Bax and downregulates the expression of Bcl-2, resulting in an increased Bax/Bcl-2 ratio.  Moreover, we also  found, that p33ING2 promotes Bax translocation to the mitochondria, alters the mitochondrial membrane potential, and induces cytochrome c release, thus activating caspases 9 and 3 upon UV irradiation. In addition, we show that under non-stressed conditions, p33ING2 upregulates Fas expression and activates caspase 8. These results indicate that p33ING2 cooperates with p53 to regulate apoptosis via activation of both the mitochondrial/intrinsic and death-receptor/extrinsic apoptotic pathways.  To investigate if p33ING2 is also involved in DNA repair in melanoma cells, we transfected the p33ING2 expression vector into melanoma cells.  W e report that  overexpression of p33ING2 enhances the repair of UV-damaged DNA, and that this process requires the activity of functional p53. W e also show that p33ING2 is not directly recruited to UV-induced DNA lesions, both cyclobutane pyrimidine dimers (CPDs) and 6-4  photoproducts  (6-4PPs), suggesting that p33ING2 is not a  component of the repair core complexes.  Furthermore, we found that p33ING2  enhances p53 acetylation and upregulates the expression of X P A , which is involved in DNA damage recognition. These observations suggest that p33ING2 cooperates with p53 in mediating nucleotide excision repair, and that X P A may be involved in this repair process. In conclusion, we have elucidated in this thesis, the role of p33ING2 in cellular stress response to UV irradiation, and the importance of this gene in the context of tumour suppression.  TABLE OF CONTENTS Abstract Table of Contents List of Tables List of Figures List of Abbreviations Acknowledgements C H A P T E R 1 INTRODUCTION 1.1  1.2  1.3  1.4  Skin Cancer 1.1.1 Types of Skin Cancer 1.1.2 The Aetiology of Skin Cancer p53, The Guardian of The Genome 1.2.1 Gene and Protein Structures 1.2.2 Cell Cycle Arrest 1.2.3 Apoptosis 1.2.4 DNA Repair 1.2.5 Post-translational Modifications of p53 The ING Tumour Suppressor Family 1.3.1 p33ING1b 1.3.2 p33ING2 1.3.2.1 Gene and Protein Structures 1.3.2.2 Expression Profile 1.3.2.3 Biological Functions 1.3.2.4 Modes of Action Objectives and Hypotheses  C H A P T E R 2 MATERIALS AND METHODS 2.1  2.2  Materials 2.1.1 Cell lines and Cell Culture 2.1.2 Plasmids 2.1.3 Antibodies Methods 2.2.1 Transfection 2.2.2 Determination of Transfection Efficiency 2.2.3 UV Irradiation 2.2.4 Light Microscopy 2.2.5 S R B Cell Survival Assay 2.2.6 Trypan Blue Exclusion Assay 2.2.7 Propidium Iodide Staining iv  2.2.8 2.2.9 2.2.10 2.2.11 2.2.12 2.2.13 2.2.14 2.2.15  Flow Cytometry Western Blot Analysis Detection of the Mitochondrial Membrane Potential Subcellular Fractionation Host-cell-reactivation Assay Immunofluorescent Staining Detection of p53 Acetylation Immunoprecipitation  C H A P T E R 3 p33ING2 E N H A N C E S UVB-INDUCED A P O P T O S I S IN HUMAN MELANOMA CELLS 3.1 3.2  3.3  37 37 38 39 40 41 42 43 44  Rationale and Hypothesis 44 Results 47 3.2.1 p33ING2 Enhances UVB-induced Apoptosis in Melanoma 47 Cells 3.2.2 p33ING2 Enhancement of UVB-induced Apoptosis Is p53- 53 dependent 3.2.3 p33ING2 Increases the Ratio of Bax/Bcl-2 in Melanoma Cells 58 3.2.4 p33ING2 Alters Mitochondrial Membrane Potential after U V B 60 Irradiation 3.2.5 p33ING2 Promotes Bax Translocation and Cytochrome c 61 Release after UVB Irradiation 3.2.6 p33ING2 Induces the Activation of Caspase 9 and Caspase 3 63 after UVB Irradiation 3.2.7 p33ING2 Activates the Death-receptor Apoptotic Pathway 65 Discussion 67  C H A P T E R 4 p33ING2 E N H A N C E S T H E R E P A I R OF U V - D A M A G E D DNA IN HUMAN MELANOMA CELLS  74  4.1 4.2  74 77 77 77 79 84 84 85  4.3  Rationale and Hypothesis Results 4.2.1 p33ING2 Enhances the Repair of UV-damaged DNA 4.2.2 p33ING2 Does not Colocalize with UV-induced DNA Lesions 4.2.3 p33ING2 Enhancement of DNA Repair Is p53-dependent 4.2.4 p33ING2 Enhances p53 Acetylation upon UV Irradiation 4.2.5 p33ING2 Upregulates X P A Discussion  CHAPTER 5 GENERAL CONCLUSIONS  92  5.1 5.2  92 94  Summary Future Directions  REFERENCES  96  v  LIST OF TABLES  Table 1.1  Relative percentages of transfection efficiencies of cell lines used in the study  vi  34  LIST OF FIGURES  Figure 1.1  Comparison of the amino acid sequences of p33ING1b and P33ING2  20  Figure 1.2  Schematic diagrams of the p33ING1band p33ING2  21  Figure 1.3  Biological functions of p33ING1 b and p33ING2  28  Figure 3.1  p33ING2 enhances UVB-induced cell death in melanoma M M R U cells  49  Figure 3.2  p33ING2 enhancement of UVB-induced apoptosis is p53dependent  55  Figure 3.3  p33ING2 upregulates Bax and downregulates Bcl-2 expression  59  Figure 3.4  p33ING2 alters mitochondrial membrane potential after U V B irradiation  62  Figure 3.5  p33ING2 promotes Bax translocation and cytochrome c release  64  Figure 3.6  p33ING2 induces the activation of caspases 9 and 3  66  Figure 3.7  p33ING2 activates the death-receptor apoptotic pathway  68  Figure 4.1  p33ING2 enhances the repair of UV-damaged DNA  78  Figure 4.2  p33ING2 does not colocalize with UV-induced DNA lesions  80  Figure 4.3  p33ING2 enhancement of DNA repair is p53-dependent  82  Figure 4.4  p33ING2 enhances p53 acetylation and upregulates X P A  86  vii  protein structural  features  of  LIST OF ABBREVIATIONS  6-4PPs aa AFP Apaf-1 AS ATM ATR BCC BER BH-3 CAT CBP Cdk Chk CPDs CS DDB DISC FADD FAT FISH GADD45 HAT HDAC HRP lAPs ING JNK LOH MAPK MDM2 mut NER NLS NTS p53AIP1 PBS PCAF PCNA PHD PI PIAS1 PtdlnsPs  6-4 photoproducts Amino acid oc-fetoprotein Apoptotic protease activating factor-1 Antisense Ataxia telangiectasia mutated kinase ATM-related kinase Basal cell carcinoma Base excision repair Bcl-2 homology 3 Chloramphenicol acetyltransferase C R E B binding protein Cyclin-dependent kinase Checkpoint kinase Cyclobutane pyrimidine dimers Cockayne syndrome Damaged DNA binding protein Death-inducing signalling complex FAS-associated death domain Factor acetyltransferase Fluorescent in situ hybridization Growth arrest and DNA damage protein Histone acetyltransferase Histone deacetylase complex Horseradish peroxidase Inhibitors of apoptosis Inhibitor of growth Jun amino-terminal kinase Loss of heterozygosity Mitogen-activated protein kinase Mouse double minute 2 Mutant Nucleotide excision repair Nuclear localization sequence Nucleolar translocation sequence p53-regulated apoptosis-inducing protein 1 Phosphate buffered saline p300/CBP-associated factor Proliferating cell nuclear antigen Plant homeodomain Propidium iodide Protein inhibitor of activated S T A T 1 Phosphoinositides  viii  PTP PUMA PVDF Rb ROS RPA SAID SAP30  see  Smac SRB TH TPEN TR TNF UV wt XP  Permeability transition pore p53-upregulated modulator of apoptosis Polyvinylidene diflouride Retinoblastoma protein Reactive oxygen species Replication protein A SAP30-interacting domain Sin3-associated protein 30 Squamous cell carcinoma Second mitochondria-derived activator of caspase Sulforhodamine B Thyroid hormone N N,N ,N'-tetrakis(2-pyridyl-methyl)ethylenediamine zinc chelator Thyroid receptor Tumour necrosis factor Ultraviolet Wild-type Xeroderma pigmentosum ,  )  ix  ACKNOWLEDGEMENTS  I would like to express my gratitude and sincere appreciation to my supervisor, Dr. Gang Li, for giving me the opportunity to set my first step in the research career, for your intellectual stimulation and continual support.  To Dr. Vincent Ho, Dr. Paul  Rennie and Dr. Chris Ong, I want to thank you for being the members of my committee, and for your suggestions as well as feedback on my research work. To my dear friends and the members of the Li lab, those who have graduated as well as those who are still persevering, your encouragement, support and help are greatly appreciated. Without you I could not have done this. To my family, I am so grateful for your understanding and moral support.  Your unconditional love has kept me  going through the ups and downs of these years. Last but not least, I would also like to acknowledge University Malaysia Sarawak for sponsoring my study.  x  CHAPTER 1. INTRODUCTION  1.1  Skin Cancer  Skin cancer is the most common cancer among all types of cancers (Boni et al., 2002). The annual incidence of skin cancer in the U.S. is estimated at about one million, accounting for about 4 5 % of all human malignancies (Landis et al., 1999). In Canada, approximately 76,000 and 4,250 new cases of non-melanoma skin cancer and melanoma respectively were diagnosed for 2003 (National Cancer Institute of Canada: Canadian Cancer Statistics 2004, Toronto, Canada, 2004).  1.1.1  Types of Skin Cancer  Skin cancers are categorized into two main types based on the cellular origin: nonmelanoma skin cancer, including basal cell carcinoma (BCC) and squamous cell carcinoma (SCC) that derives from keratinocytes, and melanoma that derives from melanocytes. Non-melanoma skin cancer is the most common skin cancer, with B C C accounting for 60% of total skin cancer cases, and S C C accounting for 30-35%. Both B C C and S C C are found mainly on sun-exposed parts of body, and their incidence correlates well with cumulative sunlight exposure.  Both B C C and S C C  show low fatality, as B C C rarely metastasize and likewise S C C shows a very low rate of metastasis. Furthermore, surgical therapy and radiotherapy are highly effective in treating these diseases.  Mutations of the p53 tumour suppressor gene have been  found frequently in B C C and S C C , and the majority of these mutations are U V specific C-VT and CC-»TT transitions. These mutations are also observed in B C C  1  precursor (basal cell nevus syndrome) and S C C precursor (actinic  keratosis),  indicating that p53 mutation may be an early event in the pathogenesis of these cancers (Kanjilal et al., 1995; Ponten et al., 1997; Sim et al., 1992; McGregor et al., 1992). In B C C , allelic loss of the patched gene (PTC) locus on chromosome 9q22 has been reported and may be responsible for basal cell carcinogenesis (Gailani et al., 1996).  Recent studies of sporadic basal cell carcinomas showed loss of  heterozygosity (LOH) on chromosome 4q32-35, mapped to tumour suppressor p33/A/G2 and SAP30 genes, the gene product of which is involved in transcription repression (Sironi et al., 2004). In contrast to the predominance of L O H on specific chromosomes, S C C shows more widespread L O H with frequent loss of markers from chromosome 3p, 9p, 13q, 17p and 17q (Quinn etai, 1994). Melanoma contributes to 2-5% of total skin cancer cases.  Melanoma  development is believed to be associated with intermittent acute sunlight exposure causing severe sunburns particularly during childhood or adolescence (Holman and Armstrong, 1984; Kricker et al., 1995).  Melanoma is associated with one of the  highest mortality rates due to its rapid metastasis, and the 5-year survival rate of patients with metastatic melanoma remains less than 10% (Roses et al., 1991). There is no effective treatment for metastatic melanoma. Unlike non-melanoma skin cancers, p53 mutation is observed in only 1-5% of primary melanoma and 11-25% of metastatic melanoma (Hussein et al., 2003a). Deletion of the chromosomal region 9q21, harbouring tumour suppressors p'lG "* * 1  4  and p 1 4 ^ , is among the most RF  common known genetic alterations detected in about 50% of familial melanoma, and is also observed in 10 to 40% of sporadic melanoma (Healy et al., 1996; Rizos et al.,  2  2001). However, loss of p1 Q  INK4a  rather than p14 ~ appears to be a more consistent AHI  target in familial melanoma (FitzGerald ef al., 1996).  Methylation in the promoter  region of INK4a that could potentially silence transcription has also been reported in 10% of melanoma cases (Gonzalgo et al., 1997). In addition, the protein expression of p'\4  ARF  is decreased from benign melanocyte nevi to metastatic melanoma,  suggesting that inactivation  of p 1 4 ^  is important in melanoma  RF  (Dobrowolski ef al., 2002). Inactivation of p16  ,  INK4a  progression  an inhibitor of cyclin-dependent  kinase 4 (Cdk4), leads to deregulation of the activity of Rb and E2F-1 transcription factor and thus stimulates cell cycle progression; whereas loss of function of p 1 4 ^ , RF  an inhibitor of mdm-2, disrupts the stability of p53 (de Grujil et al., 2001). Other genes, such as Apaf-1 and BRAF have also been implicated in the development of melanoma. Apaf-1 (apoptotic protease activating factor-1) is a proapoptotic protein that forms complexes with cytochrome c and pro-caspase 9 to mediate apoptosis.  Allelic deletion of Apaf-1 is observed in 4 0 % of malignant  melanoma cases (Hussein et al., 2002a). Loss of Apaf-1 expression is associated with resistance to chemotherapy in metastatic melanoma as cancer cells are able to escape from p53-dependent apoptosis (Soengas et al., 2001). More recent findings reported that missense mutations of the B R A F oncogene are detected in 66% of cultured and primary melanoma cells (Davies et al., 2002). The V 5 9 9 E mutation of B R A F is thought to mimic phosphorylation and activation of B R A F , resulting in a protein with high basal kinase activity, leading to constitutive activation of the R A S / E R K survival pathway (Mercer and Pritchard, 2003).  3  1.1.2  Aetiology of Skin Cancer  Sunlight is a continuous spectrum of electromagnetic radiation that can be divided into three major regions of wavelength: infrared, visible and ultraviolet (Jhappan et al., 2003). The ultraviolet portion of the solar spectrum, lying in the wavelength range of 200-400 nm, is undoubtedly the major factor in skin cancer. UV radiation can be subdivided into three wavelength ranges: U V A (320-400 nm), U V B (280-320 nm) and U V C (200-280 nm).  Absorption of UV radiation by stratospheric ozone greatly  attenuates these wavelengths, preventing U V radiations with a wavelength shorter than 300 nm (UVC and most UVB) from reaching the earth's surface. Although U V A and UVB light constitute a minute portion (0.0000001%) of the emitted sunlight, they are primarily responsible for the sun's pathological effects (Cleaver and Mitchell, 2001). U V A is the predominant component of sunlight to which humans are exposed. However, its role in skin cancers, including melanoma, is not as well documented as the role of U V B . Since U V A has a longer wavelength than U V B , it can penetrate deeper into the skin, reaching melanocytes in the basal and suprabasal layers of the skin.  U V A is photo-carcinogenic and is involved in photo-aging, but it is weakly  absorbed in DNA and protein. U V A primarily produces reactive oxygen species and free radicals, which react with DNA to form base damages, strand breaks and DNAprotein cross-links, and ultimately mutations (Tyrrell and Keyse, 1990). An example of UVA-specific mutation, 8-hydorxyguanine, is produced though oxidation of guanine (Runger et al., 1995).  UVB is largely responsible for skin cancer through direct  photochemical damage to DNA, which absorbs UV radiation within the wavelength  4  range of 245-280 nm (Tornaletti and Pfeifer, 1996). Although U V C is not supposed to be present in ambient sunlight, depletion of the ozone layer allows residual U V C radiation to reach the earth's surface. Besides, U V C is also readily produced by lowpressure mercury sterilizing lamps.  Therefore its biological consequences are  gaining more attention. Both UVB and U V C preferentially cause DNA damage at the sites of neighbouring pyrimidine bases in a DNA strand, forming dimers between these bases.  The two major photoproducts of such are the cyclobutane pyrimidine dimer  (CPD), formed specifically between adjacent thymine (T) or cytosine (C) residues, and the 6-4 photoproduct (6-4PP), generated between adjacent pyrimidine residues. C P D s and 6-4PPs can be removed from the genome through nucleotide excision repair (NER) pathway, which is involved in the removal of bulky DNA adducts induced by UV irradiation.  N E R is a complex process with more than 30 gene  products participating in multiple step enzymatic reaction: recognition of DNA lesion, single strand incision at both sides flanking the lesion, excision of the lesions containing  24-29  nucleotides,  DNA repair  synthesis to  replace the  excised  nucleotides, and ligation of the remaining single-stranded nick (Ichihashi et al., 2003; Hanawalt et al., 2003). Both types of photolesions, if not removed from the genome, can cause UV-specific mutations such as C-»T and CC->TT transitions, leading to cancer development.  C P D s are considered to be more carcinogenic than the 6-  4 P P s , forming almost three times as often and being repaired less  efficiently  (Rosenstein and Mitchell, 1987; You etal., 2001). For instance, 6-4PPs are the more rapidly repaired: 50% of 6-4PPs are removed from DNA in 2 to 6 hours, whereas  5  C P D s are much more slowly repaired: half of C P D s are removed from DNA in 12 to 24 hours (Freeman, 1988; Cleaver, 1989). The different rates of repair may be due to the fact that 6-4PPs cause considerably more distortion in DNA structure, and that they are preferentially located in the inter-nucleosomal regions of DNA, which are more accessible for damage recognition proteins.  C P D s are distributed  more  randomly but are preferentially formed on DNA at the opposite side of the DNAprotein contact surface wrapped around the nucleosomes (Gold and Smerdon, 1990; Mitchell era/., 1990).  