G2 Phase Cell Cycle Regulation by E2F4 Following Genotoxic Stress
by
MEREDITH ELLEN CROSBY
Submitted in partial fulfillment of the requirements for the Degree of
Doctor of Philosophy
Thesis Advisor: Dr. Alex Almasan
Department of Environmental Health Sciences
CASE WESTERN RESERVE UNIVERSITY
May, 2006 CASE WESTERN RESERVE UNIVERSITY
SCHOOL OF GRADUATE STUDIES
We hereby approve the dissertation of
______
candidate for the Ph.D. degree *.
(signed)______(chair of the committee)
______
______
______
______
______
(date) ______
*We also certify that written approval has been obtained for any proprietary material contained therein. TABLE OF CONTENTS
TABLE OF CONTENTS………………………………………………………………….1
LIST OF FIGURES……………………………………………………………………….5
LIST OF TABLES………………………………………………………………………...7
ACKNOWLEDGEMENTS……………………………………………………………….8
LIST OF ABBREVIATIONS……………………………………………………………10
ABSTRACT……………………………………………………………………………...15
CHAPTER 1. INTRODUCTION
1.1. CELL CYCLE REGULATION: HISTORICAL OVERVIEW………………...17
1.2. THE E2F FAMILY OF TRANSCRIPTION FACTORS……………………….22
1.3. E2F AND CELL CYCLE CONTROL
1.3.1. G0/G1 Phase Transition………………………………………………….28
1.3.2. S Phase…………………………………………………………………...28
1.3.3. G2/M Phase Transition…………………………………………………..30
1.4. CELL CYCLE MISREGULATION BY GENOTOXIC
STRESS: IONIZING RADIATION
1.4.1. The cellular response to ionizing radiation (IR) ….…………………….32
1.4.2. DNA damage response: cell cycle arrest versus apoptosis……………...35
1.4.3. Cell cycle specific arrest mechanisms………………………………...... 37
1.5. OBJECTIVES AND THESIS OVERVIEW……………………………………38
1 CHAPTER 2. PHYSIOLOGIC TARGETS OF P53 IDENTIFIED THROUGH
CHROMATIN IMMUNOPRECIPITATION (CHIP)
2.1. ABSTRACT……………………………………………………………………..42
2.2. INTRODUCTION………………………………………………………………43
2.3. MATERIALS AND METHODS
2.3.1. Cell culture and treatment…………………………………………………45
2.3.2. MOLT-4 expression profiling with cDNA arrays………………………...46
2.3.3. Chromatin Immunoprecipitation Assay (ChIP) …………………………..48
2.4. RESULTS
2.4.1. Expression profiling identifies radiation responsive genes………………..52
2.4.2. Identification of p53 binding sites in radiation-responsive
genes implicated in apoptosis and cell cycle control………………………55
2.4.3. Chromatin immunoprecipitation (ChIP) of p53 with regulatory
regions of radiation-responsive genes implicated in apoptosis
and cell cycle control………………………………………………………57
2.5. DISCUSSION…………………………………………………………………...64
CHAPTER 3. BEYOND P53: THE OPPOSING ROLES OF E2FS IN CELL CYCLE
PROLIFERATION AND DEATH
3.1. ABSTRACT……………………………………………………………………..68
3.2. INTRODUCTION………………………………………………………………68
2 3.3. E2F1: A MULTITASKING MEDIATOR………………………………………70
3.4. E2F4: A REMARKABLE REPRESSOR……………………………………….72
3.5. CONCLUSIONS……………………………………………………………..…75
CHAPTER 4. THE ROLE OF E2F4 IN PROMOTING THE G2 ARREST RESPONSE
TO IR IN PROSTATE CARCINOMA
4.1. ABSTRACT……………………………………………………………………..77
4.2. INTRODUCTION………………………………………………………………78
4.3. MATERIALS AND METHODS
4.3.1. Cell culture and treatment…..……………………………………………..81
4.3.2. Flow cytometry
4.3.2.1. Propidium Iodide staining……………………………………….82
4.3.2.2. Multiparametric staining ………………………………………..82
4.3.2.3. BrdU/PI labeling………………………………………………...83
4.3.2.4. Caspase activation assay………………………………………...83
4.3.3. siRNA……………………………………………………………………..84
4.3.4. Clonogenic assay …………………………………………………………84
4.3.5. Determination of surviving fraction……………………………………….85
4.3.6. Western Blot detection…………………………………………………….85
4.3.7. Confocal microscopy ……………………………………………………..86
4.3.8. Comet assay……………………………………………………………….86
4.3.9. C4-2 expression profiling oligonucleotide arrays…………………………87
3 4.3.10. RTQ-PCR………………………………………………………………...88
4.3.11. Chromatin Immunoprecipitation Assay (ChIP) …………………………88
4.4. RESULTS
4.4.1. C4-2 prostate carcinoma cells undergo a G2/M arrest
following IR……………………………………………………………….90
4.4.2. E2F4/p130 complexes are formed following IR…………………………..98
4.4.3. A physiological role for E2F4 following IR in the G2/M
phase control of the cell cycle……………………………………………102
4.4.4. Identification of putative E2F4 target genes involved in
the IR response….………………………………………………………..113
4.5. DISCUSSION………………………………………………………………….119
CHAPTER 5. FUTURE DIRECTIONS: MANIPULATING CELL CYCLE
CHECKPOINTS FOR TARGETED THERAPIES FOR CANCER
TREATMENT
5.1. INTRODUCTION……………………………………………………………..125
5.2. E2FS AS MOLECULAR TARGETS IN CANCER THERAPY……………...126
5.3. SUMMARY AND FUTURE DIRECTIONS………………………………….127
BIBLIOGRAPHY………………………………………………………………………129
4 LIST OF FIGURES
CHAPTER 1
Fig. 1-1. Phases of the cell cycle…………………………………………………………18
Fig. 1-2. Temporal expression of cyclins………………………………………………...21
Fig. 1-3. Members of the E2F family of transcription factors…………………………...24
Fig. 1-4. The electromagnetic spectrum…………………………………………………33
CHAPTER 2
Fig. 2-1. Model for p53-dependent regulation of cell cycle arrest and
apoptosis following IR.…………………………………………………………53
Fig. 2.2. General scheme for performing chromatin immunoprecipitation
experiments in MOLT-4 cells …………………………………………………59
Fig. 2-3. p53 binding is differentially regulated following IR…………………………...61
CHAPTER 4
Fig. 4-1. Radiation induces a G2/M cell cycle arrest independently of p53
function in prostate carcinoma C4-2 cells……………………………………..91
Fig. 4-2. The radiation-induced arrest is specific to the G2 phase of
the cell cycle…………………………………………………………………...95
Fig. 4-3. E2F4 levels are sustained after IR and E2F4 co-localized with
p130 at the time of G2 arrest……………………………………………………99
Fig. 4-4. E2F4 downregulation causes an inappropriate G2 cell cycle arrest.…………103
5 Fig. 4-5. Lack of BrdU incorporation indicates that formerly arrested
G2 cells are not recycling…………………………………………………….107
Fig. 4-6. E2F4 knock-down by siRNA initiates caspase activation and
DNA strand breakage…………………………………………………………110
Fig. 4-7. Clonogenic assays indicated that E2F4 knock-down by
siRNA can radiosensitize cells………………………………………………..112
Fig. 4-8. IR induces the downregulation of E2F4 targets………………………………116
6 LIST OF TABLES
CHAPTER 2
Table 2-1. ChIP sense (S) and antisense (AS) primers used for detecting p53
bound to chromatin …………..………………………………….…………..51
Table 2-2. Expression profiling reveals radiation responsive genes containing
putative p53 binding sites in their regulatory regions………………………..54
Table 2-3. Basal levels of p53 binding validate putative binding sites in regulatory
regions within individual genes……………………………………………...63
CHAPTER 4
Table 4-1. E2F4 target genes identified by HU-95 Affymetrix array…………………..114
Table 4-2. Putative E2F4 target genes and their respective primer sequences…………115
7 ACKNOWLEDGEMENTS
Life moves us in ways that are unpredictable and gives us challenges that demand
coping strategies, which in turn promote growth in character. To steer through moments
of joy and self-doubt, one ultimately needs to adapt to new situations, accept
uncontrollable events, and take time to reflect upon experiences. Meeting challenges directly and growing from them pervade every facet of life. In this experience, we arrive at a true sense of humility and curiosity, which are cultured through evaluating ideas and learning to accept the criticism of others. The pursuit of obtaining the Ph.D. represents a culmination of maturity on multiple of levels. Likewise, a mentor accepting students shares in this responsibility by enabling the student to mature as a scientist by providing valuable experiential knowledge and guidance. From this vantage point, I thank my
advisor, Dr. Alex Almasan, whom has encouraged my creativity and scientific
independence. I have enjoyed working and studying in his laboratory.
The support of people in the laboratory is an important element of succeeding in
science, as their input and criticism have been very useful to my work. On a personal
level, these people have also nurtured my understanding of what a world citizen truly is. I am grateful to the members of the Almasan laboratory for their support on a professional and personal level. I thank my labmates Dr. Erica DuPree, Dr. Damodar Gupta, Dr.
Suparna Mazumder, Marcela Oancea, Dragos Plesca, and Dr. Subrata Ray for their scientific insight and companionship through my graduate education. My thanks also go to my thesis committee members Drs. Jacobberger, Silverman, Stavnezer, and Veigl, for their constructive criticism. I thank the Departments of Environmental Health Sciences
8 and Cancer Biology for giving me the opportunity to pursue a Ph.D. and for supporting my scientific growth at conferences.
Finally, I thank my parents, Walter and Ethel, my sister, Melissa, my brother,
Jonathan, and my dearest Alex for their sustained encouragement. Because they believed in me, they gave me the strength and the ability to face difficulties. I am where I am today because of their endless love and support.
9 LIST OF ABBREVIATIONS
14-3-3σ tyrosine 3-monooxygenase/tryptophan 5-monooxygenase activation
protein (sigma isoform)
APAF-1 apoptotic protease activating factor 1
APC Anaphase Promoting Complex
AS antisense (primer direction)
ATM ataxia-telangiectasia mutated
ATP adenosine triphosphate
ATR ataxia-telangiectasia and rad3 related
Bax Bcl-2-associated X protein
Bcl-2 B-cell CLL/lymphoma 2
BER Base Excision Repair
Bik Bcl-2-interacting killer
Bim Bcl-2-interacting protein
bp base pair
Brca1 breast cancer 1 gene
BrdU bromodeoxyuridine
BSA bovine serum albumin
Caspase-3 apoptosis-related cysteine protease 3
Caspase-6 apoptosis-related cysteine protease 6
Caspase-7 apoptosis-related cysteine protease 7
Caspase-9 apoptosis-related cysteine protease 9
10 Ccd25A cell division cycle 25A
Ccd25C cell division cycle 25C
CDE/CHR cell cycle-dependent element, CDF-1 binding site
Cdk cyclin-dependent kinase
cDNA complimentary deoxyribonucleic acid
Chek1 cell cycle checkpoint kinase
Chek2 checkpoint kinase 2
ChIP chromatin immunoprecipitation
CKI cyclin-dependent kinase inhibitors
CRM1 required for chromosome region maintenance
Cy 3 cyanine 3 dye
Cy 5 cyanine 5 dye
Cyt C cytochrome c
DB DNA binding
DHFR dihydrofolate reductase
DN dominant negative
DNA deoxyribonucleic acid
DP1/DP2 dimerization protein 1/2
DSB double-strand break dUTP 2’-deoxyuridine 5’triphosphate
E2F E2 factor, cellular factor required for E2 viral promoter activation
EDTA ethylenediaminetetraacetic acid
EST expressed sequence tag
11 EMSA electrophoretic mobility shift assay
G1 phase gap 1 phase of the cell cycle
G2 phase gap 2 phase of the cell cycle
Gadd45 growth arrest- and DNA damage-inducible gene
GAPDH glyceraldehydes-3-phosphate dehydrogenase
Gy Gray (dose unit of IR)
HDAC histone deacetylase
HR homologous recombination
IR ionizing radiation
kD kilo-Dalton
LZ leucine zipper motif
M phase mitosis phase of the cell cycle
MB marked box motif
MCM minichromosome maintenance
MDM2 mouse double minute 2 homolog
MEFs mouse embryonic fibroblasts
MMR mismatch repair
MPF maturation promoting factor
MPM-2 mitotic protein monoclonal-2 mRNA messenger ribonucleic acid
NER nucleotide excision repair
NES nuclear exportation signal
NHEJ non-homologous end-joining repair
12 NLS nuclear localization signal
NOXA phorbol-12-myristate-13-acetate-induced protein 1
NPAT nuclear protein, ataxia-telangiectasia locus
NP-40 Nonidet P-40, non-ionic detergent p21 cyclin-dependent kinase inhibitor-1A (CDKN1A) p53 tumor suppressor protein p53 p53MH p53 Mouse/Human ortholog algorithm p107 retinoblastoma-like 1; RBL1 p130 retinoblastoma-like 2; RBL2
PA26 p53-activated gene 26
PBS phosphate buffered saline
PCR polymerase chain reaction
Pig3 p53-induced gene 3
PIPES piperazine-1,4-bis(2-ethanesulfonic acid)
PML promyelocytic leukemia tumor suppressor protein
PMSF phenylmethylsulphonylfluoride
Pttg1 pituitary tumor-transforming gene
PUMA p53-upregulated modulator of apoptosis
PI propidium iodide
Plk polo-like protein kinase
Rb retinoblastoma 1
Ring1 ring finger protein 1
RNA ribonucleic acid
13 RPA RNase protection assay
PCR polymerase chain reaction
ROS reactive oxygen species
RT-PCR reverse transcription PCR: cDNA synthesis from mRNA
RTQ-PCR real-time quantitative PCR
S sense (primer direction)
S phase synthesis phase of the cell cycle
SDS-PAGE sodium dodecyl sulfate polyacrylamide gel electrophoresis siRNA small interference RNA
SSB single-strand break
SSC sodium chloride-sodium citrate
TE Tris-EDTA buffer
TK thymidine kinase tRNA transfer RNA
TS thymidylate synthetase
YY1 transcription factor of the GLI-Kruppel class of zinc finger proteins
14 G2 Phase Cell Cycle Regulation by E2F4 Following Genotoxic Stress
Abstract
by
MEREDITH ELLEN CROSBY
The E2F family of transcription factors is comprised of nine active members to date, which activate or repress genes. Well-studied members, such as E2F1 and E2F4, are known to play key roles in G1/S transition, S-phase progression, and mitosis. E2F1 is generally regarded as a transcriptional activator and E2F4 is a transcriptional repressor; the balance of their expression levels may promote apoptosis or cell cycle arrest, respectively. Functioning together with or independently of p53 in cell cycle regulation,
E2F1, has been implicated in G1 cell cycle arrest following DNA damage; they share similar target genes.
Retinoblastoma (Rb) and related pocket proteins p130 and p107 play important roles in cell cycle control. However, because of multiple interacting partners, their specific roles have not been clear. Following genotoxic stress, p130 interacts with E2F4.
An ionizing radiation-induced G2-phase arrest was characterized by decreased expression of MPM-2, a mitosis marker, and Cyclins A2 (early G2) and B1 (late G2 and M).
Concomitant with this G2-arrest, E2F4 cellular localization was redirected to the nucleus, similar to that of p130. Knock-down of E2F4 by siRNA elicited persistent cellular DNA damage and sensitization following irradiation. Flow cytometry analyses revealed an
15 increased population of cells with an apparent S-phase content following this treatment, but these cells were not actively dividing. Downstream E2F4 targets potentially involved in the exit from G2 arrest were identified by expression-profiling. Chromatin immunoprecipitation localized E2F4 at promoter regions of the Bub3 and Pttg1 genes following irradiation.
This work indicates that E2F4 expression, nuclear localization, and target gene repression following irradiation play crucial roles in mediating the exit from G2 arrest in cells that sustain irreparable DNA damage. As the E2F4/p130 complex has been implicated in G0 control, its translocation leading to the repression of genes with G2/M function is novel. In addition to increasing the knowledge for understanding cell cycle arrest, new insights into the treatment of prostate cancers, which in aggressive forms harbor comparatively high levels of E2F4 indicate that the regulation of E2F4 functions in a manner that can lead to resistance to therapies that traditionally target cell cycle events.
16 CHAPTER 1. INTRODUCTION
1.1. CELL CYCLE REGULATION: HISTORICAL OVERVIEW
The cell cycle is comprised of 4 stages, Gap 1 (G1), Synthesis (S), Gap 2 (G2), and Mitosis (M) (Fig. 1-1). These phases are defined by an ordered series of events that allow for cell proliferation and lead to the creation of two daughter cells through cell division. The first three phases of the cell cycle are referred to as interphase. At the G1 phase, the cells either prepare for S phase or exit the cell cycle to differentiate.
Additionally, there is also a DNA checkpoint, which assesses the integrity of the DNA prior to its synthesis. After the G1 phase, DNA replication occurs in S phase. At the G2 phase, the cells prepare for the M phase. As in the G1 phase, in G2, there is a DNA damage checkpoint. In the presence of unreplicated or damaged DNA, a cell may undergo a cell cycle arrest to repair the DNA damage, or, may undergo other events that lead to apoptosis. After the G2 phase, nuclear and cytoplasmic division occurs in the M phase. The M phase is further subdivided into prophase, prometaphase, metaphase, anaphase, and telophase. In these subdivisions, the chromatin is condensed, sister chromatids are aligned, separated at their kinetochores, and pulled to opposite sides of the cell by spindle fibers. After the chromosomes have been pulled to opposite sides of the cell, the cell undergoes telophase and then cytokinesis. Cells that leave the cell cycle from G1 to differentiate are in G0. This so-called quiescence may be temporary or permanent.
17 T
EL
M A O N PH G1 A
P A M H E A SE DIFFERENTIATION TA S checkpoint PH E AS PR E OPH ASE
INTERPHASE
DNA checkpoint replication
G2 S
Fig. 1-1. Phases of the cell cycle. The cell cycle is made up of 4 phases: G1, S, G2, and
M. Phases G1 and G2 prepare the cell for DNA replication in the S phase and for mitosis,
respectively. Checkpoints to ensure DNA integrity occur at the G1 and G2 phases of the
cell cycle. Cells divide after M phase. Cells that leave the cell cycle may go on to differentiate.
18 The tight regulation of the cell cycle is achieved through the temporal expression of cyclins, their cyclin-dependent kinase (Cdk) binding partners, the cyclin-dependent kinase inhibitors, the anaphase promoting complex (APC), and accessory proteins involved in their stability and degradation. The critical cell regulator, Maturation
Promoting Factor (MPF), the oscillatory activity of CyclinB-Cdk1 provided the first
molecular insight into understanding the mechanisms underlying cell cycle regulation
(Masui and Markert, 1971). Cyclin proteins were originally identified by observing that
certain proteins in the eggs of the sea urchin Lytechinus pictus were synthesized only
after fertilization and degraded at various times during cell division (Evans et al., 1983).
That is, the factors involved in cell cycle regulation were initially understood through the
synchronization of cell division and protein expression.
As mitosis was explored through understanding the correlation between cyclin
regulation and cell cycle stage, early studies of cyclin regulation and Cdk regulation were
first understood at the interphase/mitosis boundary—that is, the G2/M transition. In
coordination with the interphase to mitotic transition, Cyclins A and B were found to
oscillate and promote the kinase activity of Cdk1 (Minshull et al., 1990). Cyclin A is
expressed earlier in the cell cycle compared to Cyclin B1 and interacts with Cdk2.
Cyclins C, D, and E were later found based on homology screening, subtractive
hybridization following stimulation with colony-stimulating factor-1 (Matsushime et al.,
1991) and complementation assays that resulted in a regain of cell cycle function in yeast
(Koff et al., 1991; Lew et al., 1991; Xiong et al., 1991). The D cyclins (e.g. D1, D2 and
D3) are expressed during the G1 phase of the cell cycle and may interact with either
Cdk4 or Cdk6. The activity of the G1 cyclins may be blocked by the cyclin-dependent
19 kinase inhibitors (CKIs), which belong to the p21 and p16 families. Cyclin E was found
to interact with Cdk2, which promotes the S-phase transition. Cdc25A positively regulates the activities of Cyclin A/Cdk2 and Cyclin E/Cdk2 by removing the inhibitory
phosphorylation on Cdk2 (Smits and Medema, 2001). At the onset of mitosis, members
of the polo-like protein kinase (Plk) family function in ensuring a proper bipolar spindle
and a proper passage through cytokinesis (Smits and Medema, 2001).
The control of cyclin expression is regulated through transcriptional regulation in
the cell cycle (Fig. 1-2). The cyclins’ transcriptional regulation is balanced by their
degradation. The protein degradation pathway is best understood for Cyclin B. Cyclin B
contains a destruction-box that is targeted by the APC ubiquitin ligase (Glotzer, 1995).
The APC catalyzes the poly-ubiquitination of Cyclin B1, leading to its proteasomal
destruction. Cdks are expressed throughout the cell cycle, but require activating
phosphorylation events and removal of inhibitory phosphorylation for their activity.
Checkpoints may be activated throughout the different phases of the cell cycle and serve
to maintain genomic stability, allow repair of DNA damage, and ensure proper DNA
replication and progression through mitosis (Andreassen et al., 2003).
20 p15/p16 p21 p18/p19 p27/p57
Cyclin D Cyclin E Cyclin A1, 2 Cyclin B Cdk4,6 Cdk2 Cdk2,1 Cdk1
Relative Protein Expression
G0/G1 S G2 M
Cell Cycle Phase
Fig. 1-2. Temporal protein expression of cyclins. Specific interactions of cyclins and their
respective Cdks promote progression through the cell cycle, but are expressed constitutively throughout the cell cycle. Cyclin D is important for the G1 phase of the cell
cycle, Cyclins E and A are critical for S-phase progression, and Cyclins A and B are
implicated in the G2/M transition. Cyclin-dependent kinase inhibitors, such as p15 and
p21 block cell cycle progression by inhibiting cyclin and Cdk interaction.
