THE FUNCTIONAL SIGNIFICANCE OF AN

ALTERNATELY SPLICED PRODUCT OF THE HDM2 GENE

A dissertation presented to

the faculty of

the College of Arts and Sciences of Ohio University

In partial fulfillment

of the requirements for the degree of

Doctor of Philosophy

Martin J. Schmerr

March 2007

This dissertation entitled

THE FUNCTIONAL SIGNIFICANCE OF AN

ALTERNATELY SPLICED PRODUCT OF THE HDM2 GENE

by

MARTIN J. SCHMERR

has been approved for

the Department of Biological Sciences

and the College of Arts and Sciences by

Susan C. Evans

Assistant Professor of Chemistry and Biochemistry

Benjamin M. Ogles

Dean, College of Arts and Sciences

SCHMERR, MARTIN J., Ph.D., March 2007, Molecular and Cellular Biology

THE FUNCTIONAL SIGNIFICANCE OF AN ALTERNATELY SPLICED PRODUCT

OF THE HDM2 GENE (192 pp.)

Director of Dissertation: Susan C. Evans

MDM2, the primary protein product of the mdm2 gene, negatively regulates the p53 tumor suppressor protein to allow normal cellular growth and division. Intriguingly, the mdm2 gene generates many other alternatively spliced transcripts, yet their function and whether they are translated remains largely unknown. While most of these transcripts or their potential translation products are most likely non-functional or they may promote tumorigenesis, several reports have indicated that some possess physiological functions.

In this study, the hdm2ALT1 splice variant was transiently expressed in the NIH 3T3 cell

line. The HDM2ALT1 protein was predicted to induce growth inhibition by blocking

MDM2’s activity and stabilizing p53. Protein analysis via immunoprecipitation and

immunoblotting demonstrated that HDM2ALT1 interacted with MDM2, disrupted

MDM2’s interaction with p53, and increased the levels of p53 protein. In addition,

HDM2ALT1 expression elevated the protein levels of the proapoptotic gene, bax. The

presence of HDM2ALT1 also stimulated p53’s transcriptional activity in the absence of any stress. HDM2ALT1 sequestered MDM2 in the cytoplasm and facilitated the entry of p53 into the nucleus. HDM2ALT1 expression reduced the proliferation rate of NIH 3T3

cells, which flow cytometric cell cycle analysis and cell death assays confirmed was a

result of apoptosis. These findings strongly indicate that expression of the HDM2ALT1 protein represents a novel mechanism by which the p53-HDM2 interaction is disrupted.

Approved:

Susan C. Evans

Assistant Professor of Chemistry and Biochemistry

Acknowledgements

I would like to extend my genuine appreciation to the many individuals who assisted in my research endeavors. My advisor, Dr. Susan Evans, permitted me a great deal of latitude to explore and implement my own ideas throughout the duration of this venture. I wish to thank my committee members, Drs. Soichi Tanda, Jennifer Hines and

Donald Holzschu, for their guidance throughout my research. I am very grateful of the massive effort put forth by Edward Frebault, Tammy Mace and Eric Johnson in the care of my transgenic animals. I am forever indebted to current and past graduate students in the lab: Dr. Min Liang, Dr. Radhika Iyer, Swapna Vemula, Yan Liu, and especially Drs.

Chrisanne Dias and Bernard Ayanga. I am exceedingly thankful to all of the undergraduate students, including Sara Maxfield, Dan Gorbett, Andrew Dittenhofer, Josh

Harbert, Sarah Smith, Susan Spotts, Steven Powell, and Nicole Stadelman, who assisted with numerous experiments throughout this project. I would like to thank the following colleagues and friends for their insightful and stimulating conversations: Tom Radzio,

Drs. Liang Huang, Kelly McCall, Leonard Kohn, Dawn Holliday and Lorie Lapierre. I greatly appreciate the endless encouragement and constant scientific discourse my family members offered. I especially wish to thank my mother, Dr. Mary Jo Schmerr, for sharing her expertise with immunoblotting and immunofluorescence. I will continually strive to emulate the standards and ethics of scientific investigation both her and my father,

Professor Lester Schmerr, have practiced for over 30 years. Most of all, I am eternally beholden to my benevolent wife, Lynn Kneile. There is no word in any language that properly conveys my deepest heartfelt appreciation for her patience and fortitude to endure these long years with me so that I may accomplish one of my dreams.

vi

Table of Contents Page

Abstract...... iii

Acknowledgements ...... v

List of Tables ...... ix

List of Figures...... x

Chapter 1: Introduction ...... 1

1.1 A dilemma of multicellularity...... 2

1.2 Underlying causes of cancer ...... 3

1.3 Loss of p53 function in tumors ...... 4

1.4 Protective mechanisms of the p53 network ...... 5

1.4.1 Activation of the p53 circuit ...... 7

1.4.2 Downstream effectors of the p53 circuit...... 10

1.5 Control of p53 activity...... 15

1.6 The p53-HDM2 negative feedback loop...... 18

1.7 Oscillations in the p53-HDM2 feedback loop ...... 20

1.8 The hdm2/mdm2 gene...... 21

1.8.1 Genomic organization...... 23

1.8.2 Transcriptional regulation...... 23

1.8.3 Post-transcriptional regulation...... 26

1.8.4 Post-translational regulation ...... 31

1.8.5 p53-independent functions...... 33

1.9 Contradictory effects of HDM2ALT1 expression ...... 35 vii

1.10 Specific aims of the present study ...... 39

1.11 HDM2 and p53 as therapeutic targets in tumors ...... 39

Chapter 2. Materials and Methods...... 45

2.1 The Tet-On system...... 46

2.2 Constructs ...... 48

2.3 Generation of transgenic mice ...... 50

2.4 Tail DNA isolation...... 50

2.5 Genotyping...... 51

2.6 Cell culture conditions ...... 53

2.7 Transient transfection...... 53

2.8 Stable transfection...... 53

2.9 Antibodies...... 54

2.10 Immunofluorescence...... 56

2.11 Western blot analysis ...... 56

2.12 Immunoprecipitation...... 57

2.13 TUNEL ...... 58

2.14 Cell death detection assay...... 58

2.15 Cell viability assay...... 59

2.16 Flow cytometry ...... 60

2.17 Luciferase assay...... 60

Chapter 3. Results...... 61

3.1 Identification of transgenic founders ...... 62

3.2 Expression of the transgenes in murine tissues...... 64 viii

3.3 Establishment of transgenic lines...... 65

3.4 Development of a stable NIH 3T3 cell line with the Tet-On system...... 66

3.5 HDM2ALT1 expression does not affect the levels of MDM2...... 69

3.6 HDM2ALT1 interacts with MDM2 ...... 71

3.7 HDM2ALT1 expression decreases the p53-MDM2 interaction...... 73

3.8 HDM2ALT1 expression increases the levels of p53 and Bax, but not

p21WAF1/CIP1 ...... 75

3.9 HDM2ALT1 stimulates the transcriptional activity of p53 ...... 77

3.10 The subcellular localization of HDM2ALT1, MDM2 and p53 ...... 79

3.11 MDM2 and HDM2ALT co-localize in the cytoplasm...... 88

3.12 The presence of HDM2ALT decreases the proliferation rate of NIH 3T3

cells ...... 90

3.13 HDM2ALT1 expression triggers apoptosis ...... 96

Chapter 4. Discussion ...... 104

Chapter 5. Future Studies ...... 115

References...... 121

Appendix: List of Abbreviations ...... 173

ix

List of Tables

Page

Table1. Upstream stress mediators and their respective post-translational

modifications of the p53 protein...... 8

Table 2. p53 target genes and their functions ...... 11

Table 3. Forward and reverse primers used to genotype transgenic mice...... 52

Table 4. Description of the antibodies used to detect MDM2, HDM2ALT1, and p53 ...... 55

Table 5. Summary of the transgenic founder mice...... 63

Table 6. Cell cycle analysis of HDM2ALT1 and mock transfected cells...... 95

x

List of Figures

Page

Figure 1. Signaling within the p53 network...... 6

Figure 2. Schematic representation of the domains of the HDM2 protein ...... 17

Figure 3. Schematic representation of the domains of the hdm2 and mdm2 splice

variants...... 27

Figure 4. Proposed model of HDM2ALT1’s function...... 38

Figure 5. The Tet-On system ...... 47

Figure 6. Map of the constructs containing hdm2ALT1 inserts ...... 49

Figure 7. Inducible control of luciferase activity in stable cell lines...... 67

Figure 8. Western blot analysis of MDM2 and HDM2ALT1 protein in transfected

cells ...... 70

Figure 9. Interaction of HDM2ALT1 with MDM2...... 72

Figure 10. HDM2ALT1 disrupts the interaction of p53 and MDM2 ...... 74

Figure 11. Western blot analysis of p53 and Bax protein in transfected cells...... 76

Figure 12. Transcriptional activity of p53 in transfected cells ...... 78

Figure 13A. Subcellular localization of HDM2ALT1, MDM2, and p53 at 24 hours

after mock transfection ...... 80

Figure 13B. Subcellular localization of HDM2ALT1, MDM2, and p53 at 48 hours

after mock transfection ...... 81

Figure 13C. Subcellular localization of HDM2ALT1, MDM2, and p53 at 72 hours

after mock transfection ...... 82 xi

Figure 13D. Subcellular localization of HDM2ALT1, MDM2, and p53 at 96 hours

after mock transfection ...... 83

Figure 14A. Subcellular localization of HDM2ALT1, MDM2, and p53 at 24 hours

after transfection with HDM2ALT1 ...... 84

Figure 14B. Subcellular localization of HDM2ALT1, MDM2, and p53 at 48 hours

after transfection with HDM2ALT1 ...... 85

Figure 14C. Subcellular localization of HDM2ALT1, MDM2, and p53 at 72 hours

after transfection with HDM2ALT1 ...... 86

Figure 14D. Subcellular localization of HDM2ALT1, MDM2, and p53 at 96 hours

after transfection with HDM2ALT1 ...... 87

Figure 15. HDM2ALT1 co-localizes with MDM2 in the cytoplasm...... 89

Figure 16. HDM2ALT1 expression decreases the growth rate of NIH 3T3 cells...... 91

Figure 17. HDM2ALT1 expression decreases the viability of NIH 3T3 cells...... 94

Figure 18. Levels of apoptosis in transfected cells...... 97

Figure 19A. TUNEL staining of fragmented DNA at 24 hours post-transfection...... 98

Figure 19B. TUNEL staining of fragmented DNA at 36 hours post-transfection...... 99

Figure 19C. TUNEL staining of fragmented DNA at 48 hours post-transfection...... 100

Figure 19D. TUNEL staining of fragmented DNA at 60 hours post-transfection...... 101

Figure 19E. TUNEL staining of fragmented DNA at 72 hours post-transfection...... 102

Figure 19F. TUNEL staining of fragmented DNA at 96 hours post-transfection ...... 103

1

Chapter 1: Introduction

2 1.1 A dilemma of multicellularity

The appearance of multicellular organisms represents a significant milestone in

biological organization. Key attributes of multicellularity, such as specialization,

programmed cell death, stigmergy (Anderson 2002; Holland and Melhuish 1999), and

dynamic intercellular communication, provide these organisms with selective advantages

(e.g., division of labor, increased size, and emergent behaviors and properties) over their

unicellular counterparts. Many complex systems are responsible for the development and homeostasis of multicellular organisms. One such example is the elaborate pattern of

cellular proliferation and cellular suicide that generates and maintains the adult human body; this structure consists of an estimated 1015 cells that are derived from a single cell

(Bertram 2000). In spite of their sophistication, metazoans often experience a widespread

problem observed in complexity—out of control. A common manifestation of this chaotic

phenomenon is uncontrolled cellular proliferation. Because the onset of neoplasms

threatens the overall survival of the host, specialized regulatory systems, such as the cell

division cycle, have evolved to precisely coordinate every cell’s decision to live, die or

multiply. These processes balance the rates of cellular proliferation and death to curtail

renegade cell growth. As these systems are also complex in regulation, their failure can

lead to diseases, such as cancer. For instance, several DNA tumor viruses utilize this flaw

for their survival; they cripple the factors restricting cell division in order to promote

rampant proliferation—an event that increases the risk of developing certain cancers

(O'Shea 2005). As with any complex system, dissection of the individual constituents is

crucial to understanding the entire process and deciphering which components of the

growth-regulatory machinery that malfunction and lead to cancer is no exception. 3 1.2 Underlying causes of cancer

Tumorigenesis is thought to require the acquisition of certain genetic mutations or

alterations in the cancer-associated genes that regulate cellular proliferation and

programmed cell death. Dysfunction of these genes may provide the affected cell with a

selective growth advantage over its normal cellular counterparts (Bertram 2000); the unrestricted and continuous clonal expansion of this abnormal cell facilitates neoplasia.

Because cancer cells incessantly divide and cannot be eliminated by programmed cell

death, they destroy the highly ordered architecture of the surrounding normal tissues and

perturb their physiological functions, ultimately leading to death of the host.

Despite the gamut of abnormal genetic and epigenetic events detected in tumors,

the cellular transformation process is thought to almost always involve genetic alterations

that affect the expression or function of two important classes of genes: proto-oncogenes

and tumor suppressor genes (Cavenee and White 1995; Fearon and Vogelstein 1990;

Weinberg 1996). Malfunction of these genes may allow a cell to evade the homeostatic

mechanisms governing cellular division and suicide. Consequently, the affected cell

acquires a selfish, unregulated proliferative capability (e.g., evasion of cell death

pathways, increased growth rate, escape from replicative senescence, decreased

dependency on exogenous growth factors, diminished intercellular communication,

invasion into the surrounding tissues, and loss of differentiation) (Compagni and

Christofori 2000). Thus, understanding how the complex array of proto-oncogenes and

tumor suppressor genes govern cellular proliferation and programmed cell death is

critical to assess how their dysfunction may contribute to the initiation, maintenance, or

progression of cancer (Pieretti et al. 1995; Simpson and Camargo 1998). 4 1.3 Loss of p53 function in tumors

Of all the tumor suppressor genes and proto-oncogenes disrupted in the cellular

transformation process, dysfunction of the p53 tumor suppressor alone greatly increases

the likelihood of developing cancer. For example, p53 germline mutations, which are

found in Li-Fraumeni syndrome patients and Trp53tm1Tyj mutant mice, cause numerous and many types of tumors to develop early in life (Donehower 1996b; Malkin et al.

1990). In contrast, constitutive activation of p53 in transgenic mice enhanced their resistance to spontaneous tumor formation (Garcia-Cao et al. 2002; Tyner et al. 2002).

Strikingly, approximately 50% of all human cancers possess deleterious mutations in the

p53 gene (Hollstein et al. 1991; Levine et al. 1991) although other non-mutational

mechanisms are known to perturb its functions. For instance, certain viral oncoproteins,

such as the human papilloma virus-16 E6 protein, directly repress and target the p53

protein for destruction (Scheffner et al. 1990; Vousden 1995). Improper cytoplasmic

localization of p53, caused by defects in its nuclear localization pathway, abolishes p53

activity (Kim et al. 2000). Loss of p53 activity also results from the deregulation of hdm2 gene, whose protein product (HDM2) restrains p53’s anti-proliferative functions. HDM2 overexpression, which leads to total loss of endogenous p53 activity, is observed in numerous types of human cancers, predominately sarcomas and, to a lesser extent, in other tumor types (Momand et al. 1998; Oliner et al. 1992). Altogether, many human tumors are thought to possess deleterious p53 mutations or harbor other non-mutational events rendering p53 non-functional (Levine 1997; Sherr and McCormick 2002).

Because the abrogation of endogenous p53 activity is almost obligate for tumor growth, its physiological functions are of paramount importance for suppressing tumorigenesis. 5 1.4 Protective mechanisms of the p53 network

Cellular stresses (e.g., genotoxic and non-genotoxic) may cause damage that

deregulates the cell cycle and triggers cellular transformation. The p53 protein safeguards against the emergence of neoplastic cells by either arresting or eliminating damaged these cells. Through a plethora of activities, p53 mediates an adaptive response to a variety of intrinsic and extrinsic stress conditions, such as hypoxia, nutrient depletion, temperature

shock, oncogenic activation, DNA damage, chromosomal aberrations, and telomere

erosion (Figure 1) (Amundson et al. 1998; Giaccia and Kastan 1998). Depending upon

the type and duration of a particular stress, activation of the p53 pathway primarily

invokes cell cycle arrest or apoptosis (Figure 1) (Sionov et al. 1999). For example, under

conditions of low stress (e.g., minor DNA damage), p53 initiates a transient growth arrest

that allows a sufficient pause to repair stress-induced damage and restore homeostasis

(Alarcon-Vargas and Ronai 2002). On the other hand, during an extremely damaging and

sustained stress (e.g., a persistent oncogenic stimulus), p53 triggers an apoptotic cascade

to eliminate a cell potentially at risk of undergoing transformation (Alarcon-Vargas and

Ronai 2002). Aside from these primary responses of the p53 network, other less-

characterized outputs involved in tumor suppression include senescence, differentiation,

inhibition of angiogenesis, crosstalk with other cellular networks, feedback modulation of p53 activity, and intercellular communication with unstressed neighboring cells (Figure

1) (Levine et al. 2006). In addition to its well-studied tumor suppressive activities, p53

also regulates other biological processes, such as aging, development, chromosomal

segregation, and DNA recombination; however, very little is known about the

mechanisms by which p53 influences these processes (Hofseth et al. 2004). 6

p53

Figure 1. Signaling within the p53 network

Stress-induced damage (input) activates the p53 signaling network. Damage sensors detect and relay these signals to p53 through a host of upstream mediators. Upon activation, p53 mediates a response to the damage by primarily inducing the transcription of its target genes. The phenotypic outcomes (output) of these downstream effectors appropriately respond to the stress conditions (Modified from Levine et al. 2006).

7 1.4.1 Activation of the p53 circuit

In response to stress conditions, numerous sensors recognize and communicate

the nature and extent of the stress-induced damage to the p53 protein (Figure 1). These

sensors induce various signal transduction cascades, which, in turn, trigger various upstream mediators to activate p53 through a myriad of post-translational modifications

(Figure 1) (Jayaraman and Prives 1999; Levine et al. 2006; Meek 1998; Sakaguchi et al.

1998). These covalent modifications, which mainly consist of phosphorylation and acetylation, stabilize and activate the latent p53 protein (Table 1) (Brooks and Gu 2003;

Chuikov et al. 2004; Levine et al. 2006; Xu 2003). Other post-translational modifications of p53 include methylation, sumoylation (Gostissa et al. 1999; Muller et al. 2000;

Rodriguez et al. 1999), ribosylation (Wesierska-Gadek et al. 1996a; Wesierska-Gadek et al. 1996b), and glycosylation (Figure 1) (Shaw et al. 1996), yet any data indicating which residues on p53 that are modified or the functional implications of these modifications is extremely limited and inconclusive (Kwek et al. 2001; Levine et al. 2006). The types and numbers of post-translational modifications of the p53 protein vary with the different forms of stress stimuli (Figure 1 and Table 1). These unique combinations of modifications are thought to relay essential information about the stress condition to p53.

Moreover, the relevant stress response pathways that are activated by p53 may also depend upon the idiosyncratic combinations of post-translational modifications (Figure 1 and Table 1). Therefore, each post-translational modification likely acts as a molecular code for p53 to incorporate and then decipher about the status of the stressed cell (Levine et al. 2006). The extensive covalent modifications of p53 efficiently allow this protein to integrate and trigger the response to a wide range of cellular stress conditions. 8 Table 1. Upstream stress mediators and their post-translational modifications of the p53 protein (Modified from Bode et al. 2004).

Mediators Activating stress input Modification site Response Kinases ATM DNA damage Ser15 Apoptosis ATR γ radiation, UV radiation Ser15, Ser37 Apoptosis AURKA Overexpression of AURKA Ser315 Ubiquitination of p53 CDK UV radiation Ser315 ↑ p53-mediated (CDC2/CDK2) transcription CHK1/CHK2 Ionizing radiation Ser20 Disrupt p53-MDM2 complex Casein kinase 1 DNA damage and Ser6, Ser9, Thr18 Inhibition of MDM2 topoisomerase inhibitors activity CSN-associated Unstressed conditions Thr150, Thr155, p53 degradation kinase complex Ser149 DNA-PK DNA damage Ser15, Ser37 Disrupt p53-HDM2 complex ERKs UV-radiation Ser15 Apoptosis ERK2 Doxorubicin Thr55 p53 activation FACT-CK2 UV radiation Ser392 ↑ p53 activity GSK3β Endoplasmic reticulum Ser315, Ser376 Block p53-mediated stress apoptosis HIPK2 UV radiation Ser46 Cell cycle arrest and apoptosis JNK UV radiation Ser20 Apoptosis JNK DNA damage Thr81 p53 stabilization MAPKAPK2 UV radiation Ser20 Apoptosis p38 kinase UV radiation Ser15, Ser33, Ser46 p53 stabilization and apoptosis p38 kinase UV radiation and DNA Ser392 ↑ p53’s DNA damage binding activity PKC Unstressed conditions Ser372, Ser378 Ubiquitination of p53 and ↑ p53’s DNA binding activity PKR Interferon Ser392 Unknown

TAFI Constitutively Thr55 Degradation and phosphorylated stabilization of p53 9 Table 1. (Continued). Mediators Activating stress input Modification site Response Acetyltransferases p300/CBP Unknown Lys370, Lys372, ↑ p53-mediated Lys373, Lys381, transcription, Lys382, Lys386 cell cycle arrest p300/PCAF Unknown Lys320 ↑ p53-mediated transcription, apoptosis p300 Unknown Lys305 ↑ p53-mediated transcription Deacetyltransferases mSin3A/HDAC Hypoxia Unknown Repress p53- mediated transcription PID/HDAC Unknown Unknown Repress p53- mediated transcription Sir2/HDAC Unknown Unknown Repress p53- mediated transcription HDAC1 Unknown Unknown Repress p53- mediated transcription Sumoylase SUMO1 UV radiation Lys386 ↑ p53-mediated transcription Neddylase MDM2 Unknown Lys370, Lys372, Unknown Lys373, Lys381, Lys382, Lys386 Ubiquitin ligases MDM2 Unstressed conditions Lys370, Lys372, Ubiquitination Lys373, Lys381, of p53 Lys382, Lys386 COP1 Unstressed conditions Unknown Ubiquitination of p53 Pirh1 Unstressed conditions Unknown Ubiquitination of p53 JNK Unstressed conditions Unknown Ubiquitination of p53 10 1.4.2 Downstream effectors of the p53 circuit

p53 mainly regulates the stress-response pathways via its function as a potent

transcription factor. Once activated, p53 mediates an appropriate response to the stress

conditions by primarily transactivating or repressing target downstream genes (el-Deiry

1998; Hupp et al. 1992). Not surprisingly, most mutations that perturb p53 function in

tumors are found within its DNA binding domain and thus attenuate p53’s transcriptional

activity (Lane and Lain 2002; Soussi and Beroud 2001; Soussi et al. 1994; Soussi and

Lozano 2005). cDNA microarray analysis of p53-regulated gene expression indicated that the p53 protein may bind to the promoter elements of at least 400 different genes (Table

2) (Zhao et al. 2000). Differential transactivation or repression of these p53-reponsive genes is controlled by many regulatory factors: sequence variation and location of the p53-response DNA elements in target genes (el-Deiry 1998; Resnick-Silverman et al.