1.2  p53, The Guardian of The G e n o m e  p53 is often referred to as "the guardian of the genome" or "the cellular gatekeeper", revealing its importance in maintaining the integrity of the genome.  In response to  genotoxic stresses, p53 is stabilized and activated to (1) promote cell cycle arrest, which inhibits the replication of the damaged genomes, (2) repair damaged DNA, and (3) induce apoptosis or senescence, eliminating defective cells. p53 achieves these goals via its sequence-specific transcriptional activation and repression activity, as well as via interaction with other proteins (Janus er al., 1999; Hussein et al., 2003a; Brooks and G u , 2003; Ho and Benchimol, 2003).  The significance of the tumour  suppressive functions of p53 is reflected by the fact that p53 is the most frequently mutated gene known to date, with over 50% of human malignancies harbouring mutations and allelic deletions of p53 (Vogelstein and Kinzler, 1992; Lee et al., 1994; Soussi et al., 1994). Furthermore, germline mutations of p53 result in Li-Fraumeni  6  syndrome, a hereditary cancer susceptibility syndrome that predisposes individuals to various cancers (Malkin era/., 1990; Srivistava et al., 1990).  1.2.1  Gene and Protein Structures  The p53 gene is located on chromosome17q13.1. This gene encodes a 393-amino acid, 53 kDa nuclear phosphoprotein. The protein can be divided structurally and functionally into different domains. transcriptional  activation  transcriptional  co-activators  The N-terminus of p53 protein contains a  domain, which (Balint  and  is  essential for  Vousden,  the  2001).  recruitment  of  Following  the  transactivation domain is a proline-rich domain, which is required for transactivationindependent pro-apoptotic functions of p53 and transcriptional repression (Chipuk et al., 2004; Ho and Benchimol, 2003).  The DNA binding domain (DBD) of p53  spanning 200-aa residues is found after the proline-rich domain. The p53 consensus binding  site  consists  of  two  copies  of  the  10-bp  motif,  5'-  PuPuPuC(A/T)(T/A)GPyPyPy-3', separated by 0-13 bp (El-Deiry et al., 1992). The C-terminus of p53 contains a nuclear localization signal, an oligomerization domain, which is required for the tetramerization function of p53, and a basic C-terminal regulatory domain. p53 is normally a short-lived protein and is maintained at low levels within the cells under non-stressed conditions. Mdm-2, a transcriptional target of p53, plays a major role in this process by regulating p53 protein stability. Mdm-2 contains an E3 ligase domain that targets both p53 and itself for ubiquitination and degradation. Mdm-2 also inhibits the p53 functions through binding to the N-terminal of p53 and  7  thus inhibiting p53 transcriptional activity. The ubiquitin ligase activity of mdm-2 also contributes to the efficient nuclear export of p53 (Balint and Vousden, 2001; X u , 2003).  Therefore, p53-mdm-2 interaction presents a negative  autoregulatory  mechanism to regulate p53 stability and activity.  1.2.2  Cell Cycle Arrest  It is well documented that the involvement of p53 in cell cycle regulation depends on its transcriptional activation activities. One of the many p53 target genes, o2V , af1  a  cyclin-dependent kinase (Cdk) inhibitor, is believed to play an important role in the induction of both G  r  and G -phase cell cycle arrests (Balint and Vousden, 2001). 2  For instance, upon a DNA damage event, p53 upregulates p 2 1  w a f 1  to inactivate  Cdk2/cyclin E complexes and prevent phosphorylation of Rb, resulting in the inhibition of cell cycle progression from G i to S phase (Harper et al., 1993; El-Deiry er al., 1993). Alternatively, but not mutually exclusively, p 2 1  w a f 1  binds to the proliferating  cell nuclear antigen (PCNA), preventing association of P C N A with the  major  replicative enzyme DNA polymerase 8, thereby inhibiting DNA replication (Prosperi, 1997; Smith et al., 2000). It is believed that p53 induces G i arrest to provide more time for the repair of DNA damage before entry into S phase for D N A replication. Another target gene of p53, 14-3-3a is involved in p53-induced G arrest, by 2  binding to and sequestering cdc2-cyclin B1 complexes in the cytoplasm, and thus preventing cells from entering mitosis (Hermeking et al., 1997; Muslin and Xing, 2000). 14-3-3a can also bind to p53 and activate its sequence-specific DNA binding, presenting a positive feedback regulation of p53 to prevent cell cycle progression in  8  damaged cells (Waterman et al., 1998). Cdc2 can also be inhibited by other p53 downstream targets, such as p 2 1  waf1  , GADD45 (growth arrest and DNA damage  protein) and Reprimo (Wang et al., 1999; Taylor and Stark, 2001; Ohki et al., 2000).  1.2.3  Apoptosis  p53 induces apoptosis when DNA damage in a cell is beyond repair.  Evidence  indicates that p53 can mediate apoptosis by both transcription-dependent and transcription-independent mechanisms (Prives and Hall, 1999; Chipuk et al., 2004). The importance of p53 transcriptional activities in mediating apoptosis is reflected by the findings that cells in which the wt p53 gene was replaced by a transcriptionally inactive mutant showed loss of both cell cycle and apoptotic functions (Chao et al., 2000; Jimenez et al., 2000). p53 is capable of transactivating numerous apoptotic genes including those responsible for the mitochondrial/intrinsic pathway and deathreceptor/extrinsic pathway. The first p53 apoptotic target gene identified was the Bax gene, a pro-apoptotic member of the Bcl-2 family (Miyashita and Reed, 1995). Other pro-apoptotic members from this family, P U M A (p53-upregulated modulator of apoptosis) and Noxa have since been identified as p53 downstream targets (Oda et al., 2000a; Nakano and Vousden, 2001; Yu et al., 2001). These proteins, together with another p53 target, p53-regulated apoptosis-inducing protein 1 (p53AIP1), localize to the mitochondria and promote changes of the mitochondrial membrane potential and the release of cytochrome c, which complexes with Apaf-1 to form the apoptosome (the functional apoptotic unit) and activate the caspase cascade  9  (Hengartner, 2000; Hussein et al., 2003b).  More recently, Apaf-1 has also been  found to be a transcriptional target of p53 and E2F1 (Moroni er al., 2001). Besides transcriptional activation of pro-apoptotic genes, p53 transcriptional repression of anti-apoptotic genes also contributes to the apoptotic effects of p53. Bcl-2, Bcl-X , Survivin, MAP4 and PIK3CA (the p110a subunit of PI3K) are some of L  the genes identified as repression targets of p53 during p53-mediated apoptosis (Ho and Benchimol, 2003; Oren, 2003). There is also evidence indicating that p53 itself can localize to the mitochondria, and promote Bax translocation to the mitochondria as well as oligomerization of Bax to initiate apoptosis, presenting an additional transcriptional-independent way of mediating apoptosis (Marchenko et al., 2000; Schuler er al., 2000; Moll and Zaika, 2001; Chipuk et al., 2004). In addition, p53 is also involved in the death-receptor pathway by upregulating death receptors F a s / A P 0 1 and DR5/KILLER, a death receptor ligand, FasL, and a death domain-containing protein, PIDD (Owen-Schaub et al., 1995; W u et al., 1997; Lin et al., 2001).  The DR5 promoter is a direct target of p53, while cell surface  expression of Fas is enhanced by p53 through promotion of its trafficking from the Golgi apparatus to the plasma membrane (Bennett et al., 1998; Takimoto and E l Deiry, 2000). Activation of death receptors by their ligands results in trimerization of the receptors and recruitment of FAS-associated death domain (FADD), which further recruits pro-caspase 8 to form death-inducing signalling complex (DISC).  Pro-  caspase 8 in DISC is activated by autocatalytic cleavage and the active caspase 8 activates effector caspases such as caspase 3 to initiate apoptosis (Ashkenazi and Dixit, 1998; Igney and Krammer, 2002).  10  1.2.4  D N A Repair  In addition to preventing replication of cells with damaged genomes via the induction of cell cycle arrest and apoptosis, p53 also participates in the repair of damaged DNA. Evidence indicates that p53 is involved in both N E R that removes UV-induced DNA lesions and chemical-induced bulky DNA adducts, and base excision repair (BER) that removes bases damaged by alkylating agents, oxygen free radicals and hydrolysis (Ford and Hanawalt, 1995; Wani et al., 1999; Offer ef al., 2001; Zhou et al., 2001). While the exact molecular mechanism by which p53 enhances N E R is not fully understood, there is evidence indicating both direct and indirect participation of p53 in N E R . For instance, the C-terminus of p53 directly binds to different forms of damaged DNA, including single-stranded breaks, ends of double-stranded breaks and insertion/deletion mismatches (Steinmeyer and Deppert, 1988; Bakalkin et al., 1995; Lee et al., 1995; Balint and Vousden, 2001). Furthermore, p53 can directly interact with several components of the repair machinery in vitro, including X P B , X P D , p62 subunit of TFIIH, C S B , replication protein A (RPA), and Ref-1 (Wang ef al., 1995; Leveillard et al., 1996; Troelstra et al., 1990; Janus ef al., 1999). possesses exonuclease  other  biochemical activities, such as DNA reannealing  activity,  that  might  also  play  a  role  in  its  repair  p53 also and  3'-5'  functions  (Mummenbrauer et al., 1996; Janus et al., 1999; Albrechtsen et al., 1999; McKay ef al., 1999; Gila etal., 2003). The indirect participation of p53 in DNA repair is mainly achieved through upregulation of p53-responsive genes that are involved in DNA repair. For instance, p53 sustains the expression of the p48 component of the damaged DNA binding  11  protein (DDB2) complexes and XPC gene, both of which are involved in damage recognition (Hwang et al., 1999; Adimoolam et al., 2002). Another p53 downstream target involved in DNA repair, G A D D 4 5 , is shown to bind to P C N A and inhibit DNA replication, thus allowing DNA repair to proceed (Smith et al., 1994). Although the exact mode of action of GADD45 in N E R is not fully understood, the significance of G A D D 4 5 in N E R is supported by the findings that GADD45-null fibroblasts have defects in N E R similar to those seen in p53-null fibroblasts, and that G A D D 4 5 deficient mice show increased radiation carcinogenesis and genomic instability comparable to that seen in p53-deficient mice (Smith et al., 2000; Hollander et al., 1999). More recently, p53 has been found to mediate UV-induced global chromatin relaxation by recruiting p300 to the sites of N E R and inducing histone acetylation, thus acting as a chromatin accessibility factor for N E R (Rubbi and Milner, 2003).  1.2.5  Post-translational Modifications of p53  The choice of p53 whether to undergo cell cycle arrest, apoptosis, senescence or DNA repair upon genotoxic stimuli depends on multiple factors, such as cell type and magnitude of the cellular stress, post-translational modifications and conformation of p53, and the presence of co-factors in p53 transcriptional activation and repression. Accumulating evidence indicates that post-translational modifications of p53 are critical in making such choices, by modulating its protein stability, sequence-specific transcriptional activation and interaction with other proteins (Appella and Anderson, 2001; X u , 2003; Oren, 2003; Brooks and G u , 2003).  p53 phosphorylation and  acetylation are the two major post-translational modifications of p53 that have been  12  extensively studied for the past decade.  More recently, other post-translational  modifications such as sumoylation and ribosylation of p53 have been demonstrated (Wesierska-Gadek ef al., 1996; Gostissa ef al., 1999; Rodriguez ef al., 1999). Generally, the N-terminus of p53 is heavily phosphorylated while the C-terminus contains phosphorylation, acetylation and sumoylation sites. p53 is phosphorylated at multiple sites by a number of kinases upon genotoxic stimuli.  For instance, phosphorylation of p53 at Ser15 by ataxia telangiectasia  mutated (ATM) kinase upon exposure to ionizing radiation is found to disrupt p53 interaction with mdm-2, leading to p53 stabilization (Shieh ef al., 1997). Additional studies suggest that phosphorylation of Ser15 prevents p53 nuclear export (Zhang and Xiong, 2001). Phosphorylation at Ser20 and Thr18 by Checkpoint kinase 1 and 2 (Chk1 and Chk2), downstream targets of A T M kinase and ATM-related (ATR) kinase, and Thr81 by Jun amino-terminal kinase (JNK) are also reported to contribute to p53 stabilization by disrupting p53-mdm-2 interaction (Chehab ef al., 2000; Shieh ef  al.,  2000;  Hirao  ef  al.,  2000;  Buschmann ef  al.,  2001).  UV-induced  phosphorylation at Ser33 and Ser37 is important for p53 activation (Sakahgushi ef al., 1998).  There is also evidence suggesting that phosphorylation at Ser15 or  Ser33/Ser37 promotes an interaction between p53, CBP/p300 and P C A F , and this interaction is crucial for acetylation at the C-terminus (Sakahgushi ef al., 1998; Dumaz and Meek, 1999).  Phosphorylation of Ser46, probably by p38 mitogen-  activated protein kinase (MAPK), seems to be specifically important in UV-induced apoptosis through the activation of p53AIP (Bulavin ef al., 1999; Oda ef al., 2000b).  13  The initial observations that CBP/p300, possessing histone acetyltransferase (HAT) activity, is a co-activator of p53 transcriptional activity led to the discovery of a novel factor acetyltransferase (FAT) function of CBP/p300 on p53 (Gu and Roader, 1997).  CBP/p300 specifically acetylates p53 at multiple lysine residues (Lys370,  Lys371, Lys372, Lys381 and Lys382) at the C-terminal regulatory domain in response to DNA damage (Gu and Roader, 1997; Sakahgushi era/., 1998). P C A F , a p30G7CBP-associated protein, is also found to acetylate p53 at Lys320 (Liu et al., 1999).  Acetylation at these sites of p53 activates sequence-specific D N A binding  activity of p53, recruits CBP/p300 to p53-dependent promoters, and stabilizes p53 by preventing mdm-2 ubiquitination at the regulatory domain for degradation (Gu and Roader, 1997; Balev et al., 2001; Rodriguez et al., 2000; Li er al., 2002). p53 is also modified by conjugation of Lys386 to a small ubiquitin-like protein, S U M O - 1 , in response to DNA damage (Gostissa et al., 1999).  While the knowledge on p53  sumoylation is still limited, it is found that sumoylation of p53 activates p53 transcriptional activities, and that the protein inhibitor of activated S T A T 1 (PIAS1) promotes sumoylation of p53 (Rodriguez er al., 1999; Kahyo er al., 2001).  1.3  The ING T u m o u r S u p p r e s s o r Family  The ING (Inhibitor of growth) family consists of five members, ING1, p33ING2, p47ING3, p29ING4 and p28ING5. ING1, through alternative splicing, encodes three isoforms, namely p47ING1a, p33ING1b and p24ING1c.  p33ING1b is the most  abundant form of ING1, and most tumour suppressive functions have been credited to this isoform.  ING proteins are found to be differentially involved in various  14  biological activities including the regulation of cell cycle arrest, apoptosis, gene transcription, and DNA repair.  Various studies have shown that ING proteins  mediate their tumour suppressive functions in a p53-dependent manner.  ING  proteins cooperate with p53 via at least two mechanisms, the direct association of ING proteins with p53 and ING-mediated p53 acetylation (Campos et al., 2004a). For instance, p33ING1b, p29ING4 and p28ING5 are known to co-precipitate with p53, while p33ING2 and p47ING3 do not.  All ING proteins, except p47ING3, are  found to facilitate p53 acetylation at Lys373/382 in different cell types (Nagashima et al., 2001; Nagashima et al., 2003; Kataoka et al., 2003; Shiseki et al., 2003). Evidence also indicates that the biological functions of ING family genes may at least in part result from interactions of ING proteins with histone acetyltransferase (HAT) and histone deacetylase (HDAC) complexes, and phosphoinositides.  1.3.1  p33ING1b  ING1, the founding member of the ING tumour suppressor family, was isolated by Garkavtsev and colleagues (1996) using subtractive hybridization between c D N A from normal human mammary epithelial cells and seven breast cancer cell lines. It contains three exons, numbered as 1a, 1b and 2, and two introns, producing three alternatively spliced forms, encoding protein variants p47ING1a, p33ING1b, and p24ING1c, all of which possess a highly conserved plant homeodomain (PHD) zinc finger motif, nuclear localization sequence (NLS) and two nucleolar translocation sequences (NTS) (Saito ef al., 2000; Gunduz ef al., 2000; Cheung and Li, 2001). p33ING1b and p24ING1c have previously been reported to mediate growth arrest,  15  apoptosis,  DNA  repair,  senescence,  anchorage-dependent  growth  and  chemosensitivity. p33ING1b, the most extensively studied isoforms, are found to induce cell cycle arrest at Gi-phase, by increasing the activation of the p21  waf1  p53-dependent manner (Garkavtsev et al., 1998). p 2 1  waf1  promoter in a  , a well-known downstream  target of p53, is a cyclin-dependent kinase (Cdk) inhibitor involved in negative regulation of cell growth by inhibiting the kinase activity of Cdk2 and thus the phosphorylation of the retinoblastoma (Rb) protein (Harper et al., 1993; El-Deiry et al., 1993). Therefore, increased expression of p 2 1 cells prevents cell proliferation.  