21
At the same time, the transcriptional regulation of genes implicated in cell division has been correlated with the E2F family of transcription factors, leading to a model in which E2F-containing complexes are critical to the cellular proliferation. In the case of synchronized NIH 3T3 cells, levels of active E2F were seen to increase with the release from a thymidine block, suggesting that it played a role in cell cycle progression
(Mudryj et al., 1991). The retinoblastoma tumor suppressor protein (Rb), in its unphosphorylated form, was found to interact with E2F (Chellappan et al., 1991). E2F regulation in normal cells was first understood from experiments where the adenovirus
E1A 12S product was used to dissociate E2F from pRb, leading to the sequestration of pRb and to the activation of E2F (Nevins, 1992). E2F’s transcriptional activity is important for the regulation of genes implicated in cell cycle progression and DNA synthesis, which include Cyclin E, dihydrofolate reductase (DHFR), Cdc6, and minichromosome maintenance (MCM) (Dyson, 1998). Insights into misregulation of the cell cycle came from other experiments in which E2F complexes were dissociated by
E1A, which resulted in a loss of cellular proliferation control (Chellappan et al., 1991).
1.2. THE E2F FAMILY OF TRANSCRIPTION FACTORS
The E2F family of transcription factors is comprised of nine different members
(E2F1-3a & 3b-8), which coordinately regulate the cell cycle (Attwooll et al., 2004;
Trimarchi and Lees, 2002). Based on their function, these members have been divided into two subgroups: activating (E2F 1-3a) and repressing (E2F 3b-5; 6-8) transcription factors. The E2F 1-5 members have the highest degree of homology (Attwooll et al.,
22 2004). The family has different affinities for members of the pRb family (pRb, p107, and
p130) and contains several conserved homology domains that are involved in DNA-
binding, heterodimerization, and transactivation (Helin, 1998) (Fig. 1-3). The pRb
members are phosphorylated in a cell cycle-dependent manner. In their
hypophosphorylated state, they bind to specific members of the E2F family and cyclins, and block proliferation.
23 Dimerization Transactivation
E2F1 & E2F2 DB LZ MB pRb Activators E2F3a DB LZ MB pRb
E2F3b DB LZ MB pRb
E2F4 & E2F5 DB LZ MB pRb
Repression Repressors
E2F6 DB LZ MB
E2F7 DB DB
E2F8 DB DB
NLS, nuclear localization signal DB DNA binding
NES, nuclear exportation signal pRb pRb family binding
LZ MB Leucine zipper and marked box motifs, DP dimerization domain
Fig. 1-3. Members of the E2F family of transcription factors. Subgroups of E2Fs contain conserved homology domains, which include their respective DNA, DP, and pRb binding
proteins. A schematic cartoon indicates homology domain conservation in the E2F family. These domains include the nuclear localization signal (NLS), the nuclear exportation signal (NES), the dimerization domain (DP) consisting of the leucine zipper
(LZ) and marked box motifs (MB) DNA binding (DB), and the pRb family binding (pRb) domains. (Adapted from Trimarchi and Lees, 2002.)
24 Despite a high degree of structural homology and some functional redundancy,
members of the E2F family have biological specificity for carrying out their activities in
gene transcription or repression, respectively (DeGregori, 2002). Subcellular localization
and transcriptional regulation are important for different members of the E2F family to function as transcription factors, as E2F activities vary with the cell cycle (Lindeman et al., 1997). Activation and repression is carried out through E2F interaction with DNA via the DNA binding domain(s). Most E2F activity is dependent upon its dimerization with a
DP protein, as well as its interaction with the pocket proteins, pRb, p107, or p130
(Stevaux and Dyson, 2002).
The activating E2Fs, 1-3a, bind only to pRb and are regulated in a cyclic manner in the cell cycle (Helin, 1998). These activating E2Fs contain nuclear localization signals.
Interactions with pRb prevent premature E2F transactivation in the cell cycle, thereby providing control beyond transcription. Gene transactivation occurs when E2F heterodimerizes with its DP partner, as this complex has a high affinity for binding DNA.
The E2F consensus DNA-binding site was first determined through DNase I footprinting to have a consensus sequence of “TTTSSCGC,” where “S” may be C or G (Blake and
Azizkhan, 1989). As chromatin remodeling is important for gene accessibility by transcription factors, GCN5 acetyltransferase activity and its cofactor, TRRAP, were also found to be important for E2F transcriptional activity (Lang et al., 2001).
E2F3b, unlike 3a, is constitutively expressed and has recently been implicated in gene repression during quiescence (Dimova and Dyson, 2005). The classical repressors,
E2F4 and E2F5, either interact with all three pocket proteins or p130, respectively; they are regulated by subcellular localization and posttranslational modifications (Lindeman et
25 al., 1997; Trimarchi and Lees, 2002). The expression of E2F4 and E2F5 is relatively stable through the cell cycle (Sardet et al., 1995). They are considered to be important for transcriptional repression perhaps because they are poor transcriptional activators. In the transcriptional repression model, Rb/E2F recruits histone deacetylases (HDACs)
(Ferreira et al., 2001; Luo et al., 1998) and SWI/SNF chromatin remodeling complexes
(Trouche et al., 1997) to the promoters of genes that are not expressed in certain phases of the cell cycle. In addition to their repressive roles in the cell cycle, E2F4 and E2F5 are critical for cell cycle exit and differentiation (Gaubatz et al., 2000).
E2F6 does not bind with the pRb members and represses transcription through its carboxyl terminus, which recruits Ring1 and YY1 binding protein with the polycomb complex of proteins (Trimarchi and Lees, 2002). E2F6 has been implicated in gene silencing in G0 at some cell cycle-related target promoters, however it does not seem to interact with the pocket proteins (Ogawa et al., 2002; Trimarchi et al., 2001). This is likely because the E2F6 protein structure is unique in that it lacks the C-terminal transactivation domain (Kherrouche et al., 2001). Additionally, E2F6 may provide insight into the repression of G1/S-associated genes versus G2/M-associated genes, as it interacts selectively with G1/S-associated E2F target genes during S phase of the cell cycle, whereas E2F4 is able to bind target gene promoters in all phases of the cell cycle
(Giangrande et al., 2004).
E2F7 and E2F8 do not have a transactivation domain and lack the pocket protein- binding domain (Dimova and Dyson, 2005). These closely related proteins do not interact with DP. Instead, they have two distinct DNA-binding domains and bind DNA as a homodimer (Christensen et al., 2005; Dimova and Dyson, 2005).
26 E2Fs (1-5) have been knocked out in mice and their effects have had diverse and
distinct roles in development and physiology. E2F1-/- and E2F2-/- mice have an increased
probability of tumorigenesis (Yamasaki et al., 1996) (DeGregori, 2002). E2F3-/- mice have partially penetrant embryonic lethality (Wu et al., 2001). E2F4-/- mice are smaller
than normal mice and have craniofacial defects (Humbert et al., 2000). E2F5-/- mice are perinatal lethal with hydrocephalus (DeGregori, 2002). The combination of E2F4-/- and
E2F5-/- is embryonically lethal (Gaubatz et al., 2000).
The E2F family of transcription factors elicits signaling that either promotes cellular growth, cell cycle exit, or terminal differentiation (Trimarchi and Lees, 2002).
Although important for the G0/G1 transition and S-phase progression, it has been recently suggested that E2Fs may also play a role in the G2/M phase transition in the mammalian cell cycle (Ren et al., 2002). Indeed, the key experiment including a triple
knockout of E2F1-/-, E2F2-/-, and E2F3-/- resulted in cells being blocked at all stages of
the cell cycle (Wu et al., 2001).
The combinatorial effects of transcriptional activators and repressors recruited to
the regulatory regions of genes affect the pattern of gene expression and this is further
complicated by the regulation of their expression in the cell cycle. Continued interest and
appreciation of the E2F family of transcription factors has resulted from the fact that E2F
family members are capable of regulating genes involved in mitotic function (Muller et
al., 2001), although the molecular mechanism is not clear (Cam and Dynlacht, 2003). The
pattern of gene expression may be matched with a specific member of the E2F family
(Black et al., 2005). Interestingly, the E2F family may control the G1/S transition and
27 G2/M transition in order to control an overall proliferation rate, such that the length of
one phase may be kept in balance with the other (Dimova and Dyson, 2005).
1.3. E2F AND CELL CYCLE CONTROL
1.3.1. G0/G1 phase transition
Based on their association with pRb, p107, and p130, the classical repressors,
E2F4 and 5 are highly expressed in the nucleus during G0/G1, making them the
predominant players during quiescence and in early G1. In comparison to E2F5, E2F4 is
expressed at higher levels and E2F4 binding to either p107 or p130 controls the majority
of E2F in vivo gene repression. In quiescent cells, the E2F4/p130 heterodimer is the
major E2F complex bound to promoter regions of growth-regulated genes (Takahashi et
al., 2000), although E2F6 has also been noted to be present at these promoters (Stevaux
and Dyson, 2002). As cells pass into the late G1, the presence of this complex greatly
diminishes at these promoters, with the activating members, E2F 1-3a becoming the
predominant E2F complexes (Takahashi et al., 2000).
1.3.2. S phase
Although expressed constitutively through the cell cycle, E2F4 exerts gene
repression during G0/G1, when it is localized in the nucleus. As the G1/S phase transition
occurs, E2F4 is exported from the nucleus by CRM1 and remains in the cytoplasm
predominantly bound to p107 until the next G1 phase (Gaubatz et al., 2001; Stevaux and
Dyson, 2002). During the G1/S transition, Cdk4/CyclinD1 and Cdk6/CyclinD3
phosphorylate members of the pRb family. Phosphorylation of Rb ensures S-phase
28 progression at the G1 restriction point in the cell cycle. Upon Rb phosphorylation, the
Rb/E2F complexes dissociate. Free E2F functions as an activator of transcription in genes
that are associated with proliferation (Sala et al., 1994). Additionally, E2F/Rb complexes
with a heterodimeric partner: DP1 or DP2. These proteins, in coordination with Cdk2, are
required for DNA synthesis (Girard et al., 1991). This regulation of Rb and E2F is
important for maintaining the temporal integrity of the cell cycle, as in the absence of Rb, mRNA and protein levels of E2F target genes are upregulated untimely, indicating that
E2F complexes containing Rb protein are rate-limiting at the G1/S transition (Almasan et al., 1995). Thus, the untimely release of E2F from pRb can promote inappropriate S- phase entry (Almasan et al., 1995).
Expression of E2Fs 1-3a peaks in the late-G1/S phase of the cell cycle and this is
important for the transcription of critical genes implicated in S-phase proliferation (e.g.
DHFR, TK, and TS) (Rayman et al., 2002; Trimarchi and Lees, 2002; Wu et al., 2001).
The regulation of E2F1 is carried out in an E2F-dependent manner, where multiple E2F binding sites in the promoter may be bound by activating and repressing E2Fs, indicating
that regulation is a dynamic process that involves the summation of bound factors’
activities (Araki et al., 2003). Expression of Cyclins A and E, are transcriptionally
regulated during S phase by the activating E2Fs (Henglein et al., 1994; Pagano et al.,
1992). NPAT, an in vivo substrate of Cyclin E-Cdk2 kinase, is a critical gene implicated
in S-phase regulation. Thus, NPAT plays a key role in histone expression and its regulation provides a direct example of E2F-dependent cell cycle control (Gao et al.,
2003). Interestingly, chromatin immunoprecipitation (ChIP) experiments have elucidated
some E2F transcription target gene differences, where E2F1 has a preference for the TK
29 promoter (Wells et al., 2000). In contrast, E2F2 has a preference for the DHFR promoter
(Wells et al., 2000). However, such preferences are limited to particular genes, as the promoters of other genes do not have such E2F specificity and multiple E2F complexes may be found at individual promoters (Zhu et al., 2004).
1.3.3. G2/M phase transition
The regulation of the Cyclin B1/Cdk1 activity regulates the G2/M transition.
Whereas other cyclins are synthesized and localize to the nucleus until their degradation,
Cyclin B1 is held in the cytoplasm until it is needed for M phase and then translocates to the nucleus. Phosphorylation on Thr161 of Cdk1 is essential for the Cyclin B1/Cdk1 complex’s activity, which is also regulated by the Cdk activating kinase (CAK: Cyclin
H/Mat1/Cdk7). The activity of Cdk1 is blocked through inhibitory phosphorylation on two other residues, Thr14 and Tyr15; the latter phosphorylation is important for preventing premature entry into mitosis. The phosphorylation of Cdk1 on Thr15 is balanced by the phosphorylation activity of Wee1/Mik1 and by the dephosphorylating
(mitosis-inducing) activity of Cdc25C (Smits and Medema, 2001).
Earlier data from the Drosophila melanogaster model system suggested that E2F gene regulation extends beyond the S phase and plays a role in mitosis (Neufeld et al.,
1998). As in S phase, the E2F family members regulate genes implicated in the G2/M transition. This was first noted through experiments that searched for new E2F target genes through a global ChIP-DNA microarray hybridization approach, which identified genes involved in M phase, as well as DNA repair, cell cycle checkpoints, and chromatin assembly (Ren et al., 2002). Using synchronized cells and a high-density DNA
30 microarray, the E2F-induced genes identified were implicated in the G1/S transition, as
well as at the G2 phase of the cell cycle (Ishida et al., 2001). Cdk1 and Cyclin B1 are the key mediators of the progression through the G/2M phase of the cell cycle. Their respective gene transcripts were identified to be targets of E2F through these combined
ChIP-array experiments.
Both Cdk1 and Cyclin B1 contain activating and inhibitory E2F binding sites within their promoters (Zhu et al., 2004). Therefore, and as expected, both activator and repressor E2Fs, E2F1 and E2F4, bind to these genes’ promoter regions. Although activator or repressor E2Fs use the same E2F sequence interchangeably, secondary elements in the binding region lend specificity. Transcriptional activation occurs through
CCAAT and B-Myb binding sites and inhibition occurs through the cell cycle-dependent element, CDF-1 binding site (CDE/CHR) control. Indeed, the presence of E2F and B-
Myb appears to be important for the cell cycle-dependent regulation of Cyclin B1 and
Cdk1 (Zhu et al., 2004). E2F7 also appears to play a role in gene repression when other
E2F members are no longer in repressive complexes in the G2 phase (Dimova and
Dyson, 2005). Finally, a link between the E2F regulation at the G1/S transition and the
G2/M transition embraces the fact that E2F3 orchestrates DNA synthesis with centrosome duplication (Dimova and Dyson, 2005).
.
31 1.4. CELL CYCLE MISREGULATION BY GENOTOXIC STRESS: IONIZING
RADIATION
1.4.1. The cellular response to ionizing radiation (IR)
The electromagnetic spectrum is made up of rays that have wavelengths extending from 10-14 through 104 m (Fig. 1-4). IR is comprised of the shortest wavelengths ~10-14-10-9. Unlike other wavelengths of energy, IR is capable of removing outer orbital electrons from molecules, which creates a positively ionized and unstable molecule that can further react with other target molecules, such as DNA and protein.
32 t s t h y g a h i e r g l v i a a l d s e w m le r r t y a a r a ib r m r f d o a - V is a M h M In V A G X U V R F T S
10-14 10-12 10 -10 10-8 10-6 10-4 10-2 1 102 104
Wavelength (m)
IONIZING RADIATION
Fig. 1-4. The electromagnetic spectrum. Ionizing radiation is comprised of Gamma rays and X-rays, which have the shortest wavelengths of the spectrum (10-14 – 10-9).
33 Current understanding of how IR affects human tissues has been documented as a
result of clinical use and occupational exposure in mines, nuclear weapon testing, and
usage in war. Dose, fractionation, shielding, dose rate, and quality of IR affect how DNA
damage is tolerated. IR absorption leads to DNA damage, where the rays cause DNA
strand breakage upon interacting with DNA directly, or through the production of
reactive oxygen species (ROS). In general, the IR-response can be divided into three
phases: physical, chemical, and biological. In general, the physical and chemical phases
occur very fast after IR and result in the generation of free radicals (Hall, 2000). The
biological phase is less clear and the effects of free radical damage on DNA affects
various signaling events, including metabolic, cell cycle, and death pathways. These effects may not be evident until years after exposure.
Because cells proliferate at different rates, cells exhibit differences in their radiosensitivity patterns. In general, cells that proliferate rapidly are sensitive and those that proliferate slowly are relatively radioresistant. Radiosensitive cells lining the GI tract exhibit an acute response, which occurs within several hours after IR and can lead to death after two weeks following exposure. In other cells that are comparatively radioresistant, such as neurons and muscle cells, the response to IR does not appear
immediately after exposure (Hall, 2000). In this case, the DNA damage may not be as easily repaired as in the acute response.
Although DNA damage may result from DNA base covalent modification, mismatch between the bases, DNA crosslinking (inter- and intra-strand covalent
linkages), the most harmful damage following IR is strand breakage in the DNA
backbone. Single-strand breaks (SSBs) are the most common breaks, but double-strand
34 breaks (DSBs) are the most lethal. Once DNA damage is recognized by the cellular sensing machinery, the cell may try to reverse the damage by photoreactivation or by single strand ligation involving DNA ligase. Alternatively, the cell may remove the damage and replace a mismatched base, damaged base, or a nucleotide section by mismatch repair (MMR), base excision repair (BER), or nucleotide excision repair
(NER). DSBs are resolved by the non-homologous end-joining (NHEJ) or homologous recombination (HR) pathways (Rothkamm et al., 2003).
Inefficient repair can lead to persisting mutations, which can be amplified and lead to misregulated cellular proliferation. DNA misrepair occurs as a result of “hot spots” in the DNA and particularly in the coding regions, which are susceptible to mutation. Although DNA is damaged in a non-preferential manner, hot spots in the DNA are a result of problematic repair, which arises in instances where the repair machinery is unable to process through the damaged regions of DNA. Misrepair of DNA can lead to base pair substitutions (transversions and transitions), V(D)J recombination, meiotic exchange, frameshift mutations during later rounds of DNA synthesis and DNA repair. In some cases, the DNA damage is tolerated and the cell does not repair the damage, which can lead to mutations during further rounds of DNA replication.
1.4.2. DNA damage response: cell cycle arrest versus apoptosis
In recognizing DNA damage, normal cells need to stop cellular proliferation. Cell cycle checkpoints occur at all phases of the cell cycle, allowing the cells to halt proliferation. The decision to undergo apoptosis versus cell cycle arrest is of great interest and is still under investigation. Following DNA damage, a molecule that may link global
35 chromatin changes and accessory scaffold proteins with DNA repair is γ-H2AX (Redon et al., 2002). Phosphorylation of γ-H2AX is used to detect and quantify the number of
DSBs following IR (Wang et al., 2005). The sensor molecules for detecting DNA damage are the ataxia-telangiectasia mutated (ATM) and the ataxia-telangiectasia and rad3 related
(ATR) protein kinases. ATM is required for proper γ-H2AX foci formation. These proteins are part of the initial signaling pathway that leads to the activation of DNA damage-sensing proteins, which are involved in cell cycle checkpoints, genome maintenance, senescence, and apoptosis. Checkpoints were initially identified following
IR through their ability to halt cell cycle progression and to allow for DNA damage to be repaired (Zhou and Bartek, 2004).
Two key effector proteins, Chek1 and Chek2 are downstream of ATR and ATM and play key roles in transducing the DNA damage checkpoint response (Zhou and
Bartek, 2004). Chek1 is the checkpoint kinase that is cell cycle regulated, with expression in the S and the G2/M phase transitions, which ensures proper progression through DNA replication and mitosis. A downstream effector molecule of Chek1 includes Cdc25A, which is phosphorylated and thus targeted for ubiquitin-mediated degradation; is thereby unable to remove the inhibitory phosphorylation on Cdk1 and 2 and blocks cell cycle progression (Zhou and Bartek, 2004). This provides a mechanism by which cells can be arrested in late G1 or S phase. In the case of the G2 arrest, Chek1 can phosphorylate residues on Cdc25C, which prevents Cdk1 activation.
Chek2 appears to have overlapping activities with Chek1, but is considered to be the critical kinase for regulating the DNA-damage response with regard to DSBs and
DNA-damage induced apoptosis (Zhou and Bartek, 2004). However, the role of Chek2
36 has been best defined by its role in mediating the p53 apoptotic and cell cycle arrest response, as it phosphorylates and is required for the IR-induced transcription of p53 apoptotic target genes. Interestingly, the E2Fs have been implicated in a model of p53- dependent and –independent apoptosis following DNA damage. E2F1 is thought to affect p53 stability through the transcriptional regulation of the E2F target gene, p14(p19)/Arf, which neutralizes the MDM2 (Dimova and Dyson, 2005). Chek2 is also thought to induce apoptosis through a p53-independent mechanism following DNA damage through direct phosphorylation of E2F1 or promyelocytic leukemia tumor suppressor protein
(PML), which transcribes genes involved in apoptosis. E2F1 also seems to play a role in upregulating proapoptotic genes, such as p53-upregulated modulator of apoptosis
(PUMA), phorbol-12-myristate-13-acetate-induced protein 1 (NOXA), and Bcl-2- interacting protein (BIM) (Hershko and Ginsberg, 2004). Additionally, ATM can also be activated by E2F2, but the commitment step for initiating apoptosis is the stabilization of
Chek2 and induction of Chek2 by E2F1 (Rogoff et al., 2004).
1.4.3. Cell cycle specific arrest mechanisms
Cell cycle arrest may occur during each of the phases of the cell cycle, but so far,
there have been two well-characterized DNA damage checkpoints at the G1 and S phases. However, there appear to be distinct mechanisms for how arrest takes place, which differentiate G1 arrest from G2 arrest. As mentioned above, ATM and ATR are implicated in the early response to DNA damage and initiate downstream signaling to
Chk 1 and 2. Arrest at the G1 phase prevents S-phase progression. The G1 arrest is
37 carried out through the repression of Cdk4/6 activity by p16 and p21. p21 is primarily regulated following the stabilization of p53 (Zhou and Bartek, 2004).
The initiation of a G2 phase arrest, unlike the G1 arrest is not dependent on p53
(Taylor and Stark, 2001). G2 checkpoints can also function in tandem with prior checkpoints by allowing additional time to repair DNA damage, which may have not been repaired in preceding cell cycle phases. A checkpoint cell cycle arrest at G2 can be maintained by the inactivation of the Cyclin B1/Cdk1 complex by inhibitory Cdk1 phosphorylation and increased levels of p21 (Andreassen et al., 2003; Smits and
Medema, 2001), as well as cytoplasmic localization of Cyclin B1. As Cdk1 activity is required for normal G2/M phase transition, blocking Cdk activity effectively prevents cell cycle progression. Interestingly, the induction of p53 and p21 have also been implicated in models of gene repression that involve members of the pRb family, but no models beyond overexpression systems have shown this to occur in vivo (Jackson et al.,
2005). Finally, the transcriptional regulation of Cdk1 may be repressed by E2F/p130 complexes in the context of upregulated p21, suggesting that the E2Fs also play a functional role in mediating the G2/M phase of the cell cycle and in modulating the DNA damage response (Taylor et al., 2001).