1998), levels of nuclear p53 protein (Chen et al. 1996a; Oren 1999; Ronen et al. 1996), and post-translational modifications that either directly influence its DNA-binding specificity (Appella and Anderson 2001; Ashcroft et al. 2000; Bulavin et al. 1999; Chao et al. 2000; Dumaz and Meek 1999) or regulate p53’s association with other components of the transcriptional machinery (Adler et al. 1997; Ashcroft et al. 2000; Fogal et al.

2000; Hirao 2000; Prives and Hall 1999; Samuels-Lev et al. 2001). For instance,

acetylation of residue Lys320 only allows p53 to transactivate cycle arrest genes, such as

p21WAF1/CIP1, while acetylation of residue Lys373 directs p53 to interact with the

promoters of pro-apoptotic genes, such as bax (Knights et al. 2006). Differential transcriptional control by p53 is vital to determine which of the various cellular processes is elicited in response to a specific type of cellular stress. 11 Table 2. p53 target genes and their functions (Modified from Bode et al. 2004). Gene Function Cell cycle inhibition p21WAF1/ CIP1 Cyclin-dependent kinase inhibitor

14-3-3σ Tethers cyclin B1-CDK1 in the cytoplasm; G2 cycle arrest

GADD45β Growth arrest and apoptosis; activation of stress responsive MTK/MEKK4/MAPKKK BTG2/TIS21 General transduction; post-translational modifications; anti- proliferative protein; transcription regulation through ESR1 MIC-1 Growth factor inhibitor

IGFBP3 Growth factor inhibitor

MDM2 Inhibits p53 and p73-mediated cell cycle arrest; apoptosis; E3 ubiquitin ligase TGF-α Growth factor inhibitor

Cyclin B1 Cell cycle control (G2/M transition)

Cyclin D1 Cell cycle control (G1/S transition)

Cyclin E Cell cycle control (G1/S, start, transition)

Cyclin B2 Cell cycle control (G2/M transition)

Cyclin D2 Cell cycle control (G1/S transition)

Cyclin A Cell cycle control (G2/M transition)

CDK4 Cell cycle control

CDK2 Cell cycle control; interacts with cyclins A, B3, D, or E

DCK1 Calcium-signaling neuronal migration TOPO-IIα Topological states of DNA by transient breakage and subsequent rejoining of DNA strands

PLK Cell division and G1 or S phase PISSLRE Cdc2-related protein kinase

BRCA-1 DNA repair and E2-dependent ubiquitination

Cdc6 DNA replication GOS2 Possibly involved in translation 12 Table 2. (Continued). Gene Function Apoptosis Bax Programmed cell death

Apaf-1 Activation of pro-caspase-9

PUMA Mitochondrial cytochrome c release

P53AIP1 Loss of ΔΨm and apoptosis

PIDD Mitochondrial cytochrome c release

NOXA Mitochondrial cytochrome c release KILLER/DR5 Receptor for the cytotoxic ligand TNFSF10/TRAIL; pro-apoptotic

Bid Pro-apoptotic TRAF4 Activation of NF-κB and JNK

Fas/Apo1 Receptor for TNFSF/FASL

BAK Binding and antagonizing the repressor Bcl-2 or E1B protein

Bcl-6 Transcriptional regulator with an important role in lymphogenesis

Wig1 p53-dependent growth regulatory pathway

Caspase 2 Activation cascade of caspases for apoptosis Caspase 3 Apoptosis execution

Caspase 9 Binding to Apaf-1 leads to activation of the protease cascade

MIHB Apoptosis inhibitor MIHC Movement of organelles along actin filaments PDCD2 DNA-binding protein with a regulatory function Cell-cell signaling EDN2 Endothelins are endothelium-derived vasoconstrictor peptides Protocadherin-1 Cell to cell adhesion and communication; extracellular matrix Angiogenesis and metastasis inhibitors Mapsin Serine protease inhibitor KAI1 Metastasis suppressor protein 13 Table 2. (Continued). Gene Function Oxidative Stress PIG3 Oxidative stress Transcription ATF3 Repress transcription Protein Metabolism

LOXL1 Lysyl oxidase family Signal transduction DUSP5 Displays phosphatase activity DGKA Converts diacylglycerol into phosphatidate and alters PKC activity

CDC25C Inducer of mitotic control

DNA repair P53R2 DNA damage repair

PCNA Auxiliary protein of DNA polymerase δ; control of DNA replication

DDB2 Repair of UV-damaged DNA; binds to pyrimidine dimers

LIG1 Seals nicks in double-stranded DNA ERCC5 DNA excision repair

RPA1 Required for SV40 DNA replication in vitro XRCC9 Post-replication repair RFC4 Elongation of multiprimed DNA template Senescence Ras Transduction of mitogenic signals

Raf Transduction of mitogenic signals p19ARF Regulator of MDM2 and E2F activity MAPK Phosphorylates microtubule-associated protein-2 (MAP2), myelin basic protein and Elk-1

E2F1 Control of cell cycle progression from G1 to S phase p16 Cyclin-dependent kinase inhibitor

PML Cell proliferation and apoptosis 14 Besides its role as a sequence-specific transcription factor, p53 modulates the

DNA repair process, which includes nucleotide excision repair (NER), base excision

repair (BER), non-homologous end-joining, homologous recombination, and mismatch repair (MMR), through its other activities (Dianov et al. 2003; Friedberg 2001;

Hoeijmakers 2001; Lieber et al. 2003; Sancar et al. 2004; Sengupta and Harris 2005;

Tang and Chu 2002). For example, p53 may participate in the DNA repair process via its intrinsic biochemical properties: a 3'Æ5' exonuclease activity (Mummenbrauer et al.

1996), DNA reannealing and DNA strand transfer (Janus et al. 1999; Sturzbecher et al.

1996), rejoining DNA with double-stranded breaks (Tang et al. 1999; Yang et al. 1997) and its interaction with the recombination protein RecA (Janus et al. 1999; Sturzbecher et al. 1996) as well as other DNA repair proteins (Adimoolam and Ford 2002; Adimoolam and Ford 2003). p53 directly senses DNA damage by associating with different DNA structures: double-stranded and single-stranded DNA, DNA mismatches, Holliday junctions (Dudenhoffer et al. 1998; Janz et al. 2002; Restle et al. 2005), and DNA adducts (Bakalkin et al. 1995; Lee et al. 1997; Lee et al. 1995; Steinmeyer and Deppert

1988). In addition to its transcription-independent role in DNA repair, p53 also transactivates several genes involved in NER, BER, and MMR (Sengupta and Harris

2005). Through these multiple activities, p53 is able to integrate considerable information from its upstream mediators that are stimulated by stress signals and subsequently activate the correct downstream processes that repair the stress-induced damage. The singular, yet integral, role of p53 in the stress response allows it to function as the central node in these pathways. This centralized architecture of the stress response network also explains why the disruption of p53’s activities greatly accelerates tumorigenesis. 15 1.5 Control of p53 activity

Once the p53 circuit is activated, a cell is irrevocably committed to an anti- proliferative state. Therefore, proper regulation of p53’s activities is essential to allow normal cellular proliferation. Otherwise, uncontrolled p53 activity would cause a complete growth arrest or inappropriately induced apoptosis (McMasters et al. 1996).

Moreover, unrestricted p53 activity is lethal during murine embryogenesis (Jones et al.

1995; Montes de Oca Luna et al. 1995) and accelerates aging-associated phenotypes in vivo, such as reduced body weight, osteoporosis, lordokyphosis, decreased longevity, and lethargy (Tyner et al. 2002). Furthermore, mice possessing a hypomorphic mdm2 allele

(i.e., express ~30% of the endogenous levels of mdm2 mRNA) displayed defects in hematopoiesis and increased radiosensitivity in certain tissues due to elevated p53 activity (Mendrysa et al. 2003). Owing to its powerful growth suppressive activities, p53 is only activated in response to damage. As these conditions are mostly transient, p53 activity is tightly constrained by maintaining its protein levels at basal levels in the absence of stress-induced damage (Moll and Petrenko 2003).

Stringent control over p53 activity is mediated by the HDM2 protein. Several domains of HDM2 block p53’s transcriptional activity and promote the destruction of p53—an event necessary for cellular recovery after completion of the stress response

(Figure 2) (Ashcroft and Vousden 1999). Under normal conditions, Akt phosphorylates

MDM2 on residues Ser166 and Ser186 (Mayo and Donner 2001). Phosphorylation of these residues promotes nuclear entry of MDM2, possibly by activating its nuclear localization signal (Figure 2) (Chen et al. 1995; Mayo and Donner 2001). Once in the nucleus, the p53-binding domain of HDM2 masks the transcriptional transactivation 16 domain of p53 (Figure 2) (Chen et al. 1994; Kussie et al. 1996; Lin et al. 1994; Lu and

Levine 1995; Oliner et al. 1993; Picksley et al. 1994; Thut et al. 1997). HDM2 also recruits histone deacetylase 1 (HDAC1) to deacetylate key lysine residues of p53 that activate its transcriptional function (Ito et al. 2002). HDM2 earmarks p53 for degradation

by monoubiquitinating all of the C-terminal p53 lysine residues via the E3 ubiquitin-

conjugating ligase activity residing in its RING finger domain (Figure 2) (Haupt et al.

1997; Kubbutat et al. 1997; Lai et al. 2001; Lohrum et al. 2001). Monoubiquitination of the lysine residues reveals the NES of p53, which associates with the CRM1 nuclear export machinery (Lohrum et al. 2001). MDM2-mediated export of p53 into the cytoplasm further restricts p53’s transcriptional activities in the nucleus (Boyd et al.

2000b; Freedman and Levine 1998; Geyer et al. 2000; Roth et al. 1998; Smart et al.

1999). Polyubiquitination of the monoubiquitinated residues by the transcriptional coactivator p300/CBP, which possesses an E4 polyubiquitin-conjugating ligase activity, mediates rapid turnover of the p53 protein (Grossman et al. 2003; Grossman et al. 1998).

Because the 26S proteasomes are abundant in both the nucleus and cytoplasm (Brooks et al. 2000; Palmer et al. 1996; Reits et al. 1997), ubiquitination of p53 results in its destruction in both subcellular compartments (Joseph et al. 2003; Shirangi et al. 2002;

Yu et al. 2000). Several reports have indicated that the acidic domain of MDM2 is required for the efficient degradation of p53 (Figure 2) (Argentini et al. 2001; Kawai et al. 2003; Kubbutat et al. 1999; Zhu et al. 2001a). The central acidic domain may interact directly with the proteasomes (Xie and Varshavsky 2000) or possibly through an adaptor molecule (Kleijnen et al. 2000). This mechanism would allow MDM2 to transport ubiquitinated p53 directly to the proteasome complexes for degradation. 17

(23-108) (179-185) (190-202) (243-301) (299-328) (438-479)

Figure 2. Schematic representation of the domains of the HDM2 protein.

The HDM2 protein consists of 491 amino acids. The shaded boxes represent the location of the various domains of HDM2 (NLS, nuclear localization signal; NES, nuclear export signal; Zn, C4-type Zinc finger; RING, Ring finger). The bold numbers indicate the coding exons. The numbers in parentheses indicate the range of amino acid residues that comprise the respective domains (Modified from Evans et al. 2001).

18 1.6 The p53-HDM2 negative feedback loop

Unbalanced activity of either p53 or HDM2 has severe consequences for the cell.

Excessive p53 activity improperly promotes the destruction of normal cells (McMasters

et al. 1996) while loss of p53 function accelerates the onset of tumors (Donehower

1996b; Malkin et al. 1990). HDM2 or MDM2 overexpression promotes the formation of

soft tissue tumors in humans (Oliner et al. 1992) and mice (Jones et al. 1998),

respectively, whereas MDM2 deficiency results in embryonic lethality due to massive

p53-mediated apoptosis (Chavez-Reyes et al. 2003; Jones et al. 1995; Langheinrich et al.

2002; Montes de Oca Luna et al. 1995). Thus, proper coordination of both p53 and

HDM2 activity is essential to mediate the stress response and permit normal cellular

proliferation during homeostatic conditions.

To orchestrate a rapid response to the damage caused by stresses and eventually

restore homeostasis, the activity of p53 and HDM2 is coordinated by a common

transcriptional regulatory motif: a negative feedback loop. In the homeostatic

environment, the HDM2 protein is produced constitutively, albeit at basal levels, to

curtail p53’s growth suppressive functions (Brown et al. 1999; Freedman et al. 1999).

Hence, the p53 protein is maintained in an inactive state and is highly unstable (i.e., a short half-life of approximately 5-30 minutes) (Moll and Petrenko 2003; Reich et al.

1983). In response to the damage caused by stress conditions, several mechanisms impede HDM2 from restricting p53’s activity and stability to rapidly increase the levels of p53 protein (Alarcon-Vargas and Ronai 2002; Wu et al. 1993). Numerous post- translational modifications of both p53 and HDM2 down-regulate HDM2’s activity by disrupting the p53-HDM2 interaction, attenuating the E3 ligase activity of HDM2, and 19 altering the subcellular distribution of p53 and HDM2 (i.e., entry of p53 into the nucleus and export of HDM2 to the cytoplasm) (Appella and Anderson 2001; Bode and

Dong 2004; Buschmann et al. 2000; Canman 1998; Hirao 2000; Khosravi 1999; Zhang and Xiong 2001). For example, phosphorylation on residue Thr18 of p53 blocks its interaction with HDM2 (Lai et al. 2000; Sakaguchi et al. 1998; Schon et al. 2002; Schon et al. 2004). Interestingly, several reports have demonstrated that cellular stresses cause the levels of hdm2 or mdm2 mRNA and protein to decrease (Chandler et al. 2006; Dias et al. 2006; Momand et al. 1992; Trinh et al. 2001; Wu and Levine 1997; Zhu et al. 2002).

After HDM2’s restriction on p53 activity is overcome, subsequent activation of the p53 protein enables it to transactivate its downstream targets and respond to the cellular stress

(Freedman et al. 1999; Lane 2001; Vousden and Lu 2002). Once the damage caused by the stress is repaired, p53 decreases its own activity by inducing expression of the hdm2 gene, which is also a transcriptional target of p53 (Chen et al. 1994; Perry et al. 1993;

Picksley and Lane 1993; Wu et al. 1993). Certain regulatory mechanisms precisely control the timing of p53-induced hdm2 transcription to ensure p53 has sufficient time to activate its respective stress response effectors prior to initiating its own degradation

(Kim et al. 1999). As the amount of HDM2 protein increases due to enhanced hdm2 transcription, HDM2 ultimately reduces the amount of p53 protein to pre-stress levels

(Wu et al. 1993). Presumably, after p53 decreases to basal levels, the diminished levels of p53-mediated hdm2 transcription restores the amount of HDM2 protein to pre-stress concentrations. This auto regulatory feedback loop properly balances p53 and HDM2 activity by tightly constraining p53’s growth suppressive effects during homeostatic conditions, yet also rapidly activates the p53 protein in response to cellular stresses. 20 1.7 Oscillations in the p53-HDM2 feedback loop

The autoregulatory loop between HDM2 and p53 predicts that the amount of p53 protein steadily increases to high concentrations while the levels of HDM2 progressively decrease in response to stress-induced damage. In contrast to this proposed analog model of the p53-HDM2 feedback loop, several reports have indicated that the levels of HDM2 and p53 protein oscillate after irradiation (Collister et al. 1998; Fu et al. 1996; Ghosh et al. 2000; Lev Bar-Or et al. 2000; Ohnishi et al. 1999). Lahav et al. (2004) and Geva-

Zatorsky et al. (2006) also demonstrated that DNA damage triggered repeated pulses of

HDM2 and p53 at regular time intervals. The authors also noted a significant correlation between the number of p53 oscillations and the dosage of radiation administered (Lahav et al. 2004). Several theoretical models have attempted to explain the oscillatory behavior of p53 and HDM2 (Ciliberto et al. 2005; Geva-Zatorsky et al. 2006; Lev Bar-Or et al.

2000; Tyson 2004; Tyson 2006). For example, stress signals may generate sufficiently large fluctuations in the pre-stress, steady state levels of p53 or MDM2 protein to cross a threshold necessary for p53 activation (Tyson 2006). The number of pulses of p53 protein may selectively determine which p53-repsonsive genes undergo transcriptional activation or repression (Lahav et al. 2004; Tyson 2006). This digital behavior of the p53 oscillations might also establish a critical threshold to ensure activation of irreversible biological processes, such as apoptosis, only occur when the stress-induced damage is chronic and beyond repair (Chen et al. 1996a; Ma et al. 2006; Ronen et al. 1996; Zhao et al. 2000). As the kinetics and dynamics of these current models of p53 oscillations are highly oversimplified and do not take into account the complex regulation of hdm2 gene expression, further investigation of this intriguing phenomenon is required. 21 1.8 The hdm2/mdm2 gene

The murine double minute 2 (mdm2) gene was discovered during a differential screen of the amplified genes residing on double minutes found in a spontaneously- transformed NIH 3T3 cell line (Cahilly-Snyder et al. 1987; Fakharzadeh et al. 1991).

Subsequent reports indicated that mdm2 deregulation induced tumorigenesis. For

example, subcutaneous injection of non-transformed rodent cells that overexpress

MDM2 into athymic nude mice lead to the formation of tumors (Fakharzadeh et al.

1991). Ubiquitous or tissue-specific overexpression of MDM2 in transgenic mice also caused a higher incidence of spontaneous tumor formation (Jones et al. 1998; Lundgren et al. 1997). Moreover, elevated MDM2 activity triggers cellular transformation;

MDM2 overexpression transformed NIH 3T3 fibroblasts, immortalized primary cultures of rat embryos fibroblasts and, in cooperation with an activated ras oncogene, transformed these primary rodent cells (Fakharzadeh et al. 1991; Finlay 1993). Thus,

MDM2’s transforming potential is activated by its overexpression. The oncogenic activities of the MDM2 protein primarily result from its complete inactivation of p53’s activities (Chen et al. 1996b; Deb 2002; Haupt et al. 1996; Kondo et al. 1995; Momand et al. 1992; Oliner et al. 1992).

Overexpression of the human orthologue of mdm2 (hdm2) has also been shown to play a role in tumorigenesis. In 3889 tumor samples examined, hdm2 was amplified at an overall frequency of 7%, with a higher incidence (20-40%) observed in soft tissue tumors (Berberich et al. 1999; Leach et al. 1993; Momand et al. 1998; Oliner et al.

1992). Aside from gene amplification (Leach et al. 1993; Marchetti et al. 1995), other

mechanisms are known to cause HMD2 overexpression (McCann et al. 1995). 22 Chromosomal translocation and enhanced transcription and translation are responsible

for HDM2 overexpression in certain tumors (Berberich and Cole 1994; Bueso-Ramos et al. 1993; Landers et al. 1997; Landers et al. 1994; Momand et al. 1998; Watanabe et al. 1996). A single nucleotide polymorphism (SNP) within the promoter region of hdm2 markedly increased the levels of hdm2 transcription and caused tumors to develop at an early age (Bond et al. 2004). As the overexpression of HDM2 and

MDM2 stimulates their tumorigenic potential, they are classified as oncogenes.

Conflicting evidence from several reports has challenged the hypothesis that

hdm2 and mdm2 function exclusively as oncogenes. For example, Brown et al. (1998)

and Kubbutat et al. (1999) observed that transient expression of hdm2 cDNA in several

normal human and murine cell lines resulted in growth arrest rather than cellular

transformation. Deletion mapping of the HDM2 protein identified two growth-

inhibitory domains responsible for this anti-proliferative phenotype (Brown et al.

1998). Other reports showed that HDM2 triggered apoptosis when overexpressed in

human medullary thyroid carcinoma cells (Dilla et al. 2002; Dilla et al. 2000). Targeted

overexpression of MDM2 in the wing imaginal disc of Drosophila melanogaster

resulted in extensive apoptosis (Folberg-Blum et al. 2002). MDM2 may also have a

role in regulating differentiation (Mayo and Berberich 1996). In support of this

hypothesis, endogenous MDM2 is highly expressed in certain differentiated tissues,

including muscle, testes, brain and skin (Dazard et al. 1997; Dazard et al. 2000;

Fakharzadeh et al. 1991; Fiddler et al. 1996; Piette et al. 1997; Shvarts et al. 1997).