w a f 1  in p33ING1b-overexpressing  p33ING1b involvement in cell cycle arrest is further  supported by the finding that the antisense construct of p33ING1b promotes cell transformation (Garkavtsev et al., 1996). Evidence also indicates the involvement of p33ING1b in G cell cycle arrest. It has more recently been found that cyclin B1 is a 2  down-regulatory target of p33ING1 b, and that overexpression of p33ING1 b enhances adriamycin-induced G arrest in a p53-dependent manner (Takahashi et al., 2002; 2  Tsang er al., 2003). In addition to its involvement in cell cycle regulation, various lines of evidence indicate that p33ING1b is also involved in apoptosis; its dependency on p53 is also well documented.  Increased expression of p33ING1b enhances serum starvation-  induced cell death in p19 mouse teratocarcinoma and NIH 3T3 cells (Helbing et al., 1997).  Co-expression of p33ING1b and c-myc dramatically enhances serum  starvation-induced apoptosis, suggesting that p33ING1b-mediated cell death may be synergistic with the c-myc apoptotic pathway.  16  p33ING1b has also been shown to  sensitize cells to chemotherapeutic agents and radiation, such as etoposide, UV irradiation and y—irradiation, in the presence of wt p53 (Garkavtsev et al., 1998). Adenoviral delivery of both p33ING1b and p53 has been shown to synergistically induce apoptosis in glioma and human oesophageal carcinoma cells (Shinoura etal., 1999; Shimada et al., 2002). Involvement demonstrated.  of  p33ING1b  in  UV-induced  response  has  also  been  The expression of p33ING1b is increased in a dose-dependent  manner in normal human keratinocytes, keratinocyte HaCat cell line, and human melanoma M M R U cell line after UV irradiation (Cheung et al., 2000; Cheung et al., 2001).  Overexpression of p33ING1b in M M R U cells upregulates the expression of  the pro-apoptotic protein Bax, thus enhancing UV-induced apoptosis via activation of the mitochondrial apoptotic pathway, and this process requires the presence of wt p53 (Cheung and Li, 2002).  Another study has demonstrated that p33ING1b  physically binds to P C N A via its PCNA-interacting protein (PIP) domain following UV exposure and enhances apoptosis (Scott et al., 2001a).  UV irradiation induces  p33ING1b translocation to nucleolus and this facilitates the apoptosis of human primary fibroblasts (Scott et al., 2001 b). p33ING1b has also been shown to be involved in DNA repair. Overexpression of p33ING1b enhances N E R of UVC-damaged exogenous plasmid DNA and U V B damaged genomic DNA in a p53-dependent manner (Cheung et al., 2001). recent studies have reported that mutations at codons 102 and 260 of  More  p33ING1b,  which are detected in human cutaneous melanoma biopsies, as well as deletion of the P H D finger motif abolish p33ING1b enhancement in N E R , suggesting the  17  requirement of this motif for proper DNA repair functions (Campos et al., 2004b). Several findings  have clarified the  involvement  machinery. p33ING1b may compete with p 2 1 in p 2 1  w a f 1  waf1  of p33ING1b in DNA repair  for the binding with P C N A , resulting  dissociation with P C N A , and thus favour DNA repair over D N A replication  (Waga et al., 1994; Scott et al., 2001a).  Furthermore, the observations that  p33ING1b can interact with chromatin, and physically interact with p300 and P C N A , potentially link chromatin remodelling to DNA repair (Vieyra et al., 2002). p33ING1b may also be involved in the repair of double strand break generated by a topoisomerase II inhibitor, adriamycin, in a p53-dependent manner (Tsang et al., 2003).  1.3.2  p33ING2  p33ING2,  also know as p33ING1L  (ING1-like), was originally isolated by Shimada  and colleagues (1998), by a homology search with the p33ING1b c D N A sequence from the Otsuka cDNA database containing randomly selected cDNA from human placental, aortal and foetal brain cDNA libraries.  It is found to be located on  chromosome 4q35.1 by R-banding FISH, confirmed with radiation-hybrid mapping (Shimada era/., 1998). xp32ING2,  the first frog homologue of p33ING2, was isolated from X. laevis  tadpole tail and brain by using a gene-specific primer from the conserved region 5' to the region encoding the P H D finger of human ING1 (Wagner et al., 2001).  18  1.3.2.1 G e n e and Protein Structures The full length of human p33ING2 cDNA contains an open reading frame of 840 bp, encoding for a 280-amino acid polypeptide with a molecular weight of 32.8 kDa. The gene products of p33ING2  show 58.9% homology with p33ING1's,  and  the  nucleotide sequences between these two genes shows 60% identity (Figure 1.1). The C-terminus of p33ING2 (amino acid residues 213-260) contains a plant homeodomain (PHD) zinc-finger motif (C4HC3-type), suggesting that p33ING2 may function as a transcription regulator and/or chromatin remodelling factor (Aasland ef al., 1995; Nouman et al., 2003).  The amino acid sequence of this motif is highly  conserved between p33ING1 and p33ING2, suggesting that they may be competing for the same targets (Shimada et al., 1998).  Like p33ING1b, p33ING2 contains a  nuclear localization sequence (NLS) motif, suggesting that p33ING2 is a nuclear protein. It also contains a unique leucine zipper domain, which promotes interaction with other leucine-zipper-containing proteins and hydrophobic interaction (Hai and Hartman, 2001; Feng ef al., 2002) (Figure 1.2).  p33ING2 proteins are mainly  localized in the nucleus (74% in the chromatin-enriched/nuclear matrix and 9% in the nucleoplasm), with a diffuse distribution in the cytosol (17%) in human fibrosarcoma cells (Gozani ef al., 2003). xp32ING2, consists of 834 nucleotides, 6 nucleotides shorter than the human p33ING2, encoding for a 277-aa (32 kDa) protein.  xp32ING2 is 68% identical to  human p33ING2, 67% identical to murine p33ING2, 3 1 % identical to  human  p47ING1a, 54% identical to human p33ING1b and 58% identical to  murine  p33ING1b. The C-terminal P H D finger domain is almost completely conserved  19  Figure 1.1 C o m p a r i s o n of the amino acid s e q u e n c e s of p33ING1b a n d p33ING2. The amino acid sequences of p33ING1b and p33ING2 share 6 0 % homology. The highly conserved plant homeodomain (PHD) zinc finger motif at the C-terminus of these proteins is indicated by shaded bar below this domain. Asterisks indicate the conserved cysteine and histidine residues in this domain.  p33INGlb p33ING2  1 1  p33INGlb p33ING2  12 51  p33INGlb p33ING2  72 101  p33INGlb p33ING2  132 151  p33INGlb p33ING2  192 201  p33INGlb p33ING2  252 251  MLSPAN GEQLHfflVN- YV E DYL DSI ESLP FgL QRNVS MLGQQQQQLYSSAALLT GERSRfflLTC YV DYL ECV ESLP H_M QRNVS  S  |I|A KYQE I LKE LgEC YE RFSF TBGABBRS MLHCVI " L I N KYQE TLKE I_DV YE KYKF D_LN_2K_ LQQLLS a  rafflsisv BBBTSBT,  100  ELVENRWRQ VDS j VEL JEAQQgLGDTVGNSGraVGADR|NGDA ELVENRBRQ MELISQCSQDPAH-SERASDKABMDSSQI  VAQSDKPNR ERR  BNNHNSENAS SNH DH DBGAS GT PKEK HA STS KKK ITS ns SDLCHMANGIEKCDDQP PKEK as SSA KKK  188 190  EREASP^^IDPNEPTYCLCNQVSYGEMIGCDNMCPIEWFHFj EREASP___flIDPNEPTYCLCNQVSYGEMIGCDN_iCPIEWFHF  :  SCVgLteKPKGKWYCPKCRG SCvHLfflKPKGKWYCPKCRG  20  SE HAYNR  JgSTg  11 RDSRSR ST  138 145  238 240  279 280  Figure 1.2 S c h e m a t i c diagrams of the protein structural features of p33ING1b and p33ING2. Both p33ING1b and p33ING2 contain a nuclear localization sequence (NLS) and a plant homeodomain (PHD) zinc finger motif, which is a putative domain of transcription regulators and chromatin-remodelling factors. p33ING1b contains a PCNA-interacting domain (PIP) and a SAP30-interacting region at its N-terminus. p33ING2 contains a unique leucine zipper at its N-terminus.  SAP30-interacting region 45  279  125  NLS  PHD  PIP  280  NLS Leucine Zipper  21  PHD  between xp32ING2 and human p33ING2.  Although it is still unclear that other  members of ING family, except ING1, express different isoforms, it is probable that the xp32ING2  transcript can be alternatively spliced in a tissue-specific manner  (Wagner et al., 2001). Jager and colleagues (1999) also identified  an ING1 -related gene by  screening breast cancer c D N A library using ING1 as a probe, designated ING2 (GenBank accession number: AF149724). The full-length c D N A is 771 bp in size. This gene shows 76% homology to ING1 in its 5' two-thirds of the sequence (nucleotide 1-480).  It contains 203 bp of 5' untranslated region and 439 bp of 3'  untranslated region, and the longest open reading frame of this gene would encode a 42-amino acid polypeptide. However, this variant is distinct from p33ING2, and the functional relevant of this variant between p33ING2 and other ING family members is unknown as there has been no further study carried out to investigate the function of this variant.  1.3.2.2 E x p r e s s i o n Profile p33ING2  m R N A expression levels were found to be ubiquitous in normal human  tissues in the form of two major bands (1.5 kb and 1.3 kb) by Northern blot analysis (Shimada et al., 1998). Most human tissues were found to express p33ING2 except for the spleen, and the expression levels were variable in different tissue types. For example the m R N A expression levels of p33ING2 was high in the testis but low in the lung and kidney, in which p33ING1b  expression was undetectable.  Like human  p33ING2, xp32ING2 was found to be expressed in all adult Xenopus tissues, with  22  variable expression levels in different tissues. xp32ING2  Similarly, the m R N A expression of  was abundant in the testis, but low in muscle and liver (Wagner et al.,  2001). It was also reported that human p33ING2 m R N A expression was significantly higher in colon cancers, where p53 abnormalities occur very frequently compared to the adjacent normal tissues. Importantly, its expression levels increased with tumour progression as poorly differentiated carcinoma expressed higher p33ING2 levels than differentiated adenocarcinoma (Shimada et al., 1998).  Unlike p33ING1b, protein  levels of p33ING2 in cancer cell lines were found to be highly variable, with undetectable expression in some cell lines. It is reported that the expression profile of p33ING2 did not correlate with the mutational status of p53. However, three out of twelve cell lines examined, expressing the most abundant p33ING2, contained either null or mutant p53 (Nagashima et al., 2001). Recent studies on loss of heterozygosity in sporadic basal cell carcinomas (BCCs) reported a high frequency of LOH on chromosome 4q32-35, which is mapped to p33ING2  and SAP30 (Sin3-associated protein), both of which are believed to be  involved in chromatin remodelling and gene regulation (Sironi et al., 2004). p33ING2 upregulates p 2 1  w a f 1  Since  and Bax, L O H of p33ING2 could impair the proper  regulation of cell cycle and apoptosis, resulting in cellular transformation and tumour progression.  In fact, this is supported by previous findings that reported loss of  expression of p21  w a f 1  in B C C s (Ahmed et al., 1997).  These findings suggest a  possible involvement of p33ING2 in the formation of basal cell carcinoma.  23  1.3.2.3 B i o l o g i c a l Functions So far, there are three studies reporting the biological functions of p33ING2 as a tumour suppressor. p33ING2 has been found to regulate cell cycle, apoptosis and gene transcription, and the presence of functional p53 is required in these processes (Figure 1.3) (Nagashima et al., 2001; Gozani et al., 2003; Kataoka et al., 2003). The first evidence indicating p33ING2 as a tumour suppressor is that p33ING2 negatively regulates cell growth of colorectal carcinoma R K O cells. Nagashima and colleagues (2001) demonstrated that overexpression of p33ING2 completely inhibits colony formation of R K O cells that carry wild-type p53, through the induction G r phase cell cycle arrest and apoptosis.  However, this is not observed in p53-  deactivated R K O - E 6 cells (Nagashima et al., 2001).  The role of p33ING2 as a  tumour suppressor is further supported by the findings that the expression of p33ING2  is  specifically  neocarzinostatin,  but  not  induced  by  by gamma  DNA  damage  irradiation,  agents,  doxorubicin,  etoposide  or  c/'s-platinum  or  bleomycin in the normal lymphoblastoid C 3 A B R cell line (Nagashima et al., 2001). This induction of p33ING2 is independent of A T M or p53 function (Nagashima et al., 2001). Taken together, these observations suggest that p33ING2 negative regulation of cell growth is p53-dependent, indicating that p33ING2 is a potential regulator in p53-mediated cellular processes (Nagashima er al., 2001). In addition to its involvement in growth control, p33ING2 has been shown to play a role in the regulation of gene transcription.  Like p33ING1b, p33ING2 was  found to enhance the promoter activities of p 2 1  and Bax in wild-type p53 R K O  w a f 1  cells, but not in R K O - E 6 cells (Nagashima et al., 2001; Kataoka et al., 2003).  24  Moreover, p33ING2 was significantly more effective than p33ING1b in enhancing these p53 transcriptional-transactivation activities (Nagashima et al., 2001). contrast to upregulation of the p 2 1  waf1  In  promoter, p33ING2 strongly repressed the  promoter activity of human a-fetoprotein (AFP), a tumour-specific marker of the hepatocellular carcinoma, possibly through binding to A F P ' s AT-motif, in p53 wildtype hepatocellular carcinoma HepG2 cells (Kataoka et al., 2003). This repressive effect of p33ING2 on the A F P promoter was much weaker in the Hep3B p53-null cell line. When co-transfected with wild-type p53, the repressive effect was restored but to a lesser extent than in the HepG2 cells, suggesting plausible p53-dependent and p53-indendpent regulatory mechanisms. It was also shown that p33ING2 inhibited the A F P promoter by antagonizing deacetylation effects of hSir2 on p53, but p33ING2 was not detected in the co-immunoprecipitate with hSir2 (Kataoka et al., 2003). The role of ING2 in hormone-dependent transcriptional response during developmental events was studied using Xenopus  tadpole metamorphosis as a  model system, in which a single stimulus thyroid hormone (TH) initiates tissuespecific proliferation, apoptosis and remodelling simultaneously. Thyroid receptors (TRs) selectively repressed or activated gene transcription in response to absence or presence of T H . It is shown that xp32ING2 transcript levels increased in response to TH-induced metamorphosis and during spontaneous metamorphosis in X. tadpole tissues (Wagner et al., 2001).  laevis  It is interesting to note that in the tail and  brain, which apoptose extensively during metamorphosis, xp32ING2  induction  followed the T R s expression pattern; whereas in the leg, which proliferate and grow,  25  xp32ING2 expression was not induced, similar to that of T R s . This indicates the role of xp32ING2 in hormonally regulated nuclear transcription factor-mediated apoptotic response.  1.3.2.4 M o d e s of A c t i o n Although p33ING2 is involved in various biological functions, it possesses no known enzymatic activity. facilitating  It is therefore reasonable to speculate that p33ING2 acts by  specific  phospholipid  protein-protein,  interactions.  In  possible  fact,  protein-DNA,  p33ING2  is  known  and  even  to  protein-  interact  with  phosphoinositides and Sin3-histone deacetylase (HDAC) complexes (Gozani et al., 2003; Kuzmichev et al., 2002). The p53 dependency of p33ING2 in exerting its tumour suppressive functions has led to investigation of the interaction between p33ING2 and p53.  Interestingly,  p33ING2 was not detected in a physical interaction with p53, unlike three other ING family members (p33ING1b, p29ING4 and p28ING5) (Nagashima et al., 2001; Shiseki et al., 2003).  However, overexpression of p33ING2 led to an increased  acetylation of p53 at Lys382 in colorectal carcinoma R K O cell line (Nagashima et al., 2001; Nagashima et al., 2003). It was reported that p53 acetylation at Lys382 in the normal lymphoblastoid C 3 A B R cell line after exposure to DNA-damaging agents, etoposide or neocarzinostatin, correlated with an increased p33ING2 expression levels (Nagashima et al., 2001). Furthermore, It was shown that antisense-p33ING2 expression in osteosarcoma O s A C L cells reduced the level of endogenous p33ING2 and acetylated p53 proportionally following exposure to etoposide (Nagashima er al.,  26  2001).  However, p33ING2 expression did not alter the phosphorylation of p53 at  either Ser15 or Ser392.  Post-translational modifications such as acetylation and  phosphorylation within the C-terminal region of p53 have been shown to facilitate p53 activation and stimulate p53 sequence-specific DNA-binding activity (Gu and Roeder, 1997; Sakagushi et al., 1998).  These observations lead to the speculation that  p33ING2 involvement in the regulation of cell cycle, apoptosis and transcription may be the results of p33ING2-mediated p53 acetylation that stabilizes p53 and enhances p53 cellular responses to genotoxic stresses and apoptotic stimuli. Gozani and colleagues (2003) were the first and the only group to identify p33ING2 as a phosphoinositides-binding module through a library expression screening using phosphoinositides (PtdlnsP)-affinity resins.  