1.5. OBJECTIVES AND THESIS OVERVIEW
Although much progress has been made through understanding the role of the
E2F family in the cell cycle over the past 30 years, new questions involving cell cycle misregulation are becoming apparent in the field. In the course of my studies, I have become aware of the impact of global gene expression profiling and how this tool may be
38 used in coordination with ChIP to investigate E2F regulated genes. This ultimately led to
the identification of new E2F target genes, which opened up the E2F/cell cycle field to
encompass cell cycle regulation at all phases of the cell cycle. As new target genes were found to be involved in the G2 and M phases of the cell cycle, E2F control of the cell cycle moved beyond the G1 and S phases to have a role in the G2/M phase transition.
Understanding cell cycle regulation in the context of E2F expression and regulation provides the backdrop against which its misregulation may be better studied in
genetically inherited diseases and cancer. Given the importance of previous work in the lab, I focused my thesis work to encompass 2 Aims. Initially, my studies focused on the regulation of physiological targets of p53, which mediated the effect of IR. I gained insights into understanding cell cycle misregulation and the limits of the p53 network, which enabled me to focus on a project that encompassed E2F4 in the context of IR.
Interestingly, the E2Fs share common transcriptional target genes with p53 and therefore provide and underlying theme for my studies.
As our lab has just identified novel E2F4/p130 complexes induced following IR in prostate carcinoma cells, I was interested in determining whether there was a role for these complexes in the cell cycle arrest after IR. Thus, Aim 1 was to determine the role and regulation of E2F4, as it served as a mediator of IR in the cell cycle response. I proposed the hypothesis that E2F4 protein levels, in contrast to those of E2F1, are maintained following IR and serve as a key determinant in regulating cell cycle control in prostate carcinoma cells. To test this hypothesis, I characterized the cell cycle distribution of cells subjected to IR, which was used as a prototype of genotoxic stress. I also used
multiparametric flow cytometry (for DNA and cyclins), investigated the cell cycle
39 dependence on E2F4 using siRNA-mediated knock-down of E2F4, and performed assays
to analyze the cellular response to down-regulated E2F4 levels. Aim 2 was to determine the role of E2F4 in the regulation of critical target genes associated with cell cycle
regulation following IR. Determining E2F4’s capacity to repress target genes following
IR tested the hypothesis. To do this, I investigated the regulation of E2F4 target genes’
expression following IR using real-time quantitative PCR (RTQ-PCR) and analyzed the levels of some E2F4 target gene products following IR using immunoblotting.
The novelty of my work consists of key observations that I have made, which have several implications for understanding the radiation-induced cell cycle arrest and more specifically, the role E2F4 plays in it. Following IR, the cells are stably arrested in
G2 and they no longer incorporate BrdU, as cycling cells do. Cells have a DNA content that, based on flow cytometry studies, corresponds to arrest at the G2 phase. Clearly, this arrest is before the M phase, as the cells were devoid of MPM-2 expression. Moreover, levels of Cyclin A2 and B1 are lower than what would be expected for G2. With the same kinetics observed for this G2 arrest, there was a colocalization of E2F4 and p130, both predominantly found in the cytoplasm, to the nucleus. Prior studies have indicated a similar role for the pRb family, especially p130. However, pRb family members have multiple interacting partners. Upon diminished E2F4 expression, these cells have an apparent S-phase DNA content, undergo a sustained G2 arrest, and do not incorporate
BrdU. In comparison to control cells, cells with knocked-down of E2F4 expression are sensitized to irradiation and undergo apoptosis. Potential E2F4 target genes were identified and the ability of E2F4 to bind to these genes’ promoter regulatory regions was demonstrated. Taken together, these findings indicate a novel role for E2F4 to mediate
40 the IR-induced cell cycle response leading to a sustained functioning in the G2 cell cycle arrest and thus providing a molecular mechanism that contributes to radioresistance.
41 CHAPTER 2. PHYSIOLOGIC TARGETS OF P53 IDENTIFIED THROUGH
CHROMATIN IMMUNOPRECIPITATION (CHIP)
2.1. ABSTRACT
Although radiation therapy has been an important modality for cancer treatment,
the molecular mechanisms underlying the overall genomic response of mammalian cells
to radiation are not well characterized. The success of radiation therapy using ionizing
radiation (IR) relies upon both cell cycle and apoptotic regulation, as conferred by the
activation of DNA damage-responsive genes. To better understand the key players
involved in this response, expression-profiling experiments were performed using
custom-made cDNA microarrays. In MOLT-4 lymphoma tumor cells, induction of p53
and transcriptional target gene products following IR supports a major role for p53 as a
transcriptional activator, but also invokes questions regarding conditional DNA binding
following IR. Using chromatin immunoprecipitation (ChIP), p53 binding was examined
following IR using primers specific for p53 binding sites in target genes. PCR analysis
indicates dynamic target gene binding. Thus, at 8 h following IR treatment, the p21 and
puma promoter sites were characterized by relative increases in chromatin precipitation,
while the Bax site was not. Since the binding of p53 to these sites only changed modestly
following radiation, other studies were conducted to characterize the presence of
constitutive binding to putative p53 DNA binding sites in several other genes.
42 2.2. INTRODUCTION
One of the most mutated genes in cancer, p53, functions as a gatekeeper in
regulating downstream events leading to responses to DNA damage and oncogenic transformation (Wahl and Carr, 2001). Genotoxic stress, such as ionizing radiation (IR),
induces a cell type-specific damage response that is dependent on, at least, the
phosphorylation and activation of p53 and downstream transactivation of its target genes.
Ionizing radiation induces p53 phosphorylation (Lakin and Jackson, 1999) at several serine and threonine residues and acetylation at several lysine residues. The phosphorylation of p53 at amino-terminal residues may inhibit MDM2 binding, promote p53 stabilization, and stimulate acetylation of residues at the p53 carboxyl terminus
(Haupt et al., 2002). Consequential to these post-translational modifications, it is believed
that p53 plays a major role in the damage response through its transcriptional
transactivation of target genes (Prives and Hall, 1999). Although it has been reported that
p53 can induce apoptosis in a transactivation-independent manner (Caelles et al., 1994),
the current work aimed to explore the most prevalent transactivation-dependent
mechanism. Target genes include mediators of cell cycle arrest, such as the cyclin-
dependent kinase inhibitor, p21, as well as mediators of apoptosis, such as Bax.
Following its activation, Bax participates in the induction of apoptosis by homodimerizing, translocating to the mitochondria, and thus promoting the release of
Cytochrome c (Cyt c) from the mitochondria into the cytoplasm (Chen et al., 2003). In the presence of adenosine triphosphate (ATP) this cytoplasmic Cyt c binds to the apoptotic protease activating factor 1 (APAF-1) adaptor molecule and recruits and
43 activates Caspase-9 through proteolytic cleavage, leading to activation of the caspase
cascade and apoptosis (Chen et al., 2000).
Expression profiling of multiple genes following ionizing radiation provides
insight into identifying radiation-responsive genes within the context of the entire genome (Perou et al., 2000). Furthermore, the kinetics of their response following
radiation can suggest potential dynamic regulation at the level of transcription. The development of DNA microarray techniques has allowed the global analysis of gene expression, providing information about changes in the transcription profile of a cell in response to different stimuli. As recent reports indicate, the response to ionizing radiation is complex and heterogeneous, and depends on cell type. There are both p53-dependent and p53-independent modes, involving multiple signaling pathways. (Amundson et al.,
1999; Amundson et al., 2001; Park et al., 2002). Although some pathways proceed through activation of AP-1 and NF-κB, p53 seems to be the major player involved in the
DNA damage response (Wahl and Carr, 2001).
DNA binding by p53 occurs in a sequence-specific manner, with transcriptional activation having an important role for its tumor suppressor function through cell cycle arrest and apoptosis regulation. Classically, the description of these sites has been limited to the regulatory region of the gene within upstream promoter sequences (Agarwal et al.,
1998). However, more recent studies suggest the presence of response elements that also occur within intronic sequences (Thornborrow et al., 2002).
Following identification of radiation-responsive genes through expression profiling, chromatin immunoprecipitation (ChIP) experiments were designed to examine the selectivity of the p53 response under physiological conditions. For the p53 target
44 genes identified in these experiments, putative p53-responsive DNA elements were
determined either based on the mouse and human ortholog p53 (p53MH) algorithm (Hoh
et al., 2002) or adapted from previously reported ChIP experiments (Kaeser and Iggo,
2002; Szak et al., 2001). PCR was then used to identify levels of binding for PUMA, p21, and Bax. As previous reports indicated, levels of p21 and PUMA were upregulated modestly following irradiation (Kaeser and Iggo, 2002). Interestingly, levels of p53- binding to the Bax promoter did not seem to change at all. As a result, other experiments focused on the degree of constitutive p53 binding in the absence of radiation to explore selectivity between multiple putative binding sites within individual genes and their promoters. Oligonucleotides designed around target sites indicated that there is sequence specificity of p53 binding to chromatin.
2.3. MATERIALS AND METHODS
2.3.1. Cell culture and treatment
Human T lymphoblastic MOLT-4 cells were obtained from the American Type
Culture Collection (Rockville, MD) and maintained in a humidified incubator at 37° C,
5% CO2. The cells were grown in RPMI-1640 with 10% (v/v) heat-inactivated fetal
bovine serum, 50 U/ml penicillin, and 50 mg/ml streptomycin (Invitrogen, Carlsbad,
CA).
Exponentially growing cells were adjusted to a density of 1 x 105 cells/ml 24 h
prior to irradiation. Cells were treated with 4 or 10 Gy of ionizing radiation and incubated
for 8 h at 37° C, 5% CO2. Irradiation was performed at 25 oC using an X-ray source
(Pantak HF320: 320 kVP, 20 A, half-value layer 2mm Cu, East Haven, CT) at a fixed
45 dose rate of ~1 Gy/min or a 137Cs γ-ray source, at a fixed dose rate of 2.8 Gy/min, as
previously described (Gong et al., 1998).
2.3.2. MOLT-4 expression profiling with cDNA arrays
The cDNA microarrays were generated based on data obtained from our previous
oligonucleotide gene chip experiments (Affymetrix HGU-95Av1 GeneChip®), with the
cDNAs that showed a significant (> 5 fold increase) or decrease in expression levels after
exposure to IR being selected. This set of five GeneChip probe arrays (A-E), contains
~63,000 probe sets and examines 54,000 UniGene clusters derived from Build 95 of
UniGene. Based on this specific UniGene build, the HG-U95Av2 array accounts for
~10,000 full-length genes and arrays B-E represent expressed sequence tag (EST)
clusters. Other cDNAs were chosen according to their role in apoptosis and cell cycle
control (Oancea, M, Stanhope-Baker, P, Frevel, M., Agarwal, M, Williams, B, and
Almasan, A., unpublished).
For hybridization probe generation, which was carried out by M. Oancea, RNA
was isolated after specific time intervals following IR (4 and 8 h). Double-stranded
cDNA was prepared from the PolyA+ mRNA using the GIBCO-BRL SuperScript Choice
System (GIBCO-BRL, Rockville, MD). An in vitro transcription reaction was used to
produce biotin-labeled cRNA using the Enzo BioArray High Yield RNA Transcript
Labeling Kit (Affymetrix, Santa Clara, CA). The sample was then purified using the
QIAGEN RNA MiniKit (Valencia, CA) and then fragmented. A hybridization mixture
was prepared including the fragmented cRNA, the Affymetrix recommended control
cocktail and herring sperm DNA. For fluorescent labeling of samples and hybridization
46 to glass slides, 100 µg of total RNA was annealed to oligo(dT) and reverse transcribed in
presence of Cy3- labeled dUTP or Cy5 labeled dUTP. The resulting cDNA was purified
using GFXTM PCR, DNA and Gel Band Purification Kit (Amersham Pharmacia Biotech
Inc., Piscataway, NJ) resuspended in 20 µl hybridization buffer (20 x SSC, tRNA, polyA,
5% SDS), heat denatured and applied to the slide, which was then placed in a sealed
humidified hybridization chamber at 65°C over night. The unbound probe was removed from the slide in a three-step wash with 2 x SSC/0.1% SDS, 2 x SSC, and 0.2 x SSC for 5
min each using Affymetrix Fluidics Station 400, FlexGE:WS2v3 Protocol (Affymetrix,
Santa Clara, CA).
For scanning and data analysis, performed by M. Oancea and Dr. J. Hissong, the
slides were dried by centrifugation then scanned using an Affymetrix Agilent 2500
Scanner. For the scanning, the pixel size was 3 microns, the filter was 570 nm, and there
were 2 scans performed using the original calibration. The Affymetrix Microarray Suite
5.0 with statistical algorithm was used for analysis (Corner+ Avg:618, Count:32 Corner-
Avg:29779, Count:32 Background Avg:442.78, Stdev:7.80, Max:459.7, Min:425.1 Noise
Avg:13.72, Stdev:0.65, Max:15.2, Min:12.4). Additionally, the Microarray Suite 5 output data was further processed with Silicon Genetics' GeneSpring(R) version 6, build 1333.
Each chip (A-E) was normalized separately. Each measurement on the chip was divided by the 50th percentile of all measurements (the median) in that sample. The percentile was calculated with all normalized measurements above 0. Values below 0.0 were set to
0.0. The scanning program calculates the fluorescence intensity as a ratio of the medians green and red fluorescent signals from each spotted cDNA sequence on the array. The background intensity of each color is also measured for each spot and subtracted from the
47 target fluorescence to give the final intensity values. The ratio is interpreted as the ratio of
concentration for its corresponding mRNA in the two cell populations (treated and
control). GeneSpring(R) version 6scaled the data so that the median expression level on
each chip was equal to 1.0 and normalized each gene by the corresponding level in the
control sample.
2.3.3. Chromatin Immunoprecipitation Assay (ChIP)
Irradiated and control cells were treated with formaldehyde and incubated in
culture media containing formaldehyde (1%) for 10 min at room temperature, with
rotation, to crosslink the protein to the DNA. The reaction was stopped with glycine
(final concentration 0.125 M, 5 min incubation) and cells were collected by gentle centrifugation at room temperature. Cell pellets were washed once with 1x cold phosphate buffered saline (PBS) and gently centrifuged at 4° C. Cells were incubated on ice for 10 min and were lysed with 5 mM PIPES (pH 8.0), 85 mM KCl, 0.5% NP-40, 10
µg/ml each of PMSF, leupeptin, and pepstatin. Nuclei pellets were collected by centrifugation and resuspended in 50 mM Tris-Cl (pH 8.1), 10 mM EDTA, 1 % SDS, and
10 µg/ml each of PMSF, leupeptin, and pepstatin. Nuclei were incubated on ice for 10 min and were sonicated 6 x 10 s/sample using a microtip (Sonifier Cell Disruptor model
W140D, Heat Systems, Ultrasonics, Inc., Plainview, N.Y.). Shearing efficiency was
optimized to achieve an average chromatin length of ~600 bp. The chromatin was then
centrifuged and the supernatant was removed. Chromatin was aliquotted and quick frozen
(-80° C) for later use.
48 Chromatin was pre-cleared with pre-blocked Staph A cells (10-15 µl/1x107 cells) (Pansorbin/Staph A cells, Calbiochem, San Diego, CA). Staph A cells were incubated with chromatin for 15 min at room temperature and samples were centrifuged.
Pre-cleared chromatin from treated and untreated cells was incubated overnight (1
µg/sample condition) with an anti-p53 (DO-1; Calbiochem) monoclonal antibody. As a negative or mock control, the antibody was incubated with 1x dialysis buffer [2 mM
EDTA and 50 mM Tris-Cl (pH 8.0)]. Chromatin complexes were immunoprecipitated with Staph A (50 µl), incubated for 15 min at room temperature, centrifuged, and the supernatants used for immunoblots. In addition, 10% of the volume of the untreated and treated samples was saved as total input chromatin. Total input chromatin served as a positive control, indicating that the chromatin, prior to immunoprecipitation, contained amplifiable DNA. Pellets were washed for 3 min/wash at room temperature with rotation: once with 1 x dialysis buffer and four times with wash buffer (pH 8.0: 100 mM Tris-Cl,
500 mM LiCl, 1% NP-40, and 1 % deoxycholate).
Samples obtained after immunoprecipitation were prepared in 30 µl of 1 x
Laemelli buffer and boiled for 10 min. Samples were loaded (20 µl/well) and electrophoresed on an 8% SDS-PAGE gel. Alternatively, total protein was estimated from whole cell lysates and 25 µg was loaded/well. Proteins were electrotransferred onto nitrocellulose membranes; membranes were probed with anti-p53 (DO-1; 1:2000 dilution) and visualized with sheep anti-mouse (Amersham, Piscataway, NJ: secondary antibody; 1:2000 dilution) and LumiGLO Chemiluminescent reagents (KPL,
Gaithersburg, MD) as previously described (Gong and Almasan, 1999; Mazumder et al.,
2000).
49 Elution buffer (50 mM NaHCO3, 1 % SDS) was added to the samples to
remove complexes of antibody/protein/chromatin. Samples were placed on a shaking
platform for 15 min, centrifuged, and supernatants were collected. The process was
repeated and the supernatants from both elution steps were combined and centrifuged to
remove residual Staph A cells. Elutions were treated with RNase A (10 mg/ml) (Gentra
Systems, Minneapolis, MN) and 5 M NaCl (final concentration 0.5 M). For reverse
crosslinking, samples, including 10% total input chromatin, were incubated for 4 h at 67 °
C. Samples were precipitated with ethanol (2.5 volumes) overnight at -20 °C, centrifuged
and pellets air-dried. Pellets were redissolved in Tris-EDTA (pH 7.5), 25 µl of 5 x
Proteinase K buffer [1.25% SDS, 25 mM EDTA, 50 mM Tris-Cl (pH 7.5)], and
Proteinase K (10 mg/ml) (Gentra Systems). Samples were incubated at 45 ° C for 1 h and
extracted with phenol:chloroform (1:1). Samples were precipitated overnight at -20 ° C with 30 µl of 5 M NaCl, 5 µg glycogen, and 750 µl ethanol. Samples were centrifuged and resuspended in 30 µl of TE buffer. The following ChIP sense (S) and antisense (AS) primers were synthesized by Integrated DNA Technologies, Inc. (Coralville, IA) and are as described in Table 2-1.
50
Table 2-1. ChIP sense (S) and antisense (AS) primers used for detecting p53 bound to chromatin
51 2.4. RESULTS
2.4.1. Expression profiling identifies radiation responsive genes
The exposure of human cells to IR induces a DNA damage response that may
result in cell cycle control or apoptosis (Fig. 2-1). We have examined radiation-regulated
gene expression in MOLT-4 cells, which contain wild-type, functional p53 and serve as a
model hematopoietic cell line (Gong and Almasan, 1999). We have previously studied
expression of several genes in the context of the cell cycle and the apoptotic response to
IR in this cell line (Gong and Almasan, 1999). To further explore potential radiation- response genes on a genome-wide scale, expression-profiling experiments were conducted with MOLT-4 cells. Based upon our custom-made cDNA arrays, our experiments revealed the presence of 137 genes that were either up-or down-regulated 2- fold. Representative genes and their levels of expression, as fold-induction with respect to control cells, are presented in Table 2-2. These data represent the kinetics of gene expression at 4 and 8 h following 10 Gy of IR, which is a clinically relevant dose of radiation (Gong and Almasan, 1999). Consideration of various time-points exposes the fluidity of gene regulation following DNA damage. From these data, we identified several novel putative p53-target genes, as well as those that had been previously reported. In particular, these included p21, Bax, PUMA, and PA26.
52
Cell cycle arrest
P P Ccyclinyclin DNA p53 p21 A Cdkcdk
Apoptosis Time (h) 0 8 Kd
p53 53 Baxbax PUMA Caspasepuma 6
caspase 6 β-actin 42
Molt-4 total lysate (4 Gy IR)
Fig. 2-1. Model for p53-dependent regulation of cell cycle arrest and apoptosis following
IR. IR increases levels and activity of p53 (inset shown as immunoblot with an anti-p53
antibody of MOLT-4 cell lysates isolated at 0 or 8 h following IR (4 Gy). As a tumor
suppressor, p53 is thought to induce cell cycle arrest or apoptosis through downstream
effectors. As a regulator of the cell cycle, p53 binds to the promoter of and transactivates
p21, which then interacts with Cyclin/Cdks to inhibit cell cycle progression. In addition, p53 is thought to mediate apoptosis through binding and transcriptionally regulating
PUMA, Caspase-6, and Bax.
53
Table 2-2. Expression profiling reveals radiation responsive genes containing putative p53 binding sites in their regulatory regions
54 We have previously shown that following DNA damage, MOLT-4 cells undergo p53-dependent cell cycle changes, characterized by p21 upregulation (Gong and
Almasan, 1999). Increased p21 levels lead to its binding of and interaction with cyclin–
cyclin dependent kinase complexes (Mazumder et al., 2000). Additionally, other targets, such as the Bcl-2 family genes Bax (Gong and Almasan, 1999) and PUMA (Oancea and
Almasan, unpublished); (Yu et al., 2001) are key mediators of apoptosis. Following IR (4
Gy), RNase protection assays (RPA) have demonstrated that steady-state p21 mRNA levels in MOLT-4 cells were upregulated maximally at 8 h with respect to untreated cells
(Gong and Almasan, 1999). Our previous work reported increased Bax steady-state transcript levels at 8 h following 4 Gy IR (Gong and Almasan, 1999). In addition, nuclear run-off assays combined with RPA, showed that the transcriptional rates increased for both p21 and Bax, after IR (Gong and Almasan, 1999). Furthermore, examining post- transcriptional and post-translational mechanisms of regulation strongly indicated that the induction was due, primarily, to transcriptional regulation of these genes. Interestingly,
RPA data indicated that p21 induction occurred before Bax upregulation, which is in
concordance with the dynamic transcriptional events identified through expression profiling experiments.