Altogether, these observations indicate that MDM2 is involved in growth-inhibitory

processes in addition to its well-known oncogenic activities. 23 1.8.1 Genomic organization

The structure of the mdm2 gene dictates its exquisite regulation and thus fine-

tunes the ability of MDM2 to powerfully and precisely restrict p53’s functions. Although

p53 homologous genes are widespread throughout the metazoan phylum, to date, mdm2

homologous genes have only been identified in certain higher vertebrates, such as

chickens, frogs, rats, and zebrafish (LaFleur et al. 2002; Marechal et al. 1997; Nelson et

al. 2006; Slee et al. 2004; Thisse et al. 2000; Veldhoen et al. 1999). Thus, the emergence

of MDM2-mediated control of p53 appears to be a recent evolutionary development in

the regulation of p53. The mdm2 gene consists of 25 kb on mouse chromosome 10,

region C1-C3 (Grier et al. 2002; Jones et al. 1996; Steinman and Jones 2002). The hdm2

gene approximately spans 32 kb on chromosome 12q13–14 (Bartl et al. 2003; Liang et al.

2004). Both genes consist of 11 introns and 12 exons, of which exons 3-12 encode their

respective proteins (Fakharzadeh et al. 1991; Oliner et al. 1992). The first two exons and

the 3’ end of exon 12 are untranslated regions (Brown et al. 1999; Oliner et al. 1992).

1.8.2 Transcriptional regulation

The gene expression of hdm2 and mdm2 is primarily controlled at the

transcriptional level by two promoters, P1 and P2 (Haines et al. 1994). With the exception of the 5’ untranslated regions, both promoters yield equivalent transcripts with

the same coding capacity (Haines et al. 1994). The P1 promoter produces either mdm2

transcripts that include exons 1 and 2 or hdm2 transcripts that contain only exon 1 (Juven

et al. 1993; Zauberman et al. 1995). Transcription initiated from the P1 promoter occurs 24 independent of p53 (Juven et al. 1993; Zauberman et al. 1995) while the P2 promoter

requires p53 binding for transcriptional activation (Barak et al. 1993; Wu et al. 1993).

Constitutive transcription from the P1 promoter maintains the basal levels of

HDM2 or MDM2 that restrain p53 activity (Brown et al. 1999; Freedman et al. 1999).

The DNA sequence of exon 1 is highly divergent between humans and mice, suggesting

that differences in the regulation of gene expression exist between the species (Oliner et

al. 1992). However, both human and mouse transcripts contain upstream open reading frames (uORF) in exon 1 that reduce the rate of translation (Brown et al. 1999). Due to the presence of the uORFs, these transcripts are poorly translated and only produce protein at basal levels (Brown et al. 1999). As a result of the inefficient translation of the

P1-derived transcripts, the regulation of this promoter has been greatly ignored (Barak et al. 1994; Berberich et al. 1999; Brown et al. 1999; Jones et al. 1996; Michalowski et al.

2001; Okumura et al. 2002; Phelps et al. 2003; Saucedo et al. 1999). Thus far, none of

the cis regulatory elements of the P1 promoter have been identified.

p53 decreases its own activity by inducing expression of HDM2 or MDM2 from

the internal P2 promoter (Barak et al. 1993; Juven et al. 1993; Wu et al. 1993). This

negative feedback loop restores the basal, pre-stress levels of p53 protein, thereby

allowing the cell to resume its normal pattern of growth and division (Wu et al. 1993).

Two tandem and imperfect p53-response DNA elements (PRE) within the first intron of

the hdm2 or mdm2 gene regulate the transcriptional activity of the P2 promoter (Juven

et al. 1993; Zauberman et al. 1995). p53 only interacts with the PRE of the hdm2 or

mdm2 gene when the DNA topology of the P2 promoter favors p53 binding (Kim et al.

1999). The lack of nucleosomes in the P2 promoter readily allows architectural proteins 25 that associate with p53, such as HMG1, to remodel this region into a suitable

conformation for p53 interaction (Uramoto et al. 2003; Xiao et al. 1998). This delayed induction of p53-mediated transcription ensures sufficient time for p53 to transactivate its downstream target genes that mediate the stress response before triggering it own destruction (Kim et al. 1999). Surprisingly, transcripts derived from the P2 promoter were detected in non-stressed tissues and arose independent of p53-mediated transcription (Mendrysa and Perry 2000). Additional evidence in support of a p53- independent role of the P2 promoter is the presence of other transcriptional elements within this promoter region: a thyroid hormone response element (Qi et al. 1999) and a composite AP1-ETS site (Ries et al. 2000; Shaulian et al. 1997). However, the functional significance of p53-independent regulation of the P2 promoter is unknown.

Certain cellular stresses have been shown to reduce the rate of transcription of the

mdm2 and hdm2 gene through unknown mechanisms (Freedman et al. 1999). For

instance, hypoxia leads to a down-regulation of mdm2 transcription, potentially through

activation of p38 MAPK (Zhu et al. 2002). mdm2 transcript levels are reduced in U2OS

cells after UV irradiation (Momand et al. 1992). In addition, mdm2 transcript levels

decrease after K+ depletion in cerebellar granule neurons (Trinh et al. 2001). Recently,

Dias et al. (2006) and Chandler et al. (2006) reported that the levels hdm2 and mdm2

transcripts decreased after exposure to certain genotoxic agents. The authors suggested

that alternative splicing of hdm2 or mdm2 pre-mRNA decreased the levels of the full-

length transcripts (Chandler et al. 2006; Dias et al. 2006). Therefore, other gene

expression processes (e.g., post-transcriptional, translational, or post-translational) may

contribute to the down-regulation of HDM2 activity following stress-induced damage. 26 1.8.3 Post-transcriptional regulation

RNA splicing is the most studied post-transcriptional regulatory process (e.g.,

polyadenylation, nucleocytoplasmic transport, and 5’ capping) that affects hdm2 and

mdm2 gene expression. This process is known to expand the protein diversity of the

mdm2 and hdm2 gene; over 40 different transcripts have been identified, although their

function and whether they are translated remains unknown (Figure 3) (Bartel et al. 2002).

Several mRNA species are generated via aberrant splicing at caused by cryptic splice

donor and acceptor sites within exons or introns. Alternative splicing of the hdm2 and mdm2 genes also produces many different-sized transcripts (Figure 3) (Bartel et al.

2002). As most of these alternatively and aberrantly spliced transcripts contain premature stop codons in their shifted open reading frames, it is highly unlikely that they are translated into proteins (Bartel et al. 2002). Nevertheless, Sigalas et al. (1996) demonstrated that five human splice variants (e.g., HDM2-A, -B, -C, -D, and -E) could be translated into protein. A prominent feature of most mdm2 and hdm2 splice isoforms is the lack of portions of the p53-binding domain and the acidic domain, as well as the

complete loss of the nuclear localization and export signals (Figure 3) (Bartel et al.

2002). The lack of the coding sequence for these key regulatory domains in most variant

transcripts and the resulting effect of these missing domains on the function of the

corresponding protein isoforms is not understood. Because nearly all mdm2 and hdm2

splice products have only been identified in tumors, they are thought to trigger cellular

transformation and accelerate tumor formation. However, an accumulating body of

evidence indicates that some of these alternate transcripts regulate endogenous

physiological processes (Bartel et al. 2002; Harris 2005). 27

Figure 3. Schematic representation of the domains of the hdm2 and mdm2 splice variants.

The relative size of the splice variants are shown in comparison with full-length MDM2. The shaded boxes denoted by an asterisk indicate the region most frequently deleted in these splice forms. The other shaded boxes represent the location of the various domains of MDM2 retained by the splice isoforms (Modified from Bartel et al. 2002). 28 A detailed analysis of hdm2 and mdm2 expression patterns in various tissues and tumors revealed that most splice variants were only detected in neoplasms (Bartel et al. 2002). Most alternate transcripts are likely non-functional because a loss of fidelity in

RNA splicing has been associated with cellular transformation (Caballero et al. 2001).

However, some reports indicate that several splice variants possess oncogenic functions that promote the formation and progression of tumors. In support of this hypothesis, the presence of the aberrantly spliced hdm2 transcripts in tumors has been correlated with a poorer prognostic outcome and enhanced malignancy (Bartel et al. 2001; Bueso-Ramos et al. 1995; Evdokiou et al. 2001; Lukas et al. 2001; Matsumoto et al. 1998; Pinkas et al.

1999; Sigalas et al. 1996). Yet, other groups did not observe a correlation between the presence of alternatively spliced variants and shortened overall patient survival (Bartel et al. 2001; Lukas et al. 2001). Nonetheless, five hdm2-derived alternatively spliced forms were shown to cause cellular transformation when expressed in NIH 3T3 cells (Sigalas et al. 1996). Steinman et al. (2004) also demonstrated a similar phenotype when either

HDM2-B or MDM2-B (the murine counterpart of HDM2-B) was transduced into NIH

3T3 cells. Tissue-restricted expression of MDM2-B within the brain of transgenic mice caused the formation of myeloid sarcomas and lymphomas (Steinman et al. 2004).

Injection of hematopoietic stem cells (HSCs) that ectopically expressed mdm2 splice variants, which were previously isolated from lymphomas in Eμ-myc mice (Eischen et al.

1999; Garcia et al. 2002), into lethally-irradiated syngeneic mice also accelerated lymphogenesis and increased the aggressiveness of the lymphomas (Fridman et al. 2003).

Many splice variants retain the domains of the full-length protein that stimulate proliferation independent of p53, which may explain their oncogenic activity. 29 In addition to the oncogenic activities and non-functional properties of the

aberrantly splice variants, some studies have suggested that the alternatively spliced

hdm2 and mdm2 transcripts possess physiologic functions (Harris 2005). A correlation

between the presence of alternatively spliced hdm2 splice variants and increased

expression of wild-type p53 was observed in primary cultures of human glioblastomas

and adult soft tissue sarcomas respectively (Bartel et al. 2001; Kraus 1999). Fridman et

al. (2003) indicated that expression of mdm2 splice variants lacking the p53-binding domain decreased the growth rate of oncogenically transformed MEFs after exposure to doxorubicin. Dang et al. (2002) also demonstrated that expression of several N-terminal truncated murine isoforms in NIH 3T3 cells and MEFs resulted in a p53-dependent growth inhibition; an intact RING finger domain was required for these isoforms to cause their anti-proliferative effects (Dang et al. 2002).Expression of p76MDM2, a bona fide

protein product of the mdm2 gene detected in several endogenous murine tissues,

antagonized MDM2’s ability to destabilize p53 (Perry et al. 2000). These authors noted

that p76MDM2 does not interact with MDM2 (Chen et al. 1993; Saucedo et al. 1999), but

interfered with MDM2’s activity through a dominant-negative mechanism. Evans et al.

(2001) reported a similar dominant-negative effect, except the splice variant, HDM2ALT1

(also known as HDM2-B), bound to HDM2. Transient expression of HDM2ALT1 in U2OS cells also increased the transcriptional activity of p53 (Evans et al. 2001). Moreover, the presence of hdm2ALT1 transcripts in mammary tumors was correlated with a better

prognosis and was also detected in the surrounding normal breast tissue (Lukas et al.

2001). Exposure to various genotoxic agents induced expression of hdm2ALT1 and mdm2-

b (murine homologue of hdm2ALT1) transcripts in several cell lines (Chandler et al. 2006; 30 Dias et al. 2006). These authors also showed that hdm2ALT1 and mdm2-b expression

decreased the levels of hdm2 and mdm2 transcripts, respectively, and increased p53

protein levels. These results strongly indicate that certain hdm2 and mdm2 splice variants

regulate cellular processes.

Alternative splicing of hdm2 and mdm2 transcripts also controls the rate of

translation of these transcripts. For example, inclusion of exon 1 in P1-derived

transcripts decreased their translational rate because the uORFs reduced the efficiency

of ribosome attachment to the nascent transcripts (Brown et al. 1999). Veldhoen et al.

(1999) also identified exon-α, which altered the product of translation. Inclusion of

exon-α caused a re-initiation of translation at a downstream start codon, which

truncates the N-terminus of the resulting protein (Veldhoen et al. 1999). This novel

mechanism may also be responsible for producing the p76MDM2 protein described above

(Perry et al. 2000). In all, these examples demonstrate how alternative splicing modifies

HDM2 or MDM2 activity via downstream processes, such translational control.

Another intriguing alternatively spliced product hdm2 product, hdm365, was

recently described by Bartl et al. (2003). hdm365, which consists of exons 1-5 and lacks polyadenylation, was induced by irradiation (Bartl et al. 2003). Moreover, p53-

mediated expression of hdm365 occurred prior to full-length hdm2 mRNA expression

(Bartl et al. 2003). Due to its structural similarities with small nuclear RNAs, which are key regulatory components of the spliceosome complex, and its exclusive subcellular localization at the site of the hdm2 transcription, hdm365 may regulate the rate of p53- dependent hdm2 transcription and post-transcriptional processing of transcripts, yet further investigation is required to elucidate the function of this unique transcript. 31 1.8.4 Post-translational regulation

HDM2 and MDM2 undergo post-translational modifications that modulate their

activity during stress and normal conditions. Phosphorylation has been the most

extensively studied covalent modification (Henning et al. 1997; Meek and Knippschild

2003; Zhang and Xiong 2001). As more than 20% of the amino acids in the HDM2 or

MDM2 protein are either serine or threonine residues, the potential number of sites that may be covalently modified is considerable (Meek and Knippschild 2003). Evidence from phosphopeptide mapping and mass spectrometry confirmed that numerous phosphorylation modifications of these two proteins occur (Meek and Knippschild

2003). Many of the upstream stress response mediators that covalently modify p53 also modify MDM2 or HDM2 to coordinate their activities in various cellular environments.

Stress-induced damage attenuates the activity of both MDM2 and HDM2. For

example, the pattern of MDM2 hyperphosphorylation observed under non-stressed

conditions was altered after irradiation (Blattner et al. 2002). Hypophosphorylation of

unknown residues in the acidic domain of MDM2 may disrupt the ability of MDM2 to

promote entry of ubiquitinated p53 into the proteasomes (Argentini et al. 2001; Blattner

et al. 2002; Kubbutat et al. 1999; Zhu et al. 2001a). Several stress-activated kinases,

such as ATM, DNA-PK, the cyclin A-cdk complex, and c-Abl, phosphorylate MDM2

to reduce its interaction with p53 (Freedman et al. 1997; Mayo et al. 1997; McCoy et

al. 2003; Zhang and Xiong 2001), prevent nuclear export of p53 (Mayo and Donner

2001), increase MDM2’s interaction with the p19ARF tumor suppressor (Zhang and

Prives 2001), and inhibit the E3 ligase activity of MDM2 (Goldberg et al. 2002;

Khosravi 1999; Maya et al. 2001). Buschmann et al. (2000) also demonstrated that UV- 32 induced phosphorylation of MDM2 by the p38 kinase increased MDM2’s auto-

ubiquitination. p38-mediated phosphorylation decreased sumoylation of MDM2, which resulted in self-ubiquitination and subsequent degradation of MDM2 (Buschmann et al.

2000; Fang et al. 2000; Honda et al. 1997). In contrast, sumoylation under non-stressed conditions directs the E3 ligase activity of MDM2 towards p53 (Buschmann et al.

2001). However, others have indicated that sumoylation does not control the E3 ligase activity of MDM2 (Fuchs et al. 2002). Other evidence suggests that acetylation may regulate the E3 ligase activity of MDM2 (Michael and Oren 2003; Wang et al. 2004).

In addition to the covalent modifications that restrict MDM2 or HDM2’s activity

during the stress response, other modification sites are essential for its p53-dependent and

independent functions under homeostatic conditions. Mitogen stimulation of PI3K

triggers Akt to phosphorylate MDM2, thereby allowing MDM2 to enter the nucleus and

inhibit p53 (Gottlieb et al. 2002; Mayo and Donner 2001; Mayo and Donner 2002; Zhou

et al. 2001). Akt-mediated phosphorylation of MDM2 also increased MDM2’s

association with the androgen receptor and initiated MDM2-dependent ubiquitination of

the receptor (Gaughan et al. 2005; Lin et al. 2003; Lin et al. 2002). Phosphorylation of

MDM2 by CK2 partially stimulated its E3 ligase activity to mediate p53 turnover (Guerra

et al. 1997; Guerra and Issinger 1998; Hjerrild et al. 2001; Siemer et al. 1999). In

contrast, CK2 shifted its kinase activity towards p53 during the stress response, which

may partially reduce the ubiquitination and subsequent degradation of p53 after exposure

to UV radiation (Keller et al. 2001). Altogether, the numerous post-translational

modifications of MDM2 and HDM2 described above strongly indicate that these two

proteins, like p53, are highly interconnected to many regulatory networks within the cell. 33 1.8.5 p53-independent functions

Ever-increasing information indicates that HDM2 and MDM2 alter proliferation

independent of p53. For example, the growth properties of p53-null cell lines were

altered by MDM2 overexpression (Dubs-Poterszman et al. 1995). MDM2 overexpression in p53-deficient Mv1Lu cells, a TGFβ1-sensitive mink lung epithelial cell line, blocked

the growth-inhibitory effects of TGFβ1 (Sun et al. 1998). MDM2 also displays p53-

independent phenotypes in vivo. Transgenic mice overexpressing MDM2 and lacking p53

activity, in addition to transgenic mice that overexpress MDM2 alone, developed

different tumor types in comparison with the spectrum of neoplasms observed in p53-null

mice (Donehower 1996a; Jones et al. 1998). Tumors harboring both mdm2 amplification

and p53 mutations are rare, but behave more aggressively than tumors with either

abnormal event alone (Almog and Rotter 1997; Cordon-Cardo et al. 1994; Florenes et al.

1994; Marks et al. 1996). Targeted overexpression of MDM2 in the mammary epithelium of p53-null and E2F-1/DP1-null mice uncoupled S phase from mitosis (Lundgren et al.

1997; Reinke et al. 1999). When MDM2 was overexpressed in the murine epidermis, differentiation was inhibited in a p53-independent manner (Alkhalaf et al. 1999; Ganguli and Wasylyk 2003). MDM2’s inhibition of terminal differentiation might explain the increased levels of proliferation because the undifferentiated cells could reenter the cell cycle. On the contrary, others have reported that elevated MDM2 expression is necessary for terminal differentiation (Dazard et al. 1997; Dazard et al. 2000; Fakharzadeh et al.

1991; Mayo and Berberich 1996; Piette et al. 1997; Shvarts et al. 1996; Weinberg et al.

1995) and does not increase the predisposition to tumor formation when expressed in

undifferentiated layers of the skin (Alkhalaf et al. 1999; Ganguli and Wasylyk 2003). 34 Another mechanism that potentially enhances p53-independent proliferation is MDM2 or HDM2’s inhibition of the growth-restrictive functions of several cell cycle regulatory proteins: pRb, p73, E2F-1, PML, p300/CBP, PCAF, and p19ARF (Balint et al. 1999;

Dobbelstein et al. 1999; Gu and Roeder 1997; Hsieh et al. 1999; Jin et al. 2002; Jin et al.

2004; Kobet et al. 2000; Loughran and La Thangue 2000; Martin et al. 1995; Ongkeko et

al. 1999; Pomerantz et al. 1998; Wadgaonkar and Collins 1999; Wei et al. 2003; Xiao et al. 1995; Zeng et al. 1999). MDM2 might also mediate its p53-indpendent, growth- promoting effects by inducing anti-apoptotic factors, such as NF-κB (Gu et al. 2002).

Furthermore, MDM2 stimulates entry into S phase by triggering DNA synthesis through its interaction with the catalytic subunit of DNA polymerase ε (Asahara et al. 2003;

Vlatkovic et al. 2000), upregulation of cyclin A levels (Leveillard and Wasylyk 1997;

Wasylyk and Wasylyk 2000), and transactivation of E2F-1/DP1 (Martin et al. 1995).

Yeast two-hybrid screening of various cDNA libraries identified other novel proteins, such as MTBP, MDM-X, and the cell cycle regulator Numb (Boyd et al. 2000a; Juven-

Gershon et al. 1998; Sharp et al. 1999; Tanimura et al. 1999; Yogosawa et al. 2003).

Aside from its negative regulation on p53, the RING finger domain may alter cellular growth through other protein-protein, protein-RNA, and possibly protein-DNA interactions (Borden 2000; Elenbaas et al. 1996; Haupt et al. 1997; Kubbutat et al. 1997).

Aside from their growth-promoting effects, MDM2 and HDM2 are known to

regulate other cellular processes independent of p53. For instance, MDM2 may be

involved in cytonucleoplasmic ribosomal transport and ribosomal biogenesis as a result

of its interaction with the ribosomal L5 protein (Guerra and Issinger 1998; Marechal et

al. 1994). HDM2 and MDM2 activate or repress transcription via their association with 35 components of the transcriptional machinery and certain transcription factors: the

transcription factor Sp1 (Guo et al. 2003; Johnson-Pais et al. 2001), E2F1/DP1 (Martin et

al. 1995), the transcriptional coactivator TAFII250 (Leveillard and Wasylyk 1997;

Wasylyk and Wasylyk 2000), and the small subunit of the TBP/TFIIE complex (Thut et

al. 1997). As the RING finger domain binds to secondary structures and specific sequences in RNA, HDM2 and MDM2 could have a role in translational regulation

(Elenbaas et al. 1996; Yin et al. 2002). MDM2 may also be linked to DNA repair through

its association with Nbs1, which is a component of the Mre11-Nbs1-Rad50 complex that

repairs double-stranded DNA breaks (Alt et al. 2005). In all, these results strongly

indicate that the p53-independent functions of HDM2 and MDM2 regulate many other

molecular processes through many activities more complex than previously considered.