It was found that  p33ING2 interacted with Ptdlns(3)P and Ptdlns(5)P through its P H D finger motif, which alone was sufficient for this binding. The importance of the P H D finger motif in this interaction was further evidenced by the findings that the zinc chelator T P E N [N,N,N',N -tetrakis(2-pyridyl-methyl)ethylenediamine] and the mutations of the basic ,  residues at the C-terminal of the P H D finger of p33ING2 greatly reduced the binding affinity of p33ING2 with Ptdlns(3)P and Ptdlns(5)P. In fact, it was shown that the P H D finger motif was a general PtdlnsPs binding domain, and that the general structure for PtdlnsPs binding was conserved. Therefore, we speculated that other ING family members might be phosphoinositide-interacting partners as well, with different specificity and binding affinity. Ptdlns(5)Ps can modulate the subcellular localization of p33ING2. This is evidenced by the finding that generation of Ptdlns(5)Ps at the plasma membrane by  27  Figure 1.3 B i o l o g i c a l functions of p33ING1b and p33ING2. p33ING1b physically associates with p53 while p33ING2 induces p53 acetylation. Both p33ING1b and p33ING2 induce Gi-phase cell cycle arrest, promote apoptosis and interact with Sin3, a component of transcription co-repressor complexes, to regulate gene transcription. p33ING1b downregulates cyclin B1 to induce G -phase cell cycle arrest and enhances DNA repair through interaction with G A D D 4 5 and P C N A . 2  Transcription Regulation  28  lipid  phosphatase  IpgD, which  dephosphorylates  Ptdlns(4,5)P2  to  Ptdlns(5)P,  recruited p33ING2 P H D finger to plasma membrane and caused loss of diffuse p33ING2 cytoplasmic distribution (Gozani et al., 2003).  Furthermore, reducing  nuclear Ptdlns(5)P levels by overexpressing PIKlip, which phosphorylates Ptdlns(5)P to form Ptdlns(4,5)P2, reduced p33ING2-association with the  chromatin/nuclear  matrix and increased p33ING2 levels in the cytosol, without changing the total p33ING2 protein levels (Gozani et al., 2003). In addition, mutations in the P H D finger motif that abrogated Ptdlns(5)P binding disrupted the ability of p33ING2 to enhance p53 acetylation and p53-dependent apoptosis. Overexpression of a P H D zinc finger demonstrated dominant-negative effects on p33ING2 functions.  Taken together,  these observations suggest that Ptdlns(5)P recruits or stabilizes p33ING2 in the chromatin, and/or interaction with Ptdlns(5)P might lead to allosteric activation of p33ING2. The interaction between p33ING2 and Sin3-HDAC complexes was reported by Kuzmichev and colleagues (2002), who found that immunoaffinity  purification of  HeLA cell extract using anti-SAP30 antibodies yielded a set of abundant polypeptides corresponding to the core H D A C complexes, composed of H D A C 1 ,  HDAC2,  RbAp46, RbAp48, Sin3 and S A P 3 0 , and less abundant polypeptides that may associate indirectly with the core Sin3-HDAC complex. Mass spectrometry analysis confirmed the presence of p33ING1b and p33ING2 in this immunopurified cell extract. A unique N-terminal domain of p33ING1b, SAID (SAP30-interacting domain) was identified as a domain required for interaction with S A P 3 0 , a component of the core Sin3-HDAC complex (Kuzmichev et al., 2002).  29  A computational search  revealed that p33ING2 also possesses SAID domain, suggesting that the presence of p33ING2 in Sin3-HDAC complexes may be achieved by direct interaction with SAP30.  Since Sin3-HDAC complexes are involved in transcription  repression,  interaction between p33ING2 and Sin3-HDAC complexes may be important for the growth inhibitory activity of p33ING2.  1.4  Objectives and Hypotheses  The primary objective of this study is to further understand the tumour suppressive role of the p33ING2 novel tumour suppressor in cellular stress response to UV irradiation, and the molecular mechanisms of p33ING2 involvement in this stress response.  W e hypothesized that p33ING2 enhances UVB-induced apoptosis and  the repair of UV-damaged DNA in a p53-dependent manner in melanoma cells. Using melanoma cell lines as an experimental model system, we have investigated the role of p33ING2 in UV-induced apoptosis (Chapter 3) and the repair of UVdamaged DNA (Chapter 4), and the molecular mechanisms involved in these stress responses.  30  CHAPTER 2. MATERIALS AND METHODS  2.1  2.1.1  Materials  Cell lines and Cell Culture  Two melanoma cell lines with known p53 status were used. The M M R U cell line was a kind gift from Dr. H.R. Byers (Boston University, School of Medicine, Boston, MA) (Byers et al., 1991); the MeWo cell line was a kind gift from Dr. A . P . Albino (Memorial Sloan-Kettering Cancer Centre, New York, NY). The p53 status in the melanoma cell lines has previously been determined by DNA sequencing: M M R U contains wild-type (wt) p53 (Li et al., 1995), while MeWo contains mutant (mut) p53 (Tilgen et al., 1983; Li et al., 1995; Bae et al., 1996). Eagle  All melanoma cell lines were maintained in  Dulbecco's  Modified  Medium  (DMEM) supplemented  (Invitrogen,  Mississauga, O N , Canada), 100  U/ml  with  penicillin, and  10% F B S 100  uxg/ml  streptomycin (Invitrogen) at 37°C in a humidified atmosphere of 5% C 0 . 2  2.1.2  Plasmids  Plasmids used for transfection  included pcDNA3-vector (Invitrogen), pcDNA3-  p33/A/G2, pcDNA3-antisense (AS)-p33//VG2, pCI-p33/A/G7ib (kind gifts from Dr. C . C . Harris, National Cancer Institute, National Institutes of Health, Bethesda, MD), p3XFLAG-vector (Sigma-Aldrich, Mississauga, ON), p3XFLAG-ING2 (a kind gift from Dr. O. Gozani and Dr. J . Yuan, Harvard Medical School, Boston, MA), pphwtp53 which contains human wild-type p53, and pED-1 which contains a point mutation in the human p53 cDNA that changes Cys135 to Serine (Johnson et al., 1991) (kind  31  gifts from Dr. S. Benchimol, University of Toronto, Toronto, O N , Canada),  PCMVCAT  (a kind gift of Dr. L. Grossman, Johns Hopkins University, Baltimore, MD), and p G F P N2 (Clontech, Windsor, O N , Canada).  2.1.3  Antibodies  Antibodies used for Western blotting were anti-p33ING2 rat monoclonal antibody (a kind gift from Dr. O. Gozani and Dr. J . Yuan, Harvard Medical School, Boston, MA), anti-CPD and anti-6-4PP primary antibodies (kind gifts from Dr. T. Matsunaga from Kanazawa University, Japan) (Mori et al., 1991), anti-/?-actin goat  monoclonal  antibody, anti-p53 (DO-1) mouse monoclonal, anti-Bax (N-20) rabbit polyclonal, antiGADD45  mouse monoclonal, anti-XPA rabbit  polyclonal, and anti-XPB  rabbit  polyclonal antibodies (Santa Cruz Biotechnology, Santa Cruz, CA), anti-Bcl-2, anticaspase 3, anti-caspase 8, anti-caspase 9, anti-acetylated polyclonal antibodies  Lys382 p53  (Cell Signalling, Beverly, MA), anti-cytochrome  rabbit  c mouse  monoclonal antibody (BD Pharmingen, Mississauga, O N , Canada), and anti-Fas (Ab1) mouse monoclonal antibody (Calbiochem, San Diego, C A ) , secondary IgG (Santa Cruz), Cy2-conjugated goat anti-rabbit  and Cy3-conjugated donkey  anti-mouse  secondary antibodies (Jackson ImmunoResearch Laboratory, West Grove, PA).  32  2.2  2.2.1  Methods  Transfection  Cells were transfected at 40-50% confluency with Effectene reagent (Qiagen, Mississauga, O N , Canada) at a ratio of 1 u.g DNA to 25 ul Effectene. An empty vector was used as control DNA.  2.2.2  Determination of Transfection Efficiency  Transfection efficiency of a particular cell line was determined by transfecting the cells with GFP-containing plasmid, p G F P - N 2 with Effectene. Twenty-four hours after transfection, the number of green fluorescence-emitting cells was assessed under a fluorescent microscope.  Transfection efficiency was calculated by the equation  number of green fluorescence-emitting cells / the total number of cells (fluorescent and non-fluorescent) X 100%.  Table 1 shows the percentages of transfection  efficiencies of the cell lines used in this study.  33  Table 1.1 Relative percentages of transfection efficiencies of cell lines used in the study. Percentages were determined by counting the number of cells emitting green fluorescence / total number of cell counted X 100%. Experiments were repeated three times.  Cell Line  Transfection Efficiency (%)  MMRU  60-70  MeWo  50 - 60  34  2.2.3  Ultraviolet Irradiation  Medium was removed and the cells (at 70% confluency) were exposed to U V B (280320 nm) using a bank of four unfiltered FS40 sunlamps (Westinghouse, Bloomfield, NJ). The petri-dish cover was left on to filter possible U V C emissions from the U V B bulb. Medium was replaced and cells were incubated in a 5% C 0 incubator at 37°C 2  for desired time periods after UVB irradiation.  The intensity of the UV light was  measured by the IL 700 radiometer fitted with a W N 320 filter and an A127 quartz diffuser (International Light, Newburyport, MA).  2.2.4  Light Microscopy  The morphology of M M R U cells transfected with empty vector, pcDNA3-p33/A/G2 or pcDNA3-AS-p33/A/G2 were subjected to microscopic assessment 24 h after U V B irradiation at 600 J / m , using an inverted microscope (Zeiss, Chester, VA). Images 2  were taken using a cooled mono 12-bit Retiga-Ex camera (Q-imaging, Burnaby, B C , Canada).  2.2.5  SRB Cell Survival Assay  Cells were grown in 24-well plates and transfected with empty vector, pcDNA3p33/A/G2 or pcDNA3-AS-p33/A/G2 at 50% confluency.  They were irradiated with  UVB at 600 J / m 24 h after transfection. Twenty-four hours after UV irradiation, cell 2  survival was determined with the sulforhodamine B (SRB) (Sigma) assay as described previously (Skehan et al., 1990; Li et al., 1998).  S R B is a pink  aminoxanthene dye with two sulfonic groups that bind to basic amino acid residues in  35  proteins in cells fixed with trichloroacetate (TCA). S R B provides a sensitive index of cellular protein content.  Briefly, the medium was removed and the cells were fixed  with 500 uJ of 10% T C A for 1 h at 4°C after treatment. The cells were then washed five times with tap water and the excess water was removed by flicking. The cells were air-dried and then stained with 500 uJ of 0.4% S R B (dissolved in 1% acetic acid) for 30 min at room temperature, washed four times with 1 % acetic acid, and air-dried. The cells were then incubated with 500 uJ of 10 mM Tris (pH 10.5) on a shaker for 20 min to solubilize the bound dye. Spectrophotometric readings were then taken at 550 nm for 100 uJ aliquots.  2.2.6  Trypan Blue Exclusion Assay  M M R U cells were grown in 24-well plates to 50% confluency before transfection. Twenty-four hours after transfection, the cells were irradiated with U V B at 600 J / m . 2  After incubation in fresh medium for 24 h, attached cells were trypsinized, pooled with the floating dead cells, and the percentage of the dead cells were determined using a hematocytometer in the presence of 0.4% of trypan blue reagent (Sigma).  2.2.7  Propidium Iodide (PI) Staining  Cells grown on coverslips in 6-well plates were transfected and then irradiated with UVB at 600 J / m 24 h after transfection. 2  Twenty-four hours later, cells were fixed by  gently overlying the media with 6 ml of ice-cold 95% ethanol and incubated for 3 min at room temperature.  The media and 95% ethanol were mixed by pipetting gently  and then incubated for 3 min at room temperature before the mixture was removed.  36  The fixed cells were stained with the mixture of 5 mg/ml PI, 0.1% Triton X-100 and 500 jag/ml RNase A at room temperature for 2 min. Coverslips were washed twice with phosphate buffered saline (PBS) before mounted onto slides with mounting media (Fisher Scientific, Nepean, Ontario, Canada).  The slides were visualized  under a fluorescent microscope (Zeiss) for apoptotic bodies.  Images were taken  with a cooled mono 12-bit Retiga-Ex camera.  2.2.8  Flow Cytometry  Cells were grown in 6-well plates for 24 h before transfection at 50% confluency. They were irradiated with UVB at 600 J / m 24 h after transfection. Twenty-four hours 2  later, cells were collected by trypsinization and pelleted by centrifugation at 2,000 g for 2 min.  Next, cell pellets were resuspended in 1 ml of hypotonic fluorochrome  buffer (0.1% Triton X-100, 0.1% sodium citrate, 25 ng/ml RNase A , 50 ug/ml PI). After overnight incubation at 4°C, the samples were then analyzed by flow cytometry to determine the percentage of sub-diploid DNA. Samples were run on a Coulter E P I C S X L - M C L flow cytometer (Beckman Coulter, Mississauga, ON) and analyzed with E X P 0 3 2 A D C analysis software.  2.2.9  Western Blot Analysis  Cells were grown in 100-mm plates, transfected with p3XFLAG-vector or p 3 X F L A G ING2 expression vector at 50% confluency and irradiated with U V B at 600 J / m 24 h 2  after transfection. Twenty-four hours later, cells were washed with P B S , harvested by scraping on ice and pelleted by centrifugation at 2,000 g for 2 min. Cell pellets  37  were lysed in 80 u.1 of triple detergent buffer (50 mM Tris-CI [pH 8.0], 150 mM NaCI, 0.02% N a N , 0.1% S D S , 1% NP-40, 0.5% sodium deoxycholate) containing freshly 3  added protease inhibitors (100 u,g/ml phenylmethylsulfonyl fluoride, 1 ug/ml aprotinin, 1 u.g/ml leupeptin, 1 fig/ml pepstatin A). The samples were then sonicated, incubated on ice for 20 min, and centrifuged at 12,000 g for 10 min at 4°C. The supernatants were collected and a Bradford assay was performed to determine the protein concentration.  Proteins (50 u.g/lane) were separated on 12% SDS-polyacrylamide  gels and blotted onto polyvinylidene difluoride Mississauga, O N , Canada).  (PVDF) membranes (Bio-Rad,  The P V D F was blocked with 5% skim milk for 1 h at  room temperature before incubating with primary antisera prepared in 5% bovine serum albumin for 1 h at room temperature. Blots were washed three times in P B S T (PBS containing 0.04% Tween-20) for 5 min each and then incubated with horseradish peroxidase (HRP)-conjugated secondary antisera for 1 h at room temperature.  The signals were detected with the enhanced chemiluminescence  detection kit (Amersham Bioscience, Baie d'Urfe, Quebec, Canada).  Protein  expressions on Western blots were quantified by densitometry using the Quantity One software  (Bio-Rad).  The fold-induction  or reduction was corrected for  differences in the actin loading control.  2.2.10 Detection of the Mitochondrial Membrane Potential Disruption  of  the  mitochondrial  membrane  potential  MitoCapture™ Apoptosis Detection Kit (Calbiochem). according to the manufacturer's specifications.  38  was  detected  using  a  The assay was performed  Briefly, cells were grown on  coverslips in 6-well plates, transfected with p3XFLAG-vector or p3XFLAG-/A/G2 expression vector at 50% confluency and exposed to UVB at 600 J / m 24 h after 2  transfection. Twenty-four hours after UV irradiation, the medium was removed and the cells were incubated with 2 ml of diluted MitoCapture™ solution at 37°C in a 5% C O 2 incubator for 15 min. The dye solution was then removed and the cells were washed twice with 1 ml of the pre-warmed incubation buffer.  The cells were then  examined immediately under a fluorescent microscope. Images were taken with a cooled mono 12-bit Retiga-Ex camera.  2.2.11  Subcellular Fractionation  Cells were grown in 100-mm plates, transfected with p3XFLAG-vector or p 3 X F L A G ING2 expression vector at 50% confluency and irradiated with U V B at 600 J / m 24 h 2  after transfection. Twenty-four hours later, cells were washed with P B S , harvested by scraping on ice, and pelleted by centrifugation at 2,000 g for 2 min. Subcellular fractionation was performed as described previously (Kim et al., 2001), with minor modifications.  Briefly, cell pellets were resuspended in 200 uJ of buffer A (20 mM  H E P E S - K O H [pH 7.5], 10 mM KCI, 1.5 mM MgCI , 1 mM sodium EDTA, 1 mM 2  sodium E G T A , 1 mM dithiothreitol, 250 mM sucrose) containing a freshly added mixture of protease inhibitors (100 u.g/ml phenylmethylsulfonyl fluoride, 1 u.g/ml aprotinin, 1 ug/ml leupeptin, 1 u.g/ml pepstatin A).  Cell suspensions were then  passed through a 26-gauge needle eight times to lyse the cells.  Under a light  microscope, lysed cells appear dull while intact cells have shiny rings around the nuclei. Unbroken cells, large plasma membrane pieces, and nuclei were removed by  39  centrifugation at 1,000 g for 10 min at 4°C.  The resulting supernatant was  centrifuged at 10,000 g for 20 min at 4°C to obtain crude cytosolic fraction (supernatant) and the mitochondria (pellet). The pellet was washed with buffer A and then solubilized in 50 ul of T N C buffer (10 mM Tris-acetate [pH 8.0], 0.5% NP-40, 5 mM CaCI ) containing freshly added protease inhibitors to obtain mitochondrial 2  fraction.  The cytosolic fraction was then generated by centrifugation  of the  supernatant at 100,000 g for 1 h at 4°C. Protein concentrations were determined by the Bradford assay.  Samples were then separated on 15% SDS-polyacrylamide  gels, blotted onto P V D F , and probed with anti-Bax (N-20), anti-cytochrome c and anti-^-actin antibodies.  2.2.12 Host-cell-reactivation Assay (CAT Assay) The pCMVc/\r plasmid that contains the gene encoding for chloramphenicol acetyltransferase (CAT) at 50 u.g/ml in 10 mM Tris-CI (pH 8.5) was irradiated with U V C at 400 J / m using an electronic ultraviolet cross-linker (Ultralum, Claremont, CA) 2  and used for transfection.  M M R U cells were grown in 6-well plates, and co-  transfected with p C M V c ^ r and pcDNA3-vector, pcDNA3-p33/A/G2 or pcDNA3-ASp33/A/G2 at 40% confluency.  Forty hours after transfection, C A T assays were  performed as previously described (Cheung et al., 2001).  