2.4.2. Identification of p53 binding sites in radiation-responsive genes implicated in apoptosis and cell cycle control
The details of the mechanisms underlying the transcriptional activation of genes
involved in cell cycle control and apoptosis remain unclear. It is generally recognized that
the process involves p53 binding to specific DNA response elements and that this binding
55 is important for transcription to occur. Previous reports recognized the importance of the
amino-terminal region of p53, which contains a transcriptional activation domain (Szak et
al., 2001). In addition, this domain’s coordination with a central p53 core domain is
important for DNA sequence binding specificity (Thornborrow et al., 2002).
Electromobility shift assay (EMSA) data provide information regarding sequence binding
and indicated p53 binding to a synthetic p53 consensus sequence (Gong and Almasan,
1999). To validate transcriptional activation, luciferase expression from reporter templates containing p53-binding sites has been utilized. Indeed, following irradiation,
MOLT-4 cells transfected with a construct harboring a synthetic p53 binding site were
able to increase luciferase expression by ~2-fold (Gong and Almasan, 1999).
p53 may exert transcriptional activation through direct DNA binding or indirectly,
through its recruitment with other DNA-binding cofactors. Moreover, this binding may
occur within regulatory regions residing in gene promoters as well as in intronic regions
(Thornborrow et al., 2002). The chromatin immunoprecipitation (ChIP) technique is
powerful in its ability to detect transcription factor binding under physiological
conditions. Several groups (Kaeser and Iggo, 2002; Takahashi et al., 2000; Weinmann et
al., 2001) have described methods to validate transcriptional target sites as well as to
identify new sites through cloning techniques (Weinmann et al., 2001).
Recently, an algorithm has been developed to identify putative p53 binding sites,
including those located within genes associated with regulation of cell cycle control and
apoptosis (Hoh et al., 2002). Although this algorithm may identify potential p53 binding
sites, actual binding of p53 to these sites must be confirmed under physiological
conditions, when the structural integrity of the chromatin is maintained. Moreover, our
56 previous studies (Gong and Almasan, 1999) and more recent expression profiling experiments (Wang et al., 2001; Zhao et al., 2000) have shown that p53 may activate a different set of genes in different cell types.
In light of the data obtained through expression profiling and RPA assays, which identified radiation responsive genes and transcriptional activation of these genes, chromatin immunoprecipitation experiments were carried out to examine these putative p53-binding sites in these relevant genes conferring apoptosis or cell cycle control.
Oligonucleotide primer sets were designed around these sites. These regulatory sites in
Bax, p21, PUMA, and PA26 and their respective homologies with the consensus p53- binding sites are shown in Table 2-2.
2.4.3. Chromatin immunoprecipitation (ChIP) of p53 with regulatory regions of radiation-responsive genes implicated in apoptosis and cell cycle control
The current study modified previously published ChIP protocols to optimize its use with the MOLT-4 cell line and IR. First, anti-p53 antibodies were tested for their ability to detect the p53 protein. Immunoblotting demonstrated antibody specificity and a
2-fold increase in p53 protein levels following treatment with IR (Fig. 2-1). This is supported by previous data, which indicated that alterations in p53 protein levels were paralleled by changes in DNA binding activity in untreated versus irradiated (4 Gy)
MOLT-4 cells using EMSA (Gong and Almasan, 1999). However, there have been questions raised regarding allosteric gene regulation and the validity of using short fragments of DNA and or naked DNA for these EMSAs (Kaeser and Iggo, 2002). Thus, we were interested in utilizing ChIP to ask when and to what degree p53 bound the
57 chromatin of its transcriptional target genes. An overview of the ChIP protocol is presented in Fig. 2-2. Following irradiation (4 Gy), cells were treated, lysed, and chromatin was isolated using an anti-p53 antibody (DO-1). Following the immunoprecipitation of p53/chromatin complexes, DNA was isolated and purified. PCR reactions then amplified the immunoprecipitated DNA.
58 Chromatin immunoprecipitation
+ 4 Gy IR, 8 h incubation
Crosslink with formaldehyde
Lyse cells; sonicate nuclei
Immunoprecipitate chromatin/protein complexes
Remove non-specific complexes **
Dissociate antibody from immunoprecipitated complexes Immunoblot
Time (h) 0 8 Uncrosslink chromatin and protein p53 -
Digest protein and RNA IgG light chain
Precipitate and purify DNA
PCR
Fig. 2-2. General scheme for performing chromatin immunoprecipitation experiments in
MOLT-4 cells. Using the primers described in Table 2-1, DNA was recovered and analyzed by PCR. Alternatively, immunoprecipitated complexes were immunoblotted.
(See inset.) **Following immunoprecipitation, supernatants were collected and saved to determine the antibody’s binding capacity. After the removal of nonspecific chromatin/protein complexes by a series of five washing steps, the immunoprecipitated p53 was detected by immunoblot with a p53-specific antibody.
59 Initial experiments were designed to investigate p53 binding site occupancy 8 h
after exposure to IR. PCR data indicate that p53 binding to both PUMA and p21 promoter regions increased after IR. Specifically, signal intensity appeared to increase ~2-3-fold with respect to untreated cells, as quantified by RTQ-PCR. Products were then visualized
by agarose gel electrophoresis (Fig. 2-3.). Strikingly, we did not see any significant
changes in p53 binding to the Bax promoter region or the putative regulatory site in
PA26.
60 t u p n i k l c GeneGene p21 p21 pumaPUMA pa26 PA26 baxBax ta o o T M 08 08 08 08 TimeTime (h) (h)
Fig. 2-3. p53 binding is differentially regulated following IR. ChIP experiments were performed in MOLT-4 cells that were treated with IR (4 Gy). The amount of p53 binding is determined in the absence or presence of IR using different oligonucleotide primers for p21, PUMA, PA26, or Bax. The mock sample served as a negative control. Following 40 cycles of PCR, DNA products were visualized on a 1% agarose gel.
61 Because p53 binding increased only modestly, constitutive levels of binding to p53 of several other genes were also investigated. The genes studied included Caspase-6,
Bik, Pig3, and Bax. Total input chromatin was used as an internal positive control for all primer sets, with β-actin serving as a negative control for nonspecific background
binding. Basal binding of p53 to the promoter region of the Caspase-6 gene gave a positive signal with respect to β-actin. A similar result was demonstrated with
oligonucleotides designed around predicted p53 binding sites in the Bik, but not in the
Pig3 or the Bax regulatory regions.
Additionally, several oligonucleotide primer sets were designed to detect the
physiological significance of p53-binding sites within individual genes. Three to four
primer sets were designed around putative p53 binding sites in the Bax and Caspase-6
genes, respectively, as previously suggested by the p53MH algorithm (Hoh et al., 2002).
Results indicating the presence of positive or undetectable signal following 40 PCR
cycles are summarized with regard to their localization within each gene in Table 2-3.
p53 binding was detected to 2 out of 3 putative regulatory regions in Bax. Specifically,
p53 appeared to be bound to the promoter region that was proximal to the transcription
start site, as well as to the site in intron 4. In contrast, p53 binding was undetectable to the
more distal Bax promoter region. Out of the 4 primer sets corresponding to putative p53-
binding sites in the Caspase-6 regulatory region, 3 sites were found to be positive for p53
binding.
62
+++ strong signal, ++ moderate signal, + weak signal, ND not detectable
Table 2-3. Basal levels of p53 binding validate putative binding sites in regulatory regions within individual genes
63 2.5. DISCUSSION
Previous in vitro and in vivo studies demonstrated that post-translational
modification of p53 leading to its accumulation allows the protein to function as a
gatekeeper, which is critical for regulating cell cycle arrest and apoptosis. Ionizing
radiation is a prototypical DNA damaging agent, which is ideally suited to model the
downstream effects of p53 accumulation and activation. Previous data from our lab
indicated that IR promoted the transcriptional upregulation of p21 and other cell cycle
regulatory genes, such as cyclin e (Mazumder et al., 2000) in MOLT-4 cells in vitro.
Thus, transcriptional transactivation of genes involved in the mediation of apoptosis and
cell cycle arrest is considered to be the major mechanism underlying the biological function of p53.
The pattern of transcriptional response to p53 is itself very heterogeneous and also depends upon cell type, the nature of the inducing signal, levels of p53 expression, and p53 functionality (Yu et al., 1999; Zhao et al., 2000). Also, most of the genes activated by p53 are codependent on other transcription factors and their presence or absence depends on the genetic background of the cell (Yu et al., 1999). The experiments carried out in NCI-H1299 (a human lung carcinoma cell line lacking p53, but expressing temperature sensitive p53) using cycloheximide (an inhibitor of protein synthesis) helped to distinguish between primary and secondary p53 target genes. p53 directly upregulates pro-apoptotic genes and cell cycle inhibitors and downregulates anti-apoptotic genes and cell cycle promoters (Kannan et al., 2001). These genes seem to be upregulated in many of the cell types analyzed (Yu et al., 1999; Zhao et al., 2000). Secondary targets of p53
64 are genes involved in many other cellular functions, and are more specific to the cell type
analyzed (Kannan et al., 2001).
Several radiation responsive genes were identified through expression profiling
experiments in MOLT-4 cells, which revealed upregulation and downregulation of
multiple genes. These experiments served to both confirm the expression pattern for
genes previously known and to identify the patterns of expression other genes, which had
not been previously known to be regulated by genotoxic stress. There are two general
models to explain the transcriptional function of p53, as it relates to the specificity of
binding target genes implicated in apoptosis and or cell cycle arrest (Prives and Hall,
1999). In one model, p53 transactivation is thought to occur independently at genes either
associated with cell cycle arrest or apoptosis, suggesting that p53 binding promotes a
transcriptional event that is selective for eliciting a pattern of transcription, such as one
promoting cell cycle arrest, over another, promoting apoptosis. In a second model, genes associated with both cell cycle arrest and apoptosis are transactivated by p53.
Additionally, there may be dynamic p53 occupancy in regulatory regions of its transcriptional target genes, which would preferentially favor cell cycle arrest or apoptosis, as pertaining to the level of genotoxic stress (Kaeser and Iggo, 2002).
Recognizing the existence of basal physiological levels of transcription for most genes, the latter model seems to provide the most likely mechanism for considering transcriptional transactivation.
Following IR, initial ChIP experiments demonstrated dynamic p53 binding to
DNA sequences in the puma and p21 target genes. However, with consideration of constitutive levels of binding, these increases were not striking, supporting the second
65 model and indicating a basal binding occupancy of p53. These trends were in agreement with those determined through studies performed recently in HCT 116 human colorectal
cancer cells (Kaeser and Iggo, 2002). Interestingly, there was no change following IR in
p53 binding to the Bax or PA26 regulatory regions. Reasons for this finding may be
related to p53 binding sequence specificity—which may also be dependent upon cell type, recruitment of other cofactors to the p53 sequence, and general chromatin accessibility. Indeed, selective recruitment of p53 to putative binding sites is most likely dependent upon the association of protein complexes and a localized environment that
supports chromatin accessibility (Kaeser and Iggo, 2002; McKinney and Prives, 2002;
Thornborrow et al., 2002). Recently, the Manfredi lab (Thornborrow et al., 2002) further explored the putative p53-binding site in the Bax promoter (Miyashita and Reed, 1995) and found that this site was not responsive to p53 transcriptional transactivation. Instead,
they identified another p53 response element in intron 1, which was essential for p53-
dependent transcription by EMSA (Thornborrow et al., 2002). Although we did not
design primers for this particular site, our data, which focused on intron 4, suggests that
intronic regions of Bax may be important for transcription.
Because there were no robust changes in p53 target gene binding following IR,
emphasis was placed on identifying and characterizing the putative p53 binding sites in
Bik, Pig3, and within the Bax and Caspase-6 regulatory regions. There were two and
three sites in Bax and Caspase-6, respectively that appeared to be bound to p53. In
particular, p53 binding to a site located in intron 3 of Caspase-6 positively correlated
with previous data indicating that p53 regulates transactivation through this intronic
sequence (MacLachlan and El-Deiry, 2002). However, our data indicate a binding site
66 within this intron that is different from the one very recently proposed (MacLachlan and
El-Deiry, 2002). We have also identified an additional p53-binding site residing in the
Caspase-6 promoter region. Undetectable binding to the residual putative p53 binding sites invokes the possibility of differential chromatin remodeling events. Such events that constrict chromatin may deny target site accessibility for the transcriptional complexes, leading to the lack of detectable binding.
The data presented in this report substantiate previous studies linking radiation responsive genes critical for cell cycle arrest and apoptosis to p53 accumulation and transcription factor chromatin binding. These data support a model of radiation-induced p53 binding to chromatin that is dynamically regulated. Furthermore, these studies define differential binding of p53 within several target genes. However, it remains unclear as to what mechanism(s) dictates the in vivo selectivity of p53 for a given target gene under physiological conditions. The question as to whether or not a transcription factor is bound
or recruited to a particular DNA regulatory sequence and the cofactors involved is a
complex question, which is dependent upon various aspects that may involve
posttranslational modifications as well as chromatin remodeling, which may be
dependent on other cofactors. Future studies will explore the differential regulation of
these and other apoptotic and cell cycle p53-regulated genes.
67 CHAPTER 3. BEYOND P53: THE OPPOSING ROLES OF E2FS IN CELL
CYCLE PROLIFERATION AND DEATH
3.1. ABSTRACT
Progression through the cell cycle is dependent upon the temporal and spatial regulation of the various members of the E2F family of transcription factors. Two of these members, E2F1 and E2F4 have opposing roles in cell cycle progression, which were defined over a decade ago. While E2F1 is an activator of cell cycle progression,
E2F4 functions as a transcriptional repressor. Recent data indicate that these transcription factors also play a role in the cellular response to DNA damage. In the case of E2F1, its overexpression leads to apoptosis. In contrast, the decreased expression of E2F4, in response to siRNA-mediated knockdown or to certain therapeutic agents, can induce apoptosis. Conversely, increased levels of E2F4 may confer resistance to apoptosis- inducing therapies used in the clinic. The balance between the activities of these two proteins in tumor cells is of great interest. Directed control of E2F1 and E2F4 action may lead to better diagnosis of disease and improved therapeutic modalities.
3.2. INTRODUCTION
E2F1 and E2F4 are spatially and temporally regulated in development, with E2F1 and E2F4 opposing each other in their function as a transcription activator or repressor, respectively. The tight regulation of E2Fs is responsible for the timely expression of genes promoting DNA synthesis, cell cycle progression, and mitosis (Polager et al.,
2002). Many new target genes have been recently discovered through gene expression
68 profiling arrays combined with chromatin immunoprecipitation (ChIP) assays. Specificity
for the regulation of these genes occurs through the E2Fs’ subcellular localization and
respective interactions with different pocket proteins and chromatin modifiers, such as
Suv93H, HDAC, BRG1, and BRG2, throughout the cell cycle (Lindeman et al., 1997;
Rayman et al., 2002; Wells et al., 2000).
The classical model for E2F regulation indicates that E2F4/p130 and HDAC
complexes are predominant during the G0/G1 phase of the cell cycle and serve to repress
E2F target genes that are important for cell cycle progression (Takahashi et al., 2000). As
cells enter the cell cycle and transit into the G1/S phase, E2F1/pRb becomes the
predominant complex. After being phosphorylated by Cdks, Rb leaves E2F1 to function
as a transcriptional activator and promote cell cycle progression. Despite its known interaction with the E2Fs, experiments indicate that Rb is rarely found bound to the chromatin, or interacts with chromatin in a transient manner (Stevaux and Dyson, 2002).
In contrast, E2F4/p130 and E2F/p107 complexes may persist on the chromatin until complexes containing E2F1 and other activator E2Fs may bind to promoters during S phase (Attwooll et al., 2004). However, earlier work stresses that pRb is extremely important for the temporal expression of target genes and that without its presence, mouse embryonic fibroblasts (MEFs) exhibit deregulated cell cycle kinetics, E2F1 target
gene expression (e.g. DHFR, thymidylate synthase), and became sensitized to DNA
damaging agents (Almasan et al., 1995). Recent data also suggest that E2Fs may bind to
multiple promoter elements, thereby invoking the ability to dynamically regulate
promoter elements in either positive or negative manners (Zhu et al., 2004). In addition to
the genes that are regulated in the cell cycle, ChIP analyses have also revealed that
69 dynamic gene regulation by the E2Fs encompasses genes that are important for the DNA
damage checkpoint and repair pathways, chromatin assembly/condensation, chromosome
segregation, and the mitotic spindle checkpoint (Blais and Dynlacht, 2004; Ren and
Dynlacht, 2004).
Although a great deal of attention has been placed on understanding the pattern of in vivo E2F activity in development, questions entailing the pattern of gene regulation and specificity have yet to be fully answered. This is especially true in the context of characterizing molecular mechanisms implicated in cancer progression and treatment.
Thus, we hope to provide an overview of the current literature, with a particular focus on the dynamics of two opposing E2F family members, E2F1 and E2F4.
3.3. E2F1: A MULTITASKING MEDIATOR
A member of the activator E2Fs, E2F1 was the founding member of the E2F family and is essential for cellular proliferation (Wu et al., 2001). Although the activities of E2F1 have been defined in development, the downstream molecular complexities related to its deregulation are currently under investigation. Other work has shown that
E2F1 may function as an oncogene (Xu et al., 1995) or as a tumor suppressor (Phillips et
al., 1999). The nature of this duality is likely to be based on the degree to which E2F1 is
expressed in the context of the cell cycle and/or following DNA damage and the
transactivation of its target genes.
As an oncogene, the overexpression or untimely expression of E2F1 can advance
cells from the G0 phase to the S phase (Johnson et al., 1993; Kowalik et al., 1995). E2F1
overexpression has been identified in HEL erythroleukemia cells, as a result of the
70 amplification of the E2F1 gene (Saito et al., 1995). E2F1 overexpression is also known to
cause neoplastic transformation in astrocytes in vitro (Miyajima et al., 1996). In colon
cancer patients, the levels of E2F1 and a target gene, thymidylate synthase were elevated,
effectively promoting cell cycle misregulation and oncogenesis (Kasahara et al., 2000).
However, in other circumstances, the overexpression of E2F1 can lead to apoptosis through p53-dependent (Hsieh et al., 2002; Kowalik et al., 1995; Wu and
Levine, 1994) and –independent pathways (Nahle et al., 2002). Early studies indicated that increased levels of E2F1 resulted in increased stability of p53 and apoptosis, which
could be blocked by mdm2 induction (Kowalik et al., 1998). Following treatment with
the DNA damaging agent, etoposide, Chek2 induction stabilizes E2F1 levels promotes
the transcriptional induction of target genes, such as p73, p16, APAF-1, and Caspase-7,
which may lead to apoptosis (Muller et al., 2001; Stevens et al., 2003). A positive
feedback loop involving the induction of Chek2 by E2F1 and increased Chek2 stability
by Atm and Nbs1 enhances p53 stability through its phosphorylation on Ser15 (Rogoff et
al., 2002) and promotes apoptosis (Rogoff et al., 2004). Alternatively, TNFα increases
the levels of E2F1, leading to degraded and decreased TRAF2 levels; this results in a loss
of JNK/SAPK activity and anti-apoptotic signaling (Phillips et al., 1999).
Reasons for this heterogeneity may result from different threshold levels of E2F1
required for differential gene transactivation of its target gene promoters, which may
favor either apoptosis or survival. Since its promoter contains sites for both activation and
repression, E2F1 levels are dynamically regulated during the cell cycle (Araki et al.,
2003). The cellular response to DNA damage adds another level of complexity to E2F1’s transcriptional regulation and downstream target effectors. However, although both E2F1
71 and E2F2 are able to cause quiescent cells to enter S phase, only E2F1 has been shown to
promote apoptosis, which delineates its function from other activating E2Fs that could
otherwise cause aberrant cell cycle regulation.
Increases in levels of E2F1 may result in deregulated gene expression that
commits cells to undergo apoptosis (Stanelle et al., 2002). Certainly post-translational
modifications, as in the case of E2F1 acetylation, which promote the induction of p73
have already been identified (Pediconi et al., 2003). Additionally, an E2F transactivation-
independent mechanism was proposed in which increased levels of E2F protein could complex with either p53 or Cyclin A, resulting in apoptosis or survival, respectively
(Hsieh et al., 2002).
3.4. E2F4: A REMARKABLE REPRESSOR
E2F4 serves as a member of the repressor E2F subfamily and is known to function in growth suppression and differentiation (Landsberg et al., 2003) Although less
is known about the activities of E2F4, it is clear it that represses genes during quiescence
(Trimarchi and Lees, 2002) and heterodimerizes with p130 after cells undergo cell cycle exit and thereby induces differentiation in neurons (Persengiev et al., 1999). E2F4 is unique compared to E2F1 in that it is primarily cytoplasmic, contains a nuclear export signal, and is dependent on CRM1 for its cytoplasmic localization (Gaubatz et al., 2001).
Its heterodimerization with the pocket proteins pRb, p107, or p130 (Moberg et al., 1996) is responsible for nuclear import.
Aside from functioning during quiescence and differentiation, E2F4, like E2F1, appears to act outside of these conventional roles. E2F-4 functions as an oncogene when
72 it is introduced into untransformed cells in vitro (Souza et al., 1997). In tumors, E2F4 and
Rb loss, blocks inappropriate gene expression and cellular proliferation and functions as a
tumor suppressor (Lee et al., 2002). Indeed, E2F-4 mutations have been identified in
gastric adenocarcinomas, ulcerative colitis-associated neoplasms, colorectal carcinomas,
endometrial cancers, and prostatic carcinomas, indicating that E2F4 plays a key role in
tumorigenesis (Souza et al., 1997). Coding repeats mutations within the E2F4 gene, are critical targets of microsatellite instability in many kinds of cancers, including childhood
and adult leukemias (Komatsu et al., 2000).
E2F4 does not appear to be necessary for cell cycle progression, but it is
important for the pocket protein-mediated G0/G1 arrest of cycling cells, as E2F4-/- MEFs fail to arrest in response to p16 (Gaubatz et al., 2000). In addition, E2F4 contributes to the DNA damage response and ensuing cell cycle arrest following exposure to ionizing radiation (IR) during the G2/M phase of the cell cycle in the C4-2 prostate carcinoma cells (Crosby and Almasan, unpublished data). In the C4-2 cells, the levels of E2F4 increase in response to IR and seem to confer cell survival (DuPree et al., 2004).