1.9 Contradictory effects of HDM2ALT1 expression

Expression of HDM2ALT1 in vivo and in vitro exhibits pleiotropic effects. The

conflicting phenotypes of HDM2ALT1 expression include p53-dependent growth

inhibition and p53-independent growth promotion. Sigalas et al. (1996) and Steinman et

al. (2004) showed that expression of HDM2ALT1 in NIH 3T3 cells increased foci formation and accelerated the rate of growth independent of p53. As the HDM2ALT1

protein cannot interact with p53, these growth-promoting effects may result from other

protein interactions of HDM2ALT1. Notably, the HDM2ALT1 protein retains the domains of

HDM2 that are known to stimulate cellular proliferation independent of p53 (Borden

2000; Elenbaas et al. 1996; Yin et al. 2002). Furthermore, hdm2ALT1 transcripts are the most frequently detected splice variants in many types of tumors (Bartel et al. 2001; 36 Evans et al. 2001; Evdokiou et al. 2001; Lukas et al. 2001; Matsumoto et al. 1998;

Sigalas et al. 1996; Tamborini et al. 2001). In contrast, Lukas et al. (2001) detected hdm2ALT1 transcripts in the normal mammary tissue surrounding breast tumors and the

presence of hdm2ALT1 transcripts in the mammary neoplasms correlated with a better

prognosis for these breast cancer patients. Evans et al. (2001) identified a physiological

function of hdm2ALT1 transcripts in MEFs and U2OS cells; HDM2ALT1 sequestered HDM2

within the cytoplasm and enhanced the transactivation of two p53-responsive promoters

(Evans et al. 2001). Chandler et al. (2006) and Dias et al. (2006) further demonstrated

that certain genotoxic stresses induced endogenous expression of hdm2ALT1 transcripts in numerous tumorigenic cell lines (e.g., MCF-7, RKO, H1299, U2OS, and SJSA-1), non- transformed immortalized cell lines (e.g., NL-20, NIH 3T3, and FHC), and early passage

MEFs. Although these conflicting effects may have resulted from expression of

HDM2ALT1 in different cellular backgrounds, MDM2-B (the murine homologue of

HDM2ALT1) displayed both contradictory phenotypes when expressed in the NIH 3T3 cell

line (Chandler et al. 2006; Steinman et al. 2004). Moreover, Fridman et al. (2003)

showed that expression of several murine splice variants of similar size and structure in

comparison with HDM2ALT1 significantly accelerated tumorigenesis in vivo while their in vitro expression decreased the growth rate of E1A and ras transformed MEFs. Therefore, other factors aside from the different cellular backgrounds might be responsible for producing these dissimilar growth effects. One explanation for these contradictory phenotypic effects of HDM2ALT1 may be the timing of its expression. For example,

induction of endogenous hdm2ALT1 was only detected after exposure to genotoxic insults

and expression of hdm2ALT1 terminated after 72 hours post-treatment (Chandler et al. 37 2006; Dias et al. 2006). Moreover, the p53-dependent growth-inhibitory effects of

ectopic HDM2ALT1 expression have only been observed when HDM2ALT1 was transiently

expressed (Dias et al. 2006; Evans et al. 2001). On the contrary, constitutive ectopic

expression of HDM2ALT1 via a retroviral-mediated delivery system resulted in the

growth-promoting phenotypes (e.g., inhibition of apoptosis, increased rate of growth

independent of p53, upregulation of cellular survival signals, and loss of contact

inhibition) (Steinman et al. 2004).

A model of HDM2ALT1’s function was proposed to explain these conflicting

phenotypes. Cellular stresses, such as DNA damage, induce alternative splicing of

hdm2 pre-mRNA to generate hdm2ALT1 mRNA, which reduces the levels of full-length

hdm2 transcripts available for translation into HDM2 protein. When the hdm2ALT1 transcripts are translated into HDM2ALT1 protein, HDM2 activity is further decreased

through the sequestration of the HDM2 protein by HDM2ALT1 in the cytoplasm. These

events perturb HDM2’s negative regulation of p53, thus resulting in stabilization and activation of p53, which, in turn, triggers the proper response to the damage. After the damage is repaired, alternative splicing of hdm2 pre-mRNA ceases. On the other hand, inappropriately sustained expression of HDM2ALT1 may activate its putative growth- promoting domains. Because the HDM2ALT1 protein also contains functional domains of HDM2 that have been implicated in stimulating proliferation and cell survival, it might also possess these p53-independent activities of HDM2 and MDM2. Therefore, constitutive HDM2ALT1 expression may act as an oncogenic stimulus that initiates

cellular transformation when constitutively overexpressed.

38

ALT1 HDM2 HDM2

Cellular stresses hdm2

Cellular p53 stresses

Figure 4. Proposed model of HDM2ALT1’s function.

The regulatory events occurring under non-stressed condition are indicated by the blue lines while the red lines specify the events that take place during the stress response. In the absence of stress, HDM2 mediates the destruction of p53. If any transcriptionally active p53 escapes degradation, p53 induces transcription of the hdm2 gene, which produces more HDM2 protein to further reduce the lingering p53 activity. During the stress response, numerous mechanisms impede the interaction between HDM2 and p53. The model of HDM2ALT1’s function predicts that cellular stresses shift the protein production of the hdm2 gene from HDM2 to HDM2ALT1, causing the levels of HDM2 protein to decrease. Then, the HDM2ALT1 protein further reduces the activity of the remaining HDM2 protein through its sequestration of HDM2 in the cytoplasm.

39 1.10 Specific aims of the present study

Due to the preeminent role of the p53 tumor suppressor in many of the molecular

pathways governing proliferation and apoptosis, as well as its frequent inactivation in tumors, a voluminous body of research has focused on p53. On the other hand, far few studies have examined the regulation and function of HDM2 in spite of its powerful control over the activity and stability of p53. Most reports investigating the regulation of

HDM2 have predominately examined how various covalent modifications of this protein

influence its ability to restrict p53’s growth suppressive activities. However,

accumulating data strongly indicates that other mechanisms of gene expression, such as

post-transcriptional processing, profoundly regulate HDM2 activity. Numerous

alternatively spliced transcripts of the hdm2 gene have been identified, but their function

and whether they are translated remains largely unknown. By using various genetic and

molecular approaches, the objective of this study is to characterize the protein function of

the most frequently detected alternatively spliced hdm2 transcript, hdm2ALT1. The specific

aims of the present study are as follows: 1) investigate the effects of inducible HDM2ALT1 expression in vivo, 2) examine the ability of HDM2ALT1 to bind to MDM2, 3) determine

whether HDM2ALT1 disrupts the p53-MDM2 interaction, 4) elucidate the molecular

mechanism by which HDM2ALT1 triggers a growth-inhibitory phenotype.

1.11 HDM2 and p53 as therapeutic targets in tumors

Surgery, chemotherapeutic drugs, and high-energy radiation are the standard

modalities used to treat neoplasms. Unfortunately, each of these treatments has

significant shortcomings that limit their usefulness in the clinical setting. For example, a 40 major drawback of surgery is the invasiveness of the procedure and complete resection

of most tumors is often complicated. Many chemotherapeutic drugs and radiotherapy

cause extensive DNA damage that triggers apoptosis in cancer cells; however, these

therapeutic agents are also highly cytotoxic and genotoxic to surrounding normal cells because they indiscriminately affect both normal and tumors cells. Due to these harmful effects on normal cells, only small doses of these agents can be safely administered, which limits their efficacy in killing the cancer cells. Furthermore, many tumor cells develop resistance to most chemotherapeutic agents. There are also concerns that the genotoxic treatments cause mutations in normal cells that may trigger cellular transformation and eventually lead to secondary tumor formation. Consequently, much attention has been focused on developing non-genotoxic therapeutic treatments.

Activation of p53 in tumors is a promising non-genotoxic target because of its

potent growth suppressive and pro-apoptotic activities (Chene 2003; Freedman et al.

1999). Moreover, most chemotherapeutic drugs and radiation treatments eliminate tumors cells through activation of the p53 network (Fischer and Lane 2004; Komarov et al.

1999; Vousden and Lu 2002). Although nearly half of all human tumors possess deleterious mutations in the p53 gene (Hainaut and Hollstein 2000), many other tumors retain p53 in its wild-type form. Therefore, activating p53 function in conjunction with conventional chemo- and radiotherapy may represent a new generation of highly- efficacious modalities used to successfully eradicate tumors that retain functional p53

(Chene 2003). An appealing approach to activate p53 activity in tumors is by inhibiting the p53-HDM2 interaction (Picksley and Lane 1993; Wu et al. 1993). Evidence from genetic studies demonstrated that disruption of this interaction potently activates p53’s 41 growth suppressive activities under stressed and non-stressed conditions (Chavez-

Reyes et al. 2003; Jones et al. 1995; Langheinrich et al. 2002; Mendrysa et al. 2003;

Montes de Oca Luna et al. 1995). Crystallographic information and biochemical analyses have defined the key residues necessary for the hydrophobic interaction between HDM2 and p53 (Bottger et al. 1997a; Bottger et al. 1997b; Bottger et al. 1996; Chen et al. 1993;

Freedman et al. 1997; Kussie et al. 1996). In addition, utilizing this therapeutic approach would be highly effective at restoring p53 activity in tumors that overexpress HDM2

(Klein and Vassilev 2004; Momand et al. 1998).

A growing body of evidence from various in vitro and in vivo studies has validated the p53-HDM2 interaction as a viable therapeutic target. For example, microinjection of monoclonal antibodies, which bound to HDM2 and blocked its association with p53, into tumor cell lines resulted in nuclear accumulation of p53

(Blaydes et al. 1997; Bottger et al. 1997a; Midgley and Lane 1997). Phage display analysis identified several natural peptides that bound to MDM2 with higher affinity than the native p53 sequence (Bottger et al. 1997a; Bottger et al. 1996; Kanovsky et al. 2001).

Intracellular expression or microinjection of chimeric mini proteins fused in frame with these peptides into several cancer cell lines stimulated the transcriptional activity of p53

(Bottger et al. 1997b; Kanovsky et al. 2001; Wasylyk et al. 1999). Derivatization of these peptides generated synthetic peptides with even higher avidity for HDM2 (Bottger et al.

1997b; Garcia-Echeverria et al. 2000), yet these derivatives only marginally induced growth arrest or apoptosis in various tumor cell lines (Chene et al. 2000; Chene et al.

2002). Natural inhibitors of the p53-HDM2 interaction have been identified from various biological sources (Duncan et al. 2003; Duncan et al. 2001; Stoll et al. 2001). However, 42 due to several undesirable properties of these compounds, such as side effects, low potency, complex chemical structures, and high molecular mass, these compounds are unsuitable for therapeutic use (Duncan et al. 2003; Duncan et al. 2001; Stoll et al. 2001).

Derivatives of these natural inhibitors, in addition to other polycyclic compounds, displayed anti-proliferative effects in tumor cells, but it remains unknown if their effects are solely exerted through inhibition of the p53-HDM2 interaction (Kumar et al. 2003;

Zhao et al. 2002). Intriguingly, a styrylquinazoline compound, designated CP-31398, was shown to reactivate both wild-type and mutant p53 by inducing a conformational change in the DNA binding domain of p53 (Wang et al. 2003b). Furthermore, the E3 ligase activity of HDM2 was found to be inhibited by several small molecule compounds (Lai et al. 2002). Therefore, other activities of p53 and HDM2 may also be desirable therapeutic targets (Issaeva et al. 2004). In all, these studies have provided essential clues for the development of drugs capable of competitively inhibiting the p53-HDM2 interaction. On the other hand, all of these compounds displayed marginal anti-tumor activity in vivo.

Because the contact surface between HDM2 and p53 is relatively small (Chene

2003), several groups have investigated whether small molecule inhibitors can disrupt this interaction. Small molecules have distinct advantages, such as greater stability, improved delivery, and enhanced bioavailability, over larger macromolecules, (e.g., peptides, antisense oligonucleotides, or antibodies) (Fotouhi and Graves 2005). Recently,

Vassilev et al. (2004) identified a class of small molecule antagonists (Nutlins) that disrupted the p53-HDM2 interaction. The Nutlins only exerted their growth suppressive effects in tumor cells that retained functional p53, which indicates that they activate the p53 stress response pathway (Vassilev et al. 2004). Remarkably, oral administration of 43 the Nutlins to athymic nude mice bearing established human tumor xenografts over a

three week period inhibited 90% of tumor growth without any discernable toxicity to the surrounding normal tissues (Vassilev et al. 2004). Furthermore, the Nutlins induced a reversible growth arrest, but not apoptosis, when applied to human and mouse normal diploid fibroblasts (Vassilev et al. 2004). The desirable pharmacological properties of the

Nutlins (e.g., cell membrane permeability, potent displacement of p53 from HDM2, prolonged stability, low toxicity to normal cells, and high selectivity for the p53-HDM2 interaction) indicate they have significant value as a broad-spectrum non-genotoxic therapeutic for tumors retaining p53 activity (Vassilev et al. 2004).

Another approach to disrupt the p53-HDM2 interaction is by down-regulating the

levels of HDM2 protein. Knockdown of HDM2 levels in tumors that overexpress this

protein may reactivate p53 function and subsequently trigger apoptosis. Furthermore,

reduced levels of HDM2 protein may also trigger an anti-tumor effect in neoplasms with

non-functional p53. Antisense oligonucleotides against hdm2 mRNA have been shown to

inhibit HDM2 expression in tumor cell lines (Chen et al. 1998; Chen et al. 1999; Geiger

et al. 2000; Grunbaum et al. 2001; Prasad et al. 2002; Wang et al. 2002a; Wang et al.

2003a; Zhang et al. 2004b) as well as in human xenografts injected into athymic nude

mice (Tortora et al. 2000; Wang et al. 2001; Wang et al. 2002b; Wang et al. 2003a;

Wang et al. 1999; Zhang and Wang 2000; Zhang et al. 2003). In these in vitro and in vivo

models, the reduced levels of HDM2 lead to stabilization and activation of p53.

Interestingly, these anti-MDM2 antisense oligonucleotides, in combination with various

genotoxic agents, reduced the number and size of multiple tumor types in the xenografts

models regardless of p53 status (Tortora et al. 2000; Wang et al. 2001; Wang et al. 44 2002b; Wang et al. 2003a; Wang et al. 1999; Zhang and Wang 2000; Zhang et al.

2003). A similar synergistic effect was observed in several tumor cell lines (Kondo et al.

1995; Meye et al. 2000; Sato et al. 2000). Mendrysa et al. (2003) further validated this combinatorial treatment approach by demonstrating that a decline in MDM2 activity increased radiosensitivity in several murine tissues. Down-regulation of MDM2 expression though an antisense approach may impede its p53-independent oncogenic activities, such as its inhibition of the cyclin kinase inhibitor p21WAF1/CIP1 (Klein and

Vassilev 2004; Zhang et al. 2004a). In support of this hypothesis, anti-HDM2 antisense oligonucleotides decreased mutant p53 levels and reduced tumor mass and numbers in a human soft tissue tumor xenograft model (Wurl et al. 2002). An antisense-mediated approach is unlikely to have severe side effects in normal tissues because genetic studies performed by Mendrysa et al. (2003) showed that decreased levels of MDM2 only triggers a small subset of endogenous normal tissues to undergo p53-mediated apoptosis.

These antisense oligonucleotides may also be used to down-regulate the expression of oncogenic splice variants detected in tumors, yet this approach has not been formally evaluated. However, targeting these splice variants may not be suitable because most reports examining the therapeutic implications of alternatively spliced mdm2 and hdm2 transcripts indicate that they generally enhance p53 activity (Harris 2005). 45

Chapter 2: Material and Methods

46 2.1 The Tet-On system

The Tet-On system (Clontech, Mountain View, CA) is a binary inducible system based on two transcriptional regulatory elements from the Tn10 transposon found in

Escherichia coli: the Tet repressor protein (TetR) and the operator, tet-O (Gossen and

Bujard 1992). The rtTA transactivator protein of the Tet-On system is a fusion protein

composed of a mutant TetR DNA-binding domain and the potent VP16 transactivation

domain from the herpes simplex virus VP16 transcription factor (Gossen and Bujard

1992). The tet-O site is a DNA-binding element recognized by the mutant and native

TetR DNA-binding domain (DBD) (Gossen and Bujard 1992). In the presence of doxycycline (a derivative of tetracycline), rtTA binds to a transcriptional regulatory DNA element consisting of seven tandem tet-O sequences (TRE) and transactivates hdm2ALT1 expression under the control of a minimal cytomegalovirus (CMV) promoter (Figure 5).

The levels of gene expression depend upon the concentration of doxycycline present. The tTS repressor protein eliminates transcriptional leakiness (i.e., rtTA has a residual affinity for the TRE site in the absence of doxycycline) of the Tet-On system (Zhu et al. 2001b).

tTS is a fusion protein composed of the TetR DNA-binding domain and the KRAB-AB silencing domain of the Kid-1 transcriptional repressor (Zhu et al. 2001b). In the absence

of doxycycline, tTS binds to the TRE site and represses initiation of the basal

transcriptional machinery, while addition of doxycycline dissociates tTS from the tet-O

site, allowing rtTA to bind and transactivate transcription (Figure 5) (Freundlieb et al.

1999). The TRE site co-regulates the expression of a firefly luciferase gene in addition to

hdm2ALT1 (Figure 5). These two promoters under the control of single TRE site allow

synchronized expression of HDM2ALT1 and luciferase (Figure 5). 47

Figure 5. The Tet-On system.

1) When doxycycline is absent, the TetR DBD of the tTS protein (Repressor) assumes a conformation that allows it to bind to the TRE site while the KRAB-AB domain suppresses the ability of RNA polymerase to induce transcription of the luciferase and hdm2ALT1 genes. The conformation of the mutant TetR DNA-binding domain of the rtTA protein prevents its association with the TRE site. 2) When doxycycline is present, the TetR DBD of the tTS fusion protein undergoes a conformational change that reduces its avidity for the TRE site. On the other hand, structural changes in mutant TetR DBD of the rtTA protein greatly increase its affinity for TRE while the VP16 domain powerfully stimulates RNA polymerase to initiate transcription of luciferase and hdm2ALT1.

48

2.2 Constructs

pcDNA3 ALT1

The hdm2ALT1 cDNA in the pCR 2.1 vector (Invitrogen, Carlsbad, CA) was

excised with the restriction enzyme EcoRI. This ~750 bp fragment was then cloned into

the unique EcoRI site within the polylinker of the pcDNA3 vector (Figure 6) (Invitrogen,

Carlsbad, CA) with the Quick Ligation Kit (New England Biolabs, Ipswich, MA).

Restriction mapping with ApaI confirmed that the insert was in the correct orientation.

The pcDNA3 ALT1 construct was used to transiently express hdm2ALT1.

Tet-O hdm2ALT1

The hdm2ALT1 cDNA in the pCR 2.1 vector was excised with the restriction

enzyme EagI. The resulting ~800 bp fragment was ligated into the unique EagI site of the pBI-L vector (Figure 6) (Clontech, Mountain View, CA). The correct orientation of the insert was confirmed by restriction digestion with ApaI. The Tet-O hdm2ALT1 construct

was utilized, in combination with the other components of the Tet-On system, pTet-On

(rtTA, Activator) and pTet-tTS (tTS, Repressor) (Clontech, Mountain View, CA), to

control the inducible expression of hdm2ALT1.

p53 luciferase reporter

The p53 luciferase reporter vector (Panomics, Fremont, CA) was used to examine

the transcriptional activity of p53. This plasmid contains a p53-responsive DNA element

upstream of a minimal thymidine kinase promoter that drives expression of firefly luciferase reporter gene. When p53 is transcriptionally active, it is able to bind to the PRE site and induce expression of firefly luciferase. 49

Tet-O hdm2ALT1

pcDNA3 ALT1

Figure 6. Map of the constructs containing hdm2ALT1 inserts

The hdm2ALT1 cDNA was excised from the pCR 2.1 vector and then was ligated into the polylinker sites of either pBI-L or pcDNA3. The size and other transcriptional regulatory components of the vectors are shown. Selected restriction enzymes sites are also indicated on the plasmids and inserts. The vector maps of pBI-L and pcDNA3 were modified from www.clontech.com and www.invitrogen.com, respectively. 50 2.3 Generation of transgenic mice

All transgenic constructs were linearized with appropriate restriction enzymes to

remove the vector sequences. The DNA fragments were then purified with the QIAquick

PCR Purification Kit (Qiagen, Valencia, CA) and resuspended at a concentration of 4

ng/μl in microinjection buffer (0.5 mM Tris-Cl, 25 mM EDTA, pH 7.5). All solutions and

chemicals described throughout this study were obtained from Sigma (St. Louis, MO) unless otherwise indicated. Constructs were microinjected into pronuclei derived from

SJL x C57BL/6J F1 zygotes (Jackson Laboratories, Bar Harbor, ME) and implanted into

pseudopregnant females (Jackson Laboratories, Bar Harbor, ME) by the staff at the

Edison Biotechnology Institute Transgenic Facility. Mice were housed in microisolation

cages containing corn cob bedding (The Andersons, Maumee, OH). Cages were kept in a

room with a 12 hr light cycle and mice were provided with distilled water and food ad

libitum (LabDiet 5P00 Prolab RMH 3000, PMI Nutritional International, Richmond, IN).