Briefly, cells were  harvested and the cell pellet was resuspended in a 50 u,l solution of 0.25 M Tris-CI (pH8.0) and 5 mM EDTA. Cell-free extracts of the transfected cells were obtained by three repeated freeze-thawings (liquid-nitrogen to freeze and 37°C to thaw), heated to 65°C for 10 min, centrifuged at 12,000 g for 10 min, and the cleared supernatant  40  was used for the C A T assay. The assay reaction mixture contained 7.5 uJ of 5 mM of chloramphenicol, 50 uJ of cell-free extract, 1 uJ of 2.5 mM [ H]acetyl-CoA, and 16.5 uJ 3  of double-distilled water (ddH 0). The reaction mixture was incubated with the cell2  free extract at 37°C for 90 min. Following incubation, 200 uJ of ice-cold ethyl acetate was added, tubes were vortexed, and centrifuged at 12,000 g for 4 min. After quickfreezing the aqueous phase in liquid-nitrogen, the organic phase was removed and extracted with 200 uJ of d d H 0 . 2  radioactivity  was  determined  The organic phase was dried to completion and in  a  scintillation  Determinants were performed in triplicates.  counter  (Beckman  Coulter).  Controls included transfection with  undamaged plasmid DNA and mock transfection without plasmid DNA.  2.2.13 Immune-fluorescent Staining M M R U cells were grown on coverslips in 6-well plates and transfected with p3XFLAG-/A/G2 at 50% confluency. After 24 h, the medium was removed and an isopore polycarbonate filter with 5 urn pores (Millipore, Nepean, Ontario, Canada) was placed on top of the cell monolayer on the coverslip. Cells were irradiated with U V C at 200 J / m using an UV cross-linker (Ultralum). The filter was gently removed 2  and cells were cultured for desired time periods before fixation. Cells were fixed and labelled as described previously (Wang et al., 2003) with slight modifications. Briefly, cells were washed twice with phosphate buffered saline (PBS) for 5 min each, fixed with 2 ml of fixation solution (2% paraformaldehyde, 0.5% Triton X-100 in P B S ) for 30 min at 4°C. Cells were then washed three times with P B S for 5 min each, incubated with 2 M HCI for 10 min to denature DNA, followed by three washes with P B S for 5  41  min each. Cells were blocked with goat normal serum for 1 h at room temperature, followed by three washes with P B S . Coverslips containing fixed cells were then inverted onto 80 uJ of 1:30 dilution of anti-CPD or anti-6-4PP primary antibody and incubated in a humid chamber for 1 h, followed by three washes with P B S for 5 min each. Coverslips were then incubated with 100 uJ of 1:150 dilution of Cy3-conjugated donkey anti-mouse secondary antibody (Jackson ImmunoResearch) for 1 h and washed three times with P B S for 5 min each. Cells were then double-stained with 1:30 dilution of anti-FLAG rabbit polyclonal antibody (Sigma), followed by staining with 1:150 dilution of Cy2-conjugated goat anti-rabbit secondary antibody (Jackson ImmunoResearch).  Finally, coverslips were incubated with 1:3000 dilution of stock  Hoechst 33258 (20 mM) for 10 min, washed three times with P B S for 5 min each and mounted onto slides with mounting medium (Fisher Scientific).  Slides were then  visualized under a fluorescent microscope. Images were taken with a cooled mono 12-bit Retiga-Ex camera.  2.2.14 Detection of p53 Acetylation Cells were grown in 100-mm plates, transfected with p3XFLAG-vector or p 3 X F L A G ING2 expression vector at 50% confluency and irradiated with U V B at 200 J / m 24 h 2  after transfection.  The histone deacetylase (HDAC) inhibitor trichostatin A (TSA)  (Sigma) was added at a final concentration of 5 uJVI 3 h prior to harvest to aid in the detection of p53 acetylation (Nagashima et al., 2001; Saito et al., 2002). Cells were then harvested at the indicated time-points after UV irradiation, and lysed for Western  42  blot analysis. M M R U cells treated with 0.2 u.g/ml of doxorubicin (Sigma) for 8 h were used as a positive control for p53 acetylation.  2.2.15 Immnoprecipitation M M R U cells were grown in 100-mm plates for 24 h and transfected with p 3 X F L A G ING2 plasmid at 50% confluency. Twenty-four hours after transfection, cells were lysed with 1% triton-X solubilization buffer containing freshly added protease inhibitors (100 ug/ml phenylmethylsulfonyl fluoride, 1 ij.g/ml aprotinin, 1 ug/ml leupeptin, 1 jug/ml pepstatin A). The samples were then sonicated, incubated on ice for 20 min, and centrifuged at 12,000 g for 10 min at 4°C. The supernatants were collected  and  concentration.  a  Bradford  assay  was  performed  to  determine  the  protein  Protein (500-1000 ug) was incubated with anti-FLAG  mouse  monoclonal antibody (Sigma) or a control normal mouse IgG (Santa Cruz) at 4°C overnight, and then with protein G sepharose (Amersham Bioscience) at 4°C for 1 h. The beads were washed three times with solubilization buffer prior to boiling for 3 min.  The precipitates were then resolved by electrophoresis, followed by Western  blot analysis.  43  CHAPTER 3. p33ING2 ENHANCES UVB-INDUCED APOPTOSIS IN HUMAN MELANOMA CELLS  3.1  Rationale and Hypothesis  Cutaneous melanoma is a type of skin cancer resulting from uncontrolled proliferation of the epidermal melanocytes.  Epidemiological studies strongly implicate UV  irradiation as the primary aetiological factor for melanoma (Mackie 1998; Gilchrest et al., 1999). Studies on the carcinogenic effects of different UV wavelengths identified UVB as the major cause of the carcinogenic effects of sun exposure (Yuspa et al., 1991). UV irradiation induces the formation of cyclobutane pyrimidine dimers and 6-4 photoproducts which may lead to mutations and carcinogenesis if the lesions are not removed from the genome promptly (Beukers and Berends, 1960; Setlow and Carrier, 1966; Alcalay et al., 1990). The incidence of melanoma is increasing most rapidly in the Caucasian population among all malignancies except lung cancer in women (Howe et al., 2001; Bronchez and Naeyaert, 2000). Malignant melanoma is associated with one of the highest mortality rates due to its rapid metastasis, and the 5-year survival rate of patients with metastatic melanoma remains less than 10% (Howe et al., 2001; Roses et al., 1991).  The reason for the poor prognosis of  melanoma is due to its resistance to conventional radiotherapy and chemotherapy. It is well accepted that apoptosis is a common mode of action employed by various anticancer drugs, and that the expression of apoptotic genes mediates chemosensitivity in cancer cells.  The tumour suppressor p53 is an important  transcription factor of various apoptotic genes such as Bax, PUMA, and Noxa (Henry  44  et al., 2002). It is the most frequently mutated gene known to date, with the mutation rate of more than 50% in human malignancies (Hussein et al., 2003a).  However,  mutational studies revealed that the p53 gene is altered in only 1-5% and 11-25% of primary and metastatic melanoma, respectively (Hussein et al., 2003a).  In fact,  protein expression profile of p53 evidenced that p53 is overexpressed in the late stage of melanoma tumourigenesis (Hussein et al., 2003a). The lower frequency of p53 mutation in melanoma suggests that other tumour suppressor genes and/or apoptotic genes may also play key roles in the pathogenesis of melanoma. Several studies have provided evidence that both p33ING1b and p33ING2 require the activity of p53 in exiting tumour suppressive functions such as regulation of transcription, growth arrest and apoptosis (Campos et al., 2004a). p33ING1b and p33ING2 modulate p53 activity via distinct mechanisms.  p33ING1b physically  associates with p53 while p33ING2 promotes p53 acetylation, resulting in the stabilization of p53 and thus enhancement of the p53 cellular responses to genotoxic stresses and apoptotic stimuli.  Furthermore, recent studies indicated that the  interaction of p33ING2 with phosphoinositides, Ptdlns(3)P and Ptdlns(5)P, through its P H D zinc finger motif is critical for its ability to enhance p53 acetylation and mediate p53-dependent etoposide-induced apoptosis (Gozani et al., 2003). To date, there are a few reports indicating that p33ING1b plays a significant role in stress response to UV irradiation.  Previously, we found that p33ING1b  cooperates with p53 to enhance UVB-induced apoptosis via activation of the mitochondrial apoptotic pathway in melanoma cells (Cheung and Li, 2002). Another group has demonstrated that p33ING1b possesses two distinct nucleolar targeting  (  45  sequences (NTS) within the nuclear localization domain, which promotes  the  translocation of p33ING1b to the nucleolus after UV irradiation (Scott ef al., 2001b). It was also reported that the PCNA-interacting-protein (PIP) domain at the N-terminus of p33ING1b binds competitively to the interdomain connector loop of P C N A upon UV irradiation (Scott ef al., 2001a). However, study on the role of p33ING2 in the stress response to UV irradiation is lacking. Due to the structural similarities between p33ING1b and p33ING2, we investigated the role of p33ING2 in cellular stress response to U V B irradiation and the molecular mechanisms of p33ING2 involvement in this stress response by using melanoma cells as an experimental model system. We  hypothesized that p33ING2 enhances UVB-induced apoptosis in a p53-  dependent manner.  46  3.2  Results  3.2.1  p33ING2 Enhances UVB-induced Apoptosis in Melanoma Cells  To investigate the role of p33ING2 in UVB-induced apoptosis, we transfected a melanoma cell line, M M R U , with either  pcDNA3-vector, pcDNA3-p33//VG2 or  pcDNA3-AS-p33/A/G2 expression vector.  Western blot analysis confirmed the  expression of p33ING2, suggesting successful transfection of M M R U cells (Fig. 3.1 A).  To determine if p33ING2 enhances UV-induced cell death, the transfected  cells were irradiated with UVB at various doses (0, 200, 400, and 600 J/m ). Cells 2  were fixed and stained with S R B 24 h after UV irradiation. The amount of S R B dye that remains bound to the surface of the fixed cells is a determinant of cell survival. W e determined the cell survival rate of UV-irradiated p33ING2-overexpressing M M R U cells in comparison to cells transfected with empty vector alone or with antisense p33/A/G2. W e found that overexpression of p33ING2 consistently reduced cell survival at various doses of UVB (Fig. 3.1 B). After U V B irradiation at 200, 400 and 600 J / m , the survival rates of M M R U cells overexpressing p33ING2 were 2  79.4%, 64.9% and 53.2% compared to the vector control with 94.4%, 87.9% and 78.0%, respectively (P<0.001, P<0.0001, and P<0.00001, t-test). The microscopic images show that there were significantly more dead cells (round, shrunk, and detached) in M M R U cells overexpressing p33ING2 24 h after U V B irradiation compared to the control cells (Fig. 3.1C). To further confirm these observations, trypan blue exclusion assay was carried out to quantitate dead cells, which were stained blue due to their inability to exclude the dye by the cell membrane, while live  47  cells were unstained. The results in Fig. 3.1 D show that M M R U cells overexpressing p33ING2 had a significantly higher rate of cell death (39.8%) after U V B irradiation at 600 J / m compared to 16.6% in the vector control cells (P<0.0001, t-test). W e also 2  observed that overexpression of p33ING2 alone, without exposure to cellular stress, induced 10.4% of cell death compared to 5.0% in the control cells (P<0.001, t-test). p33ING1b was used as a positive control in this experiment as we previous reported that p33ING1b enhances UVB-induced apoptosis in melanoma cells (Cheung and Li, 2002).  Consistent with the previous findings, p33ING1b significantly induced cell  death after U V B irradiation ( P O . 0 0 0 1 , t-test). To confirm that p33ING2-mediated cell death after U V B irradiation is the result of apoptosis, not necrosis, cells were stained with propidium iodide (PI), which binds stoichiometrically to DNA.  Cells undergoing apoptosis are characterized by  chromatin condensation, formation of apoptotic bodies, and DNA fragmentation (Wylie, 1997; Hengartner, 2000).  The representative images taken under the  fluorescent microscope show that p33ING2-transfected M M R U cells underwent apoptosis after UVB irradiation (Fig. 3.1 E).  Quantitative data of PI staining in Fig.  3.1 F indicates that there were significantly more apoptotic cells in p33ING2overexpressing M M R U cells (43.8%) compared to the control cells (20.0%) (P<0.001, t-test).  Consistent with the results in Fig. 3.1 D, overexpression of p33ING2 alone  caused 17.1% of cells to undergo apoptosis compared to 5.1% in the control cells (P<0.01, t-test). The enhancement of apoptosis induction by p33ING2 was further confirmed by flow cytometry.  DNA fragmentation, a hallmark feature of apoptosis,  results in a hypodiploid DNA content represented by the Sub-Gi population in the  48  Figure 3.1 p33ING2 e n h a n c e s U V B - i n d u c e d cell death in m e l a n o m a M M R U c e l l s . (A) Western blot analysis of p33ING2 expression in M M R U cells transfected with pcDNA3-vector (V), pcDNA3-p33/A/G2 (ING2) or pcDNA3-antisense-p33/A/G2 (AS) expression vector. An anti-p33ING2 monoclonal antibody was used to detect p33ING2 and /5-actin was used as a loading control. (B) Cell survival rate by S R B assay of UVB-irradiated M M R U cells transfected with pcDNA3-vector (V), pcDNA3p33/A/G2 (ING2), or pcDNA3-AS-p33/A/G2 (AS) expression vector. Data represent means ± S D from triplicates. The experiment was repeated twice with similar results. (C) Microscopic images of M M R U cells transfected with pcDNA3-vector (V) or pcDNA3-p33/A/G2 (ING2) irradiated with U V B at 0 or 6 0 0 J / m . Photographic images were taken 24 h after U V B irradiation. Magnification, 2 0 0 x . (D) Cell death assessment by trypan blue exclusion assay on M M R U cells transfected with pcDNA3-vector (V), pcDNA3-p33/A/G2 (ING2) or pCI-p33/A/G7 (ING1b) after U V B irradiation. Data represent means ± S D from triplicates. The experiment was repeated three times with similar results. (E) Representative PI staining images of apoptotic cells after U V B irradiation. M M R U transfected with pcDNA3-p33//VG2 were irradiated with U V B at 6 0 0 J / m and images were taken under fluorescent microscope 24 h after UV irradiation. White arrowheads indicate cells with condensed nucleic and apoptotic bodies. (F) Quantitative data from PI staining of M M R U cells in E. Data represent means ± S D from triplicates. 2  2  49  V  ING2  50  AS  c  52  DNA cell cycle histogram.  Similarly, without UV treatment, cells overexpressing  p33ING2 had 20.7% apoptotic cells compared to 6.4% in the control cells (P<0.01, ttest) (Fig. 3.2A). After UVB irradiation, p33ING2 induced apoptosis in 46.2% of cells compared to 22.3% in the control cells ( P O . 0 0 1 , t-test) (Fig. 3.2A).  Clearly, cells  overexpressing p33ING2 displayed a larger Sub-Gi population compared to the control cells under both non-UV- and UV-irradiated conditions. The slight variation of the percentage of dead cells obtained by trypan blue exclusion assay and flow cytometry (Fig. 3.1 D and Fig. 3.2A) is possibly due to variability in the technical approaches. Collectively, we demonstrate that p33ING2 enhances UVB-induced apoptosis in melanoma cells.  3.2.2  p33ING2 Enhancement of UVB-induced Apoptosis Is p53-dependent  It was shown in various reports that functional p53 is required for p33ING1b in exerting its tumour suppressive functions, such as growth inhibition, apoptosis, and the repair of UV-damaged DNA (Garkavtsev ef al., 1996; Garkavtsev ef al., 1998; Shinoura ef al., 1999; Cheung et al., 2001; Cheung and Li, 2002). Furthermore, two reports on p33ING2 have shown that p33ING2-induced growth inhibition  and  apoptosis are p53-dependent (Nagashima ef al., 2001; Gozani et al., 2003). These observations led us to hypothesize that p33ING2 and p53 may work together in the enhancement of UVB-induced apoptosis in melanoma cells. To test our hypothesis, we co-transfected the wild-type p53 M M R U melanoma cells with p33/A/G2 and mutant p53 plasmid pED-1, and exposed the cells to UVB irradiation at 600 J / m . 2  Western blot analysis using an anti-p53 antibody, which recognizes both wt and  53  mutant p53, confirmed the expression of p53 in M M R U cells, suggesting successful transfection of M M R U cells (Fig. 3.2A).  Using flow cytometry, we found that co-  expression of p33ING2 and mutant p53 significantly reduced the enhancement of apoptosis by p33ING2 even without UVB irradiation (9.5% in co-transfected cells compared to 20.7% in p33/A/G2-transfected cells; P<0.001, t-test). Consistently, the enhancement of apoptosis by p33ING2 was significantly reduced by co-expression of p33ING2 and mutant p53 after U V B irradiation (34.2%) compared to 46.2% in p33/A/G2-transfected cells (P<0.001, t-test) (Fig. 3.2B). However, co-expression of p33ING2 and mutant p53 did not completely abrogate p33ING2-mediated U V B induced apoptosis.  This may be due to the endogenous wild-type p53 not being  completely inactivated. Therefore, we used M e W o melanoma cells that carry a mutant p53 gene to further investigate the p53 dependency of p33ING2 enhancement in UVB-induced apoptosis. S R B survival assay was carried out on vector-, p33/7VG2- or transfected M e W o cells after UVB irradiation with increasing doses.  p33ING1b-  A s reported  previously, p33ING1b, the additional control of this experiment, did not enhance UVB-induced cell death in the absence of functional p53. Similarly, overexpression of p33ING2 did not enhance UVB-induced cell death in the absence of functional p53 at all doses examined (Fig. 3.2D). These results were also supported by the results from flow cytometry analysis showing similar percentages of Sub-Gi apoptotic cells in M e W o cells transfected with vector or p33/A/G2 (Fig. 3.2E).  