However, in contrast to the increased levels of E2F1 following DNA damage and activation of apoptosis, E2F4 levels in H1299 lung adenocarcinoma cells decrease after treatment with cyclin-dependent kinase inhibitors and with DNA damaging agents, but not microtubule inhibitors, and are associated with apoptosis (Ma et al., 2004). Similarly,
C4-2 cells treated with siRNA E2F4 and IR undergo apoptosis, as evidenced by the cleavage of pro-Caspase-3 and of poly(ADP-ribose) polymerase (DuPree et al., 2004).
Thus, as in the case of E2F1, aberrant changes in E2F4 expression can result in a cellular response that may promote cell death or cell survival.
73 Although the mechanism underlying E2F1-induced apoptosis has been well
characterized, the role of E2F4 in apoptosis has not. The changes in the levels of E2F4 are thought to promote increased sensitivity, as siRNA against E2F4 promoted apoptosis
(DuPree et al., 2004; Ma et al., 2004), but the underlying mechanism has not been well studied. Because of the observation that E2F4 levels are higher in E2F1-/- cells and that
levels of E2F4 decrease after levels of E2F1 increase, it is thought that E2F4 may be
transcriptionally repressed by E2F1 (Ma et al., 2004). Because the overexpression of
E2F4 does not protect cells from drug- or E2F1-induced apoptosis, it has been proposed
that E2F4 does not function to block E2F1 (Ma et al., 2004).
Decreased levels of E2F4 may promote apoptosis by vacating the E2F1 binding
sites present in the promoters of proapoptotic genes, which would otherwise block the
E2F1 transactivation and apoptosis (Ma et al., 2004). Multiple binding sites for E2Fs
have been characterized in a number of promoters for target genes, which may also add to
the complexity of gene transactivation (Zhu et al., 2004). Thus, the integration of the
relative promoter capacities or competition for discrete E2F binding sites reflects the
ability of E2F1 or E2F4 to enhance or resist the induction of apoptosis. E2F4 deficiency
sensitizes cells and E2F1 deficiency decreases sensitivity to apoptotic stimuli. Thus, it is
proposed that E2F1 and E2F4 oppose each other not just in their control of the cell cycle
function, but also in the context of the response to DNA damaging agents, which
indicates that they provide a critical balance for gene regulation.
74 3.5. CONCLUSIONS
Our current knowledge indicates that coordinate E2F regulation is associated with
the control of cell cycle. Moreover, E2F1 and E2F4 may be regulated aberrantly, as in the
case of DNA damage response. In an effort to reconcile these activities, we look to
cancer cells, which have cell cycle pathways containing E2F/Rb family mutations. These
cells have aberrant growth profiles that result from deregulated E2F expression, where
E2Fs are either induced or repressed. In the case of Rb-/-, premature S-phase entry could
lead to deregulated gene expression and apoptosis, indicating that the role of the pocket
proteins is vital to cell cycle maintenance (Almasan et al., 1995). An integral part of this
system of checks and balances, the levels of E2F1 and E2F4 have opposing roles within
normal cell cycle regulation, as well as in their response to certain therapeutics, such as
DNA damaging agents.
Increased levels of E2F1, E2F2, and E2F3a lead to apoptosis, where E2F1 is the
most potent inducer; in contrast, E2F4 expression does not promote apoptosis. However,
decreased levels of E2F4 sensitize cells to chemotherapeutic or radiation-induced
apoptosis (DuPree et al., 2004; Ma et al., 2004). In contrast, increased levels of E2F4
serve to increase the survival of cells. In E2F1-/- cells, such as mouse embryonic
fibroblasts (MEFs), levels of E2F4 are higher upon treatment with cisplatin, flavopiridol,
VP-16 (topoisomerase II inhibitor), paclitaxel, or roscovotine (Ma et al., 2004).
Therefore, agents that decrease levels of E2F4 and block its interaction with p130 can potentially enhance cancer therapeutics.
As many studies have focused on the overexpression of the E2Fs and examining its activities based on reporter constructs, the next step is to determine biologically
75 relevant circumstances where levels of the E2Fs are deregulated. Thus, future studies should focus on mutating E2F binding sites and address the overall contribution of these defined changes. Studies must also be conducted to unravel the complexity of having promoters that contain multiple E2F binding sites.
76 CHAPTER 4. E2F4 IS A KEY MEDIATOR IN PROMOTING THE G2 ARREST
RESPONSE FOLLOWING IR
4.1. ABSTRACT
Retinoblastoma (pRb) and related pocket proteins p130 and p107 play important
roles in cell cycle control. However, because of multiple interacting partners, their
specific roles have not been clear. Here, we investigated the role of E2F4 following
genotoxic stress, which we have previously identified to interact with p130. Using multi-
parametric cell cycle analyses, we defined an ionizing radiation-induced G2-phase arrest
characterized by decreased expression of MPM-2, a mitosis marker, and Cyclins A2 and
B1. Concomitant with this G2-arrest, E2F4 cellular localization was redirected to the
nucleus, similar to that of p130. Knock-down of E2F4 elicited persistent cellular DNA
damage and sensitization following irradiation, as measured by comet, caspase activation,
and clonogenic assays, respectively. Surprisingly, flow cytometry analyses revealed a markedly increased population of cells with apparent S-phase content following this treatment. BrdU-labeling studies indicated that this population of cells was not actively dividing and therefore represented G2 cells with diminished DNA content. Downstream
E2F4 targets potentially involved in the exit from G2 arrest were identified by oligonucleotide microarray expression-profiling experiments. Chromatin immunoprecipitation localized E2F4 at promoter regions of the Bub3 and Pttg1 mitotic genes following irradiation. These data indicate that E2F4 expression, nuclear localization, and target gene repression following irradiation play crucial roles in mediating entry and subsequent exit from G2 arrest in cells that sustain irreparable DNA
77 damage. As the E2F4/p130 complex has been implicated in G0 control, its surprising
activation leading to repression of genes with critical G2/M function may represent a
hallmark of genotoxic stress response.
4.2. INTRODUCTION
Progression through the cell cycle is dependent upon the integrated expression of
factors that both positively and negatively regulate the cell cycle. This complex
regulatory network may either promote transitions in the cell cycle through the activity of
cyclins and their respective cyclin-dependent kinases, or may block progression through
the initiation of checkpoints that occur during G1/S or G2/M. Whereas in response to
DNA damage an arrest may occur at the G1/S-phase transition, most common is a G2
arrest, particularly in tumor cells that largely lack an effective G1/S arrest. Extensive
experimental evidence supports the idea that the tumor suppressor gene, p53, plays a
critical role in carrying out the G1 arrest (Wahl and Carr, 2001). The G2/M arrest
appears to be much more complex and still occurs in cells lacking p53 (Chan et al., 2000;
Taylor and Stark, 2001). The G2 arrest may occur via different pathways, including the
checkpoint pathway implicating ATM/ATR and Chek1/Chek2, which converge on the
regulation of Cdc25C and its downstream Cdk1 target (Kastan and Bartek, 2004)(Ray,
Gupta, Macklis, Almasan, unpublished data). Although p53 may be playing a role in the
duration of this checkpoint, the G2/M arrest still occurs in p53-deficient cells, suggesting
that there are other critical factors that may be regulating its initiation. Thus, although
different components of the G2-phase arrest have been identified, the potential role of
78 other molecules involved in cell cycle regulation, such as those belonging to the E2F family of transcription factors, remains unclear.
The E2F family of transcription factors has long been appreciated as a master controller of the expression of genes implicated in the cell cycle. This family is composed of nine proteins, E2F1-3a, b-8. The first three of these proteins, E2F1-3a are considered to be transcriptional activators, while E2Fs 3b-8 are regarded to be transcriptional repressors (Trimarchi and Lees, 2002). These proteins are regulated spatially and temporally throughout the cell cycle and most heterodimerize with DP-1, DP-2, pRB, and the related p107 and p130 pocket proteins to promote or inhibit transcriptional potential, respectively (Takahashi et al., 2000). Cell synchronization experiments identified E2F family cell cycle-phase specific complexes. E2F4/p130 and E2F5/p130 complexes were present primarily in quiescent cells (Sardet et al., 1995). Levels of E2F4 decreased to form free E2F, E2F1, E2F4/pRb, and E2F4/p107-Cdk2 complexes during S-phase progression (Takahashi et al., 2000; Farkas et al., 2002).
Although less is known about the role of E2Fs in the G2/M-phase transition, recent work indicates that cooperative promoter occupancy by multiple E2Fs also contributes to the transcriptional control during this phase of the cell cycle (Zhu et al.,
2004). Cell cycle-specific fluctuations in the levels of E2F4 bound to the promoters of its target genes actively repressed transcription (Takahashi et al., 2000). The progression through the cell cycle is dependent on the degree of phosphorylation of pocket proteins, which regulate E2F4 association, resulting in the deactivation of repressive complexes, as well as the nuclear localization of E2F4 (Farkas et al., 2002; Gaubatz et al., 2001).
79 pRb-deficient cells show activated mRNA and protein levels of E2F target genes
(Almasan et al., 1995; Herrera et al., 1996). Moreover, doxorubicin-induced DNA
damage, coupled with the introduction of the E7 papilloma virus protein, which binds and inactivates the Rb-family members, reduced the capacity of colorectal cells to undergo
G2 arrest via derepression of Stathmin and AIM-1 (Polager and Ginsberg, 2003). Recent work has implicated p130 in the mechanism of G2 arrest following treatment with either etoposide or adriamycin (Jackson et al., 2005). Interestingly, cells that are p53-null are still capable of undergoing arrest. In contrast, cells lacking p130 and p107 cannot arrest and, as a consequence, die (Jackson et al., 2005), which is similar to the DHFR-inhibitor methotrexate-treated Rb-deficient cells (Almasan et al., 1995). Taken together, the findings support the importance of the Rb family members for the G2 arrest.
However, the pRb members interact with multiple partners and can control events in a wide range of signaling pathways, including cell cycle regulation, hypoxia (Budde et al., 2005), and cancer development (Attwooll et al., 2004; Nevins, 2001). Because of these findings, we focused our attention to the role of E2F4, which we have shown earlier to associate with p130 following irradiation (DuPree et al., 2004), in the G2 arrest. We have previously identified E2F4 binding to the hypophosphorylated form of p130 and its nuclear translocation (DuPree et al., 2004), but the significance of this interaction and how it affects cell cycle regulation remained unclear.
The current studies focused on establishing the kinetics of E2F4 and p130’s nuclear localization in response to genotoxic stress, such as E2F’s role in cell cycle control. Multiparametric cell cycle analyses clearly indicated that cells arrested at the G2 phase. This G2 arrest was sustained via E2F4 knock-down. This E2F4 depletion greatly
80 sensitized cells to irradiation-induced cell death. The identification of E2F4-repressed
target genes known to be important for the G2/M transition in cooperation with the
capacity of E2F4 to bind some target genes’ promoters under physiological conditions
suggests a role for E2F4 in the control of G2 arrest. These findings reveal a novel role for
E2F4 as a G2-arrest mediator that may play a critical role in the radiation resistance of
prostate carcinoma, which is characterized by increased E2F4 expression (Waghray et al.,
2001).
4.3. MATERIALS AND METHODS
4.3.1. Cell culture and treatment
Prostate carcinoma C4-2 cells were a gift from Dr. W. Heston (Cleveland Clinic
Foundation) and were maintained in a humidified incubator at 37° C, 5% CO2. The cells
were grown in monolayer culture in RPMI-1640 with 10% (v/v) heat-inactivated fetal
bovine serum, 50 U/ml penicillin, 50 mg/ml streptomycin (Invitrogen, Carlsbad, CA)
(Ray and Almasan, 2003).
Exponentially-growing cells were adjusted to a density of 1 x 105 cells/flask the
day before the experiment was performed. Cells were treated with 10 Gy of ionizing
radiation and incubated for specific time points at 37° C, 5% CO2. Irradiation was performed at 25 oC, using an X-ray irradiator (Pantak HF320: 320 kVP, 20 A, half-value
layer 2mm Cu, East Haven, CT) emitting at a fixed dose rate of ~2 Gy/min, which is
comparable to methods previously described (Gong et al., 1998).
81 4.3.2. Flow cytometry analyses
4.3.2.1. Propidium iodide staining
After treatment, cells were harvested at different time-points by trypsinization
(Invitrogen) and were washed with ice-cold phosphate buffered saline (PBS). The cells
were fixed with 90% methanol (–20°C) for at least 12 h. On the day of analysis, cells
were washed twice with PBS, treated with Ribonuclease A (RNase A, 10mg/ml) (Gentra
Systems, Minneapolis, MN) for 20 min, and stained with 50 µg/ml Propidium Iodide (PI)
(Sigma St Louis, MO). Samples were run on a Becton Dickinson FACScan with a 488 nm Argon laser with 3 color capabilities (Becton Dickinson, San Jose, CA). Data were
processed using FlowJo V6.1.1 software (Tree Star, Inc., San Carlos, CA).
4.3.2.2. Multiparametric staining (MPM-2, Cyclin A, Cyclin B1, DNA)
Cells were harvested and fixed as above. One million cells per time point were
incubated with antibodies that were labeled with fluorescent dyes prior to DNA staining,
as previously described (Yan et al., 2004). The fixed cells were washed twice with ice- cold PBS and once with PBS/bovine serum albumin (BSA; 20 mg/ml in PBS). Cells were
stained for 1 h with Fluorescein-5-EX, succinimidyl ester- (FSE) conjugated anti-Mpm2
(Upstate, Lake Placid, NY), Alexa 647-conjugated anti-Cyclin B1 (GNS1 clone (Yan et
al., 2004)), and Phycoerythrin (PE)-conjugated anti-Cyclin A (Beckman Coulter, Miami,
FL). Cells were washed three times with PBS/BSA and stained with Hoechst 33342
(Invitrogen) for 30 min prior to analysis. The fluorescence measurements were done on a
BD LSR II Flow Cytometer (BD Biosystems).
82 4.3.2.3. BrdU/PI labeling
Cells in the absence or presence of various small-interfering (si) RNAs were
pulse-labeled with bromodeoxyuridine (BrdU, 10 µM) (Sigma) by growing cells in the presence of BrdU for 45 min. Cells were then harvested and fixed overnight, as above.
One million fixed cells were washed twice with PBS. The cell pellet was treated with 4C
HCl, which was prepared in 0.1% Triton X-100 for 20 min at room temperature (RT).
PBS/BSA in 0.1% Triton X-100 was added to the cells, which were then pelleted by
centrifugation. After aspirating the supernatant, the pellet was treated with 0.1 sodium borate buffer (pH 8.5) for 2 min at RT and centrifuged again at 700 x g for 5 min. The supernatant was aspirated and the pellet was washed once with PBS/BSA and twice with
PBS/BSA in 0.1% Triton X-100. The cells were stained with 20 µl of FITC-conjugated anti-BrdU (BD Biosciences, San Jose, CA) for 1 h at 37°C in dark. Cells were washed twice with PBS/BSA, treated with RNAse A (10mg/ml) (Gentra Systems) for 20 min, and stained with PI (50 µg/ml) (Sigma).
4.3.2.4. Caspase activation assay
The CaspaTag™ Pan-Caspase In Situ Assay Kit, Fluorescein (Chemicon
International, Temecula, CA) was used to examine caspase activation in cells that were
treated with siRNA against E2F4 in the absence or presence of IR treatment. Cells were
incubated with the Fluorochrome Inhibitors of Caspases (FLICA) reagent for 1 h prior to
trypsinization, collected, and washed three times with the wash buffer, as described by
the manufacturer. The inhibitor, a carboxyfluorescein-labeled fluoromethyl ketone peptide inhibitor of caspase (FAM-VAD-FMK), binds covalently to activated caspases’
83 reactive cysteine residue that is localized on the large subunit of the active caspase
heterodimer. This reaction inhibits further enzymatic reactivity and the green fluorescence produced by the bound reagent is directly proportional to the amount of active caspases at the time of sample preparation. Samples were analyzed for caspase positive (+) or negative (-) cells on a BD LSR II Flow Cytometer (BD Biosystems).
4.3.3. siRNA
ds siRNA (E2F4) was prepared using sense and anti-sense RNA oligonucleotides, which were prepared according to the procedures indicated by Ambion™ and as described previously (DuPree et al., 2004). C4-2 cells, grown to ~50% confluence, were transfected with siRNA-E2F4 (100 nM) using Oligofectamine (2.6 µl/ml) (Invitrogen) and Optimem (10 µl/ml) (Invitrogen) and then irradiated with 10 Gy IR. To monitor for non-specific siRNA transfection-induced events, we included an ineffective (I) siRNA against E2F4 (siRNA E2F4 #2, I), a second siGAPDH control, as well as the vehicle control, which were the transfection reagents. Cells were harvested 24 h post-IR and mRNA levels for E2F4 target genes were analyzed by Real time quantitative-polymerase chain reaction (RTQ-PCR). In the case of cell cycle and BrdU labeling experiments, cells were transfected 24 h prior to IR and collected at the indicated time-points following IR.
4.3.4. Clonogenic assay
Exponentially growing cells (pretreated with siRNA against E2F4 as indicated above and untreated) were trypsinized, and cell concentration was determined. Known numbers of cells were inoculated in Petri dishes and exposed to graded doses (0-10 Gy)
84 of X-ray radiation. After irradiation, cells were allowed to grow until the surviving cells
produced visible colonies of at least 50 cells/colony. After 12 days, the colonies were
fixed and stained in methanol: acetic acid (75: 25, v/v) containing 0.5% crystal violet
(w/v) for (12 days).
4.3.5. Determination of surviving fraction
The colonies containing more than 50 cells were counted and the plating efficacy (PE)
and surviving fraction (SF) were determined as:
PE = (Colonies counted × 100)/Cells plated
SF = Plating efficacy of treated group/ Plating efficacy of control group
4.3.6. Western Blot detection
Total protein, whose concentration was estimated by the Bio-Rad Protein Assay
(Bio-Rad Laboratories, Hercules, CA) from whole cell lysates, was electrophoresed (25
µg) on a 10% SDS-PAGE gel, as described previously (Crosby et al., 2004; DuPree et al.,
2004). Proteins were then electrotransferred onto nitrocellulose membranes that were
probed for E2F1 (C-20; 1:500), E2F4 (C-19; 1:200), Cyclin A (H-432; 1:1000), Cyclin
B1 (GNS1; 1:1000), all from Santa Cruz Biotechnology, Inc, Santa Cruz, CA), or Pttg1
(1:1000; MBL International Corporation, Woburn, MA). Additionally, β-actin (A5441;
1:1000; Sigma) was used as a loading control. Blots were then visualized with either sheep anti-rabbit or sheep anti-mouse secondary antibodies, which were conjugated with horseradish peroxidase (HRP) (1:2000; Amersham Biosciences, Piscataway, NJ) or
LumiGLO Chemiluminescent reagents (KPL, Gaithersburg, MD).
85 4.3.7. Confocal microscopy
Cells grown on cover slips were fixed with 3.7% formaldehyde/PBS for 20 min at
RT. Following fixation, the cells were washed three times with PBS, blocked, and
permeabilized for 5 min at RT with blocking buffer (2% goat serum in 0.3% Triton X-
100 in PBS). The cells were incubated with anti-E2F4 and anti-p130 antibodies (1:100 in
Triton X-100 blocking buffer; Santa Cruz Biotechnology) for 1 h at RT. The cells were
again rinsed three times and incubated with Alexa 488- and 594-conjugated anti-
rabbit/anti-mouse secondary antibodies (1:200, in Triton X-100 blocking buffer;
Invitrogen), respectively for 45 min in dark at RT. The coverslips were rinsed three times
with blocking buffer, mounted with Vectashield (Vector Laboratories, Burlingame, CA),
and the fluorescence signals were examined using a Leica TCS-SP2 (Leica Microsystems
AG, Wetzlar, Germany) microscope at 40 x magnification.
4.3.8. Comet assay
Trypsinized cells were cryogenically frozen in freezing solution (80% FBS, 10%
RPMI, and 10% DMSO). On the next day, the cells were washed in 1x PBS and added to
pre-heated (37°C) Low-melting point (LM) Agarose (650 µL). The solution was pipetted onto slides (pre-covered with 1% agarose) and spread evenly over the slides. The samples were incubated in dark (4°C) to ensure that the LM Agarose solidified. The chilled slides were placed into pre-chilled lysis buffer and cells were allowed to lyse for 40 min (4° C).
Following lysis, the slides were immersed in alkaline solution (pH > 13). The slides were left in the solution for 30-35 min (4°C). The slides were placed into a horizontal electrophoresis chamber, aligning them equidistantly from the electrodes. A
86 rate of 1 V/cm was applied to the samples and run ~30 min (4°C). The slides were
washed with deionized H20 to remove the alkaline buffer (20 min). Samples were air- dried and kept in dark at RT. PI (50 µg/ml) was added to the samples, which were again dried, and kept in the dark prior to being viewed by microscopy.
4.3.9. C4-2 expression profiling oligonucleotide arrays (HU-95)
Double-stranded cDNA was prepared from the PolyA+ mRNA using the GIBCO-
BRL SuperScript Choice System (GIBCO-BRL, Rockville, MD). An in vitro
transcription reaction was used to produce biotin-labeled cRNA using the Enzo BioArray
High Yield RNA Transcript Labeling Kit (Affymetrix, Santa Clara, CA). The sample was
then purified using the QIAGEN RNA MiniKit (Valencia, CA) and then fragmented. A
hybridization mixture was prepared including the fragmented cRNA, the Affymetrix
recommended control cocktail and herring sperm DNA. For fluorescent labeling of
samples and hybridization to glass slides, 100 µg of total RNA was annealed to oligo(dT) and reverse transcribed in presence of Cy3- labeled dUTP or Cy5 labeled dUTP. The resulting cDNA was purified using GFXTM PCR, DNA and Gel Band Purification Kit
(Amersham Pharmacia Biotech Inc., Piscataway, NJ) resuspended in 20 µl hybridization
buffer (20 x SSC, tRNA, polyA, 5% SDS), heat denatured and applied to the slide, which
was then placed in a sealed humidified hybridization chamber at 65°C over night. The unbound probe was removed from the slide in a three-step wash with 2 x SSC/0.1% SDS,
2 x SSC, and 0.2 x SSC for 5 min each using Affymetrix Fluidics Station 400,
FlexGE:WS2v3 Protocol (Affymetrix, Santa Clara, CA).