2.4 Tail DNA isolation

Up to 5 mm of tail was cut from mice less than 12 days of age and placed into a

microcentrifuge tube. 30 μl of proteinase K solution (20 mg/ml) and 500 μl SSTE (0.1 M

Tris-Cl, pH 8; 5 mM EDTA; 0.2% SDS; 0.2 M NaCl) was added to each tube and the tail

clips were digested overnight at 55ºC with periodic shaking. Genomic DNA was isolated

by phenol/chloroform extraction followed by ethanol precipitation. The DNA pellet was

dissolved in 300 μl of 10 mM Tris-Cl pH 8.5 and stored at 4ºC. The purity and amount of genomic DNA was quantified with the BioMate3 spectrophotometer (ThermoSpectronic,

Rochester, NY). 51 2.5 Genotyping

Identification of transgenic alleles (e.g., rtTA, tTS, and hdm2ALT1, and luciferase) was accomplished with polymerase chain reaction (PCR) analysis. The components of each PCR reaction were as follows: 100 ng of genomic DNA, 2.5 mM of dTNPs, 1 U of

Taq DNA polymerase, and 2.5 μM of each primer. The Taq DNA polymerase and

primers were purchased from New England Biolabs (Ipswich, MA) and BioSynthesis

(Lewisville, TX), respectively. The sequences of the forward and reverse primers for

each of the transgenes are indicated in Table 3 and have been previously described (Dias et al. 2006; Hocker et al. 2001; Ray et al. 1997; Sigalas et al. 1996; Zhu et al. 2001b).

All PCR reactions were performed in the PTC-100 Thermal Cycler (MJ Research,

Watertown, MA). The PCR parameters for amplification of rtTA and tTS transgenes were an initial cycle at 94ºC for 5 min, followed by 30 cycles at 94ºC for 1 min, annealed at 58°C for 1 min, and elongated at 72°C for 2 min, and a final extension step at 72°C for

5 min. The PCR conditions for amplification of the hdm2ALT1 transgene was an initial

cycle at 95ºC for 10 min, followed by 30 cycles at 94ºC for 1 min, annealed at 55°C for

1 min, and elongated at 72°C for 2 min, and a final extension step at 72°C for 10 min.

The PCR parameters for amplification of the luciferase transgene was an initial cycle at

94ºC for 5 min, followed by 34 cycles at 94ºC for 1 min, annealed at 52°C for 1 min, and

elongated at 72°C for 2 min, and a final extension at 72°C for 5 min. The products of the

PCR reactions were resolved by horizontal electrophoresis with a 1.5% agarose gel

containing ethidium bromide (0.5 µg/ml) to visualize the PCR products. Amplification of

the rtTA, tTS, hdm2ALT1, and luciferase alleles generated PCR products of 531 bp, 472 bp, 657 bp, and 391 bp, respectively. 52

Table 3. Forward and reverse primers used to genotype transgenic mice

Transgene Forward primer sequence (5'Æ 3') Reverse primer sequence (5’ Æ 3’)

rtTA (Activator) GTCGCTAAAGAAGAAAGGGAAACAC TTCCAAGGGCATCGGTAAACATCTG

tTS (Repressor) GAGTTGGCAGCAGTTTCTCC GAGCACAGCCACATCTTCAA

Tet-O HDM2ALT1 (Operator) CTGGGGAGTCTTGAGGGACC CAGGTTGTCTAAATTCCTAG

Luciferase (Operator) CAGAGGACCTATGATTATGTCCGG CGGTACTTCGTCCACAAACACAAC

53 2.6 Cell culture conditions

The NIH 3T3 cell line was generously provided by Dr. Donald Holzschu, Ohio

University. This cell line was cultured in Dulbecco’s Modified Eagle Medium Reduced

Serum (DMEM-RS) supplemented with 5% bovine growth serum (BGS) and 0.1 mg/ml

penicillin G (100 U/ml)/streptomycin (100 µg/ml)/amphotericin B (0.25 µg/ml) (1%

P/S/A) at 37°C in a humidified atmosphere of 5% CO2. Cells were subcultured

approximately every 4 to 5 days when cells reached 70-80% confluency. All cell culture

media and supplemental additives were obtained from Hyclone (Logan, UT).

2.7 Transient transfection

Transient transfections were performed in either 10 cm or 24 well tissue culture

plates. Transfections were performed with Lipofectamine 2000 (Invitrogen, Carlsbad,

CA) or Lipofectamine LTX transfection reagents (Invitrogen, Carlsbad, CA) in

accordance with the manufacturer’s recommendations. For each transfection in a 10 cm

plate, 1 X 105 cells were transfected in serum free media with 12 μg of plasmid DNA and

60 μl Lipofectamine 2000. For each transfection in a 24 well plate, 2.5 X 103 cells were

transfected in serum free media with 0.5 μg of plasmid DNA and 2 μl Lipofectamine

LTX. Six hours post-transfection, the media was removed and complete media (5% BGS,

1% P/S/A) was added. All samples were analyzed within 96 hrs post-transfection.

2.8 Stable transfection

Lipofectamine LTX reagent was used to transfect NIH 3T3 cell as described

above. Six hours post-transfection, complete media (5% BGS, 1% P/S/A) was added to 54 cells. 48 hrs later, cells were cultured in complete media (5% BGS, 1% P/S/A)

containing an antibiotic (800 μg/ml G418, 800 μg/ml hygromycin B, or 6.0 μg/ml puromycin) to facilitate positive selection. Individual clones were isolated with cloning discs (Clontech, Mountain View, CA) and cultured into separate wells. Complete medium with the antibiotics was replaced every four days to ensure continuous selection.

2.9 Antibodies

The various epitopes, protein and species specificity, antibody dilutions, and the

isotype and host of the antibodies used to detect MDM2, HDM2ALT1, and p53 are

described in Table 4. The monoclonal antibodies Ab-1, Ab-3, Ab-4, and Ab-6 were

purchased from Calbiochem (San Diego, CA) while the E19 and H221 polyclonal

antibodies were acquired from Santa Cruz Biotechnology (Santa Cruz, CA). The

fluorescein-conjugated anti-rabbit, anti-mouse and Texas red-conjugated anti-rabbit, anti-

goat, anti-mouse secondary antibodies used for immunofluorescence were purchased

from Vector Laboratories (Burlingame, CA). The Texas red- and fluorescein-conjugated

secondary antibodies were diluted 1:500. The HRP-conjugated goat anti-mouse, bovine

anti-rabbit and bovine anti-goat secondary antibodies used for western blot analysis were

obtained from Santa Cruz Biotechnology (Santa Cruz, CA). The HRP-conjugated

secondary antibodies were diluted 1:20,000. The monoclonal antibody TET03 used to

detect rtTA and tTS was purchased from BocaScientific (Boca Raton, FL) and was

diluted 1:500. The monoclonal anti-β-actin antibody (Sigma, St. Louis, MO) and was

diluted 1:10,000. The monoclonal antibodies F5 and B9 used to detect p21WAF1/CIP1 and

Bax (Santa Cruz Biotechnology, Santa Cruz, CA), respectively, were diluted 1:500. 55

Table 4. Description of the antibodies used to detect MDM2, HDM2ALT1, and p53.

Species Protein Protein Antibody Western blot Immunofluorescence Antibody specificity domain Epitope recognized isotype dilution dilution Ab-1 Human N-terminus 26-169 MDM2 Mouse 1:500 1:500 monoclonal IgG2A Ab-3 Human, mouse C-terminus 383-491 MDM2, Mouse 1:500 1:250 ALT1 HDM2 monoclonal IgG2A Ab-4 Human, mouse Acidic 153-222 MDM2 Mouse 1:500 1:250 (central) monoclonal IgG2A ALT1 Ab-6 Human C-terminus 383-491 HDM2 Mouse 1:500 1:250 monoclonal IgG2A E19 Mouse N-terminus Unknown p53 Goat polyclonal 1:1000 1:250 IgG H221 Human, mouse Central 100-320 MDM2 Rabbit polyclonal 1:1000 1:500 region IgG

56 2.10 Immunofluorescence

NIH 3T3 cells were grown on glass coverslips in 10 cm plates. Transient

transfections were performed as described above. At defined time points post-

transfection, cells were washed once in 1X phosphate buffered saline (PBS) and attached

to slides by fixation with 95% ethanol/5% acetic acid for 5 min at -20ºC. After three

washes in 1X PBS, cells were blocked in 1X PBS/2% chicken serum (Hyclone, Logan,

UT) for 30 min at room temperature (RT) and then incubated for 1 hr at room

temperature with one or a combination of the following antibodies: Ab-1, Ab-3, Ab-4,

Ab-6, E19, or H221. The dilutions for each of the antibodies used are indicated in Table

4. The cells were then washed three times in 1X PBS with 0.1% Tween 20 (PBS-T) and incubated with the appropriate secondary antibody (Texas red- or fluorescein-conjugated)

diluted 1:500 at RT for 1 hr in the dark. The coverslips were mounted onto glass slides

containing anti-fade mounting medium and 4, 6-diamidino-2-phenylindole (DAPI, 1.5

μg/ml, Vector Laboratories, Burlingame, CA). Cells were photographed using the Nikon

E600 fluorescence microscope (Nikon, Japan) equipped with a digital camera (DC

Imaging, West Chester, OH) at 200X magnification. Fluorescent images were merged

with the SPOT Advanced software (Diagnostic Instruments, Sterling Heights, MI)

2.11 Western blot analysis

Total protein lysates were collected from cells at various time points. Cells were

washed twice with 1X PBS and lysed on ice with protein lysis buffer (50 mM Tris-HCl

pH 7.6, 150 mM NaCl, 0.05% SDS, 0.5% IGEPAL CA-630) containing an assortment of

protease inhibitors (Roche Applied Science, Indianapolis, IN) for one hour. The protein 57 extracts were harvested by centrifugation (13,200 RPM) at 4ºC for 15 min. The

bicinchoninic acid assay (Pierce Biotechnology, Rockford, IL) was used to determine the

protein concentration of the lysates. Equal protein concentrations from the cell lysates or

precipitates from immunoprecipitation reactions were heated at 95ºC for 5 min and

centrifuged (13,200 RPM) for 1 min at RT. The lysates were then loaded and separated

by 10% SDS-polyacrylamide gel electrophoresis (SDS-PAGE). The protein samples were

transferred onto the Immobilon-P membrane (Millipore, Billerica, MA) by electroblotting

(HEP-3 Panther Semi Dry Electroblotting System, Portsmouth, NH). The membranes

were blocked in 1X PBS-T/1% bovine serum albumin (BSA) (Pierce Biotechnology,

Rockford, IL) for 1 hr and incubated with one of the following antibodies overnight at

4ºC to detect MDM2 (Ab-4 or Ab-3), HDM2ALT1 (Ab-3 or Ab-6), p53 (E19), β-actin

(anti-β-actin), p21WAF1/CIP1 (F5), Bax (B9), or rtTA and tTS (TET03). The respective

HRP-conjugated secondary antibodies were diluted 1:20,000 in 1X PBS-T/1% BSA and

incubated for one hour. The membranes were washed four times with 1X PBS for 10 min

each wash. Protein samples were visualized on autoradiography film (Denville Scientific,

Metuchen, NJ) using the ECL-Advance Western Blotting Detection Kit (Amersham

Biosciences, Pittsburgh, PA).

2.12 Immunoprecipitation

Equal amounts (750 μg) of protein from total protein lysates were used for co-

immunoprecipitation reactions. Samples were precleared by adding 1.0 μg of the

secondary antibody corresponding to the host species of the primary antibody used (e.g.,

mouse, rabbit or goat) and 20 μl of Protein G PLUS-Agarose beads (Santa Cruz 58 Biotechnology, Santa Cruz, CA) for 30 min at 4°C. The precleared lysates were then

incubated with 1 μg of the appropriate primary antibody for 1 hour at 4°C. 20 μl of the

Protein G beads was then added to each sample and the mixture was incubated at 4°C

overnight on a rotating platform. The immunoprecipitates were collected by

centrifugation (2,500 RPM) at 4°C for 5 min and washed four times with 0.5 ml 1X PBS,

repeating the centrifugation between each wash. The pellet was resuspended in 25 μl of

protein lysis buffer (50 mM Tris-HCl pH 7.6, 150 mM NaCl, 0.05% SDS, 0.5% IGEPAL

CA-630) and analyzed by SDS-PAGE.

2.13 TUNEL

Cells were grown on four-chambered slides with a chemically modified surface

(Nalgene, Rochester, NY) at a density of 1 X 103 cells per chamber. Cells were

transfected according the transient transfection procedure for the 24 well format

described above. Apoptotic cells were detected with the DeadEnd Fluorometric TUNEL

System (Promega, Madison, WI) in accordance with the manufacturer’s instructions.

Anti-fade mounting medium containing DAPI (1.5 μg/ml) was added to each slide. Cells were photographed cells at 200X magnification using the Nikon E600 fluorescence microscope.

2.14 Cell death detection assay

Cells were transiently transfected in a 24 well format as described above. At

various time points post-transfection, the cells were washed with 1X PBS and lysed. The

extent of apoptosis in the lysates was measured with the Cell Death Detection ELISAPLUS 59 assay (Roche Applied Science, Indianapolis, IN) according to the manufacturer’s

instructions. This quantitative sandwich-enzyme immunoassay detects histone-associated

DNA fragments (nucleosomes) released into the cytoplasm during apoptosis. The

peroxidase-conjugated antibody complexes were incubated at RT for 20 min with the 2,

2’-azino-di-(3 ethylbenzthiazoline sulfonic acid) substrate. Enzymatic cleavage of this

substrate generated a green complex that was quantified photometrically at 405 nm with

the SpectraMax M2 Microplate Reader (Molecular Devices, Sunnyvale, CA). Each time

point was performed in triplicate to calculate an average and standard deviation (SD).

2.15 Cell viability assay

The cellular concentration of ATP was quantified with the CellTiter-Glo

Luminescent Cell Viability Assay (Promega, Madison, WI) according to the manufacturer’s recommendations. Luminescence was measured with the Lumat LB 9507 luminometer (Berthold Technologies, Oak Ridge, TN). A standard curve was generated to directly correlate the number of viable cells with the concentration of ATP. Cells were seeded at densities ranging from 100 to 1,000,000 cells per well in a 24 well format. Each sample was plated in triplicate to calculate an average and SD. After a direct linear relationship between the number of cells and the corresponding luminescence measured was established over several orders of magnitude, cells were grown in 24 well plates and transiently transfected as previously described. At defined time points, the amount of luminescence activity was measured to determine the number of viable cells. The samples were performed in triplicate to report an average ± SD for each time point examined. Linear regression was performed to calculate the growth rate of the cells. 60 2.16 Flow cytometry

Cells were cultured in 10 cm plates and transiently transfected as described above.

Cells were trypsinized and centrifuged (1,200 RPM) for 5 min. Cells were washed three

times in 1X PBS and fixed in 5 ml of ice-cold 70% ethanol added dropwise while

vortexing. Cells were stored at 4ºC for at least 4 hrs. Cells were then centrifuged (1,200

RPM) for 5 min and resuspended in 1X PBS containing 5 μg/ml RNase A and 50 μg/ml propidium iodide. Finally, cells were analyzed by the FACSort flow cytometer (Becton

Dickinson, San Jose, CA) and the CellQuest software (Becton Dickinson, San Jose, CA).

The resulting DNA histograms were used to calculate the percentage of cells in each

phase of the cell cycle. Each sample was performed in triplicate to determine an average

± SD. Averages of the percentage of cells populations in each cell cycle phase were

compared by the t test.

2.17 Luciferase assay

Cells were grown in 24 well plates and co-transfected with 0.25 μg of a firefly

luciferase construct (e.g., pBI-L or p53 Luciferase Reporter Vector) and 0.25 ng of the

Renilla luciferase vector pRL-TK (Promega, Madison, WI). The amount of firefly and

Renilla luminescence was quantified with the Dual Luciferase Reporter System Assay

(Promega, Madison, WI). The firefly luciferase activity was normalized against the

activity of Renilla luciferase. The activity of the Renilla luciferase acted as a control to

determine the baseline levels of transcription and translation within each cell. Cells were

washed once with 1X PBS and lysed for 30 min. Luminescence was measured with the

Lumat LB 9507 luminometer. 61

Chapter 3: Results

62 3.1 Identification of transgenic founders

The efficacy of transgenesis for the tTS construct was much higher (23.78%) in

comparison with the rtTA (6.25%) and Tet-O hdm2ALT1 (3.08%) transgenes (Table 5). Of

the 143 pups born from the zygotes microinjected with the tTS transgene expression

cassette, 34 were identified as transgenic founders (Table 5). One of the tTS transgenic

founder mice died at three months of age for unknown reasons. PCR analysis of the 128

viable pups generated from microinjection of the rtTA expression cassette identified eight

as transgenic founders (Table 5). Two of the rtTA transgenic founder mice were

sacrificed at two and four months of age due to a severe skin irritation and the

development of a tumor within the testes, respectively. Selected organs from these affected rtTA founders were snap frozen in a dry ice/ethanol bath and stored at -80ºC for future analysis. 65 viable pups were produced from microinjection of the Tet-O hdm2ALT1

expression cassette into zygotes. Genotyping for both the hdm2ALT1 and luciferase

transgene identified two mice as transgenic founders (Table 5). Once transgenic founders were identified, they were mated with C57Bl/6J mice purchased from Jackson

Laboratories (Bar Harbor, ME) to increase the percentage of the C57Bl/6J background.

Crossing transgenic founders with C57Bl/6J mice was also used to determine which of the founder mice transmitted their respective transgene to offspring.

63 Table 5. Summary of the transgenic founder mice.

Number of Efficacy of Transgene Transgene founders transgenesis transmission tTS (Repressor) 34 23.78% Four generations and ongoing rtTA (Activator) 8 6.25% Stopped after two generations hdm2ALT1 and 2 3.08% Stopped after one luciferase generation

64 3.2 Expression of the transgenes in murine tissues

After founders were identified by PCR analysis and were shown to transmit their

transgene to offspring, the protein expression of the corresponding transgenes in each of

the lines was examined. One male and female from each of the transgenic lines was

sacrificed to harvest organs (e.g., brain, lung, small intestine, kidney, liver, heart, testis,

ovary, mammary gland, muscle, and spleen). Tissues were homogenized on ice in a

protein lysis buffer (50 mM Tris-HCl pH 7.6, 150 mM NaCl, 0.5% IGEPAL CA-630) containing a mixture of protease inhibitors. The concentration of total protein in the tissue homogenates was quantified with the bicinchoninic acid assay and equal amounts of protein from the tissue extracts were then separated by SDS-PAGE. Western blot analysis with the TET03 antibody was used to detect the tTS and rtTA proteins. Several transgenic lines were identified that did not to express rtTA or tTS in every tissue (data not shown). As a result, these transgenic lines were discontinued. Only one rtTA and tTS transgenic line, each having the highest transgene expression in all of the tissues examined by western blot analysis, were used in subsequent experiments (data not shown). Because leaky hdm2ALT1 expression in the uninduced state may cause an

undesirable phenotype, the levels of hdm2ALT1 expression were examined in transgenic

mice containing the Tet-O hdm2ALT1 expression cassette. Although these transgenic mice

were predicted to not express HDM2ALT1 due to the lack of the transcriptional activator,

rtTA, the minimal CMV promoter has been shown to have varying levels of background expression in different tissues (Furth et al. 1991; Ryding et al. 2001). None of the tissues examined by western blot analysis showed detectable levels of hdm2ALT1 expression in

the two Tet-O hdm2ALT1 transgenic lines (data not shown). 65 3.3 Establishment of transgenic lines

The goal was to mate transgenic founders with C57Bl/6J mice for four

generations to increase the percentage of C57Bl/6J genetic background over 90%. The

high percentage of the C57Bl/6J background should minimize any contributing

phenotypic effects caused by genetic modifiers in the SJL background. Western blot

analysis of selected tissues from several transgenic lines identified an optimal transgenic

line for each of the three different transgenes. The tTS transgenic line has been

successfully backcrossed with the C57Bl/6J mice for four generations and transmission

of the tTS transgene is ongoing. A breeding pair of the tTS transgenic line is currently being maintained while all founders and previous generations from this line have been sacrificed. The rtTA transgenic line has been backcrossed with C57Bl/6J mice for two generations. However, mice from the F2 generation did not transmit the rtTA transgene to any offspring after repeated mating. To date, several attempts to reestablish the rtTA transgenic lines by backcrossing the remaining rtTA transgenic founders with C57Bl/6J mice have also been unsuccessful. The two Tet-O hdm2ALT1 transgenic lines have been

backcrossed with the C57Bl/6J mice for one generation. Notably, both of the two Tet-O

hdm2ALT1 founders required nearly one year to produce any viable offspring.

Backcrossing the F1 generation mice with C57Bl/6J mice produced progeny that did not

possess either the hdm2ALT1 or the luciferase transgene. Repeated mating of the Tet-O

hdm2ALT1 founders with C57Bl/6J mice only resulted in small broods of non-transgenic

ALT1 pups. All Tet-O hdm2 founders and F1 mice have been sacrificed. As the rtTA and

Tet-O hdm2ALT1 transgenic lines could not be established, the in vivo study examining

HDM2ALT1 expression was ended. 66 3.4 Development of a stable NIH 3T3 cell line with the Tet-On system

Because attempts to generate transgenic lines with the components of the Tet-On

system were unsuccessful, the phenotypic effects of inducible HDM2ALT1 expression

were examined in the NIH 3T3 cell line. The objective was to generate a stable cell line

with the regulatory components of the Tet-On system. Consecutive stable transfections,

in the order of rtTA, tTS and Tet-O hdm2ALT1, were performed to select stable clones that have low background expression of HDM2ALT1 in the absence of doxycycline and high

induction (i.e., > 500 fold) of HDM2ALT1 expression when doxycycline is added to the

growth medium. As the site of integration of the constructs can profoundly influence the

expression of the tTS and rtTA proteins, approximately 100 clones were selected and

screened after each stable transfection. Positive selection for isolating stable clones was

accomplished with the following selective antibiotics: G418, rtTA; hygromycin B, tTS;

puromycin, Tet-O hdm2ALT1. The stable clones were screened by examining the levels of

luciferase activity after the addition of doxycycline. Stable clones were transiently

transfected with the empty pBI-L reporter plasmid (Figure 6) and the Renilla luciferase

vector, pRL-TK. Six hours post-transfection, the media was removed and complete media

containing doxycycline (1 μg/ml) was added to the cells. Luciferase activity was

measured 72 hrs post-transfection with the Dual Luciferase Reporter System Assay.