To confirm  the  requirement of functional p53 in p33ING2 enhancement of UVB-induced apoptosis, we transfected MeWo cells with wild-type p53 (pphwtp53) or wild-type p53 together  54  Figure 3.2 p33ING2 enhancement of UVB-induced apoptosis is p53-dependent. (A) Western blot analysis of p33ING2 and p53 expression in p53 wild-type M M R U cells transfected with pcDNA3-vector (V), pcDNA3-p33/A/G2 (ING2) or mutant p53 pED-1 expression vector. (B) Quantitation of apoptosis rate of M M R U cells transfected with pcDNA3-vector (V), pcDNA3-p33/A/G2 (ING2), or pED-1 and pcDNA3-p33/A/G2 (mutp53+ING2). Transfected cells were irradiated with U V B at 600 J / m and cells were trypsinized and analyzed by flow cytometry 24 h after UVB irradiation. Data represent means ± S D from triplicates. (C) Western blot analysis of p33ING2 and p53 expression in p53 mutant MeWo cells transfected with pcDNA3vector (V), pcDNA3-p33//VG2 (ING2) or wild-type p53 (p53) expression vector. (D) Survival assay of MeWo cells transfected with pcDNA3-vector (V), pcDNA3-p33/A/G2 (ING2) or pCI-p33/A/G) (ING1b). Cells were irradiated with various doses of U V B and S R B survival assay was performed 24 h after UV irradiation. Data represent means ± S D from triplicates. The experiment was repeated twice with similar results. (E) Cell death assessment by flow cytometry of M e W o cells transfected with pcDNA3-vector (V), pcDNA3-p33/A/G2 (ING2), wild-type p53 pphwtp53 (p53), or pphwtp53 and pcDNA3-p33/A/G2 (p53+ING2). Transfected cells were irradiated with UVB at 600 J / m and flow cytometry was performed 24 h after UV irradiation. Number in the histograms represents the percentage of Sub-G-i population. The experiment was repeated twice with similar results. 2  2  55  A  V  ING2 pED-1 ING2  m m m  56  V  ING2 p53 ING2 p53 Actin  400  200  600  UVB Dose (J/m ) 2  ING2 5.4  p53  p53+ING2  6.4  6.9  7.5  OJ 10 •«->  c 3 o o  i  29,0%  25. )%  2 4 . 0%  J  24. 9%  600J i  1 ku4  DNA Content  57  with p33//VG2, to  see if  re-introduction  enhancement of apoptosis by p33ING2.  of wild-type  p53 could  restore  the  Western blot analysis confirmed  the  expression of p33ING2 and p53 in MeWo cells, suggesting successful transfection of M e W o cells (Fig. 3.2C). Our data indicated that neither transfection with wild-type p53 alone nor co-transfection with p53 and p33/A/G2 sensitized MeWo cells to U V B induced apoptosis.  This may be due to the presence of dominant negative p53  mutation in MeWo cells that abrogates the effect of the wild-type p53 re-introduced into the cells.  3.2.3  p33ING2 Increases the Ratio of Bax/Bcl-2 in Melanoma Cells  Recent findings of ING proteins on upregulation of Bax expression and its promoter activity (Nagashima et al., 2001; Nagashima er al., 2003; Cheung and Li, 2002) led us to hypothesize that p33ING2 enhances UVB-induced apoptosis by upregulating Bax  expression  in melanoma  cells.  To determine  if p33ING2  upregulates  endogenous Bax expression, we transfected M M R U cells with empty vector or p33/7VG2 and subjected these cells to UVB irradiation at 0 or 600 J / m . Twenty-four 2  hours after UV irradiation, proteins were extracted and Western blot analysis was performed to determine Bax expression in these cells. The levels of Bax protein were normalized with the levels of /?-actin, the loading control, accordingly.  The  normalized Bax expression level in non-UV-irradiated vector-transfected cells was arbitrarily defined as 1, and the relative fold of Bax induction of other samples was calculated and indicated below each band in the representative figure of Western blot analysis (Fig. 3.3A). Our results demonstrate that cells overexpressing p33ING2  58  Figure 3.3 p33ING2 upregulates Bax and downregulates Bcl-2 expression. (A) Representative figure of Western blot analysis of Bax and Bcl-2 proteins. M M R U cells transfected with p3XFLAG-vector (V) or p3XFLAG-//VG2 (ING2) were irradiated with U V B at 0 or 600 J / m . Cells were harvested and lysed for Western blot analysis 24 h after UV irradiation. Antibodies against p33ING2, Bax, and Bcl-2 were used. Anti-p33ING2 antibody detected both endogenous (lower bands) and transfected p3XFLAG-/A/G2 (upper bands). y?-actin was used as a loading control. The number below each band indicates the relative fold of Bax induction or Bcl-2 reduction by densitometry. (B) Ratio of Bax/Bcl-2 proteins in non-UV-irradiated and UV-irradiated M M R U cells transfected with p3XFLAG-vector (V) or p3XFLAG-/A/G2 (ING2). Data represent means ± S D from three independent experiments. 2  OJ V  600J  ING2  V  ING2 ING2  Bax 1  1.38  1.21  1.42 Bcl-2  1  0.93 0.91  0.30 __t_i_ ! , M |  w  B  i  •V • ING2  re k m re CQ  "I.  600  UVB Dose (J/m ) 2  59  A p f i n A  C  l  ,  n  have higher Bax expression compared to the vector controls without UV irradiation. However, the induction of Bax expression by p33ING2 was not enhanced after UV irradiation (Fig. 3.3A). These observations led us to examine the expression levels of anti-apoptotic protein, Bcl-2, which was found to neutralize Bax pro-apoptotic effects by forming heterodimers with Bax (Hengartner, 2002).  Interestingly, we found that  p33ING2 downregulated Bcl-2 expression, with more significant reduction of Bcl-2 protein after UV irradiation (Fig. 3.3A).  The ratio of Bax to Bcl-2 is thought to  determine the degree of heterodimer formation, and thus the sensitivity of cells to apoptosis (Adam and Cory, 1998). Therefore, we determined the ratio of Bax to Bcl2 in these samples. Our results revealed that p33ING2 significantly increased the ratio of Bax/Bcl-2, with more profound increment (4.6-fold) after UV irradiation ( P O . 0 0 0 1 , t-test) (Fig. 3.3B). These data strongly suggest that p33ING2 enhances UVB-induced apoptosis by increasing the overall active Bax expression levels.  3.2.4  p33ING2 Alters Mitochondrial Membrane Potential after UVB Irradiation  It has been shown that UV irradiation activates the mitochondrial apoptotic pathway, and that p33ING1b promotes UVB-induced apoptosis via this pathway (Cheung and Li, 2002; Antonsson et al., 2001; Green and Reed, 1998). Overall increase of Bax induced by p33ING2 suggests that p33ING2 may also mediate UVB-induced apoptosis via the mitochondrial pathway. The mitochondrial membrane potential is decreased in the early stage of apoptosis induced by many types of stimuli (Li, et al., 1999; Zhang et al., 1999a; Hengartner, 2000).  Therefore, we first investigated  whether p33ING2 activates the mitochondrial apoptotic pathway by determining the  60  mitochondrial membrane potential changes after U V B irradiation.  W e transfected  M M R U cells with empty vector or p33/A/G2 and exposed the cells to U V B irradiation at 600 J / m . Twenty-four hours later, cells were stained with a cationic dye using the 2  MitoCapture apoptosis detection kit. The cationic dye accumulated and aggregated in the mitochondria in healthy cells and emitted an orange-red fluorescence.  In  apoptotic cells, the dye remained in the cytoplasm due to alteration of the mitochondrial membrane potential and emitted green fluorescence. were visualized under a fluorescent microscope. irradiated  control  Stained cells  Fig. 3.4A shows that non-UV-  cells stained orange-red while green fluorescence staining  (apoptotic cells) was observed in UVB-irradiated cells. After UV irradiation, there is a significantly higher percentage of apoptotic cells in p33//VG2-transfected cells compared to vector control (31.8% versus 7.7%; P<0.001, t-test) (Fig. 3.4B). These data suggest that p33ING2 enhances UVB-induced apoptosis via the mitochondrial apoptotic pathway by altering the mitochondrial membrane potential.  3.2.5  p33ING2 Promotes Bax Translocation and Cytochrome c Release after  UVB Irradiation Upon activation, Bax translocates to the mitochondria where active Bax oligomerizes in the mitochondrial outer membrane to increase mitochondrial permeability, leading to cytochrome c release and apoptosis (Cory and Adam, 2002; Igney and Krammer, 2002). Since p33ING2 upregulation of Bax is not enhanced in melanoma cells after UV irradiation (Fig. 3.3A), we hypothesized that p33ING2 may also enhance apoptosis by promoting translocation of Bax to the mitochondria upon UV irradiation.  61  Figure 3.4 p33ING2 alters mitochondrial membrane potential after UVBirradiation. M M R U cells transfected with pcDNA3-vector (V) or pcDNA3-p33//VG2 (ING2) were irradiated with U V B at 0 or 6 0 0 J / m . Disruption of mitochondrial membrane potential was detected using the MitoCapture apoptosis kit 24 h after UVB irradiation. (A) Representative images of M M R U control cells and cells overexpressing p33ING2 after UVB irradiation. Live cells emit orange-red fluorescence and apoptotic cells emit green fluorescence (indicated by white arrow heads). Magnification, 4 0 0 x . (B) Quantitation of the percentage of apoptotic cells based on the changes in the mitochondrial membrane potential. A total of 500 cells were counted for each sample. Data represent means ± S D from triplicates. 2  A  OJ  600J 9  a  J  %  • -*'•-• • H* v.-  *  4  *  i'  :  tl *p  \ i:  i • :'  t-  -  ^  •  #  A <  V  m  • .  ING2  V ft  62  %  To test our hypothesis, we transfected M M R U cells with empty vector or p33/A/62 and subjected these cells to UVB irradiation at 600 J / m . Cells were harvested 24 h 2  after UV irradiation and subcellular fractionation was carried out to separate the cytosolic and mitochondrial fractions, which were then subjected to Western blot analysis.  Fig. 3.5 shows that in the non-UV-irradiated cells, Bax localized in the  cytosol and cytochrome c was mainly found in the mitochondria. Cytochrome c was partially released from the mitochondria in cells overexpressing p33ING2, consistent with the findings that p33ING2 alone can induce apoptosis (Fig. 3.1 D, 3.1 F, and 3.2B). Twenty-four hours after UV irradiation, Bax translocated from the cytosol to the mitochondria, and cytochrome c was released from the mitochondria into the cytosol.  There is more Bax translocation and cytochrome c release in cells  overexpressing p33ING2 compared to the control cells. These data indicate that p33ING2 promotes Bax translocation to the mitochondria, causing mitochondrial membrane potential changes and thus promotes cytochrome c release into the cytosol to initiate apoptosis.  3.2.6  p33ING2 Induces the Activation of Caspase 9 and Caspase 3 after UVB  Irradiation Released cytochrome c facilitates the interaction between Apaf-1 and pro-caspase 9, resulting in the cleavage and activation of caspase 9 and downstream effector caspases, such as caspases 3 and 7 (Li et al., 1997; Cryns and Yuan, 1998). To confirm if the mitochondrial apoptotic pathway is activated in p33ING2-mediated UVB-induced apoptosis, M M R U cells transfected with empty vector or p33//VG2 were  63  Figure 3.5 p33ING2 promotes B a x translocation and c y t o c h r o m e c release. M M R U cells transfected with p3XFLAG-vector (V) or p3XFLAG-//VG2 (ING2) were irradiated with UVB at 0 or 600 J / m . Subcellular fractionation was performed 24 h post irradiation and Western blot analysis was carried out on the cytosolic (c) and mitochondrial (m) fractions. Antibodies against Bax and cytochrome c were used. f3actin was used as a loading control. 2  OJ V  ING2  600J V  ING2 Bax (c) Bax (m) °y  t o  c  <) c  Cyto c (m) Actin  64  UVB-irradiated at 0 or 600 J / m . Cells were harvested 12 h after UV irradiation and 2  lysates were subjected to Western blot analysis on the cleavage of caspase 9 and caspase  3.  Fig. 3.6  showed that the  pro-caspase 9 levels in p33ING2-  overexpressing cells after UV irradiation were greatly reduced compared to the vector control cells and non-UV-irradiated p33ING2-overexpressing cells (0.73- and 0.69fold respectively, P<0.01, t-test). significant  However, without UV irradiation, there was no  cleavage of caspase 9 observed in p33ING2-overexpressing cells  compared to the vector control cells. Consistently, pro-caspase 3 (35 kDa) levels were reduced and more cleaved caspase 3 fragments (19 and 17 kDa) were produced in cells overexpressing p33ING2 compared to the vector control cells. Cells overexpressing p33ING2 showed 0.69- and 0.85-fold  of pro-caspase 3  reductions compared to the vector control cells, with and without exposure to UV irradiation respectively (P<0.01, t-test).  The cleavage was more pronounced in  p33ING2-overexpressing cells after UV irradiation compared to non-UV-irradiated control cells.  These observations strongly suggest that p33ING2 induces the  activation of caspase 9 and caspase 3 in response to UV irradiation.  3.2.7  p33ING2 Activates the Death-receptor Apoptotic Pathway  Previous studies have shown that UV irradiation also activates the death-receptor or extrinsic apoptotic pathway by activating Fas and releasing tumour necrosis factor-a (TNF-a) (Schwarz et al., 1995; Rehemtulla et al., 1997; Aragane et al., 1998). Therefore, we investigated if p33ING2 induces activation of this alternative pathway upon UV irradiation. The Western blot in Fig. 3.7 shows that p33ING2 upregulates  65  Figure 3.6 p33ING2 induces the activation of c a s p a s e s 9 a n d 3. M M R U cells transfected with p3XFLAG-vector (V) or p3XFLAG-/A/G2 (ING2) were irradiated with UVB at 0 or 600 J / m . Cells were harvested 24 h after U V irradiation and lysates were prepared for Western blot analysis using anti-p33ING2, anti-caspase 9 and anticaspase 3 antibodies. Anti-caspase 9 antibody detected pro-caspase 9 (47 kDa) and anti-caspase 3 antibody detected pro-caspase 3 (35 kDa) the cleaved fragments (19 and 17 kDa). /?-actin was used as a loading control. 2  OJ  V  ING2  600J  V  ING2 ING2 —Pro-caspase 9 —Pro-caspase 3  ^Cleaved caspase 3 Actin  66  the expression of the death receptor Fas by 12.0- and 10.6-fold in non-UV- and UVirradiated cells respectively (P<0.001, t-test). These results suggest that p33ING2 enhancement of UVB-induced apoptosis may be partially attributed to the death receptor-mediated apoptosis pathway. U V C irradiation was found to upregulate both Fas in primary keratinocytes (Leverkus et al., 1997). However, we did not observe upregulation of Fas in the control cells after UVB irradiation, possibly due to different wavelengths of UV and cell types used. To confirm the involvement of p33ING2 in the death-receptor apoptotic pathway, we examined the cleavage of caspase 8, the initiator of the caspase cascade in the death receptor apoptosis pathway.  The  majority of the pro-caspase 8 (57 kDa) was cleaved in cells overexpressing p33ING2, producing higher levels of the cleaved fragments (43, 41 and 18 kDa) compared to that in the control cells both in non-UV- and UV-irradiated cells (Fig. 3.7). The nonUV-irradiated p33ING2-overexpressing cells showed a 0.47-fold reduction of procaspase 8 levels compared to the vector control cells (P<0.01, t-test).  However,  there was not substantially more caspase 8 activation observed in UVB-irradiated p33ING2-overexpressing  cells  compared  to  the  non-UV-irradiated  p33ING2-  overexpressing cells. These data suggest that p33ING2 activates the death-receptor apoptotic pathway regardless of UVB irradiation.  3.3  Discussion  Since p33/A/G2 was cloned in 1998, there have only been three studies indicating the role of p33ING2 as a tumour suppressor candidate through regulation of gene transcription, induction of cell cycle arrest and apoptosis in human fibrosarcoma,  67  Figure 3.7 p33ING2 activates the death-receptor apoptotic pathway. M M R U cells transfected with p3XFLAG-vector (V) or p3XFLAG-/A/G2 (ING2) were irradiated with U V B at 0 or 600 J / m . Cells were harvested 24 h after UV irradiation and lysates were prepared for Western blot analysis using anti-p33ING2, anti-Fas (Ab-1), and anti-caspase 8 antibodies that detects both pro-caspase 8 (57 kDa) and the cleaved fragments (43, 41 and 18 kDa). /3-actin was used as a loading control. 2  A  OJ  V  ING2  600J  V  ING2 ING2 Fas Actin  OJ  V  ING2  600J  V  ING2 ING2 ^-Pro-caspase 8 3 = Cleaved caspase 8 Cleaved caspase 8 Actin  68  colorectal carcinoma, and hepatocellular carcinoma cell lines (Nagashima er al., 2001; Gozani et al., 2003; Kataoka et al., 2003).  Recent studies showed that  overexpression of p33ING1b appeared to enhance DNA repair and apoptosis in melanoma cells and fibroblasts (Cheung et al., 2001; Scott et al., 2001a; Scott et al., 2001b; Cheung and Li, 2002).  Furthermore, p33ING1b has also been shown to  translocate to the nucleolus and bind to P C N A after UV irradiation (Scott et al., 2001a; Scott et al., 2001b).  