87 The Affymetrix Microarray Suite 5.0 with statistical algorithm was used for analysis (Corner+ Avg:618, Count:32 Corner- Avg:29779, Count:32 Background
Avg:442.78, Stdev:7.80, Max:459.7, Min:425.1 Noise Avg:13.72, Stdev:0.65, Max:15.2,
Min:12.4). Additionally, the Microarray Suite 5 output data was further processed with
Silicon Genetics' GeneSpring(R) version 6, build 1333. Each chip (A-E) was normalized separately.
4.3.10. RTQ-PCR
PCR amplification reactions were comprised of 1 x iQ SYBR Green Supermix
(Bio-Rad Laboratories, Hercules, CA) and 4 µM each of sense and anti-sense primer.
Reactions were carried out in a 96-well optical reaction plate with optical caps (Applied
Biosystems, Foster City, CA) in an iCycler iQ Real-Time PCR Detection System spectrofluorometric thermal cycler (Bio-Rad Laboratories, Hercules, CA, USA) with an initial 3 min incubation at 95° C followed by 40 cycles of amplification: 95°C for 15 sec
and 60°C for 1 min. RTQ-PCR primers used for detecting cDNA were synthesized by
Integrated DNA Technologies, Inc. (Coralville, IA).
4.3.11. Chromatin Immunoprecipitation Assay (ChIP)
Chromatin immunoprecipitation experiments were performed as previously
described with some modifications (Crosby et al., 2004). Briefly, treated and control cells
were incubated in culture media containing formaldehyde (1%) for 10 min at RT, with
rotation, to crosslink the protein to the DNA. The reaction was stopped with glycine
(final concentration 0.125 M, 5 min incubation at RT). Cells were trypsinized and
88 collected by gentle centrifugation at RT. Cells were washed once with ice-cold PBS and gently centrifuged at 4°C prior to lysis and chromatin sonication.
Chromatin was pre-cleared with pre-blocked Pansorbin® (Calbiochem, La Jolla,
CA) (10-15 µl/1x107 cells). The Pansorbin® was incubated with the chromatin for 15
min at RT and samples were centrifuged. Pre-cleared chromatin from treated and
untreated cells was incubated overnight (1 µg/sample condition) with anti-E2F4 (C-19;
Santa Cruz Biotechnology) polyclonal antibody. As a negative or mock control, the
antibody was incubated with 1x dialysis buffer [2 mM EDTA and 50 mM Tris-Cl (pH
8.0)]. Chromatin complexes were immunoprecipitated with Pansorbin®. Elution buffer
(50 mM NaHCO3, 1% SDS) was added to the complexes to separate them from the
Pansorbin®. Two sequential elution steps were performed in which the complexes were
spun on a rotating platform for 15 min and centrifuged at 500 x g. The supernatants from the elutions were collected and combined. Adding 0.5 M NaCl and heating the complexes for 4 h at 67°C reversed the formaldehyde crosslinking.
Following DNA purification from immunoprecipitated chromatin complexes, which involved the sequential RNase A treatment (10 mg/ml), Proteinase K treatment (10 mg/ml), and DNA precipitation, 40 cycles of RTQ-PCR was performed using 2-3 µl purified ChIP DNA/reaction. RTQ-PCR primers used for detecting ChIP DNA were previously described (Ren et al., 2002) and synthesized by Integrated DNA Technologies,
Inc. The following primers were used: Bub3: (sense) 5’-GCC CAA AAT GGG ATT CTT
GTG -3’, (antisense) 5’-TCA TTT TAC CAC CGC GCT GGG-3’; Pttg1: (sense) 5’-
AAG ACC TGC GTG AGT GAA TGG-3’, (antisense) 5’-CCA GCT CTC AAA TCT
TCC AGC-3’; Cdc6: (sense) 5’-AAA GGC TCT GTG ACT ACA GCC A-3’, (antisense)
89 5’-GAT CCT TCT CAC GTC TCT CAC A-3’; β-actin: (sense) 5’-AAC TCT CCC TCC
TCC TCT TCC TC-3’, (antisense) 5’-GAG CCA TAA AAG GCA ACT TTC GG-3’.
Amplified reactions were separated on a 2% agarose gel stained with Ethidium Bromide
(Bio-Rad Laboratories, Hercules, CA).
4.4. RESULTS
4.4.1. C4-2 prostate carcinoma cells undergo a G2/M arrest following IR
Varying in their response to IR, some cells will arrest and others will undergo
apoptosis, which in some cases is under the control of the p53 transcription factor. C4-2 cells arrested at the G2/M transition of the cell cycle following IR. To determine if p53
activity was required for the radiation-induced cell cycle arrest, C4-2 cells were stably
transfected with a p53 dominant-negative (dn) harboring a C-terminal fragment
(Ossovskaya et al., 1996), such that following IR, its transcriptional activity was
abrogated, as shown by a dramatic decrease from abundant to undetectable levels of p21
(Ray, Gupta, Macklis, & Almasan, unpublished). Both parental C4-2 (Fig. 4-1A) and
derivative C4-2 cells containing the stably expressed dn p53 (Fig. 4-1B) arrested at 6 h post-IR with maximal effect at 16 h. During this time, there was a dramatic decrease in the S-phase cell population, with the G1 population remaining constant. However, recovery of the normal cell cycle phase distribution did not occur at least through 48 h post-IR. At that time, the population of G2/M-phase cells was still predominant, as compared to the proportion of S-phase cells. Taken together, these data indicate that the cells arrested predominantly in G2/M regardless of their p53 status, which argues that p53 is not required for this arrest.
90
A. 100% G2/M 90% S 80% G1
s 70%
l
l
e
c 60%
l
a t 50%
o
T
f 40%
o
% 30% 20% 10% 0% 0 6 12 24 48 Time after IR (h) C4-2 Time(h)0 6 122448 G1 50.76 34.82 42.52 33.82 35.79 S 32.54 52.7 6.3 6.14 10.24 G2/M 16.7 12.48 51.91 60.4 53.97
91 B. 100% G2/M 90% S 80% G1
s 70%
l
l
e
C 60%
l
a t 50%
o
T
f 40%
o
% 30% 20% 10% 0% 0 6 12 24 48 Time after IR (h) C4-2 dn p53 Time(h)0 6 122448 G1 35.22 18.49 34.6 16.53 24.56 S 48.34 64.16 28.86 22.4 17.88 G2/M 16.44 17.36 36.54 61.07 57.56
Fig. 4-1. Radiation induces a G2/M cell-cycle arrest independently of p53 function in prostate carcinoma C4-2 cells. Flow cytometry analyses, to determine the number of cells in the various phases of the cell cycle by staining for DNA content at different time- points following IR, were performed as described in the “Materials and Methods.” Cells were collected at the intervals indicated and fixed in 90% methanol. Cells were washed, treated with RNase A, stained with PI, and analyzed by flow cytometry. Percentages of
C4-2 (A) and C4-2 dn p53 (B) cells in the different phases of the cell cycle are presented with respect to time. Data are representative of three separate and independent experiments.
92 Distinguishing G2 from M cells is not possible using propidium iodide (PI)
staining alone, as both populations have 4C DNA content. [Rather than “4N”, “4C” is
used to distinguish DNA content independent of its ploidy (Jacobberger, J.W., personal
communication).] Therefore, a multi-parametric approach utilizing highly specific cell
cycle markers was used to delineate the cell cycle distribution. Thus, we employed a strategy to investigate the levels of Cyclin A2 and Cyclin B1, which are known to regulate the G2/M transition, as well as the proteins that are specifically phosphorylated during mitosis and are stained with MPM-2 (Yan et al., 2004). This approach is similar to the multiparametric staining that utilizes the phosphorylation status of histone 3. The three markers were combined with Hoechst staining for DNA content, thus providing a four-dimensional analysis of the cells.
Focusing on the mitotic population of cells, which is defined by DNA content and
MPM-2 positive staining, we were able to delineate the population of G2- versus M- phase cells after IR treatment. In control cells, a 3.2% mitotic population was present, which was almost completely depleted (~0.09 %) following IR (Fig. 4-2A). The decrease in the mitotic population indicates that these cells are arrested in G2, as almost no mitotic cells are found after IR. Based on the light scatter profiles shown in the upper right-hand corners of the Cyclin A2 plot, this population of A-/B- state is not apoptotic (Fig. 4-2A).
By examining the ratio of Cyclin A2-positive to Cyclin B1-positive cells, we hoped to further define this G2 arrest as being early, as characterized by higher Cyclin A2 levels or as late, as distinguished by higher Cyclin B1 levels. Our results indicate that the cells arrest at the G2 phase, as visualized by a population of cells that stains positively for both
Cyclin A2 and Cyclin B1. Positive staining for both cyclins asymptotically decreased at
93 12 – 24 h post-IR (Fig. 4-2B). While most cells are no longer positively staining for these
cyclins, there are some expressing these cyclins, perhaps suggesting that these cells are
proceeding into a Cylin A-/B- state after IR. Based on the light scatter profiles shown in
the upper right-hand corners of the Cyclin A2 plot, this population of A-/B- state is not
apoptotic (Fig. 4-2A). These results were confirmed by immunoblotting experiments for
Cyclin A2 and Cyclin B1 (Fig. 4-2C), which indicated that their levels were ~ 3-fold and
~1.5-fold lower, respectively, in irradiated cells at 12 h post-IR. Paradoxically, cells
containing high levels of Cyclin A are associated with the G2 phase, while high levels of
Cyclin B are indicative of both the G2 and the M phases of the cell cycle. Interestingly,
the cells with 4C DNA content had low levels of both Cyclin A and Cyclin B. A 4C
DNA-content has also been suggested for cells undergoing an arrest at the G1 phase, as
they undergo endoreduplication or those that proceed through mitosis inappropriately and
as a result undergo mitotic catastrophe. These cells did not stain positively for the MPM-
2-mitosis marker through the 24 h time-period of the experiment.
94 A.
Control 6 h 12 h 24 h
00 500500 1000 0 500 500 1000 00 500500 1000 00 500 500 1000 DNA DNA
95 B.
Time (h) G1 S G2 M A+/B+ CycA- 4C C 34 52 13 3.2 61.5 2.1 6 31 48 21 0.025 60.3 8 12 31 32 37 0.089 28 37 24 35 24 41 0.095 20 43
A+/B+ 70 CycA- 4C
60
s 50
40
30
Percent of Total Cell Total of Percent 20
10
0 C61224 Time (h)
96
C. Time(h)03 6122448 Cyclin B1
Cyclin A2
β -actin
Fig. 4-2. The radiation-induced cell cycle arrest is specific to the G2 phase of the cell cycle. Multiparametric staining using the mitotic marker, MPM-2 (FSE, y-axis), Cyclin A
(PE, y-axis), Cyclin B1 (Alexa 647, y-axis) and Hoescht 33342 (x-axis) for DNA content, was performed (A). The proportion of mitotic cells, highlighted with an arrow in A, was examined at the indicated times following IR (10 Gy) treatment. Bivariate analyses indicate that staining for Cyclin A and B decrease with time, yet most cells retain 4C
DNA content (B). For each staining combination and analysis, cells were collected before
(0 h) and at 6, 12, and 24 h after IR. Immunoblotting for Cyclin A2 and Cyclin B1 was also performed using the respective primary antibodies (C). β-actin was used as a loading control.
97 4.4.2. E2F4/p130 complexes are formed following IR
As we have previously reported, IR induced complex formation of E2F4, but not
E2F5 with the unphosphorylated form of p130 (DuPree et al., 2004). Our previous work also indicated that E2F4 was present in the nuclei of irradiated cells at 24 h after IR treatment. However, the kinetics of E2F4 localization following IR and its potential colocalization with p130 with respect to cell cycle arrest was not determined. To address this, we first investigated the levels of E2F4 in C4-2 cells following treatment with IR or, for comparison, with a topoisomerase II inhibitor (VP-16). Levels of E2F4 were slightly decreased at 8 h following IR, then increased above control levels, which were maintained through at least 72 h post-IR, indicating that its sustained presence could contribute to the G2 arrest (Fig. 4-3A). VP16 treatment led to a slightly decreased level at
16 h and with an additional decrease at 24 h. In great contrast, the protein levels of the
E2F1 transcription factor, which may counterbalance the activity of E2F4 (Crosby and
Almasan, 2004), were dramatically down-regulated by 24 h. Decreased E2F1 levels were sustained through at least 72 h post-IR (Fig. 4-3A).
Examining the dynamics of its subcellular localization by confocal microscopy indicates that E2F4 could be detected in the nucleus as early as 4 h post-IR (Fig. 4-3B).
The same kinetics was obtained for p130 with the fluorescence signal overlay indicating
E2F4 colocalization with p130 in the nucleus. The E2F4/p130 colocalization was time- dependent reaching a peak at 24 h post-IR. These results indicate a time-dependent subcellular localization of the E2F4/p130-containing complexes, which together with the sustained levels of E2F4, but not E2F1, support an important role of E2F4 in the IR- response.
98
A. Time ( h) 8 24 48 72 Agent C IR VP-1 6 IR VP-1 6 IR VP-1 6 IR VP-1 6
E2 F4
β? -actin
Time (h) 8 24 48 72 Agent C IR V P- 1 6 IR V P-1 6 IR V P-1 6 IR V P-1 6
E2 F1
β? -actin
99 B.
I
P
A
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p
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100 Fig. 4-3. E2F4 levels are sustained after IR and E2F4 co-localizes with p130 at the time
of G2 arrest. IR and VP-16 treatments lead to sustained levels of E2F4 protein
expression. C4-2 cells were left untreated (control), treated with 10 µM VP-16 or
irradiated (10 Gy), and harvested at the time points indicated. Cell lysates were prepared
and equal amounts of total protein were resolved by 10% SDS-PAGE. Immunoblotting
was performed with anti-E2F4 and anti-E2F-1 antibodies. β-actin was used as a loading control (A). Immunofluorescence confocal microscopy (40 x magnification) with antibodies specific for E2F4 and p130 indicated colocalization following IR. Merging the fluorescence signals provided by the two antibodies indicates their nuclear co-localization following IR (B).
101 4.4.3. A physiological role for E2F4 following IR in the G2/M phase control of the cell
cycle
To investigate the functional role of E2F4’s nuclear translocation after IR, we
examined the cell cycle arrest profile following IR in cells in which E2F4 levels were
diminished. We took advantage of siRNAs that we have previously designed and tested
to effectively knock-down E2F4 levels (DuPree et al., 2004). To address the
physiological role of E2F4 following IR, cell cycle analyses were performed in cells that
had been transfected with E2F4 siRNA. Cell treatment with vehicle control, GAPDH-
specific, or non-specific E2F4 siRNA, did not result in any substantial changes in the cell
cycle distribution (Fig. 4-4B). In contrast to the control-treated cells, those exposed to
E2F4 siRNA exhibited a substantial decrease in the proportion of the G2/M phase cell
population at 12 h post-IR (Fig. 4-4C). Moreover, there was an increase in the S-phase
proportion of cells that had been treated with siRNA E2F4 at 12 h post-IR (Fig. 4-4C).
Additionally, PI-staining identified a population of cells with a sub-G1 DNA content,
indicating that some cells were undergoing apoptosis (Fig. 4-4D). These effects were enhanced at 24 h post-IR.
102 ) (I 2 1 A. # # H l 4 o 4 D F tr F P 2 n 2 A iE o iE iG s C s s
E2F4
β -actin
B. siE2F4 [100 nM, Ineffective (I)] G2/M S 100% G1 90% 80% 70% 60% 50% 40% % of cell cycle 30% 20% 10% 0% control siE2F4 siE2F4 IR (12 ) siE2F4 IR (24 h)
siGAPDH (100 nM) G2/M S
100% G1 90% 80% 70% 60% 50% 40%
% of cell cycle 30% 20% 10% 0% control siGAPDH siGAPDH IR (12 h) siGAPDH IR (24 h)
103
C. siE2F4 (100 nM) G2/M S G1 100% 90% 80% 70% 60% 50% 40% % of cell cycle cell % of 30% 20% 10% 0% control siE2F4 siE2F4 IR (12 h) siE2F4 IR (24 h)
D. IR (12 h) + siE2F4
sub-G1
DNA DNA
104
Fig. 4-4. E2F4 downregulation causes an inappropriate G2 cell cycle arrest. Cells were treated with a vehicle control, an a siRNA against GAPDH, an ineffective siRNA against
E2F4, or a selective siRNA against E2F4 24 hours prior to collection and immunoblot processing (all on the same immunoblot), indicating that the specific siRNA against E2F4
(#1) was highly effective (A). The non-specific E2F4 siRNAs did not cause a marked decrease in the G2 arrest (B). In contrast, the selective siRNA against E2F4 caused an apparent increase in the S-phase population (C), as well as a sub-G1 population of cells
(D).
105 These results indicate an increased population of S-phase cells in the irradiated
siRNA E2F4-treated cells. There was a population of cells that were apparently formerly
G2/M cells, which could be recycling back to S-phase and undergoing apoptosis (Oancea et al., 2006). BrdU-pulse labeling was therefore used to distinguish between S-phase cells that are readily dividing and those with a decreased DNA content corresponding to what
is expected for cells in S-phase. The cells required about 12 h to achieve a robust G2-
arrest, with a considerably diminished proportion of S-phase cells. BrdU incorporation
decreased considerably in a time-dependent manner following IR, with a minimal S-
phase population left at 16 h following IR (Fig. 4-5A). Thus, in control irradiated cells,
BrdU incorporation was detected in approximately 20% of the cells. In contrast, at 16 h
following IR, there were only few cells that remained positive for BrdU. Those cells that
were BrdU-positive were part of the population of G2/M phase cells, which were
presumably cells transitioning from the S to the G2 phase during the 45 min BrdU pulse.
Surprisingly, in contrast to the control or irradiated C4-2 cells, cells that were treated with
E2F4 siRNA prior to irradiation and that appeared to be in S-phase based on their DNA
content (Fig. 4-4C), did not incorporate BrdU (Fig. 4-5B). Cells that were treated with siE2F4 or siE2F4 + IR and that were unable to incorporate BrdU were 8% and 42%, respectively.
106
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9
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6
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+
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2
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2 anti-BrdU +
(
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A
N
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5
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9
1
+
)
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A
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7
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6
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+
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3
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1
2
+
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C anti-BrdU A. B.
107
Fig. 4-5. Lack of BrdU incorporation indicates that formerly arrested G2 cells are not
recycling. Dual staining measuring the incorporation of BrdU and of DNA content
following the transfection and irradiation was performed to eliminate the possibility that
the cells were re-entering and populating the S-phase of the cell cycle. Boxes inside each
plot indicate the gating used for identifying BrdU positive cells and the percentages of
BrdU positive cells are given in the upper left-hand corner of the box. Cells were treated
with vehicle alone or with siRNA against E2F4, incubated for 24 h, irradiated, and labeled 45 min prior to collection at either 6 or 12 h following IR (A). Although control
cells show active incorporation of BrdU, the cells transfected and collected at 16 h
following IR had a BrdU-negative S-phase population (B).
108 These results indicate that the cells with diminished E2F4 expression were not recycling back to S-phase. These were most likely cells arrested at the G2 phase that had
diminished DNA content because they were undergoing cell death. Whereas cells
arrested in G2 after IR usually comprised ~ 60% of the total cell population, the G2
population in the siE2F4-treated and irradiated cells was reduced to ~ 35%. To
determine whether these cells were undergoing apoptosis, caspase activation, a hallmark
of cell death, was examined. Staining with CaspaTag™, a unique reagent that binds covalently to activated caspases through an available cysteine residue, indicated that 40% of the cells treated with siE2F4 had activated caspases at 16 h post-IR (Fig. 4-6A). There
was caspase activation by siE2F4 transfection alone in about 15-20% of cells. In addition
to caspase activation, comet assays indicated that the combination of siE2F4 and IR
increased DNA strand breakage, as seen in the more pronounced comet tails, as
compared to IR-treated alone, or siE2F4 treated cells (4-6B). To determine the long-term
impact of the E2F4 knock-down, we examined clonogenic cell survival following
treatment with different doses of ionizing radiation. Our results demonstrate that the
E2F4 siRNA-treated cells are more sensitive to radiation, as indicated by the LD50
decrease from 2.1 Gy to 1.1 Gy (Fig. 4-7). There was a significant decrease in clonogenic
survival at 4 Gy.
109 A.
Control IR (16 h)
siE2F4 (16 h) siE2F4 + IR (16 h)
110
B.
Control IR
siE2F4 siE2F4 + IR
Fig. 4-6. E2F4 knock-down by siRNA initiates caspase activation and DNA strand
breaks. The CaspaTag™ assay was used to analyze caspase activation; proteolytic
caspase activation results in free cysteine residues that covalently bind fluorescent
substrate (A). Comet assays were performed under alkaline conditions to determine the amount of single- and double-strand DNA breaks (B).
111 Colony Formation Capacity Colony Formation Capacity
Radiation Dose (Gy)
Fig. 4-7. Clonogenic assay indicates that E2F4 knock-down can sensitize cells to IR.
Cells were plated (plating efficacy 80 -80%), treated with siRNA against E2F4 or vehicle alone and irradiated. After treatment with siE2F4 (100 nM, 24 h), cells were re-plated logarithmically (100 cells/ml - 2500 cells/ml, depending on radiation dose and in a final volume of 4 ml) followed by irradiation (0-6 Gy) and incubated for 10 days, forming visible colonies. After 10 days, the colonies were fixed, stained with 1% crystal violet (in methanol: acetic acid). The capacity to form colonies was calculated with respect to the untreated control and the results are expressed as the mean ± SD. (The significance was tested between IR and siE2F4 + IR by F test by using Prism software.) Cells were seeded at an appropriate density and colony formation was determined by the colony-forming assay and modeled. Clonogenic studies were done in triplicate for each data point.
112 4.4.4. Identification of putative E2F4 target genes involved in the IR response
We next sought to identify genes whose expression was associated with the IR- induced G2 arrest by performing oligonucleotide microarray experiments on untreated
LNCaP, C4-2, and irradiated C4-2 cells. We mined our data (Buchsbaum et al., unpublished) for genes that were both > 2-fold down-regulated at 6 and 24 h following IR
(Ren et al., 2002). From this experiment, we identified genes that fulfilled these criteria
(Table 4-1). Fold changes are given with respect to their relative abundances found in control cells. We then verified that some of these transcripts, previously suggested to have a role in the G2/M checkpoint (Polager and Ginsberg, 2003; Ren et al., 2002) were indeed downregulated by performing RTQ-PCR using the primers listed in Table 4-2.