Firefly luciferase (pBI-L) activity was normalized against Renilla luciferase activity

(pRL-TK). After screening 150 rtTA stable clones, one clone was selected that possessed

more than 5000 fold induction in the presence of doxycycline and low levels of

expression in the uninduced state (Figure 7). Frozen stocks were made from each of the

optimal stable clones. Next, stable transfection with the tTS construct was performed. 67

10000 Doxycycline No doxycycline

1000

100

10

Fold induction (Relative luciferase activity) activity) luciferase (Relative induction Fold 1 rtTA rtTA/tTS rtTA/tTS/Tet-O hdm2alt1 Tet-On components

Figure 7. Inducible control of luciferase activity in stable cell lines.

Stable cell lines were generated by sequentially transfecting NIH 3T3 cells with each of the components of the Tet-On system. Stable cell lines were screened by examining the induction of luciferase activity with doxycycline (1 μg/ml) or without doxycycline. The optimal stable clones displaying the highest induction of luciferase activity in the presence of doxycycline and lowest luciferase activity in the absence of doxycycline are indicated. Values are reported as an average ± SD.

68 The tTS protein was expected to eliminate basal firefly luciferase activity in the

absence of doxycycline. Screening of 125 tTS stable clones identified one clone with

~5000 fold induction when media was supplemented with doxycycline and no induction

of luciferase activity in the uninduced state (Figure 7). Western blot analysis of protein

lysates from this double stable cell line confirmed the presence of the rtTA and tTS

proteins (data not shown). Finally, the Tet-O hdm2ALT1 was stably transfected into the

rtTA/tTS stable clone. Of the 151 clones screened, none were found to have inducible

luciferase activity when doxycycline was present in the media (Figure 7). Because the

addition of doxycycline induced simultaneous expression of the firefly luciferase and

HDM2ALT1 proteins, the screen was repeated using lower concentrations (1-100 ng/ml) of

doxycycline to avoid potential cytotoxic effects of high levels of HDM2ALT1 expression.

Again, no detectable firefly luciferase activity was observed in the lower concentrations

of doxycycline (data not shown). Concentrations of doxycycline as low as 1 ng/ml have

been reported to successfully induce gene expression with the Tet-On system (Freundlieb et al. 1997; Freundlieb et al. 1999; Gossen et al. 1995). Microscopic visualization of the cells in the presence of doxycycline revealed that the cells were healthy without any of the morphological characteristics of apoptosis (e.g., membrane blebbing and/or loss of anchorage). A frozen stock of the NIH 3T3 cell line containing both rtTA and tTS stably integrated was thawed and the stable transfection with Tet-O hdm2ALT1 was repeated. The

number of clones utilized in the screening process was increased to 314. Once again,

firefly luciferase activity was not detected after induction with doxycycline (data not

shown). As attempts to generate a stable cell line with all of the components of the Tet-

On system were unsuccessful, this experimental approach was ended. 69 3.5 HDM2ALT1 expression does not affect the levels of MDM2

Due to the failure to express HDM2ALT1 under the control of the Tet-On system

in vivo and in cell culture, the effect of transient HDM2ALT1 expression was investigated

in the NIH 3T3 cell line. Cells were transiently transfected with the pcDNA3 ALT1

vector or the empty pcDNA3 vector as previously described. To verify that the

HDM2ALT1 protein was expressed in HDM2ALT1 transfected cells and that HDM2ALT1 was not present in mock transfected cells, protein lysates from transfected cells were analyzed by immunoblotting with the Ab-3 antibody. Ab-3 recognizes MDM2 and HDM2ALT1

(Table 4), which allowed both proteins to be simultaneously detected in the same protein lysate sample. HDM2ALT1 expression was detected after 24 hrs post-transfection and

increased steadily up to 66 hrs in HDM2ALT1 transfected cells (Figure 8). After later time

points post-transfection, the amount of HDM2ALT1 protein gradually declined until its

levels were barely detectable at 96 hrs (Figure 8). An increase in the levels of MDM2

was detected between 48 and 66 hrs when the levels of HDM2ALT1 expression peaked in

the HDM2ALT1 transfected cells (Figure 8). However, the levels of MDM2 protein

remained relatively constant at all of the other time points examined (Figure 8). The

HDM2ALT1 protein was not detected in mock transfected cells (Figure 8). No other

endogenous MDM2 isoforms (e.g., post-translationally modified forms of the MDM2

protein or additional MDM2 splice variants) were detected with the Ab-3 antibody

(Figure 8). The MDM2 protein was detected in mock transfected cells and its protein levels were constant at all of the time points examined with the exception of the 0 hr control sample (Figure 8).

70

Figure 8. Western blot analysis of MDM2 and HDM2ALT1 protein in transfected cells.

NIH 3T3 cells were transiently transfected with the pcDNA3 ALT1 vector (HDM2ALT1) or the empty pcDNA3 vector (Mock). At various time points (hours) post-transfection, total protein lysates were harvested from cells. The Ab-3 antibody was used to detect the levels of MDM2 and HDM2ALT1 protein. The MDM2 protein migrated at 90 kDa while the HDM2ALT1 protein migrated at approximately 47 kDa. The levels of β-actin were used as a loading control.

71 3.6 HDM2ALT1 interacts with MDM2

The HDM2ALT1 protein cannot interact with the p53 protein because it lacks the

p53-binding domain (Evans et al. 2001; Sigalas et al. 1996); however, HDM2ALT1 has been shown to interact with the HDM2 protein in MCF-7 cells (Dias et al. 2006; Evans et al. 2001). The murine homologue of HDM2ALT1, MDM2-B, has also been demonstrated

to bind to MDM2 (Chandler et al. 2006). Because the HDM2ALT1 and MDM2-B proteins

are 82% identical at the amino acid level (Steinman et al. 2004), the ability of HDM2ALT1

to interact with MDM2 was investigated. The Ab-6 and Ab-4 antibodies were used to immunoprecipitate HDM2ALT1 and MDM2, respectively, in HDM2ALT1 or mock

transfected cells. The Ab-3 and Ab-1 antibodies were not used for immunoprecipitation

because Ab-3 binds to both HDM2ALT1 and MDM2 and Ab-1 cannot recognize either

protein. The supernatant of each immunoprecipitation was saved for further analysis. The

precipitates were immunoblotted with Ab-6, Ab-4, Ab-3, or Ab-1. The Ab-6 and Ab-4

antibodies did not display cross-reaction with MDM2 or HDM2ALT1, correspondingly

(Lanes 1 and 6, Figure 9). The Ab-1 antibody did not detect HDM2ALT1 or MDM2 (Lanes

13-16, Figure 9) as predicted. The Ab-3 antibody was able to detect both HDM2ALT1 and

MDM2 after immunoprecipitation with either Ab-6 or Ab-4 (Lanes 9 and 10, Figure 9).

Immunoprecipitation of HDM2ALT1 with Ab-6 followed by western blot analysis with

Ab-4 exclusively detected the MDM2 protein (Lane 5, Figure 9). Immunoprecipitation of

MDM2 with Ab-4 followed by western blot analysis with Ab-6 only detected the

HDM2ALT1 protein (Lane 2, Figure 9). The HDM2ALT1 protein was not detected in mock

transfected cells (Lanes 3, 4, 8, 11, and 12, Figure 9). 72

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16

Figure 9. Interaction of HDM2ALT1 with MDM2.

Protein extracts were harvested from cells 72 hrs post-transfection with pcDNA3 ALT1 (HDM2ALT1) or the empty pcDNA3 vector (Mock). Equal amounts (750 μg) of protein lysates were immunoprecipitated (IP) with Ab-6 or Ab-4 and immunoblotted (WB) with the Ab-6, Ab-4, Ab-3, and Ab-1 antibodies. The HDM2ALT1 protein was only detected in cells transfected with the pcDNA3 ALT1 vector. MDM2 was detected in both mock and HDM2ALT1 transfected cells. The MDM2 protein migrated at 90 kDa while the HDM2ALT1 protein migrated at approximately 47 kDa.

73 3.7 HDM2ALT1 expression decreases the p53-MDM2 interaction

Evans et al. (2001) demonstrated that when HDM2 was bound to HDM2ALT1,

HDM2 could not concurrently bind to p53. Expression of HDM2ALT1 also increased the

transcriptional activity of p53 in the U2OS cell line presumably by disrupting the p53-

HDM2 interaction (Evans et al. 2001). As HDM2ALT1 was shown to bind to MDM2

(Figure 9), the effect of HDM2ALT1 expression on the p53-MDM2 interaction was examined. The supernatants from the immunoprecipitation reactions described above

were immunoblotted with the E19 antibody to examine the levels of unbound p53. High

levels of p53 protein were detected in the supernatants from lysates of HDM2ALT1 transfected cells (Figure 10A). On the other hand, the levels of p53 protein were barely detectable in the supernatants from lysates of mock transfected cells (Figure 10B). The amount of p53 protein associated with MDM2 protein in mock or HDM2ALT1 transfected

cells was also examined to confirm the levels of unbound p53 protein detected in the

supernatants from the previous immunoprecipitation reactions. Immunoprecipitation of

either p53 with the E19 antibody or MDM2 with the Ab-4 antibody followed by western

blot analysis with the E19 antibody was performed. Immunoprecipitation and

immunoblotting with E19 detected the p53 protein in both mock and HDM2ALT1

transfected cells (Figure 10C). Immunoprecipitation of MDM2 with Ab-4 followed by

western blot analysis with E19 detected very low levels of p53 associated with MDM2 in

the HDM2ALT1 transfected cells (Figure 10C). Much higher levels of p53 protein were associated with MDM2 in the mock transfected cells (Figure 10C).

74

Figure 10. HDM2ALT1 disrupts the interaction of p53 and MDM2

Protein extracts were harvested from cells 72 hrs post-transfection with pcDNA3 ALT1 (HDM2ALT1) or the empty pcDNA3 vector (Mock). Equal amounts (750 μg) of protein lysates were immunoprecipitated (IP) with Ab-6 or Ab-4. (A and B) After immunoprecipitation of MDM2 and HDM2ALT1, equal concentrations of protein (30 μg) from supernatants of the IP reactions were analyzed by western blot analysis to detect the levels of unbound p53. The E19 antibody was utilized to detect the p53 protein. β-actin was used as a loading control. . The p53 protein migrated at 53 kDa. (C) MDM2 and p53 were immunoprecipitated with the Ab-4 and E19 antibodies, correspondingly. Western blot analysis with the E19 antibody was used to detect p53 in these IP reactions. 75 3.8 HDM2ALT1 expression increases the levels of p53 and Bax, but not p21WAF/CIP1

Disruption of the p53-MDM2 interaction has been shown to increase the stability of the p53 protein (Mendrysa et al. 2003; Vassilev 2004; Vassilev et al. 2004). Due to the disruption of the interaction of MDM2 and p53 when HDM2ALT1 was expressed and the

increased levels of unbound p53 protein in the IP reactions (Figure 10), the total protein

levels of p53 were examined in HDM2ALT1 transfected cells by western blot analysis. The p53 protein was detected after 24 hrs and its protein levels steadily increased up to 60 hrs post-transfection with HDM2ALT1 (Figure 11A). After 60 hrs, the levels of p53 protein

gradually decreased until undetectable at 96 hrs (Figure 11A). p53 was not detected at

any of the time points examined in mock transfected cells (Figure 11A).

Once activated, p53 primarily transactivates the expression of downstream target

genes involved in cell cycle arrest or apoptosis (el-Deiry 1998; Hupp et al. 1992; Levine

et al. 2006). To determine whether the increased levels of p53 resulted in transcriptional activation of downstream effectors of the stress response, the levels of two p53- repsonsive target genes, p21WAF1/CIP1 (cell cycle arrest) and bax (apoptosis), were

examined. The p21WAF1/CIP1 protein was not detected in either HDM2ALT1 or mock

transfected cells by western blot analysis (data not shown). The Bax protein was detected

after 30 hrs in HDM2ALT1 transfected cells and its levels increased steadily up to 42 hrs

post-transfection (Figure 11B). After 42 hrs post-transfection, the levels of Bax protein

remained constant up to 96 hrs. The Bax protein was not detected at any of the time

points examined in mock transfected cells (Figure 11B). 76

Figure 11. Western blot analysis of p53 and Bax protein in transfected cells.

NIH 3T3 cells were transiently transfected with the pcDNA3 ALT1 vector (HDM2ALT1) or the empty pcDNA3 vector (Mock). At various time points (hours) post-transfection, protein lysates were harvested from transfected cells and were analyzed by western blot. (A) The E19 antibody was used to detect p53. (B) The B9 antibody was used to detect the Bax protein. Bax migrated at approximately 21 kDa while p53 migrated at 53 kDa. The levels of β-actin were used as a loading control.

77 3.9 HDM2ALT1 stimulates the transcriptional activity of p53

An accumulating body of evidence suggests that stabilization of p53 by disrupting

the p53-MDM2 interaction is sufficient to activate p53’s transcriptional activity (Bottger

et al. 1997b; Kanovsky et al. 2001; Mendrysa et al. 2003; Wasylyk et al. 1999). In

support of these observations, the levels of Bax were increased by HDM2ALT1 expression

(Figure 11B). However, several studies have indicated that the p53 protein increases the

activity of Bax independent of p53-mediated transcriptional activation of the bax gene

(Chipuk et al. 2004; Chipuk et al. 2003; Degli Esposti and Dive 2003; Scorrano and

Korsmeyer 2003). To confirm that p53 was transcriptionally activated by HDM2ALT1 expression, a p53-responsive reporter construct was utilized. NIH 3T3 cells were transfected with the p53 luciferase reporter vector and assayed for firefly luciferase activity at various time points post-transfection. The activity of firefly luciferase from the reporter plasmid in HDM2ALT1 transfected cells increased from 24 hrs and peaked at 78

hrs post-transfection. After 78 hrs post-transfection with HDM2ALT1, the levels of firefly

luciferase activity steadily declined (Figure 12). Firefly luciferase expression from the

reporter construct was not significantly induced in mock transfected cells (Figure 12).

78

25 HDM2ALT1 Mock 20

15

10

5

Fold induction (Relative luciferase activity) 0 0 20406080100 Time (hours)

Figure 12. Transcriptional activity of p53 in transfected cells.

NIH 3T3 cells were transfected with the p53 Luciferase Reporter construct (firefly) and pRL-TK (Renilla) in addition to the pcDNA3 ALT1 vector (HDM2ALT1) or the empty pcDNA3 vector (Mock). At the time points indicated, transfected cells were lysed and analyzed for firefly and Renilla luciferase activity with the Dual Luciferase Reporter System Assay. Firefly luciferase activity was normalized against Renilla luciferase activity to determine the fold induction of firefly luciferase activity. Fold induction values were reported as an average ± SD.

79 3.10 The subcellular localization of HDM2ALT1, MDM2 and p53

The proposed model of HDM2ALT1 function predicts that HDM2ALT1 inhibits

MDM2 activity by sequestering MDM2 within the cytoplasm. HDM2ALT1 is also predicted to disrupt the p53-MDM2 interaction and subsequently allow p53 to enter the nucleus. The subcellular localization of MDM2 and p53 were analyzed in the presence or absence of HDM2ALT1 expression. The Ab-6, Ab-4, and E19 antibodies were used to examine the subcellular localization of HDM2ALT1, MDM2, and p53, correspondingly.

The Ab-6 antibody displayed cross reactivity with MDM2 at lower dilutions (1:50 and

1:100). However, after performing serial dilutions with this antibody, dilutions of 1:250 or higher were shown not to exhibit any cross-reaction (data not shown). None of the secondary antibodies displayed any background staining when incubated alone with cells

(data not shown). The Ab-3 antibody was not utilized because this antibody recognizes epitopes found in both MDM2 and HDM2ALT1 and thus cannot distinguish between the two proteins. Under normal conditions, p53 is diffused throughout the cytoplasm whereas

MDM2 is predominately restricted within the nucleus (data not shown). After examining cells at several time points post-transfection with the empty pcDNA3 vector, MDM2 remained restricted within the nucleus while p53 continued to be localized in the cytoplasm (Figures 13A, B, C and D). From 24 hrs to 72 hrs post-transfection, the

HDM2ALT1 protein was detected within the cytoplasm, MDM2 was re-localized to cytoplasm, and p53 was exclusively detected in the nucleus (Figures 14A, B, and C). At

96 hrs post-transfection, very few cells displayed positive staining for the HDM2ALT1 protein while MDM2 was predominately localized within the nucleus and p53 diffused throughout the cytoplasm (Figure 14D). 80

Figure 13A. Subcellular localization of HDM2ALT1, MDM2, and p53 at 24 hours after mock transfection.

Cells were transiently transfected with the empty pcDNA3 vector. Cells were immunostained with the Ab-6, Ab-4 or E19 antibodies. Positive staining for MDM2, HDM2ALT1, or p53 is indicated by the red color. The nucleus was visualized by staining with DAPI (blue). Cells were photographed at either 200X or 400X magnification. The images were merged and the merge color (purple) indicates the MDM2, HDM2ALT1 or p53 was localized within the nucleus.

81

Figure 13B. Subcellular localization of HDM2ALT1, MDM2, and p53 at 48 hours after mock transfection.

82

Figure 13C. Subcellular localization of HDM2ALT1, MDM2, and p53 at 72 hours after mock transfection.

83

Figure 13D. Subcellular localization of HDM2ALT1, MDM2, and p53 at 96 hours after mock transfection.

84

Figure 14A: Subcellular localization of HDM2ALT1, MDM2, and p53 at 24 hours after transfection with HDM2ALT1.

Cells were transiently transfected with the pcDNA3 ALT1 vector. Cells were immunostained with the Ab-6, Ab-4 or E19 antibodies. Positive staining for MDM2, HDM2ALT1, or p53 is indicated by the red color. The nucleus was visualized by staining with DAPI (blue). Cells were photographed at either 200X or 400X magnification. The images were merged and the merge color (purple) indicates the MDM2, HDM2ALT1 or p53 was localized within the nucleus.

85

Figure 14B: Subcellular localization of HDM2ALT1, MDM2, and p53 at 48 hours after transfection with HDM2ALT1.

86

Figure 14C: Subcellular localization of HDM2ALT1, MDM2, and p53 at 72 hours after transfection with HDM2ALT1.

87

Figure 14D: Subcellular localization of HDM2ALT1, MDM2, and p53 at 96 hours after transfection with HDM2ALT1.

88 3.11 MDM2 and HDM2ALT1 co-localize in the cytoplasm

The subcellular localization of MDM2 and HDM2ALT1 were simultaneously

observed to determine if the two proteins co-localize. At 48 hrs post-transfection with

either pcDNA3 or pcDNA3 ALT1, cells were immunostained with the H221 rabbit

polyclonal antibody and the Ab-6 mouse monoclonal antibody to detect MDM2 and

HDM2ALT1, respectively. The epitope recognized by the H221 antibody spans the central

region of MDM2 that is absent in the HDM2ALT1 protein, thus this antibody exclusively

binds to MDM2 (Table 4). The H221 antibody was also used to detect MDM2 because

the host species of this primary antibody differed from the Ab-6 antibody. Texas red-

conjugated anti-mouse and fluorescein-conjugated anti-rabbit secondary antibodies were

used to visualize the Ab-6 (red) and H221 (green) antibodies, respectively. MDM2 was

localized within the nucleus of mock transfected cells (Figure 16) while HDM2ALT1 was not detected in these cells (Figure 16). Cells transfected with HDM2ALT1 displayed

positive immunostaining for HDM2ALT1 and MDM2 in the cytoplasm (Figure 16). The

merged fluorescent images of MDM2 and HDM2ALT1 indicated that the two proteins co-

localize due to the appearance of the merge color (yellow). 89

Mock HDM2ALT1

Figure 15. HDM2ALT1 co-localizes with MDM2 in the cytoplasm.

After 48 hrs post-transfection with the pcDNA3 ALT1 (HDM2ALT1) or pcDNA3 (Mock), cells were immunostained with the Ab-6 and H221 antibodies. Mock transfected cells displayed positive staining for MDM2 (green) in the nucleus and HDM2ALT1 was not detected. Reciprocally, when HDM2ALT1 was transiently expressed in cells, both HDM2ALT1 (red) and MDM2 (green) were detected in the cytoplasm. The nucleus was visualized by staining with DAPI (blue). The images were merged. The yellow merge color indicated the MDM2 and HDM2ALT1 co-localize in the cytoplasm in HDM2ALT1 transfected cells. The aqua merge color indicated the MDM2 localizes in the nucleus in mock transfected cells. Cells were photographed at 400X magnification. 90 3.12 The presence of HDM2ALT decreases the proliferation rate of NIH 3T3 cells

Sigalas et al. (1996) demonstrated that transient expression of HDM2ALT1 in the

NIH 3T3 cell line increased foci formation. Steinman et al. (2004) also observed several

growth-promoting effects, such as increased proliferation, loss of contact inhibition, and up-regulation of cellular survival factors, after HDM2ALT1 was stably transduced into

NIH 3T3 cells. In contrast, Dang et al. (2002) and Fridman et al. (2003) observed that the transient expression of several murine splice variants of similar size in comparison with

HDM2ALT1 decreased the growth rate of MEFs. Furthermore, HDM2ALT1 has been shown

to stabilize and activate p53, which is also expected to result in an anti-proliferative phenotype (Chandler et al. 2006; Dias et al. 2006; Evans et al. 2001). The expression of

Bax and stabilization and transcriptional activation of p53 by HDM2ALT1 (Figures 10, 11

and 12) were predicted to trigger a p53-dependent inhibition of growth. Thus, the effect

of transient HDM2ALT1 expression on the growth rate of NIH 3T3 cells was investigated.