It was reported that p33ING2 protein expression was  specifically induced by DNA damaging agents such as etoposide  in  normal  lymphoblastoid C 3 A B R cells and osteosarcoma O s A - C L cells, and that this induction of p33ING2 parallels the increase of acetylation of p53 at Lys382 (Nagashima et al., 2001). However, information on the role of p33ING2 in the cellular stress response to UV irradiation is lacking. To investigate the role of p33ING2 in UV stress response, we for the first time demonstrated that p33ING2 enhances UVB-induced apoptosis in melanoma cells, and this enhancement requires the participation of functional p53. This is consistent with the findings that its homologue p33ING1b also cooperates with functional p53 in cell growth control and apoptosis (Garkavtsev et al., 1998; Shinoura et al., 1999; Cheung and Li, 2002). Recent studies have shown that other ING family members, p47ING3, p29ING4 and p28ING5, like p33ING1b and p33ING2, induce growth arrest and apoptosis in a p53-dependent manner (Nagashima er al., 2003; Shiseki et al., 2003). p33ING1b, p29ING4, and p28ING5 are now known to co-precipitate with p53, while p33ING2 and p47ING3 do not (Nagashima et al., 2001; Nagashima et al., 2003; Shiseki era/., 2003). In addition, p33ING1b, p33ING2, p29lNG4, and p28ING5  69  are found to stimulate acetylation of p53 at Lys382 and/or Lys373 (Nagashima ef al., 2001; Kataoka ef al., 2003; Shiseki ef al., 2003), resulting in activation  and  stabilization of p53, and thus promote p53 sequence-specific DNA-binding and transcription activation (Gu and Roeder, 1997; Sakagushi ef al., 1998).  Based on  these findings, it is reasonable to propose that ING proteins enhance the p53 response through physical association with p53 and acetylation of p53. Furthermore, recent studies provide evidence that p33ING1b is involved in chromatin remodelling through physical association with S A P 3 0 of the Sin3-histone deacetylase (HDAC) corepressor complexes and with T R R A P , P C A F , and p300/CBP, which  constitute  histone acetyltransferase (HAT) co-activator complexes (Skowyra ef al., 2001; Kuzmichev ef al., 2002; Vieyra et al., 2002). p53 has also been found to constitute HAT co-activator and H D A C co-repressor complexes (Feng ef al., 2002).  It was  reported that p53 could both induce and repress its downstream transcription targets coordinately, depending on the cell type and stress context (Weber and Zambetti, 2003).  For instance, during p53-dependent apoptosis, pro-apoptotic genes Bax,  PUMA,  and Noxa are induced while anti-apoptotic genes Map4 and stathmin are  selectively repressed via recruitment of p53-Sin3A or p53-PID H D A C complexes (Zhang et al., 1999b; Johnsen ef al., 2000; Hussein ef al., 2003a). Thus it is likely that p33ING2 cooperates with p53 in modulating the state of histone acetylation, which is known to affect gene expression (Feng ef al., 2002).  p33ING2 may cooperate with p53 in regulating the transcription of specific proapoptotic and anti-apoptotic genes in response to UV irradiation.  The proper  expression of pro-apoptotic and anti-apoptotic proteins dictates to the cells whether  70  to survive or undergo apoptosis. Bax, a downstream transcription target of p53, is a pro-apoptotic member of the Bcl-2 family that resides in the cytosol as a monomer. Upon induction of apoptosis, Bax undergoes conformational changes, translocates to the outer mitochondrial membrane and oligomerizes. This causes permeabilization of the outer mitochondrial membrane, possibly through the opening of permeability transition pore (PTP), therefore allowing efflux of apoptogenic factors such as cytochrome c and Smac/DIABLO (Hengartner, 2000; Cory and Adams, 2002; Hsu and Youle, 1998; Murphy et al., 1999; Gross et al., 1999; Shimizu et al., 1999). In contrast, anti-apoptotic Bcl-2, a mitochondrial membrane protein, antagonizes the pro-apoptotic effect of Bax by forming heterodimers with Bax (Bush and Li, 2003). In this study, we found that p33ING2 upregulates the endogenous expression of Bax (Fig. 3.3A). However, this upregulation is not enhanced after UV irradiation, which does not explain the enhancement of apoptosis by p33ING2 seen in Fig. 3.1 and 3.2. Nevertheless, we also found that p33ING2 significantly downregulates Bcl-2 expression and increases the ratio of Bax/Bcl-2, most drastically after UV irradiation (Fig. 3.3B). The balance between the pro-apoptotic protein Bax and anti-apoptotic protein Bcl-2 determines the sensitivity of cells to apoptosis; cells with more proapoptotic proteins are sensitive to death, while cells with an excess of anti-apoptotic Bcl-2 family members are usually resistant (Hengartner, 2000). The increase of the Bax to Bcl-2 ratio by p33ING2 increases the overall active Bax expression levels, and thus enhances cell death via the mitochondrial apoptotic pathway.  In addition, we  found that, upon UV irradiation, p33ING2 promotes Bax translocation to the mitochondria, enables Bax oligomerization in the mitochondrial outer membrane,  71  enhances mitochondrial membrane potential changes, and thus induces cytochrome c release from the mitochondria into the cytosol (Fig. 3.4 and 3.5). Once released, cytochrome c binds to the cytoplasmic scaffolding protein Apaf-1, causing ATP-dependent conformational changes that allow Apaf-1 to bind to the prodomain of an inactive initiator caspase, pro-caspase 9, to form a heptameric apoptosome (Hersey and Zhang, 2001; Igney and Krammer, 2002). Pro-caspase 9 in the apoptosome is activated by an allosteric change on Apaf-1 in the apoptosome (Li et al., 1997; Stennicke et al., 1990).  The active caspase 9 then initiates a  cascade of proteolytic activation of other effector/executioner caspases, first by cleaving pro-caspase 3 and 7, followed by cleavage of pro-caspase 6 by activated caspase 3. This parallels our observations that both caspase 9 and caspase 3 are cleaved after U V irradiation, and p33ING2 enhances the cleavage and thus activates both the initiator and effector caspases (Fig. 3.6). Surprisingly, caspase 8, an initiator caspase of the death-receptor pathway, is also found to be activated by p33ING2 (Fig. 3.7). This may have resulted from the upregulation of the death receptor Fas expression by p33ING2 (Fig. 3.7).  The observations that p33ING2 alone, without  UVB irradiation, can induce apoptosis (Fig. 3.2A) may be explained by the activation of Fas/caspase 8 pathway as a comparable amount of Fas and caspase 8 are activated in p33/A/G2-transfected cells with or without UVB irradiation (Fig. 7). Since Fas is also a transcription target of p53 (Johnstone ef al., 2002), it is possible that p33ING2 cooperates with p53 in upregulating Fas transcription. When activated, Fas homotrimerizes, leading to the recruitment of FAS-associated death domain (FADD) which further recruits pro-caspase 8 to form death-inducing signalling complex  72  (DISC). Pro-caspase 8 in DISC is activated by autocatalytic cleavage and the active caspase 8 activates effector caspases such as caspase 3, which explains the increased caspase 3 cleavage in p33ING2-overexpressing cells compared to vector control without UV irradiation (Fig. 3.6, lanes 1 and 2). The active effector caspases then cleave a number of cellular substrates, which lead to characteristic biochemical and morphological changes observed in Fig. 3.1 C and E (Igney and Krammer, 2002). Since Bid is a transcription target of p53, and cleavage and activation of Bid by active caspase 8 can activate the mitochondrial pathway after apoptosis induction through death receptors (Luo et al., 1998; Sax et al., 2002), it would be of interest to investigate the role of Bid in p33ING2-mediated apoptosis and the cross-talk between the intrinsic and extrinsic apoptosis pathways. In summary, this study demonstrates that p33ING2 cooperates with p53 to regulate apoptosis via activation of both the mitochondrial/intrinsic and deathreceptor/extrinsic apoptotic pathways.  Under non-stressed conditions, p33ING2  upregulates Fas expression and activates caspase 8.  Upon UVB irradiation,  p33ING2 significantly downregulates Bcl-2 expression and increases the Bax/Bcl-2 ratio, promotes Bax translocation to mitochondria, alters mitochondrial membrane potential, induces cytochrome c release and activation of caspase 9, and therefore enhances UVB-induced apoptosis. These findings provide a better understanding of the role of the tumour suppressor candidate p33ING2 in cellular stress response to UV irradiation, which may lead to new insights into more effective prevention and treatment regimes for human cutaneous melanoma.  73  CHAPTER 4. p33ING2 ENHANCES THE REPAIR OF UV-DAMAGED DNA IN HUMAN MELANOMA CELLS  4.1  Rationale and Hypothesis  DNA repair and apoptosis are two essential mechanisms by which cells maintain their genomic integrity.  DNA damages accumulate due to replication errors,  endogenous and environmental DNA-damaging agents such as reactive oxygen species (ROS) and ultraviolet (UV) rays. These damages can be removed by the DNA repair network, or otherwise senescence and apoptosis if the damages are beyond repair (Mitchell ef al., 2003).  One of the major DNA repair pathways is  nucleotide excision repair (NER), which eliminates a variety of DNA damage including UV-induced photolesions, cyclobutane pyrimidine dimers (CPDs) and 6-4 photoproducts (6-4PPs) (Tanaka ef al., 2001). The complete N E R reaction involves a sequential assembly of repair proteins to the damage sites to mediate damage recognition, dual incision and gap-filling DNA synthesis (Petit and Sancar, 1999). Defects in N E R lead to increased cancer incidence as observed in xeroderma pigmentosum  (XP), developmental and neurological abnormalities as seen in  Cockayne syndromes (CS) and trichothiodystrophy; all of these are characterized by a high sensitivity to sunlight (Mitchell ef al., 2003; Tanaka ef al., 2001). Several studies have reported that the expression of wild-type p53 is required for efficient N E R of UV-induced DNA damage. For instance, Li-Fraumeni fibroblasts homozygous for p53 mutation were found to be deficient in the rate and extent of N E R of the genomic DNA (Ford and Hanawalt, 1995; 1997). Furthermore, reduced  74  N E R was also observed when the wild-type p53 function was disrupted by the human papillomavirus E6 oncogene that targets p53 for degradation, or expression of a dominant-negative mutant p53 (Smith et al., 1995). Recent studies have shown that p53 is involved both directly and indirectly in N E R of UV-induced photolesions. The direct participation of p53 in this process is demonstrated by p53 binding to X P B and X P D helicases via its C-terminus (Wang et al., 1995). p53 has also been shown to regulate both the basal and UV-inducible expression levels of p48, a subunit of damaged DNA binding protein (DDB), as well as the XPC gene (Hwang et al., 1999; Adimoolam er al., 2002). More recently, Rubbi and Milner (2003) have demonstrated that p53 can act as a chromatin accessibility factor that mediates UV-induced global chromatin relaxation by recruiting p300 to the sites of N E R and inducing histone acetylation, suggesting indirect participation of p53 in N E R . p53 dependency of the tumour suppressive functions of p33ING2, as well as other members of the ING tumour suppressor family, has been well documented (Garkavtsev et al., 1998; Shinoura et al., 1999; Cheung and Li, 2002; Nagashima et al., 2001; Nagashima et al., 2003; Shiseki et al., 2003). Two main mechanisms may help explain how ING proteins enhance p53 responses: the association of ING members with p53 and ING-mediated acetylation of p53 (Campos et al., 2004a). Although p33ING2 does not physically associate with p53, it modulates the activity of p53 by enhancing the stability of p53 through acetylation at Lys382 and/or Lys373 (Nagashima et al., 2001; Kataoka et al., 2003).  Recent studies indicate that the  interaction of p33ING2 with phosphoinositides, Ptdlns(3)P and Ptdlns(5)P, through its plant homeodomain (PHD) zinc finger motif at the C-terminus is critical for its ability  75  to enhance p53 acetylation and mediate p53-dependent apoptosis (Gozani et al., 2003). Previously, we found that overexpression of p33ING1b enhances the repair of UV-damaged DNA in melanoma cells, and that this process requires the presence of functional p53.  In addition, the physical association between p33ING1b and  G A D D 4 5 was detected, suggesting the involvement of p33ING1b in G A D D 4 5 mediated N E R pathway (Cheung et al., 2001 b). More recently, we demonstrated that mutations at codon 102 or 260 as well as deletion of the P H D finger motif abolished p33ING1 b enhancement in N E R , suggesting that P H D zinc finger motif is required for proper D N A repair functions (Campos et al., 2004b).  In light of the structural and  functional similarities between p33ING2 and p33ING1b (Feng ef al., 2002; Campos ef al., 2004a), we hypothesized that p33ING2 may also enhance the repair of UVdamaged DNA in a p53-dependent manner.  76  4.2  4.2.1  Results  p33ING2 Enhances the Repair of UV-damaged DNA  To investigate the role of p33ING2 in repairing UV-damaged DNA, we performed host-cell-reactivation assay by co-transfecting a UV-damaged pCMVCAT plasmid that contains the chloramphenicol acetyltransferase reporter gene with either pcDNA3vector, pcDNA3-p33/A/G2 or pcDNA3-AS-p33/A/G2 into melanoma M M R U cells. Western blotting confirmed the expression of these plasmids, suggesting successful transfection of M M R U cells (Fig. 4.1 A). The activity of this reporter gene was used as an indicator of the extent of repair. W e found that M M R U cells overexpressing p33ING2 had a significantly higher rate of repair of the UV-damaged plasmid (59.1%) compared to the vector (24.1%; P<0.01, t-test) or antisense control (26.0%; P<0.01, t-test) (Fig. 4.1B). These results indicate that p33ING2 enhances the repair of UVdamaged DNA in melanoma cells.  4.2.2  p33ING2 Does not Colocalize with UV-induced DNA Lesions  To study if p33ING2 is a component of the N E R complex, M M R U cells transiently overexpressing p33ING2 were irradiated with U V C at 200 J / m with an isopore 2  polycarbonate filter containing 5 u.m pores overlaid on the cells. Thus, only part of the nucleus was UV-irradiated through the microfilter, while remaining parts of the nucleus covered by the filter were blocked from the radiation.  Therefore, this  technique is useful in exploring protein colocalization with UV-induced DNA lesions. The intracellular localization of p33ING2 with two major UV-induced D N A lesions,  77  Figure 4.1 p33ING2 e n h a n c e s the repair of UV-damaged DNA. (A) Western blot analysis of p33ING2 expression in M M R U cells transfected with pcDNA3-vector (V), pcDNA3-p33//VG2 (ING2) or pcDNA3-antisense-p33//VG2 (AS) expression vector. An anti-p33ING2 monoclonal antibody was used to detect p33ING2 and (3actin was used as a loading control. (B) Assessment of the repair rate of UVdamaged DNA by host-cell-reactivation assay in M M R U cells co-transfected with undamaged or damaged DCMVCAT and pcDNA3-vector (V), pcDNA3-p33//VG2 (ING2), or pcDNA3-AS-p33/A/G2 (AS) expression vector. C A T activity was determined by scintillation counting and expressed as net dpm damaged dose/net dpm zero dose. Data represent means ± S D from triplicates. The experiment was repeated thrice with similar results.  78  cyclobutane pyrimidine dimers (CPDs) and 6-4 photoproducts (6-4PPs), at different time-points after UV irradiation were examined using immunofluorescent staining. Cells double-labelled for X P B and 6-4PPs were used as a positive control as it was previously shown that X P B colocalized with UV-induced DNA lesions (Wang et al., 2003). Fig. 4.2A shows that the majority of X P B (green) colocalized with the 6-4PP sites (red) 1 h after UV irradiation. Consistent with a previous report (Gozani et al., 2003), the majority of p33ING2, labelled with green fluorescence, was detected in the nucleus with a weak diffuse distribution in the cytoplasm (Fig 4.2B and C).  The  photolesions, 6-4PPs and C P D s were labelled with red fluorescence.  The  fluorescent intensity of 6-4PPs faded at a quicker rate than that of C P D s . It became very weak 4 h after U V irradiation and was undetectable after that, whereas the fluorescent intensity of C P D s remained strong in all time-points examined.  Our  results demonstrated that p33ING2 does not colocalize with either DNA lesions, 64 P P s or C P D s at all time-points examined (Fig. 4.2B and C), suggesting that p33ING2 is not a component of the repair core complexes which are recruited to UVinduced DNA lesions.  4.2.3  p33ING2 Enhancement of DNA Repair Is p53-dependent  Since p33ING2 may be involved in the repair of UV-damaged DNA via indirect mechanisms, it is likely that p33ING2 cooperates with other factors in performing this function. Supported by the finding that its homologue, p33ING1 b, requires functional p53 to enhance N E R (Cheung et al., 2001), we investigated whether p33ING2 enhancement of DNA repair requires the presence of p53. W e disrupted the  79  Figure 4.2 p33ING2 does not colocalize with UV-induced D N A lesions. (A) M M R U cells were fixed 1 h after exposure to U V C at 200 J / m through a 5 u,m isopore polycarbonate filter cover. Cells were double-labelled for X P B (green) and 64 P P s (red). (B) and ( C ) M M R U cells transfected with p3XFLAG-/A/G2 were exposed to U V C at 200 J / m through a 5 u.m isopore polycarbonate filter cover. Cells were fixed at 0, 5, 15, 30 min and 1, 4, 8 and 24 h, and double-stained for p33ING2 (green) and 2 major U V photoproducts C P D s and 6-4PPs (red). Representative immunofluorescent staining images of 6-4PPs (B) and C P D s ( C ) . Nuclei were stained blue with Hoechst stain. Magnification, 400x. 2  2  A  C  Hoechst  XPB  6-4PP  Merge  Hoechst  °'^|  1h  •  o DB ING2  CPD  5'l  B  Hoechst 0*  ING2  m 6^4PP  Merge  5'  30'|  •  15"  30'  1h  4hl  •  8hl  im  24h 4h  80  1  Merge  endogenous wt p53 in M M R U cells by introducing a dominant-negative mutant p53 pED-1 plasmid (Cheung et al., 2001).  