Although the changes in gene expression levels obtained from RTQ-PCR were not as dramatic as those obtained from the microarray data, the transcripts were all found to be down-regulated by an average of 3-fold (Fig. 4-8A). Clearly, the overall trends of downregulation were seen in both the expression profiling data, as well as in the RTQ-
PCR experiments. At 6 h following IR, the transcript levels were downregulated at least
2-fold. At this time-point, the most dramatic repression was observed for the Cyclin E1
(5.5-fold), Cyclin B1 (6.3-fold), and Chek1 (3.5-fold) transcripts. Interestingly, the transcripts for Pttg1 and Bub3 seemed to be delayed in their downregulation, with their levels continuing to decrease in a time-dependent manner through the duration of the experiment. In comparison, other transcripts, particularly those for Cyclin B1 and Cyclin
E1 were downregulated rapidly by 6 h and then started to increase again at 24 h.
113
Time (h) Gene 0 6 24
Bub1b 1.4 -2.5 -16.1 Cenpe 0.9 -2 -24.6
Chek1 1 -8.8 -3.8 Hec 7.2 -8.7 -23.9
Mad2L1 3.5 -2.4 -3.8
Pttg1 28.7 -4.4 -10.2
Cdc6 1.3 -2.6 -2
Table 4-1. E2F4 target genes identified by HU-95 Affymetrix array
114
SENSE (5' to 3') ANTISENSE (5' to 3') GAPDH: CAC CAC TGA CAC GTT GGC AG GAA ACT GTG GCG TGA TGG C Bub3: TTG GTG TAA GTC TGA ACC CAT CTT T CAC AGT AAC TCT AAC ACA TCC CTT AGG G Cyclin E1: CCC ATC CTT CTC CAC CAA AG CCC TGT TTG ATG CCA TCC AC Cdc6: GCC TCA GCC TCC CGA GTA G TAG GGA GGC CAA GGT GGG Chek1: TGG TTG ACT TCC GGC TTT CT TTC ACC AGG ATT CCC CAG AG Cyclin B1: TGG TGA AGA GGA AGC CAT GG AAG AGC TGT TCT TGG CCT CAGT Hec: CAG AGG CAA AGA AGC GAT TGA TTT GAC AAG GCA GTT GGC AC Mad2l1: TGA GGT CCT GGA AAG ATG GC CAG TGG CAG AAA TGT CAC CG p107: ATC CAG GTA CCA CCG CCA T TGG GTT GTG ACA TCA GGC TG Pttg1: TTT GAC CTG CCT GAA GAG CA GAT TGG ATT CCC ATG GTG GA
Table 4-2. Putative E2F4 target genes and their respective primer sequences
115 A. mRNA expression Pttg1 0 Cdc6 02624p107 Mad2L1 -1 Chek1 Cyclin E1 -2 Bub3 Cyclin B1 -3
-4 Fold Change
-5
-6
-7 Time (h)
B. Time (h) 0 3 6 12 24 48
Pttg1
β-actin
C.
ChIP E2F4: Mock Total input Time(h) 0 424042404240 4240 0 bp 750 500 300 150 ββ -actin-actin bub3Bub3 Pttg1 pttg1 Cdc6 cdc6
116 Fig. 4-8. Irradiation induces the downregulation of E2F4 targets. RTQ-PCR was
performed to validate the levels of mRNA, of genes identified by the microarray (Table
4-1) at various time points following IR (A). The data are quantitatively expressed with
respect to the endogenous GAPDH control. Values reported are relative to the 0 h time-
point. Levels of the E2F4 target gene product, Pttg1, were analyzed by SDS-PAGE and immunoblotting (B). ChIP analyses confirmed dynamic target binding to Bub3 and Pttg1 genes and constitutive binding to Cdc6 (C). β-actin primers were used to confirm that
there was no non-specific E2F4 binding and to serve as a positive control for the “Total input.” In the “Mock” control, ChIP was carried out without chromatin to ensure no non-
specific binding.
117 To determine whether the transcriptional downregulation also led to a decrease in
the protein levels, we examined the effect of IR on two E2F target candidate gene
products, Cyclin B1 (Fig. 4-2C) and Pttg1 (Fig. 4-8B). Pttg1 levels decreased
approximately 3-fold compared to the untreated samples. Both proteins demonstrated
similar kinetics of downregulation following IR at 12 h post-IR. In general, the trends
examined at the protein level mimic those determined at the transcript level. However, in
the transcripts, there appeared to be some recovery in mRNA levels at the 24 h time-
point, but the protein levels did not recover through 48 h. These data suggest that
combined transcriptional and post-transcriptional control mechanisms regulate levels of
critical mitotic proteins in such a way that their expression is completely blocked after
IR.
To determine if E2F4 played a direct role in the regulation of the genes identified by the microarray and RTQ-PCR experiments, we next performed promoter-binding analyses that utilized our expertise in chromatin immunoprecipitation (ChIP) (Crosby et al., 2004). The promoter sites we examined were chosen based on having been defined previously from studies that investigated the putative role of E2F4 in various cell cycle phases, including G2/M functions, through combined cross-linking DNA immunoprecipitation and microarray experiments (Ren et al., 2002).
We were interested in examining the effect of E2F regulation in the absence of any perturbing factors, such as those encountered during cell synchronization experiments. We therefore investigated the promoter occupancy at the time when the
E2F4/p130 complexes entered into the nucleus and the cells exhibited the most prominent
G2 arrest, as determined by cell cycle and immunocytochemistry analyses. ChIP analyses
118 indicated that E2F4 is recruited to the promoter binding sites of Pttg1 and Bub3 at 4 h
after IR treatment. E2F4 localization on these promoter sites persisted through at least 24
h (Fig. 4-8C.). In the case of Cdc6, E2F4 was present both before and after IR, indicating
constitutive binding. As expected, there was no binding to the β-actin promoter region,
indicating that there was no non-specific binding of E2F4. Based upon these data, we
conclude that E2F4 has a direct role in the repression of multiple genes that encode
important mitotic regulators and is recruited to their promoters, such as those of Pttg1
and the Bub3 gene promoters following IR.
4.5. DISCUSSION
The G2 arrest represents a mechanism by which cells are able to halt DNA segregation and thereby prevent the accumulation of mutations, which would otherwise promote carcinogenesis (Kastan and Bartek, 2004). This arrest serves to allow repair of
DNA damage, with its outcome being either that cells resume their proliferation or, if the damage is too severe, remain permanently arrested at G2 (Linke et al., 1996), or become senescent (Narita et al., 2003). This mechanism is beneficial in the case of non-neoplastic tissues. However, in neoplastic cells, this same cell cycle block can promote resistance to therapies that rely on eliminating actively replicating cells. Clearly, identifying key mediators of G2 arrest is critical to sensitization of otherwise therapy-resistant cells.
Recently, an early and transient arrest leading to accumulation of cells at the
G2/M phase of the cell cycle has been described to be ATM-dependent (Xu et al., 2002).
A second G2 arrest that occurs later and is responsive to radiation caused in the earlier phases of the cell cycle, is independent of ATM (Iliakis et al., 2003). E2F4 is a good
119 candidate for regulating this second G2 arrest, as the arrest we have identified does not appear to be a fast transient response. Rather, the arrest is sustained, indicating ATM- independence. Indeed, the depletion of ATM by siRNA-mediated knockdown had no effect of the G2 arrest (Ray, Gupta, Macklis, & Almasan, unpublished).
Studies to identify several key proteins implicated in G2 arrests indicated that the arrest is dependent on the Rb family members (Chan et al., 2000; Taylor et al., 2001). In
the context of overexpression of p53 and in the absence of DNA damage, p130 was found
to interact with E2F4 and to repress the Cdk1 promoter (Taylor et al., 2001). Other
overexpression experiments indicated that p53 regulated the G2 arrest via Cyclin B1
repression (Innocente et al., 1999). Yet, the arrest in C4-2 cells seems to occur
independently of the levels of Cyclin B1 and associated kinase activities (Ray, Gupta,
Macklis, & Almasan, unpublished). In this system, Cyclin B1 levels and associated
kinase activities show minimal changes following IR in a p53-deficient C4-2 cell line and
these proteins are dramatically downregulated in C4-2 cells (Ray, Gupta, Macklis, &
Almasan, unpublished); both cell lines have a comparable G2 arrest.
Rather than focusing on E2F4 overexpression, we chose to examine a
physiological condition where E2F4 function was involved in the response to DNA
damage. Irradiation resulted in a slight E2F4 upregulation and its simultaneous nuclear
colocalization with p130. This was in direct contrast to decreased levels of E2F1
following IR (Rogoff and Kowalik, 2004; Stevens et al., 2003). As other reports indicated
that increased levels of E2F1 could lead to apoptosis (Rogoff and Kowalik, 2004), our
results suggest that E2F4 and not E2F1 regulates the G2 arrest following IR. The
decreased levels of E2F1 provide an explanation for the lack of apoptosis and the ensuing
120 radiation resistance of these cells. By downregulating the E2F4 protein levels by siRNA-
mediated knock-down prior to irradiation, the G2 arrest was greatly diminished.
Moreover, the cells became sensitized to radiation-induced cell killing, as shown by the increased sub-G1 content of cells, decreased proportion of G2 cells (sub-G2), and upregulated caspase activity. The overall cell cycle distribution reflected an increase in an apparent S-phase population, showing a clear reduction in the G2 proportion of cells. To our knowledge, this is the first demonstration that E2F4 is critical for the G2 arrest and that its inhibition leads to cell death in G2.
The E2F family of transcription factors has now been implicated in the control of all phases of the cell cycle, including the regulation of genes during the G2/M phase transition (Giangrande et al., 2004; Ishida et al., 2001). In cellular proliferation, E2F4 is bound to regulatory regions of target genes during the quiescent G0 phase of the cell cycle. Conversely, other activator E2Fs are replacing E2F4 on these and other regulatory regions during the other phases of the cell cycle, to allow for cell cycle progression.
Recent work has shown that at least two promoters that are regulated by the E2F family,
Cyclin B1 and Cdk1, contain both activator and repressor elements, which may regulate
increased and decreased promoter activities, respectively (Zhu et al., 2004). This and
another report (Giangrande et al., 2004) suggested that the E2F family also functioned
during the G2 phase of the cell cycle, leading to the idea that this family may have an
important role throughout the cell cycle (Ren et al., 2002; Zhu et al., 2004). An important
insight of this particular study was the identification of promoter regions, which
contained both positive- and negative-acting E2F elements. The mutation of critical E2F
sites abolished the ability of the activating cofactor, B-Myb to interact with the Cdk1
121 promoter, as B-Myb is known to be important for the expression of Cyclin B1 and Cdk1
(Zhu et al., 2004).
Previous ChIP studies indicated that E2F members occupied gene promoters in a cell cycle specific manner, suggesting that the different members had specific roles
during the cell cycle and were coordinately targeted to specific genes (Rayman et al.,
2002; Takahashi et al., 2000; Wells et al., 2000). Together with other reports indicating multiple promoter occupancies, leading to both positive- and negative transcriptional activation thresholds (Giangrande et al., 2004), we suggest a physiological role for E2F4 in the IR-dependent G2/M cell cycle arrest. That is, just as exogenous E2F 1-3 can promote cell cycle progression, we suggest that the presence of the inhibitory E2Fs, such
as E2F4, regulate the G2 cell cycle arrest. More specifically, we found E2F4 to be
responsible for repressing two critical mitotic genes, Bub3 and Pttg1, following IR. In support of our findings, it was shown that E2F4 was capable of binding to promoters that
were both regulated during the G2/M phase of the cell cycle, as well as to promoters that
could be occupied by the activating E2Fs (1-3) (Zhu et al., 2004). ChIP experiments
provided direct evidence for E2F4 binding to target genes following IR. Although finding
a particular target gene that is responsible for carrying out the function of E2F4 is of
interest, multiple genes are likely to be important for sustaining the G2 arrest following
IR (Buchsbaum et al., unpublished). This is further supported by a report that indicated
that 14-3-3σ and p21 have cooperative and non-redundant roles at the G2/M checkpoint
(Chan et al., 2000).
Irradiated C4-2 cells are able to remain arrested for an extended period of time in
culture, but eventually resume cell proliferation and form colonies. Because the levels of
122 the G2/M cyclins were low, one may suggest that the cells are capable of undergoing
endoreduplication. However, the DNA content profiles and BrdU-incorporation experiments indicated that the cells were not undergoing additional rounds of DNA replication beyond the G2 block. Moreover, this is no indication that the cells ever
reached mitosis based on examining specific (Cyclin A and B1) and general (MPM-2)
mitotic markers. Had the cells been recovering from arrest, abnormal mitoses should have
occurred, such as those characterized in mitotic catastrophe (Chang et al., 2000).
Interestingly, prostate cancer epithelial cells used in our investigation are known to not
undergo such events (Roninson I, personal communication). pRb-negative but not pRb-
positive cells are known to undergo endoreduplication (Niculescu et al., 1998). Since C4-
2 cells have functional Rb and p21, whose deficiency has been implicated in this process
(Chang et al., 2000), it suggests that these cells undergo cell cycle exit initiated at the G2
phase of the cell cycle that does not involve endoreduplication or mitotic catastrophe.
Interestingly, a number of E2F4-bound genes originally identified through array
analyses in human primary fibroblasts were also bound to E2F1 (Ren et al., 2002). This
observation has been extended recently to the promoters of genes that are regulated
during the G2/M phase of the cell cycle, which contain distinct E2F4 and E2F1 binding
sites (Zhu et al., 2004). Therefore, there is likely a balance between the E2Fs that occupy
these promoters, indicating that having more E2F4 bound may result in cell cycle arrest
(Crosby and Almasan, 2004). Conversely, having more E2F1 bound is expected to allow
for continued progression through the cell cycle. In the case of DNA damage, E2F1
levels may increase to promote cell death (Crosby and Almasan, 2004) (Crosby and
Almasan, 2004; Rogoff and Kowalik, 2004). In C4-2 cells subjected to genotoxic stress,
123 the expression levels of E2F1 decrease and the levels E2F4 are sustained. Whereas
increased levels of E2F1 are indicative of apoptosis, decreased levels may allow for
increased cell survival as a result of not activating E2F1 apoptotic target genes. E2F4
inactivation and E2F1 activation could promote apoptosis, as both may lead to increased
gene expression. Therefore, cells may avoid apoptosis through gene repression mediated
by E2F4’s nuclear colocalization with p130.
We have found that, E2F4, which was defined earlier to be involved in gene repression during G0, has a similar role in repressing genes after DNA damage. Whereas
E2F1 is known to have roles beyond its function in the G1/S transition to activate genes, we now provide evidence for E2F4 acting in a similar manner that goes beyond its established role in cellular quiescence. Whereas previous efforts were focused on determining E2F4 promoter binding in the different phases of the cell cycle, we now
propose that this protein also acts in the DNA damage stress response pathway.
Therefore, the implications of E2F4 translocation to the nucleus and its involvement
following DNA damage provide new insight into the G2 arrest response following
genotoxic stress. The biological role for each of the E2F4 repressed genes in the
genotoxic stress response is of great interest for our future studies.
124 CHAPTER 5. FUTURE DIRECTIONS: MANIPULATING CELL CYCLE
CHECKPOINTS FOR TARGETED THERAPIES FOR CANCER TREATMENT
5.1. INTRODUCTION
Traditional therapies have focused on taking advantage of the hallmarks of cancer cells, which include genetic instability and high proliferation rates, the alkylating agents, anti-metabolites, topoisomerase inhibitors, and radiomimetic anticancer drugs have been very successful in treating patients (Zhou and Bartek, 2004). Despite substantial success, some patients develop resistance over time and others do not respond at all. Moreover, the delicate therapeutic window for dose recommendation is often difficult to monitor, making them considerably toxic.
As cancers develop through misregulation of proliferative pathways, an insightful therapeutic strategy for trying to manage cancers is to utilize specific personalized molecular-based targeting. In recognizing that abnormal proliferation is due to misregulation in the cell cycle, recent therapies have focused on chemosensitization via checkpoint signaling pathways, which are controlled, in part, by Chek1 and Chek2. In the case of tumors harboring p53-deficiency, ablating Chek1 causes hypersensitization to IR in DT40 avian cells (Zachos et al., 2003) and a dominant negative Chek1 in humans leads to radiosensitization (Koniaras et al., 2001). However, other data suggest that an effector of Chek1, brca1, when mutated, can abrogate the G2/M arrest without causing
radiosensitivity, indicating that blocking Chek1 activation may not be sufficient for
treating all cancers with p53-deficiencies (Xu et al., 2001). Thus, identifying a cancer’s
125 molecular fingerprint of gene mutations and protein misregulation is critical to understanding which therapies will be most useful for favorable treatment.
5.2. E2FS AS MOLECULAR TARGETS IN CANCER THERAPY
Clearly, the balance of the E2F family members’ activities is necessary to maintain proper regulation of the cell cycle, differentiation, and oncogenesis. However, coupled with this direct control is the fact that misregulation by the E2Fs or pRB is enough to drive oncogenesis, which was identified through the fact that the E2F/pRb pathway was a viral oncoprotein target (Nevins, 1992). Indeed, deregulated pRb/E2F activity is well documented in cancers, which include osteocarcinomas, small lung carcinomas, and breast carcinomas and is associated with poor prognosis (Sherr, 1996).
The loss of the p16 cyclin kinase inhibitor results in a loss of G1 cyclin control, which leads to E2F accumulation and cell cycle misregulation (Nevins, 2001).
Although there have been many mutations identified in pRb, mutations in the
E2Fs have not been as striking, aside from genetic mutations occurring in the E2F4 trinucleotide repeat element (Nevins, 2001; Yoshitaka et al., 1996). Additionally, increased expression of E2F1 is correlated in non-small lung carcinomas and its activity is beyond the inhibitory control of pRb (Gorgoulis et al., 2002). Interestingly E2F1 expression is associated with apoptosis under cellular conditions that allow it to be coupled with p53 and to activate apoptotic gene targets. However, the overexpression of
E2F1 in cellular conditions containing aberrant apoptotic signaling, as in p53-deficiency, can bypass the threshold of apoptosis and, as a result, cells continue to proliferate
(Tsantoulis and Gorgoulis, 2005). Some traditional therapies target the gene products of
126 E2F1, such as methotrexate and DHFR (Kaelin, 2003). The ability to modulate E2F1’s target gene expression with traditional anticancer drugs and via modulation of E2F1 levels presents a new strategy of managing cancers through multiple mechanisms.
E2F4 is overexpressed in prostate tumor epithelial cells (Waghray et al., 2001).
Additionally, it is also overexpressed in colon cells and clinically appears to protect against apoptosis (Tsantoulis and Gorgoulis, 2005). In neurons, a similar protective mechanism includes the complexing of p130/E2F4 with HDAC1 and Suv39H1, which has been associated with condensed chromatin and is known to promote gene silencing
(Liu et al., 2005). Recent studies suggest that the interplay between E2F1 and E2F4 is important for sensitizing cells and that decreased levels of E2F4 can result in increased drug sensitivity (Ma et al., 2004). Based on this anti-apoptotic capability, E2F4 is an attractive therapeutic target for sensitizing cells.
5.3. SUMMARY AND FUTURE DIRECTIONS
In the past 10 years, there has been a growing appreciation of the role of the
E2F/pRb and p53 tumor suppressor pathways in mediating the G1 cell cycle arrest
(Almasan et al., 1995). Insights into pRb ablation by knock-down and knockout strategies have provided the cornerstone for determining E2F misregulation. As the same molecular networks that are involved in cellular proliferation mediate cell cycle arrest, we have now shown a link between E2F4/p130 repressor activity and G2 cell cycle arrest. The
E2F/p130 complex translocates to the nucleus following IR and appears to be required for the G2 arrest activation.
127 Although E2F4 may be a good target for sensitizing radioresistant cells, the genes that it is targeting for repression may be a more selective approach to sensitization.
Therefore, in future studies, we plan to study the genes whose transcripts were downregulated after IR. As we saw dynamic binding to the Bub3 and Pttg1 promoters, we believe that these candidate genes may be important effector molecules of the E2F4 repression model. By overexpressing these or relevant E2F4 transcriptional targets, we hope to induce radiosensitivity. As there are many target genes of E2F4, there may not be one gene that is responsible for mediating radioresistance. Most likely, there is a combination of genes that acts in concert to promote resistance.
The components of the DNA damage response are in delicate balance with the metabolic signaling pathways of the cells. In being able to find therapies that selectively enhance the therapeutic window for cancer treatment; we may be able to decrease the incidence of secondary cancers that can appear after traditional therapies. Finally, targeting E2F4 and/or its gene targets represents the future of molecular medicine in oncology, where a potential therapy takes advantage of striking differences in E2F4 expression in tumor cells versus normal cells. Future therapies should be based on the selective molecular targeting of misregulated proteins that promotes homeostasis.
Because cancers are in fact not exogenous to the body, as in bacterial or viral infections, the conceptual idea of killing can sometimes lead to drug resistance and create even more aggressive tumors. Therefore, our goal for understanding cell cycle regulation in cancer and in the DNA damage response should be to promote the conceptual framework of evaluating the expression of the major players of cellular proliferation, as they relate to normal levels and basing therapeutic strategies on these data.
128
BIBLIOGRAPHY
Agarwal, M. L., Taylor, W. R., Chernov, M. V., Chernova, O. B., and Stark, G. R. (1998). The p53 network. J Biol Chem 273, 1-4.
Almasan, A., Yin, Y., Kelly, R. E., Lee, E. Y., Bradley, A., Li, W., Bertino, J. R., and Wahl, G. M. (1995). Deficiency of retinoblastoma protein leads to inappropriate S-phase entry, activation of E2F-responsive genes, and apoptosis. Proc Natl Acad Sci U S A 92, 5436-5440.
Amundson, S. A., Bittner, M., Chen, Y., Trent, J., Meltzer, P., and Fornace, A. J., Jr. (1999). Fluorescent cDNA microarray hybridization reveals complexity and heterogeneity of cellular genotoxic stress responses. Oncogene 18, 3666-3672.
Amundson, S. A., Bittner, M., Meltzer, P., Trent, J., and Fornace, A. J., Jr. (2001). Induction of gene expression as a monitor of exposure to ionizing radiation. Radiat Res 156, 657-661.