Growth curves were performed with mock and HDM2ALT1 transfected cells as well as

non-transfected cells. The growth rate of HDM2ALT1 transfected cells was significantly reduced in comparison with mock transfected or non-transfected cells (Figure 22). The

growth rate increased slightly after 78 hrs post-transfection (Figure 22) when the levels of

HDM2ALT1 protein were decreasing (Figure 8). Although fewer cells were present after

54 hrs and later time points in the mock transfected cells when compared with the non-

transfected cells, the growth rates of the mock transfected cells and non-transfected cells

were not significantly different (Figure 22). Visual inspection of the plates containing

mock transfected and non-transfected cells revealed that cells were completely confluent

at 96 hrs while HDM2ALT1 transfected cells were approximately 30-50% confluent. 91

140000 HDM2ALT1 120000 Mock Untreated 100000

80000

60000 Number of cells 40000

20000

0 0 20406080100 Time (hours)

2 HDM2ALT1 y = 192.8x – 4629 R = 0.8224 Mock y = 3556.8x – 197238 R2 = 0.9477 2 Untreated y = 3481.4x – 144983 R = 0.9676

Figure 16. HDM2ALT1 expression decreases the growth rate of NIH 3T3 cells.

Cells were plated in triplicate and transfected with the pcDNA3 ALT1 vector (HDM2ALT1), the empty pcDNA3 vector (Mock) or were not transfected (Untreated). At the various time points indicated, cells were trypsinized and counted on a hemocytometer. The number of cells was reported as an average ± SD. Linear regression was performed to calculate the growth rate of the transfected and non-transfected cells between 54 hrs and 78 hrs post-transfection. The R2 values indicated the goodness-of-fit of the linear regression.

92 An additional cellular viability assay was performed to confirm the decreased

growth rate of NIH 3T3 cells transfected with HDM2ALT1. The highly sensitive CellTiter-

Glo Luminescent Cell Viability Assay was used to determine the relative number of cells

through quantification of the levels of ATP in these cells. After generating a standard

curve to directly correlate the numbers of cells with the luminescence measured, a linear

relationship between the two parameters was established over four orders of magnitude

(data not shown). The rate of proliferation was significantly decreased in the HDM2ALT1

transfected cells when compared with the mock transfected cells (Figure 17). As

observed with the growth rates described in Figure 16, the relative growth rate of

HDM2ALT1 transfected increased slightly after 78 hrs post-transfection (Figure 17) as

HDM2ALT1 protein levels declined (Figure 8). After visually inspecting the mock and the

HDM2ALT1 transfected cells at 90 hrs post-transfection, mock transfected cells were fully

confluent and HDM2ALT1 transfected were approximately 30-50% confluent.

Although the cellular viability assay and the growth curves indicated that

HDM2ALT1 expression decreased the rate of proliferation of NIH 3T3 cells, these assays

cannot distinguish between an inhibition of cellular proliferation and cytotoxicity. To determine whether HDM2ALT1 triggered apoptosis or arrested cellular growth, flow

cytometric cell cycle analysis of transfected cells was performed. HDM2ALT1 expression significantly increased the percentage of cells in the apoptotic sub G0 population and G1

phase (P < 0.01) from 24 hrs up to 96 hrs post-transfection when compared with mock

ALT1 transfected cells. The percentage of HDM2 transfected cells in the S and G2/M phases

(P < 0.01) was also significantly decreased in comparison with mock transfected cells from 24 hrs to 72 hrs post-transfection (Table 6). The percentage of HDM2ALT1 93 transfected cells in the G2/M and S phases did not significantly differ (P > 0.05) from those in mock transfected cells at 96 hrs post-transfection (Table 6). The largest percentage of cells in the sub G0 population was detected at 48 hrs (Table 6) when high levels of HDM2ALT1 protein were observed (Figure 8). The percentages of cell populations in the G1 and sub G0 phases declined at 96 hrs, but were still significantly higher (P < 0.01) in comparison with the mock transfected cells (Table 6). The highest percentage of cells arrested in the G1 phase was observed (60 hrs, Table 6) when the protein levels of HDM2ALT1 were elevated (60 hrs, Figure 8). With the exception of the percentage of mock transfected cells in the sub G0 fraction at 24 hrs (P < 0.01),the percentage of cell populations in the various cell cycle phases of mock transfected cells did not significantly differ with the non-transfected control at any of the time points tested (P > 0.05).

94

8000000

HDM2ALT1 7000000 Mock

6000000

5000000

4000000

3000000

Relative Light Units (RLU) Units Light Relative 2000000

1000000

0 020406080100 Time (hours)

HDM2ALT1 y = 40231x – 823,924 R2 = 0.9558 Mock y = 118324x – 3,000,000 R2 = 0.9891

Figure 17. HDM2ALT1 expression decreases the viability of NIH 3T3 cells

Cells were plated in triplicate and transfected with the pcDNA3 ALT1 vector (HDM2ALT1) or the empty pcDNA3 vector (Mock). At the various time points indicated, the concentration of ATP in cells was quantified with the CellTiter-Glo Luminescent Cell Viability Assay. Luminescence activity was reported as an average ± SD. Linear regression was performed to calculate the relative rate of cellular growth of the transfected cells between 30 and 72 hrs post-transfection. The R2 values indicated the goodness-of-fit of the linear regression.

95

Table 6. Cell cycle analysis of HDM2ALT1 and mock transfected cells.

Time

(hours) Sub G0 (%) G1 (%) S (%) G2/M (%) HDM2ALT1 Mock HDM2ALT1 Mock HDM2ALT1 Mock HDM2ALT1 Mock

0 2.10 ± 0.43a 22.16 ± 0.81a 34.30 ± 0.89a 36.44 ± 1.00 a

24 10.2 ± 0.83b 3.85 ± 0.34d 46.69 ± 0.95b 21.53 ± 0.73c 10.52 ± 0.92b 35.38 ± 0.60c 21.07 ± 1.36b 34.35 ± 0.89c

36 14.01 ± 1.68b 3.83 ± 0.43c 43.30 ± 1.05b 26.48 ± .83c 5.73 ± 0.42b 30.88 ± 1.20c 18.58 ± 0.79b 39.17 ± 1.18c

48 24.00 ± 1.11b 5.87 ± 1.22c 45.40 ± 0.91b 23.05 ± 2.22c 10.37 ± 1.21b 37.92 ± 1.21c 24.12 ± 1.43b 38.50 ± 0.99c

60 18.01 ± 1.39b 3.86 ± 0.71c 53.73 ± 1.59b 21.10 ± 4.34c 6.37 ± 0.56b 31.77 ± 0.90c 18.28 ± 1.47b 40.93 ± 4.45c

72 21.39 ± 1.21b 1.97 ± 0.32c 43.34 ± 1.43b 19.29 ± 1.22c 6.22 ± 0.91b 30.94 ± 1.71c 25.92 ± 2.26b 44.75 ± 5.04c

96 8.00 ± 0.77b 2.26 ± .33c 32.41 ± 0.90b 22.53 ± 2.19c 30.89 ± 5.78e 32.71 ± 5.73c 39.30 ± 6.32e 45.77 ± 2.86c

Note: Cells were plated in triplicate and transfected with the pcDNA3 ALT1 vector (HDM2ALT1), the empty pcDNA3 vector (Mock). Results are reported as averages ± SD. aNon-transfected cells bStatistically significant difference with mock transfected cells at the given time point (P < 0.01, t test). cStatistically insignificant difference with non-transfected cells at the given time point (P > 0.05, t test). dStatistically significant difference with non-transfected cells at the given time point (P < 0.01, t test). eStatistically insignificant difference with mock transfected cells at the given time point (P < 0.01, t test).

96 3.13 HDM2ALT1 expression triggers apoptosis

The cell cycle analysis of cells transfected with HDM2ALT1 indicated that a

significant percentage of cells were in the sub G0 cell population (Table 6). To confirm

that the HDM2ALT1 transfected cells were undergoing apoptosis, the Cell Death Detection

ELISAPLUS assay was utilized. This ELISA-based assay measures the extent of DNA fragmentation by detecting the nucleosomes released into the cytoplasm after Ca2+- dependent endonuclease cleavage of chromatin during the early stages of apoptosis. The levels of apoptosis in the HDM2ALT1 transfected cells were significantly higher than the

background levels of apoptosis detected in the mock transfected cells at 36 hrs. The amount of apoptosis in the HDM2ALT1 transfected cells increased up to 96 hrs post- transfection whereas minimal levels of DNA fragmentation were detected in the mock

transfected cells at all of the time points examined.

The TUNEL assay was also performed to confirm the results that indicated

HDM2ALT1 expression induced DNA fragmentation (i.e., a hallmark of apoptosis) in NIH

3T3 cells. The DeadEnd Fluorometric TUNEL System utilizes enzymatic incorporation

of fluorescein-12-dUTP at the 3’-OH ends of fragmented DNA to visualize and quantify

the amount of fragmented DNA in apoptotic cells. Untreated (non-transfected) NIH 3T3

cells did not display positive staining for DNA fragmentation (data not shown). Only a small number of HDM2ALT1 transfected cells stained positively at 24 hrs post-transfection

(Figure 19A). Beginning at 36 hrs, the number of TUNEL positive cells steadily

increased up to 96 hrs post-transfection with HDM2ALT1 (Figures 19B, C, D, E, and F).

Only a small number of mock transfected cells displayed positive staining for fluorescein

at all of the time points examined (Figures 19A, B, C, D, E, and F). 97

5 HDM2ALT1 4.5 Mock 4

3.5

3

2.5

2

1.5 Abosorbace nm) (405 1

0.5

0 0 102030405060708090100 Time (hours)

Figure 18. Levels of apoptosis in transfected cells.

Cells were plated in triplicate and transfected with the pcDNA3 ALT1 vector (HDM2ALT1) or the empty pcDNA3 vector (Mock). At the various time points indicated, the levels of DNA fragmentation were measured with the Cell Death Detection ELISAPLUS assay. Absorbance values were reported as an average ± SD.

98

Figure 19A. TUNEL staining of fragmented DNA at 24 hours post-transfection.

Cells were transiently transfected with pcDNA3 ALT1 or pcDNA3. Positive staining for DNA fragmentation is indicated by the green color. The nuclei were visualized by staining with DAPI (blue). Cells were photographed at 200X magnification. The images were merged and the intensity of the merge color (aqua) indicates extent of apoptosis in cells.

99

Figure 19B. TUNEL staining of fragmented DNA at 36 hours post-transfection.

100

Figure 19C. TUNEL staining of fragmented DNA at 48 hours post-transfection.

101

Figure 19D. TUNEL staining of fragmented DNA at 60 hours post-transfection.

102

Figure 19E. TUNEL staining of fragmented DNA at 72 hours post-transfection.

103

Figure 19F. TUNEL staining of fragmented DNA at 96 hours post-transfection. 104

Chapter 4: Discussion

105 The p53 tumor suppressor protein is the central regulatory node in the stress

response pathways. Depending on the type and extent of the stress conditions, p53 activates numerous factors that repair or eliminate the damaged cell to maintain genomic integrity. Activation of the p53 network requires stabilization of the p53 protein. Post-translational modifications of p53 and HDM2 stabilize and activate the p53 protein primarily through disruption of the p53-HDM2 interaction (Appella and

Anderson 2001; Bode and Dong 2004). However, a number of reports have indicated

that additional regulatory mechanisms other than covalent modifications of these two proteins down-regulate MDM2 or HDM2 activity in response to cellular stresses

(Momand et al. 1992; Trinh et al. 2001; Wu and Levine 1997; Zhu et al. 2002). For example, Dias et al. (2006) and Chandler et al. (2006) demonstrated that genotoxic agents (e.g., high-energy radiation and chemotherapeutic drugs) reduced the levels of

HDM2 and MDM2 protein due to decreased levels of the respective transcripts. These authors showed that the decline in the levels of hdm2 and mdm2 transcripts was caused by a shift in the post-transcriptional processing of the hdm2 and mdm2 pre-mRNA to predominately produce the hdm2ALT1 and mdm2-b transcripts, respectively. The

alterative splicing of the hdm2 and mdm2 gene products in response to certain cellular

stresses represents a novel post-transcriptional mechanism that up-regulates p53

activity. These results also support a model proposed by Kastan and colleagues that

indicated DNA damage stabilized p53 through an unidentified post-transcriptional

mechanism (Kastan et al. 1991). Although genotoxic stresses induced production of the

hdm2ALT1 and mdm2-b transcripts, which concomitantly decreased the levels of the 106 hdm2 and mdm2 transcripts and proteins, the functions of the putative protein

products of these alternatively spliced transcripts were not examined by these authors.

Evans et al. (2001) and Steinman et al. (2004) investigated the effects of

HDM2ALT1 and MDM2-B proteins in animal and cell culture models, yet reported

conflicting phenotypes: p53-dependent growth inhibition and p53-independent growth

promotion, respectively. Due to the contradictory effects of HDM2ALT1 expression in

dissimilar experimental systems (i.e., different cell lines and transient versus stable

expression of the alternatively spliced transcripts), a unifying model of HDM2ALT1’s

function was proposed to resolve these incongruous results. DNA damage alters the

splicing pattern of the hdm2 or mdm2 pre-mRNA to generate hdm2ALT1 or mdm2-b

transcripts, correspondingly. Due to a decline in the levels of full-length mdm2 and

hdm2 transcripts, the levels of the respective proteins are reduced, which, in turn,

diminishes the negative regulation of p53’s stability and activity. Subsequent

translation of the alternatively spliced transcripts produces the HDM2ALT1 and MDM2-

B proteins, which further disrupt the interaction of HDM2 or MDM2, respectively, with

the p53 protein. The resulting stabilization of the p53 protein activates the p53 stress

response network, leading to phenotypic outputs, such as DNA repair, cell cycle arrest,

or apoptosis. Once the response to the stress-induced damage is complete, alternative

splicing of the hdm2 and mdm2 pre-mRNA ceases and the normal pattern of post-

transcriptional processing resumes. Aside from their role in stabilizing p53, HDM2ALT1 and MDM2-B potentially have physiological functions in other biological processes, such as cellular survival and growth (Steinman et al. 2004). Notably, HDM2ALT1 and

MDM2-B retain several functional domains of HDM2 and MDM2, respectively, that 107 regulate other cellular mechanisms independent of p53 and under non-stressed

conditions (Ganguli and Wasylyk 2003). Although the growth-promoting effects of

HDM2ALT1 and MDM2-B may be physiologically relevant, the role of these p53-

independent mechanisms was not examined in the present study.

The initial focus of this study was to investigate the phenotypic effects of

HDM2ALT1 expression in vivo. The aim of this experimental approach was to generate a

transgenic line that possessed all of the regulatory components of the Tet-On system

controlling the inducible expression of HDM2ALT1. A transgenic line containing the tTS

repressor was successfully established. However, repeated efforts to generate transgenic

lines harboring the rtTA and Tet-O hdm2ALT1 constructs were unsuccessful.

One of the limitations of the Tet-On system is the potential toxicity of the rtTA

protein. Overexpression of the rtTA transactivator protein, which contains the potent

VP16 transcriptional transactivation domain, induced transcriptional squelching as a

result of competition for the limited components of the basal transcriptional machinery

(Baron et al. 1997; Gallia and Khalili 1998). Furthermore, expression of the rtTA

transactivator protein alone has been demonstrated to affect cellular proliferation and to

induce various morphological changes (Gallia and Khalili 1998). The observed side

effects of the rtTA protein may explain why transmission of the rtTA transgene stopped

after two generations (Table 6). Although fertile, the rtTA founders transmitted the

transgene to offspring at a very low rate (<10%). This could indicate that expression of

the rtTA transgene impinged upon gametogenesis. Chromosomal rearrangements

during meiosis at the site of integration of the rtTA transgene in the F2 generation may

have resulted in enhanced expression of the rtTA protein. As high levels of the rtTA 108 protein have been shown to cause transcriptional squelching, these potential lethal

effects on the germ cells could have rendered the F2 mice sterile. Repeated attempts to

breed the remaining rtTA founders and F1 rtTA transgenic mice with C57Bl/6J mice

likely failed as these mice were well past the optimal breeding age (>12 months).

Another limitation of the Tet-On system is transcriptional leakiness of the

artificial rtTA-responsive promoter (Freundlieb et al. 1997; Gossen et al. 1993; Gossen and Bujard 1992). Although this minimal promoter has been engineered to lack the strong enhancer elements of the CMV immediate early promoter (Gossen and Bujard

1992), the presence of nearby enhancers can powerfully induce transcription from this

artificial promoter even when the rtTA transactivator protein is not bound to the TRE

site (Freundlieb et al. 1999). Therefore, depending upon the site of integration of this

doxycycline-responsive construct, high levels of uninduced transcriptional transgene

activation may result (Freundlieb et al. 1997; Freundlieb et al. 1999; Gossen et al.

1993; Gossen and Bujard 1992). While certain levels of background expression are

considered acceptable in some experimental systems, the transcriptional leakiness is not

tolerated when the transgene product is toxic (Lee et al. 1998). In these instances, even

extremely low levels of background expression can prevent the establishment of stable

transgene expression in animal or cell culture models. These findings may explain the

low efficacy of transgenesis associated with the Tet-O hdm2ALT1 construct (3.08%,

Table 6); the staff at the Edison Biotechnology Institute Transgenic Facility reported an

average transgenesis efficacy of 12%. The low number of founders possessing the Tet-

O hdm2ALT1 construct may have resulted from many of the microinjected zygotes

undergoing apoptosis in the early stages of embryogenesis. This embryonic lethality 109 possibly resulted from the widespread background expression of HDM2ALT1

enhancing p53 activity; unrestricted p53 activity during gestation has been shown to

result in an embryonic lethal phenotype (Jones et al. 1995; Montes de Oca Luna et al.

1995). Steinman et al. (2004) also indicated that constitutive expression of MDM2-B,

the murine homologue of HDM2ALT1, was lethal during embryonic development, although the authors did not investigate the underlying cause of the lethality. The

toxicity of constitutive MDM2-B expression throughout the developing embryo may

have resulted from MDM2-B’s activation of p53. Two viable Tet-O hdm2ALT1 founders

were generated (Table 6); however, the HDM2ALT1 protein was not detected in any of

the tissues examined (data not shown) in the F1 generation, indicating that the Tet-O

hdm2ALT1 construct integrated either into a silent chromosomal locus or into a site not

adjacent to strong enhancer elements. The explanation of why the F2 generation of the

rtTA transgenic mice stopped transmitting the rtTA transgene may also account for the

ALT1 similar effect observed in the Tet-O hdm2 transgenic mice (i.e., the F1 generation of

the Tet-O hdm2ALT1 mice did not transmit the transgene to offspring); chromosomal rearrangements of the Tet-O hdm2ALT1 locus during meiosis may have increased

HDM2ALT1 expression in the developing gametes. Elevated levels of p53 protein in the gametes may inappropriately trigger apoptosis, although no studies, to date, have

formally evaluated this hypothesis. Another possible explanation for the failure of

transgene transmission from the F1 generation could be explained by the effects of genetic modifiers in the C57Bl/6J background. The zygotes used for microinjection of the Tet-O hdm2ALT1 construct consisted of the mixed SJL and C57BL/6J genetic

backgrounds. Transgenic founders were then bred with mice from the pure C57BL/6J 110 background to minimize the potential phenotypic effects caused by genetic modifiers

in the SJL genetic background. However, the increased percentage of the C57BL/6J

genetic background may have unmasked the phenotypic effects of certain genetic

modifiers in this background. These modifiers, in cooperation with basal levels of

uninduced HDM2ALT1 expression, may have exacerbated the potential toxic effects of

ALT1 HDM2 expression during gestation or in the germ line of the F1 generation. Several

attempts to reestablish the Tet-O hdm2ALT1 transgenic lines by mating founders and

C57BL/6J mice likely failed due to the old age (>15 months) of these mice.

After efforts to generate transgenic mice possessing the regulatory components

of the Tet-On system failed, the focus of this study shifted to examine the effects of

inducible HDM2ALT1 expression in a cell culture model. The NIH 3T3 cell line was chosen for this experimental model because both of the conflicting phenotypes of

MDM2-B expression were observed in this cell line (Chandler et al. 2006; Steinman et

al. 2004). Because an anti-HDM2ALT1 antibody that does not cross-react with HDM2 is

currently unavailable, human cell lines could not be used to exclusively detect the

HDM2ALT1 protein due to the presence of endogenous HDM2. Because NIH 3T3 cells

express MDM2, the use of a species-specific antibody that recognizes both HDM2 and

HDM2ALT1 would only detect the HDM2ALT1 protein in these cells. Furthermore, the

high degree of homology between HDM2ALT1 and MDM2-B indicated that they will

have similar functions (Steinman et al. 2004). The Tet-On system components were

stably integrated into NIH 3T3 cells to control inducible HDM2ALT1 expression. In the

presence of the rtTA transactivator alone, there were significant levels of uninduced

luciferase expression (Figure 7). However, when the tTS repressor protein was 111 introduced into the rtTA stable cell line, the levels of uninduced expression were significantly decreased to almost undetectable levels (Figure 7). This rtTA/tTS double

stable cell line displayed high induction and low background expression. However,

when the Tet-O hdm2ALT1 construct was stably introduced into the double stable cell

line, no induction of luciferase activity was detected after the addition of doxycycline.