Western blot analysis confirmed  the  expression of pED-1 (refer to Section 3.2.2, Fig 3.2A). Using host-cell-reactivation assay, we found that mutant p53 significantly reduced the enhancement of DNA repair by p33ING2 (42.4% in cells co-transfected with p33/A/G2 and pED-1 vs 64.7% in p33/A/G2-transfected cells; P<0.01, t-test) (Fig. 4.3A). However, mutant p53 did not completely abrogate p33ING2-mediated UV-damaged D N A repair. This may be due to the endogenous wt p53 not being completely inactivated. It has been shown that the expression levels of p33ING2 in cells transfected with pED-1 were similar to those in the vector control (refer to Section 3.2.2, Fig 3.2A), eliminating the possibility that the reduced rate of DNA repair seen in pED-1-transfected cells was due to downregulated p33ING2 expression by mutant p53. Next, we performed host-cellreactivation assay using MeWo cells that carry a mutant p53 gene to further validate the p53 dependence of p33ING2 enhancement in N E R . Our results showed that p33ING2 did not enhance the repair of UV-damaged DNA in the absence of functional p53 (Fig. 4.3B). W e also transfected MeWo cells with wt p53 (pphwtp53) or wt p53 together with p33/A/G2, to see if re-introduction of wt p53 could restore the enhancement of N E R by p33ING2.  The expression of p33ING2 and wt p53  (pphwtp53) in M e W o cells was confirmed using Western blot analysis (refer to Section 3.2.2, Fig 3.2C). Our data in Fig. 4.3B showed that neither transfection with wt p53 alone nor co-transfection with p53 and p33/A/G2 increased the DNA repair capacity in MeWo cells. This may be due to the presence of dominant negative p53 mutation in MeWo cells that abrogates the effect of the wt p53 re-introduced into the  81  Figure 4.3 p33ING2 enhancement of DNA repair is p53-dependent. (A) Hostcell-reactivation assay was performed in M M R U cells co-transfected with UVdamaged PCMVCAT and pcDNA3-vector (V), pcDNA3-p33//VG2 (ING2), or pcDNA3p33/A/G2 and mutant p53 pED-1 (ING2+mut p53). (B) Host-cell-reactivation assay was performed in MeWo cells co-transfected with UV-damaged OCMVCAT and pcDNA3-vector (V), pcDNA3-p33/WG2 (ING2), wild-type p53 (wt p53) alone or pcDNA3-p33/A/G2 and wild-type p53 (ING2+wt p53). Data represent means ± S D from triplicates. The experiment was repeated twice with similar results.  82  83  cells.  These observations suggest that functional p53 is required for p33ING2  enhancement of DNA repair.  4.2.4 p33ING2 Enhances p53 Acetylation upon UV Irradiation Recent findings that p33ING2 and other ING proteins (p33ING1b, p29ING4 and p28ING5) (Campos et al., 2004a) induced p53 acetylation in various cell lines prompted us to examine whether p33ING2 enhances p53 acetylation after UV irradiation.  W e exposed M M R U cells overexpressing p33ING2 and control cells to  UVB irradiation at 200 J / m , and examined the expression levels of acetylated p53 at 2  different time-points (Fig. 4.4A). M M R U cells treated with 0.2 jag/ml of doxorubicin for 8 h were used as a positive control for p53 acetylation.  p33ING2 induced p53  acetylation in both non-UV- and UV-irradiated cells compared to the vector control. Induction of p53 acetylation was associated with a maximal accumulation of p53 protein in cells overexpressing p33ING2 4 h after UV irradiation (Fig. 4.4A).  4.2.5 p33ING2 Upregulates XPA Induction of p53 acetylation by p33ING2 after UV irradiation led us to hypothesize that p33ING2 may upregulate the p53 downstream target G A D D 4 5 , which has been shown to be essential in repairing UV-damaged D N A (Smith et al., 2000).  We  irradiated M M R U cells transfected with empty vector or p33/A/G2 with U V B at 200 J / m and performed Western blot analysis 24 h later. W e found that p33ING2 did not 2  significantly increase G A D D 4 5 expression in both non-UV- and UV-irradiated cells (Fig. 4.4B).  To further study the pathways involved in p33ING2-mediated D N A  84  repair, we examined whether p33ING2 regulates the expression of X P A and X P B , which are part of the core complexes involved in N E R . Our results revealed that p33ING2 significantly upregulated X P A expression.  Under  non-UV-irradiation  conditions, the expression level of X P A in cells overexpressing p33ING2 was 2-fold higher than that in the vector control (P<0.01, t-test); whereas after UV irradiation, cells overexpressing p33ING2 showed 5-fold induction of X P A compared to 1.4-fold in the vector control (P<0.001, t-test) (Fig. 4.4B).  However, p33ING2 did not  upregulate the expression levels of X P B . Since p33ING1b has been shown to bind to G A D D 4 5 (Cheung er al., 2001), we investigated to see if p33ING2 physically binds to G A D D 4 5 , X P A , or X P B .  W e performed immunoprecipitation with an anti-FLAG  mouse monoclonal antibody and found that p33ING2 did not physically interact with G A D D 4 5 , X P A or X P B (Fig. 4.4C). These results suggest that X P A may be a crucial component in p33ING2-mediated repair of UV-damaged DNA.  4.3  Discussion  Previous studies have shown that p33ING2 is involved in the regulation of gene transcription, cell cycle progression, and apoptosis in various cancer cell lines including colorectal carcinoma, hepatocellular carcinoma and human fibrosarcoma cells, and that it requires the presence of wt p53 to exert these tumour suppressive functions (Nagashima et al., 2001; Kataoka et al, 2003; Gozani et al., 2003). In this study, we for the first time demonstrated that overexpression of p33ING2 enhances the repair of UV-damaged DNA, and that this enhancement requires the participation of functional p53 (Fig. 4.1 and 4.3). W e also showed that p33ING2 participates  85  Figure 4.4 p33ING2 enhances p53 acetylation and upregulates XPA. (A) Western blot analysis of acetylated p53 levels in M M R U cells transfected with p3XFLAG-vector (V) or p3XFLAG-//VG2 (ING2). Cells were irradiated with U V B at 0 or 200 J / m and harvested 2, 4, 8 and 12 h after UV irradiation. Antibodies against p33ING2, acetylated p53 at Lys382, and total p53 were used. Anti-p33ING2 antibody detected both endogenous (lower bands) and transfected p3XFLAG-/A/G2 (upper bands). M M R U cells treated with 0.2 |j,g/ml of doxorubicin for 8 h were used as a positive control (C). (B) Western blot analysis of GADD45, X P A and X P B expression levels in M M R U cells transfected with p3XFLAG-vector (V) or p 3 X F L A G ING2 (ING2) 24 h after U V B irradiation at 0 or 200 J / m . Antibodies against p33ING2, GADD45, X P A and X P B were used. /5-actin was used as a loading control. * indicates non-specific bands. ( C ) Densitometry of X P A induction in (B). (D) C o immunoprecipitation (IP) of p33ING2 with GADD45, X P A and X P B . Total cell lysates of M M R U transfected with p3XFLAG-/A/G2 were immunoprecipitated with a control antibody (Lane 2) or with anti-FLAG antibody (Lane 3). Ten percent of lysate input for immunoprecipitation (Lane 1) was used as a control and the supernatant of p33ING2 IP (Lane 4) was used to verify the IP. Antibodies against p33ING2, G A D D 4 5 , X P A and X P B were used in Western blot analysis. 2  2  200J 0J C  V  ING2 ^^HHg|j^  2h V  ING2 ^HH^^^^  4h V  ING2 ^mjujMjjjSjk^  8h V  12h  ING2 ^^^^^^  V  ING2 ING2 Acp53 p53 Actin  86  200J  OJ  87  ING2 GADD45 XPA XPB  88  indirectly in the repair of UV-damaged DNA as p33ING2 does not colocalize with either of the two major UV-induced photolesions, C P D s and 6-4PPs (Fig. 4.2). These results suggest that p33ING2 is not directly recruited to the DNA damage sites with other repair core complexes required for damage recognition ( X P C - h H R 2 3 B , X P E - D D B 1 and X P A - R P A ) , excision of the lesions (TFIIH/XPB-XPD, X P G and X P F E R C C 1 ) , gap-filling DNA synthesis and ligation (DNA polymerase 5 or e, D N A ligase I) (Wang et al., 2003; Mitchell et al., 2003). Although p53 was found to bind X P B and X P D in vitro (Wang ef al., 1995), this association was not validated in vivo and p53 too did not colocalize to the DNA damage sites in normal human fibroblasts following UV irradiation (Fitch et al., 2003). The p53-dependence of p33ING2 enhancement of UV-damaged DNA repair suggests that p33ING2 may enhance N E R by regulating the p53 D N A repair pathway. A few lines of evidence indicate that p53 is required for efficient N E R of UVinduced DNA lesions and chemical-induced bulky DNA adducts. Thus, it plays an important role in protecting cells from tumourigenesis (Rubbi and Milner, 2003). p53 has been shown to bind and modulate the activities of the NER-associated helicases X P B and X P D (Wang ef al., 1995). Furthermore, p53 can sustain the expression of the p48 component of the DDB2 complexes and XPC gene, thus participating at least indirectly in N E R (Hwang ef al., 1999; Adimoolam ef al., 2002). More recently, it has been reported that p53 acts as a chromatin accessibility factor, mediating UVinduced global chromatin relaxation by recruiting p300 to the sites of N E R and inducing histone acetylation (Rubbi and Milner, 2003). However, the exact molecular mechanism by which p53 enhances N E R is yet to be determined.  89  W e found that overexpression of p33ING2 induces p53 acetylation  in  melanoma cells after UV irradiation (Fig. 4.4A). This is in agreement with the finding that p53 acetylation at Lys382 in the normal lymphoblastoid C 3 A B R cell line after exposure to DNA-damaging agents, etoposide or neocarzinostatin, correlated with an increased expression level of p33ING2 (Nagashima et al., 2001).  Therefore, it is  reasonable to conclude that p33ING2 enhances p53 acetylation, at least at residue Lys382, in response to DNA damage.  It is well known that post-translational  modifications such as acetylation and phosphorylation of p53 play a crucial role in the stabilization and activation of p53 as well as in protein-protein interactions (Xu, 2003). Several studies have shown that p53 acetylation within the C-terminal  region  regulates p53 stability by preventing ubiquitination by mdm-2 (Xu, 2003). There is also evidence to suggest that acetylation of p53 within the C-terminal region results in a conformational change that stimulates p53 sequence-specific DNA-binding activity (Gu and Roeder, 1997; Sakagushi era/., 1998), and recruits CBP/p300 transcription co-activators  to  p53-dependent  transcriptional activity of p53.  promoters  (Xu,  2003),  thus  activating  the  Our observations support the notion that p33ING2  enhances p53 acetylation, thus increasing the stability of p53 protein, as there was more p53 accumulated in cells overexpressing p33ING2 compared to the vector control up to 4 h after UV irradiation (Fig. 4.4A). The increased p53 acetylation by p33ING2 may account for the p53-dependence of p33ING2 enhancement in N E R . The p33ING2 enhancement in N E R may also be achieved in part by upregulating the expression of the repair recognition protein X P A . W e demonstrated that p33ING2 upregulates the expression of the XPA gene, but not the XPB gene, in  90  both UV- and non-UV-irradiated cells, with more pronounced upregulation of XPA after UV irradiation (Fig. 4.4B). Since DNA damage recognition is the rate-limiting step in repair and X P A is a component of the X P A - R P A DNA damage recognition complex (Thoma et al., 2003), upregulation of X P A by p33ING2 may significantly enhance the DNA repair process. Previously, we found that p33ING1b physically interacts with G A D D 4 5 but does not transcriptionally regulate GADD45 (Cheung et al., 2001). In this study, we found that p33ING2 neither physically interacts with nor upregulates the expression of G A D D 4 5 .  These findings may reflect the distinct  molecular pathways in which p33ING1b and p33ING2 are involved in mediating DNA repair.  In summary, we demonstrate that p33ING2 enhances the repair of UV-  damaged DNA in a p53-dependent manner. It appears that p33ING2 is not directly recruited to UV-induced DNA damage sites. The p33ING2-mediated enhancement in N E R is mostly likely achieved by increasing p53 acetylation and upregulating the expression of X P A .  91  CHAPTER 5. GENERAL CONCLUSIONS  5.1  Summary  p33ING2 has been shown to share many biological functions with its homologue p33ING1b in regulating cell cycle, apoptosis and gene transcription. These functions have been shown to require the presence of functional p53. Despite their functional similarities, p33ING1b and p33ING2 modulate the activity of p53 via distinct modes of action. While p33ING1b has been shown to physically associate with p53, p33ING2 has been found to interact with phosphoinositides and induce p53 acetylation in response to cellular stress. Several studies have provided evidence that p33ING1b is involved in UV-induced responses.  However, study of the role of p33ING2 in  cellular stress response to UV irradiation is lacking. In this thesis, we have sought to investigate the role of p33ING2 in cellular stress responses to U V irradiation and the molecular mechanisms involved. W e first investigated to see if p33ING2 is involved in UVB-induced apoptosis, using  melanoma  cells as an experimental  model  system.  W e found  that  overexpression of p33ING2 enhanced UVB-induced apoptosis. W e also found that the p33ING2 enhancement of UVB-induced apoptosis required the presence of wildtype  p53.  Moreover,  we  have  demonstrated  that  p33ING2  significantly  downregulated Bcl-2 expression and increased the Bax/Bcl-2 ratio, promoted Bax translocation to mitochondria, altered mitochondrial membrane potential, induced cytochrome c release and activation of caspase 9, therefore enhancing UVB-induced apoptosis.  It is interesting, however, under non-stressed conditions, p33ING2  92  upregulated Fas expression and activated caspase 8 to induce apoptosis. Based on our findings, we therefore concluded that p33ING2 cooperates with p53 to regulate apoptosis  via  activation  of  both  the  mitochondrial/intrinsic  and  death-  receptor/extrinsic apoptotic pathways. Despite the distinct molecular mechanisms involved, similarities in the role of p33ING2 and p33ING1b in UVB-induced apoptosis prompted us to whether ING2 plays a role in the repair of UV-damaged DNA.  investigate  W e found that  overexpression of p33ING2 enhanced the repair of UV-damaged DNA, and that p53 was required for this process in melanoma cells. W e also found that p33ING2 was not  directly  recruited  to  UV-induced DNA damage sites.  Nevertheless, we  demonstrated that overexpression of p33ING2 enhanced p53 acetylation and the expression of X P A , with more profound enhancement after U V B irradiation.  These  observations suggest that p33ING2 enhances the repair of UV-damaged DNA, and that this enhancement is most likely achieved by increasing p53 acetylation and upregulating the expression of X P A . Taken together, we have elucidated in this thesis the novel functions of p33ING2 in cellular stress responses to UV irradiation and the significance of this gene in the context of tumour suppression. With further exploration, our knowledge of this gene may lead to new insights into more effective prevention and treatment regimes for human cutaneous melanoma.  93  5.2  Future Directions  Activation of the Fas death-receptor pathway leads to cleavage and activation of Bid by caspase 8, promoting oligomerization of Bax and Bak, resulting in cytochrome c release (Wei et al., 2000; Luo et al., 1998), thus providing a link between the deathreceptor and the mitochondrial pathway. Furthermore, Bid is a transcriptional target of p53 (Sax et al., 2002). It would be interesting to see if Bid is upregulated and/or activated in p33ING2-mediated apoptosis and if cross-talk between these two apoptotic pathways is involved.  Recent studies reported that S m a c (second  mitochondria-derived activator of caspase) is one of the factors released from the mitochondria during apoptosis (Martinou and Green, 2001; van Loo et al., 2002). S m a c promotes apoptosis by releasing caspase 3 from the inhibition by inhibitors of apoptosis (lAPs) and inducing cleavage of caspase 3 (Wu et al., 2000; Chai et al., 2000).  Furthermore, Smac peptides were shown to sensitize lymphoma cells to  TRAIL-induced apoptosis (Gua et al., 2002). It would be of interest to determine if Smac has a role in p33ING2-mediated apoptosis. In this thesis, we clearly show that p33ING2 upregulates the expression of XPA.  It is crucial to determine if p33ING2 transcriptionally upregulates X P A or  stabilizes the X P A protein, and if induction of X P A by p33ING2 is p53-dependent. It is reported that p53 recruits p300 to N E R sites and induces histone acetylation to relax the chromatin to allow the accessibility of other repair proteins (Rubbi and Milner, 2003). Since p33ING2 enhancement of N E R is p53-dependent and p33ING2 is not recruited to N E R sites, it will be interesting to determine if p33ING2 is required  94  to recruit p300 to the N E R sites and cooperates with p53 in acetylating histones upon UV irradiation.  95  REFERENCES  Aasland R, Gibson T J , Stewart A F . The P H D finger: implications for chromatinmediated transcriptional regulation. Trends Biol Sci 1995, 20: 56-9. Adams J M , Cory S. The Bcl-2 protein family: arbiters of cell survival. Science 1998, 281:1322-6. Adimoolam S , Ford J M . p53 and DNA inducible-expression of the xeroderma pigmentosum group C gene. 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