Several groups have reported difficulties in establishing stable cell lines with the

tetracycline inducible gene expression systems due to the cytotoxicity associated with

high expression of the transactivator protein (Baron et al. 1997; Gallia and Khalili

1998; Kenny et al. 2002). However, the rtTA protein did not cause any apparent

toxicity when stably expressed in the NIH 3T3 cells. Several possibilities could explain

the lack of induction in the stable Tet-O hdm2ALT1 clones. First, the Tet-O hdm2ALT1

construct may have integrated into a silent chromosomal locus. However, considering

the number of stable clones that were evaluated (>450), the probability of this event

occurring in all of the clones is highly unlikely. The experimental approach utilized for

stable transfection may provide another explanation for the lack of induction of

luciferase. The Tet-O hdm2ALT1 construct did not possess a eukaryotic antibiotic

selection marker, thus a plasmid containing the puromycin expression cassette

(Clontech, Mountain View, CA) was co-transfected with the Tet-O hdm2ALT1 construct to facilitate positive selection. Because two separate plasmids were used for stable transfection procedure, the probability that the puromycin-resistant clones that contained both constructs stably integrated was likely less than those that contained the puromycin expression cassette alone. In addition, when the Tet-O hdm2ALT1 construct

was initially introduced into the cells by liposome-mediated transfection, many of these 112 constructs would not have integrated into the genome. As a result, high levels of

uninduced HDM2ALT1 expression may have been generated by the non-integrated Tet-O hdm2ALT1 constructs (Sambrook and Russell 2001). Moreover, overexpression of the

rtTA protein in these cells may have also contributed to uninduced HDM2ALT1

expression. The resulting elevated levels of HDM2ALT1 might have triggered apoptosis

in these transfected cells while cells possessing only the puromycin expression cassette

would not be affected. Thus, the stable clones surviving puromycin selection may have

only possessed the plasmid containing the puromycin expression cassette.

As attempts to control HDM2ALT1 expression with Tet-On system failed, a different experimental approach was undertaken to examine HDM2ALT1’s function.

Transient expression of the HDM2ALT1 protein in NIH 3T3 cells was successful and

resulted in a growth inhibitory phenotype (Figures 16, 17, 18, and 19, Table 6).

Although the presence of HDM2ALT1 did not affect the protein levels of endogenous

MDM2, HDM2ALT1 expression prevented MDM2 from binding to p53 (Figures 8, 9,

and 10). The ability of HDM2ALT1 to disrupt MDM2’s association with p53 resulted in

sequestration of MDM2 in the cytoplasm while p53 was free to enter the nucleus

(Figures 14 and 15). The diminished p53-MDM2 interaction also resulted in increased

protein levels of p53 (Figure 11). The interaction of HDM2ALT1 (human protein) with

MDM2 (murine protein) suggests that the function of the HDM2ALT1 and MDM2-B

alternative spliced products are evolutionarily conserved between humans and mice.

Intriguingly, disruption of the p53-MDM2 interaction by HDM2ALT1 alone was

sufficient to stimulate p53’s transcriptional functions in the absence of any stress signal

(Figure 12). Other groups reported a similar effect when the p53-MDM2 interaction 113 was disrupted in various cell lines and in vivo (Bottger et al. 1997a; Bottger et al.

1997b; Bottger et al. 1996; Mendrysa et al. 2003; Vassilev et al. 2004). Thus, additional regulatory mechanisms aside from the post-translational modifications mediated by the upstream effectors of the stress response are capable of activating the transcription-dependent functions of p53. In addition, other unknown functions of

HDM2ALT1, aside from its disruption of the p53-MDM2 interaction, may stimulate the

transcriptional activity of p53. Due to the activation of p53’s transcriptional activity by

HDM2ALT1 (Figure 12), the elevated levels of Bax were presumed to be a result of p53-

dependent transcriptional activation of the bax gene (Figure 11); however, the

possibility of transcription-independent activation of Bax by p53 or other regulatory

factors cannot be excluded (Baptiste and Prives 2004).

The decreased growth rate (Figures 16 and 17) of HDM2ALT1 transfected cells was

predicted to be a result of the increased levels of Bax protein (Figure 11), which, when

properly activated, triggers the release of cytochrome c from mitochondria and induces the intrinsic apoptotic cascade (Degli Esposti and Dive 2003). The detection of apoptosis between 36 and 96 hrs post-transfection (Figure 18 and 19, Table 6) when elevated levels of Bax protein were detected (Figure 11) supports this prediction. Although overexpression of Bax alone has been shown to induce apoptosis (Bargou et al. 1996;

Fortuno et al. 1998; Guo et al. 2000; Pastorino et al. 1998; Sawada et al. 2000; Zheng et al. 2005), the amount of Bax protein detected when HDM2ALT1 was expressed likely

represents endogenous levels of Bax upon physiological induction by p53. Because

activation of Bax requires Bid-induced conformational change and mitochondrial translocation to initiate the apoptotic cascade (Desagher et al. 1999; Eskes et al. 2000; 114 Makin et al. 2001; Wang et al. 1996), HDM2ALT1 expression may have increased p53-

dependent transcriptional activation of the bid gene to induce Bax-dependent apoptosis.

However, the expression of Bid in HDM2ALT1 transfected cells was not examined. Aside

from activation of the Bax protein, other pro-apoptotic effectors may be responsible for

the induction of apoptosis when HDM2ALT1 was transiently expressed.

In summary, transient expression of HDM2ALT1 in NIH 3T3 cells triggered a p53- dependent growth inhibition. The HDM2ALT1 protein reduced the activity of the MDM2

protein by sequestering MDM2 in the cytoplasm. As a result, the levels of p53 protein

increased. Through an unknown mechanism, p53 became transcriptionally active when

HDM2ALT1 was present. HDM2ALT1 expression also decreased the rate of proliferation

and induced apoptosis, possibly via activation of Bax. Altogether, these findings strongly

suggest that HDM2ALT1 expression may represent a novel molecular process by which the

p53-MDM2 interaction is disrupted. The results presented throughout this study support

the proposed model of HDM2ALT1’s physiological functions during conditions of cellular

stress. 115

Chapter 5: Future studies

116 Several interesting questions have arisen from this study. 1) What is the

significance of the growth-promoting effects of HDM2ALT1 observed in certain

experimental systems? 2) Is induction of the upstream mediators of the stress response

required for activation of p53’s transcription-dependent and -independent activities?

3) Does HDM2ALT1 expression during embryogenesis cause lethality as observed with

MDM2-B? 4) Does HDM2ALT1 expression prevent the growth of tumors?

1) Do the growth-promoting effects of HDM2ALT1 expression have a physiological role? The contradictory growth-inhibitory and growth-promoting effects of MDM2 expression are also observed with HDM2ALT1 expression (Evans et al. 2001;

Steinman et al. 2004). HDM2ALT1 may bind to some of the proteins that associate with

HDM2 or exhibit novel interactions. For instance, the HDM2ALT1 protein retains

several domains (e.g., zinc and RING finger) of the HDM2 protein that are involved in

other cellular processes independent of p53. Thus, these unidentified interactions of

HDM2ALT1 could explain the growth-promoting effects of HDM2ALT1 expression

observed in certain experimental systems (Steinman et al. 2004). The proposed model

of HDM2ALT1 expression in this study predicts that these growth-stimulatory effects are

a result of inappropriately sustained expression as well as high concentrations of the

HDM2ALT1 protein. Overexpression of HDM2ALT1 may result in non-specific protein

interactions that inadvertently stimulate cellular proliferation. However, the timing of

HDM2ALT1 expression might also explain these protein interactions. Certain covalent modifications or unknown mechanisms may modify HDM2ALT1 to interact with other

proteins under different cellular conditions. The yeast two-hybrid screening could be

used to identify the other proteins that interact with HDM2ALT1. Plasmids encoding the 117 HDM2ALT1 protein fused in frame to the GAL4 DNA binding domain (e.g., bait

plasmid ) and human cDNA library inserts fused in frame to the GAL4 activation

domain (e.g., prey plasmid) would be generated. An appropriate yeast reporter strain would be transformed with the bait plasmid. These transformants would be mated with a yeast strain that has undergone a library-scale transformation. Western blot analysis would confirm the expression of the bait fusion protein. The strength of the protein interaction would be scored based on the levels of β-galactosidase activity. Prey

plasmids from positive colonies would be sequenced to determine the identity of the

protein(s) that interacted with the HDM2ALT1 protein.

2) How is p53 activated in the absence of a stress signal? Post-transcriptional

modifications have been predominately thought of as the key determinants of p53’s transcriptional activation. However, these conclusions have been mostly based on

transfection and biochemical studies involving nonnative conditions. Recent in vivo

evidence has shown that certain covalent modifications within the C-terminus of p53

previously thought to be required for its activation of target downstream genes (Appella

and Anderson 2001; Xu 2003) were not essential to stimulate p53’s transcriptional

activity (Krummel et al. 2005). Mutations in the serine and threonine residues that

prevent phosphorylation of p53 only marginally affected the activation of p53 (Levine et

al. 2006). Disruption of the p53-HDM2 interaction alone was sufficient to induce the full

tumor suppressive effects of p53 (Bottger et al. 1997a; Bottger et al. 1997b; Bottger et al.

1996). For example, the Nutlins, which exclusively disrupt the p53-HDM2 interaction, potently activated p53 in vivo in the absence of any stress signal (Vassilev 2004).

Moreover, reduced levels of endogenous MDM2 correlated with increased transcriptional 118 activity of p53 without exposure to any stress-induced damage (Mendrysa et al.

2003). The authors also showed that the lowered mdm2 expression alone decreased the

rate of cellular proliferation in several tissues (Mendrysa et al. 2003). These observations

challenge the hypothesis that only post-translational modifications of p53 are required for

its stabilization and activation. Therefore, other regulatory processes aside from post-

translational modifications may be necessary to stabilize and activate the p53 protein.

Uncovering these mechanisms will also further validate therapeutic approaches designed to restore p53 activity in tumors by disrupting the p53-HDM2 interaction. The potent transcriptional activation of p53 in the absence of stress-induced damage may also explain its apparent toxicity with the Tet-On system in vivo. The relatively small number of transgenic founders generated may have been caused by the leakiness of the Tet-O hdm2ALT1 construct, which would have elevated p53 activity during embryogenesis and

ultimately resulted in lethality. To circumvent this problem in vivo, the Tet-O hdm2ALT1 construct could be microinjected into embryos from the established tTS transgenic line.

Because tTS powerfully suppresses the transcriptional activation controlled by the TRE site, this should eliminate the transcriptional leakiness of Tet-O hdm2ALT1 construct and

increase the number of transgenic founders born. Once a viable transgenic line with

HDM2ALT1 under the control of the Tet-On system is established, the activation of p53 by

HDM2ALT1 in the absence of stress-induced damage could be examined in vivo. Analysis

of the post-translational modifications of p53 in cells transiently transfected with

HDM2ALT1 could determine which, if any, modifications are necessary for activation of

its transcriptional function. Examining the mRNA expression levels of several other p53- 119 target genes would determine whether HDM2ALT1 exclusively stimulated p53-

dependent transcription of the bax gene or if other p53-repsonsive genes were activated

by p53.

3) Does HDM2ALT1 expression have an effect on embryogenesis? Constitutive

expression of MDM2-B (the murine counterpart of HDM2ALT1) caused embryonic

lethality (Steinman et al. 2004). Unrestricted p53 activity has been shown to cause a

lethal phenotype during embryogenesis (Chavez-Reyes et al. 2003; Jones et al. 1995;

Langheinrich et al. 2002; Montes de Oca Luna et al. 1995). As the HDM2ALT1 protein

has been shown to block MDM2 activity, constitutive expression of HDM2ALT1 in presence of the wild-type p53 allele during embryogenesis is also expected to be embryonic lethal. Concomitant loss of p53 function in the absence of endogenous

MDM2 activity is predicted to rescue this lethal phenotype as observed with the viable p53-/- mdm2-/- mice (Jones et al. 1995; Montes de Oca Luna et al. 1995). This

hypothesis can be tested by crossing a transgenic line that expresses HDM2ALT1 under

the control of the components of the Tet-On system with 129-Trp53tm1Tyj p53-/- mice.

Doxycycline would be administered in the drinking water of pregnant females and

cross the placental barrier to induce HDM2ALT1 expression only in the developing

embryos. Pregnant females would lack HDM2ALT1 expression to avoid any maternal

effects of its expression on the developing embryos. Of the progeny generated by the

cross of HDM2ALT1 inducible mice and 129-Trp53tm1Tyj p53-/- mice, only HDM2ALT1

inducible p53-/- mice and those lacking the components of the Tet-On System would be expected to survive. If HDM2ALT1 expression increases p53 activity, then the

HDM2ALT1 inducible p53+/- embryos would die during gestation due to uncontrolled 120 p53 activity. The lethality of these pups would be detectable due to the alteration of the Mendelian ratios of pups born. The genotype of viable pups would be confirmed by

PCR analysis.

4) Is p53’s ability to suppress tumor growth increased by HDM2ALT1 expression? In BALB/c mice, which are predisposed to mammary tumors, the p53-null allele increases the incidence of breast carcinomas to 55%, indicating that a slight decrease in p53 levels accelerates mammary tumor development (Kuperwasser et al.

2000; Ullrich et al. 1996). As HDM2ALT1 expression leads to increased p53 activity and stability, HDM2ALT1 expression should decrease the frequency of mammary tumors or delay the onset of tumors in these mice. To test this hypothesis, a transgenic line with

HDM2ALT1 expression under the control of the Tet-On system would be crossed with

BALB/c p53-/- females. Sublethal exposure (5 Gy) to ionizing radiation has been shown to decrease the time to mammary tumor development in BALB/c x C57Bl/6J F1 hybrid mice (Blackburn et al. 2003), thus a similar expedited effect is expected when

HDM2ALT1 inducible p53+/- mice are similarly irradiated. HDM2ALT1 expression would then be induced by administering doxycycline in the drinking water of mice post- irradiation. If HDM2ALT1 expression increases p53 activity, the frequency of mammary tumors is expected to decline when compared with uninduced controls.

121

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173

Appendix

174 List of Abbreviations

AKT: Protein kinase B

Apaf-1: Apoptotic protease activating factor 1

AP1-ETS: Activator protein 1 – ETS transcription factors

AMPK: AMP-activated protein kinase

Ampr: Ampicillin resistance gene

ATF3: Activating transcription factor 3

ATM: Ataxia telangiectasia mutated protein

ATR: Ataxia telangiectasia related protein

AURKA: Aurora kinase

B99: G2 and S phase expressed protein 1

BAK: Bcl2-antagonist/killer 1

Bax: Bcl2-associated X protein

β-globin poly A: Beta globin polyadenylation signal sequence

BGH pA: Bovine growth hormone polyadenylation signal sequence

BGS: Bovine growth serum

BRCA1: Breast cancer early onset protein 1

BTG2/Tis21: B-cell translocation gene 2

Casp3: Caspase 3

Casp8: Caspase 8

Casp9: Caspase 9

CBP: CREB binding protein

Cdc2: Cell division control protein 2 175 Cdc6: Cell division control protein 6

Cdc25: cell division control protein 25

CDK: Cyclin dependent kinase

Cdk2: Cyclin-dependent kinase 2

Cdk4: Cyclin-dependent kinase 4 cDNA: Complementary DNA

CHK1: Checkpoint kinase 1

CHK2: Checkpoint kinase 2

CK2: Casein kinase 2

CMV: Cytomegalovirus

ColE1: ColE1 replication origin

COP1: Constitutive photomorphogenic protein 1

CRM1: Chromosome region maintenance protein 1 (exportin 1)

Cyto C: Cytochrome C

DBD: DNA-binding domain

DCK: Cyclin Dependent Kinase1

DDB2: Damage-specific DNA binding protein 2

DMEM-RS: Dulbecco’s Modified Eagle Medium Reduced Serum

DNA: Deoxyribonucleic acid

DNA-PK: DNA-dependent protein kinase dNTPs: Deoxyribonucleotide triphosphates

DR5: Death receptor 5

DGKA: Diacylglycerol kinase alpha 176 DUSP5: Dual specificity phosphatase 5

EDN2: Endothelin 2

EDTA: Ethylenediaminetetraacetic acid

ELISA: Enzyme-linked immunosorbent assay

ERCC5: Excision repair cross-complementing rodent repair deficiency, complementation group 5

ERKs: Extracellular signal-regulated kinases

Eμ-myc: c-myc oncogene under control of the immunoglobulin heavy chain enhancer

F1 ori: Filamentous phage origin of replication

FACT: Facilitates chromatin transcription complex

FHC: Immortalized, nontumorigenic human colonic epithelial cell line

Fluorescein-12-dUTP: fluorescein-6-carboxaminocaproyl-[5-(3-aminoallyl)-2'- deoxyuridine-5'-triphosphate]

GADD45: Growth arrest and DNA-damage-inducible protein 45

GADD45β: Growth arrest and DNA-damage-inducible protein 45 beta

GD-AiF: Glioblastoma derived – angiogenesis inhibitor factor

GOS1: v-SNARE protein involved in Golgi transport

GSK3β: Glycogen synthase kinase 3 beta

H1299: Lung carcinoma cell line

HAUSP: Herpes virus-associated ubiquitin-specific protease

HDAC1: Histone deacetylase 1

HDM2: Human orthologue of murine double minute 2 protein hdm2: Human orthologue of murine double minute gene or RNA 177 HDM2ALT1: Human double minute 2 alternately spliced protein isoform hdm2ALT1: Human double minute 2 alternately spliced mRNA transcript

HIPK2: Homeodomain interacting protein kinase 2

HMG1: High mobility group 1 protein

HSC: Hematopoietic stem cells

HPV: Human papilloma virus

IGF-BP3: Insulin like growth factor-binding protein 3

IP: Immunoprecipitation

JNK: c-jun amino-terminal kinases

KAI: Tumor metastasis suppressor gene kb: Kilobases kbp: Kilobase pairs kDa: Kilodaltons

KRAB-AB: Kruppel associated box A and B domain

LIG1: DNA ligase 1

LOXL1: Lysyl oxidase-like 1

MAPK: Mitogen-activated protein kinase

MAPKAPK2: Mitogen-activated protein kinase-activated protein kinase 2

MASPIN: Mammary serine protease inhibitor

MCF-7: Epithelial mammary adenocarcinoma cell line

MDM2: Murine double minute 2 protein

MDMX: Murine double minute 4 protein mdm2: Murine double minute gene or RNA 178 MEF: Mouse embryo fibroblasts

MIHB: Apoptosis protein 1

MIHC: Apoptosis protein 2

Mv1Lu: Mink lung epithelial cell line mSin3a: Homolog of transcriptional regulator SIN3A

MTBP: Microtubule binding protein

Nbs1: Nijmegen breakage syndrome 1

Nedd8: Neural precursor cell expressed, developmentally down-regulated 8

NES: Nuclear export signal

NF-κB: Nuclear factor of kappa light chain gene enhancer in B-cells

NIH 3T3: Immortalized murine embryonic fibroblast cell line

NL-20: Immortalized, nontumorigenic human bronchial epithelial cell line

NLS: Nuclear localization signal

NO: Nitric oxide

P53AIP: p53 regulated apoptosis inducing protein-1 p53-R2: p53-inducible ribonucleotide reductase

PAG 608: p53-activated gene 608

PAI: Plasminogen activator inhibitor-1

PBS: Phosphate buffered saline

PBS-T: Phosphate buffered saline with 0.1% Tween 20

PCAF: p300/CBP-associated factor

PCD2: Programmed cell death 2 pCMV: Immediate-early promoter cytomegalovirus 179 PCNA: Proliferating cell nuclear antigen

PCR: Polymerase chain reaction

PERP: p53 apoptosis effector related to Pmp22

PI3K: Phosphoinositide-3 kinase

PIAS-1: Protein inhibitor of activated STAT

PID: p53 target protein in the deacetylase complexes

PIDD: p53 induced death domain protein

PIGs: p53-induced genes

PIRH-2: p53-induced ubiquitin protein ligase 2

PKC: Protein kinase C

PKR: Protein kinase R

PLK: Polo-like kinase

PML: Promyelocytic leukemia protein

PolyA: Polyadenylation signal sequence

PP2A: Protein phosphatase 2A

PRE: p53-responsive DNA element

PISSLRE: Ets-like gene and cdc-related kinase

PTEN: Phosphatase and tensin homolog

PUMA: Bcl-2 binding component 3

RecA: Homologous DNA recombination protein

REDD1: DNA-damage-inducible transcript 4

RFC4: Replication factor C (activator 1) 4

RPA1: Replication protein A1 180 RKO: Colonic carcinoma cell line

ROS: Reactive oxygen species rNTP: Ribonucleoside triphosphates

Set9: SET domain containing lysine methyltransferase 9

SD: Standard deviation

SDS: Sodium dodecyl sulfate

Siah-1: Seven in absentia homolog 1

Sir-2: Silent information regulator 2

SIRT-1: Silent mating type information regulation 2 homolog 1

SJSA-1: Osteosarcoma cell line

SNP: Single nucleotide polymorphism

SUMO1: Small ubiquitin-like modifier 1

SV40: Simian vacuolating virus 40

TAF1: TBP-associated transcription factor 1

TAFII250: TBP-associated transcription factor 250

TBP: TATA box binding protein

TFIIE: Transcription factor IIE

TGF-α: Transforming growth factor alpha

TGFβ1: Transforming growth factor beta 1

TRAF4: TNF receptor associated factor 4

TRE: Seven tandem tet-O DNA elements

TSAP6: Tumor suppressor activated pathway 6

TSC2: Tuberous sclerosis protein 2 181 TSP-1: Thrombospondin-1

TOPO-IIα: Topoisomerase II alpha

Topors: Topoisomerase I binding, arginine/serine-rich SUMO ligase

TUNEL: TdT-mediated dUTP Nick-End Labeling

Ubc9: Ubiquitin-conjugating enzyme 9

UbcH5B/C: E2 ubiquitin-conjugating enzyme

U2OS: Human osteosarcoma tumor cell line uORF: Upstream open reading frame

UV: Ultraviolet radiation

Wig1: Wild-type p53-induced gene 1

Wip1: Wild-type p53-induced phosphatase 1

WRN: Werner syndrome protein

XRCC5: Fanconi anemia, complementation group G