Andreassen, P. R., Lohez, O. D., and Margolis, R. L. (2003). G2 and spingle assembly checkpoint adaptation, and tetraploidy arrest: implications for intrinsin and chemically induced genomic instability. Mut Res 532, 245-253.
Araki, K., Nakajima, Y., Eto, K., and Ikeda, M. A. (2003). Distinct recruitment of E2F family members to specific E2F-binding sites mediates activation and repression of the E2F1 promoter. Oncogene 22, 7632-7641.
Attwooll, C., Denchi, E. L., and Helin, K. (2004). The E2F family: specific functions and overlapping interests. Embo J 23, 4709-4716.
Black, E. P., Hallstrom, T., Dressman, H. K., West, M., and Nevins, J. R. (2005). Distinctions in the specificity of E2F function revealed by gene expression signatures. Proc Natl Acad Sci U S A 102, 15948-15953.
Blais, A., and Dynlacht, B. D. (2004). Hitting their targets: an emerging picture of E2F and cell cycle control. Curr Opin Genet Dev 14, 527-532.
129 Blake, M. C., and Azizkhan, J. C. (1989). Transcription factor E2F is required for efficient expression of the hamster dihydrofolate reductase gene in vitro and in vivo. Mol Cell Biol 9, 4994-5002.
Budde, A., Schneiderhan-Marra, N., Petersen, G., and Brune, B. (2005). Retinoblastoma susceptibility gene product pRB activates hypoxia-inducible factor-1 (HIF-1). Oncogene 24, 1802-1808.
Caelles, C., Helmberg, A., and Karin, M. (1994). p53-dependent apoptosis in the absence of transcriptional activation of p53-target genes. Nature 370, 220-223.
Cam, H., and Dynlacht, B. D. (2003). Emerging roles for E2F: Beyond the G1/S transition and DNA replication. Cancer Cell 3, 311-316.
Chan, T. A., Hwang, P. M., Hermeking, H., Kinzler, K. W., and Vogelstein, B. (2000). Cooperative effects of genes controlling the G(2)/M checkpoint. Genes Dev 14, 1584- 1588.
Chang, B. D., Broude, E. V., Fang, J., Kalinichenko, T. V., Abdryashitov, R., Poole, J. C., and Roninson, I. B. (2000). p21Waf1/Cip1/Sdi1-induced growth arrest is associated with depletion of mitosis-control proteins and leads to abnormal mitosis and endoreduplication in recovering cells. Oncogene 19, 2165-2170.
Chellappan, S. P., Hiebert, S., Mudryj, M., Horowitz, J. M., and Nevins, J. R. (1991). The E2F transcription factor is a cellular target for the RB protein. Cell 65, 1053-1061.
Chen, Q., Crosby, M., and Almasan, A. (2003). Redox Regulation of Apoptosis Before and After Cytochrome C Release. Korean J Biol Sci 7, 1-9.
Chen, Q., Gong, B., and Almasan, A. (2000). Distinct stages of cytochrome c release from mitochondria: evidence for a feedback amplification loop linking caspase activation to mitochondrial dysfunction in genotoxic stress induced apoptosis. Cell Death Differ 7, 227-233.
Christensen, J., Cloos, P., Toftegaard, U., Klinkenberg, D., Bracken, A. P., Trinh, E., Heeran, M., Di Stefano, L., and Helin, K. (2005). Characterization of E2F8, a novel E2F- like cell-cycle regulated repressor of E2F-activated transcription. Nucleic Acids Res 33, 5458-5470.
130 Crosby, M. E., and Almasan, A. (2004). Opposing roles of E2Fs in cell proliferation and death. Cancer Biol Ther 3, 1208-1211.
Crosby, M. E., Oancea, M., and Almasan, A. (2004). p53 binding to target sites is dynamically regulated before and after ionizing radiation-mediated DNA damage. J Environ Pathol Toxicol Oncol 23, 67-79.
DeGregori, J. (2002). The genetics of the E2F family of transcription factors: shared functions and unique roles. Biochim Biophys Acta 1602, 131-150.
Dimova, D. K., and Dyson, N. J. (2005). The E2F transcriptional network: old acquaintances with new faces. Oncogene 24, 2810-2826.
DuPree, E. L., Mazumder, S., and Almasan, A. (2004). Genotoxic stress induces expression of E2F4, leading to its association with p130 in prostate carcinoma cells. Cancer Res 64, 4390-4393.
Dyson, N. (1998). The regulation of E2F by pRB-family proteins. Genes Dev 12, 2245- 2262.
Evans, T., Rosenthal, E. T., Youngblom, J., Distel, D., and Hunt, T. (1983). Cyclin: a protein specified by maternal mRNA in sea urchin eggs that is destroyed at each cleavage division. Cell 33, 389-396.
Farkas, T., Hansen, K., Holm, K., Lukas, J., and Bartek, J. (2002). Distinct phosphorylation events regulate p130- and p107-mediated repression of E2F-4. J Biol Chem 277, 26741-26752.
Ferreira, R., Naguibneva, I., Mathieu, M., Ait-Si-Ali, S., Robin, P., Pritchard, L. L., and Harel-Bellan, A. (2001). Cell cycle-dependent recruitment of HDAC-1 correlates with deacetylation of histone H4 on an Rb-E2F target promoter. EMBO Rep 2, 794-799.
Gao, G., Bracken, A. P., Burkard, K., Pasini, D., Classon, M., Attwooll, C., Sagara, M., Imai, T., Helin, K., and Zhao, J. (2003). NPAT expression is regulated by E2F and is essential for cell cycle regulation. Mol Cell Biol 23, 2821-2833.
Gaubatz, S., Lees, J. A., Lindeman, G. J., and Livingston, D. M. (2001). E2F4 is exported from the nucleus in a CRM1-dependent manner. Mol Cell Biol 21, 1384-1392.
131 Gaubatz, S., Lindeman, G. J., Ishida, S., Jakoi, L., Nevins, J. R., Livingston, D. M., and Rempel, R. E. (2000). E2F4 and E2F5 play an essential role in pocket protein-mediated G1 control. Mol Cell 6, 729-735.
Giangrande, P. H., Zhu, W., Schlisio, S., Sun, X., Mori, S., Gaubatz, S., and Nevins, J. R. (2004). A role for E2F6 in distinguishing G1/S- and G2/M-specific transcription. Genes Dev 18, 2941-2951.
Girard, F., Strausfeld, U., Fernandez, A., and Lamb, N. J. (1991). Cyclin A is required for the onset of DNA replication in mammalian fibroblasts. Cell 67, 1169-1179.
Glotzer, M. (1995). Cell cycle. The only way out of mitosis. Curr Biol 5, 970-972.
Gong, B., and Almasan, A. (1999). Differential upregulation of p53-responsive genes by genotoxic stress in hematopoietic cells containing wild-type and mutant p53. Gene Expr 8, 197-206.
Gong, B., Chen, Q., and Almasan, A. (1998). Ionizing radiation stimulates mitochondrial gene expression and activity. Radiat Res 150, 505-512.
Gorgoulis, V. G., Zacharatos, P., Mariatos, G., Kotsinas, A., Bouda, M., Kletsas, D., Asimacopoulos, P. J., Agnantis, N., Kittas, C., and Papavassiliou, A. G. (2002). Transcription factor E2F-1 acts as a growth-promoting factor and is associated with adverse prognosis in non-small cell lung carcinomas. J Pathol 198, 142-156.
Hall, E. J. (2000). Radiobiology for the Radiobiologist, 5th edn (Philadelphia, Lippincott Williams & Wilkins).
Haupt, Y., Robles, A. I., Prives, C., and Rotter, V. (2002). Deconstruction of p53 functions and regulation. Oncogene 21, 8223-8231.
Helin, K. (1998). Regulation of cell proliferation by the E2F transcription factors. Curr Opin Genet Dev 8, 28-35.
Henglein, B., Chenivesse, X., Wang, J., Eick, D., and Brechot, C. (1994). Structure and cell cycle-regulated transcription of the human cyclin A gene. Proc Natl Acad Sci U S A 91, 5490-5494.
132 Herrera, R. E., Sah, V. P., Williams, B. O., Makela, T. P., Weinberg, R. A., and Jacks, T. (1996). Altered cell cycle kinetics, gene expression, and G1 restriction point regulation in Rb-deficient fibroblasts. Mol Cell Biol 16, 2402-2407.
Hershko, T., and Ginsberg, D. (2004). Up-regulation of Bcl-2 homology 3 (BH3)-only proteins by E2F1 mediates apoptosis. J Biol Chem 279, 8627-8634.
Hoh, J., Jin, S., Parrado, T., Edington, J., Levine, A. J., and Ott, J. (2002). The p53MH algorithm and its application in detecting p53-responsive genes. Proc Natl Acad Sci U S A 99, 8467-8472.
Hsieh, J. K., Yap, D., O'Connor, D. J., Fogal, V., Fallis, L., Chan, F., Zhong, S., and Lu, X. (2002). Novel function of the cyclin A binding site of E2F in regulating p53-induced apoptosis in response to DNA damage. Mol Cell Biol 22, 78-93.
Humbert, P. O., Rogers, C., Ganiatsas, S., Landsberg, R. L., Trimarchi, J. M., Dandapani, S., Brugnara, C., Erdman, S., Schrenzel, M., Bronson, R. T., and Lees, J. A. (2000). E2F4 is essential for normal erythrocyte maturation and neonatal viability. Mol Cell 6, 281- 291.
Iliakis, G., Wang, Y., Guan, J., and Wang, H. (2003). DNA damage checkpoint control in cells exposed to ionizing radiation. Oncogene 22, 5834-5847.
Innocente, S. A., Abrahamson, J. L., Cogswell, J. P., and Lee, J. M. (1999). p53 regulates a G2 checkpoint through cyclin B1. Proc Natl Acad Sci U S A 96, 2147-2152.
Ishida, S., Huang, E., Zuzan, H., Spang, R., Leone, G., West, M., and Nevins, J. R. (2001). Role for E2F in control of both DNA replication and mitotic functions as revealed from DNA microarray analysis. Mol Cell Biol 21, 4684-4699.
Jackson, M. W., Agarwal, M. K., Yang, J., Bruss, P., Uchiumi, T., Agarwal, M. L., Stark, G. R., and Taylor, W. R. (2005). p130/p107/p105Rb-dependent transcriptional repression during DNA-damage-induced cell-cycle exit at G2. J Cell Sci 118, 1821-1832.
Johnson, D. G., Schwarz, J. K., Cress, W. D., and Nevins, J. R. (1993). Expression of transcription factor E2F1 induces quiescent cells to enter S phase. Nature 365, 349-352.
Kaelin, W. G., Jr. (2003). E2F1 as a target: promoter-driven suicide and small molecule modulators. Cancer Biol Ther 2, S48-54.
133 Kaeser, M. D., and Iggo, R. D. (2002). Chromatin immunoprecipitation analysis fails to support the latency model for regulation of p53 DNA binding activity in vivo. Proc Natl Acad Sci U S A 99, 95-100.
Kannan, K., Amariglio, N., Rechavi, G., Jakob-Hirsch, J., Kela, I., Kaminski, N., Getz, G., Domany, E., and Givol, D. (2001). DNA microarrays identification of primary and secondary target genes regulated by p53. Oncogene 20, 2225-2234.
Kasahara, M., Takahashi, Y., Nagata, T., Asai, S., Eguchi, T., Ishii, Y., Fujii, M., and Ishikawa, K. (2000). Thymidylate synthase expression correlates closely with E2F1 expression in colon cancer. Clin Cancer Res 6, 2707-2711.
Kastan, M. B., and Bartek, J. (2004). Cell-cycle checkpoints and cancer. Nature 432, 316- 323.
Kherrouche, Z., Begue, A., Stehelin, D., and Monte, D. (2001). Molecular cloning and characterization of the mouse E2F6 gene. Biochem Biophys Res Commun 288, 22-33.
Koff, A., Cross, F., Fisher, A., Schumacher, J., Leguellec, K., Philippe, M., and Roberts, J. M. (1991). Human cyclin E, a new cyclin that interacts with two members of the CDC2 gene family. Cell 66, 1217-1228.
Komatsu, N., Takeuchi, S., Ikezoe, T., Tasaka, T., Hatta, Y., Machida, H., Williamson, I. K., Bartram, C. R., Koeffler, H. P., and Taguchi, H. (2000). Mutations of the E2F4 gene in hematological malignancies having microsatellite instability. Blood 95, 1509-1510.
Koniaras, K., Cuddihy, A. R., Christopoulos, H., Hogg, A., and O'Connell, M. J. (2001). Inhibition of Chk1-dependent G2 DNA damage checkpoint radiosensitizes p53 mutant human cells. Oncogene 20, 7453-7463.
Kowalik, T. F., DeGregori, J., Leone, G., Jakoi, L., and Nevins, J. R. (1998). E2F1- specific induction of apoptosis and p53 accumulation, which is blocked by Mdm2. Cell Growth Differ 9, 113-118.
Kowalik, T. F., DeGregori, J., Schwarz, J. K., and Nevins, J. R. (1995). E2F1 overexpression in quiescent fibroblasts leads to induction of cellular DNA synthesis and apoptosis. J Virol 69, 2491-2500.
134 Lakin, N. D., and Jackson, S. P. (1999). Regulation of p53 in response to DNA damage. Oncogene 18, 7644-7655.
Landsberg, R. L., Sero, J. E., Danielian, P. S., Yuan, T. L., Lee, E. Y., and Lees, J. A. (2003). The role of E2F4 in adipogenesis is independent of its cell cycle regulatory activity. Proc Natl Acad Sci U S A 100, 2456-2461.
Lang, S. E., McMahon, S. B., Cole, M. D., and Hearing, P. (2001). E2F transcriptional activation requires TRRAP and GCN5 cofactors. J Biol Chem 276, 32627-32634.
Lee, E. Y., Cam, H., Ziebold, U., Rayman, J. B., Lees, J. A., and Dynlacht, B. D. (2002). E2F4 loss suppresses tumorigenesis in Rb mutant mice. Cancer Cell 2, 463-472.
Lew, D. J., Dulic, V., and Reed, S. I. (1991). Isolation of three novel human cyclins by rescue of G1 cyclin (Cln) function in yeast. Cell 66, 1197-1206.
Lindeman, G. J., Gaubatz, S., Livingston, D. M., and Ginsberg, D. (1997). The subcellular localization of E2F-4 is cell-cycle dependent. Proc Natl Acad Sci U S A 94, 5095-5100.
Linke, S. P., Clarkin, K. C., Di Leonardo, A., Tsou, A., and Wahl, G. M. (1996). A reversible, p53-dependent G0/G1 cell cycle arrest induced by ribonucleotide depletion in the absence of detectable DNA damage. Genes Dev 10, 934-947.
Liu, D. X., Nath, N., Chellappan, S. P., and Greene, L. A. (2005). Regulation of neuron survival and death by p130 and associated chromatin modifiers. Genes Dev 19, 719-732.
Luo, R. X., Postigo, A. A., and Dean, D. C. (1998). Rb interacts with histone deacetylase to repress transcription. Cell 92, 463-473.
Ma, Y., Freeman, S. N., and Cress, W. D. (2004). E2F4 deficiency promotes drug- induced apoptosis. Cancer Bio & Ther 3, 1262-1269.
MacLachlan, T. K., and El-Deiry, W. S. (2002). Apoptotic threshold is lowered by p53 transactivation of caspase-6. Proc Natl Acad Sci U S A 99, 9492-9497.
Masui, Y., and Markert, C. L. (1971). Cytoplasmic control of nuclear behavior during meiotic maturation of frog oocytes. J Exp Zool 177, 129-145.
135 Matsushime, H., Roussel, M. F., Ashmun, R. A., and Sherr, C. J. (1991). Colony- stimulating factor 1 regulates novel cyclins during the G1 phase of the cell cycle. Cell 65, 701-713.
Mazumder, S., Gong, B., and Almasan, A. (2000). Cyclin E induction by genotoxic stress leads to apoptosis of hematopoietic cells. Oncogene 19, 2828-2835.
McKinney, K., and Prives, C. (2002). Efficient specific DNA binding by p53 requires both its central and C-terminal domains as revealed by studies with high-mobility group 1 protein. Mol Cell Biol 22, 6797-6808.
Minshull, J., Golsteyn, R., Hill, C. S., and Hunt, T. (1990). The A- and B-type cyclin associated cdc2 kinases in Xenopus turn on and off at different times in the cell cycle. Embo J 9, 2865-2875.
Miyajima, M., Nornes, H. O., Sato, K., and Neuman, T. (1996). Overexpression of E2F1 in astrocytes leads to neoplastic transformation and changes in expression of retinoblastoma family members. J Neurosci Res 46, 108-113.
Miyashita, T., and Reed, J. C. (1995). Tumor suppressor p53 is a direct transcriptional activator of the human bax gene. Cell 80, 293-299.
Moberg, K., Starz, M. A., and Lees, J. A. (1996). E2F-4 switches from p130 to p107 and pRB in response to cell cycle reentry. Mol Cell Biol 16, 1436-1449.
Mudryj, M., Devoto, S. H., Hiebert, S. W., Hunter, T., Pines, J., and Nevins, J. R. (1991). Cell cycle regulation of the E2F transcription factor involves an interaction with cyclin A. Cell 65, 1243-1253.
Muller, H., Bracken, A. P., Vernell, R., Moroni, M. C., Christians, F., Grassilli, E., Prosperini, E., Vigo, E., Oliner, J. D., and Helin, K. (2001). E2Fs regulate the expression of genes involved in differentiation, development, proliferation, and apoptosis. Genes Dev 15, 267-285.
Nahle, Z., Polakoff, J., Davuluri, R. V., McCurrach, M. E., Jacobson, M. D., Narita, M., Zhang, M. Q., Lazebnik, Y., Bar-Sagi, D., and Lowe, S. W. (2002). Direct coupling of the cell cycle and cell death machinery by E2F. Nat Cell Biol 4, 859-864.
136 Narita, M., Nunez, S., Heard, E., Lin, A. W., Hearn, S. A., Spector, D. L., Hannon, G. J., and Lowe, S. W. (2003). Rb-mediated heterochromatin formation and silencing of E2F target genes during cellular senescence. Cell 113, 703-716.
Neufeld, T. P., de la Cruz, A. F., Johnston, L. A., and Edgar, B. A. (1998). Coordination of growth and cell division in the drosophila wing. Cell 93, 1183-1193.
Nevins, J. R. (1992). E2F: a link between the Rb tumor suppressor protein and viral oncoproteins. Science 258, 424-429.
Nevins, J. R. (2001). The Rb/E2F pathway and cancer. Hum Mol Genet 10, 699-703.
Niculescu, A. B., 3rd, Chen, X., Smeets, M., Hengst, L., Prives, C., and Reed, S. I. (1998). Effects of p21(Cip1/Waf1) at both the G1/S and the G2/M cell cycle transitions: pRb is a critical determinant in blocking DNA replication and in preventing endoreduplication. Mol Cell Biol 18, 629-643.
Ogawa, H., Ishiguro, K., Gaubatz, S., Livingston, D. M., and Nakatani, Y. (2002). A complex with chromatin modifiers that occupies E2F- and Myc-responsive genes in G0 cells. Science 296, 1132-1136.
Ossovskaya, V. S., Mazo, I. A., Chernov, M. V., Chernova, O. B., Strezoska, Z., Kondratov, R., Stark, G. R., Chumakov, P. M., and Gudkov, A. V. (1996). Use of genetic suppressor elements to dissect distinct biological effects of separate p53 domains. Proc Natl Acad Sci U S A 93, 10309-10314.
Pagano, M., Pepperkok, R., Verde, F., Ansorge, W., and Draetta, G. (1992). Cyclin A is required at two points in the human cell cycle. Embo J 11, 961-971.
Park, W. Y., Hwang, C. I., Im, C. N., Kang, M. J., Woo, J. H., Kim, J. H., Kim, Y. S., Kim, H., Kim, K. A., Yu, H. J., et al. (2002). Identification of radiation-specific responses from gene expression profile. Oncogene 21, 8521-8528.
Pediconi, N., Ianari, A., Costanzo, A., Belloni, L., Gallo, R., Cimino, L., Porcellini, A., Screpanti, I., Balsano, C., Alesse, E., et al. (2003). Differential regulation of E2F1 apoptotic target genes in response to DNA damage. Nat Cell Biol 5, 552-558.
137 Perou, C. M., Sorlie, T., Eisen, M. B., van de Rijn, M., Jeffrey, S. S., Rees, C. A., Pollack, J. R., Ross, D. T., Johnsen, H., Akslen, L. A., et al. (2000). Molecular portraits of human breast tumours. Nature 406, 747-752.
Persengiev, S. P., Kondova, II, and Kilpatrick, D. L. (1999). E2F4 actively promotes the initiation and maintenance of nerve growth factor-induced cell differentiation. Mol Cell Biol 19, 6048-6056.
Phillips, A. C., Ernst, M. K., Bates, S., Rice, N. R., and Vousden, K. H. (1999). E2F-1 potentiates cell death by blocking antiapoptotic signaling pathways. Mol Cell 4, 771-781.
Polager, S., and Ginsberg, D. (2003). E2F mediates sustained G2 arrest and down- regulation of Stathmin and AIM-1 expression in response to genotoxic stress. J Biol Chem 278, 1443-1449.
Polager, S., Kalma, Y., Berkovich, E., and Ginsberg, D. (2002). E2Fs up-regulate expression of genes involved in DNA replication, DNA repair and mitosis. Oncogene 21, 437-446.
Prives, C., and Hall, P. A. (1999). The p53 pathway. J Pathol 187, 112-126.
Ray, S., and Almasan, A. (2003). Apoptosis induction in prostate cancer cells and xenografts by combined treatment with Apo2 ligand/tumor necrosis factor-related apoptosis-inducing ligand and CPT-11. Cancer Res 63, 4713-4723.
Rayman, J. B., Takahashi, Y., Indjeian, V. B., Dannenberg, J. H., Catchpole, S., Watson, R. J., te Riele, H., and Dynlacht, B. D. (2002). E2F mediates cell cycle-dependent transcriptional repression in vivo by recruitment of an HDAC1/mSin3B corepressor complex. Genes Dev 16, 933-947.
Redon, C., Pilch, D., Rogakou, E., Sedelnikova, O., Newrock, K., and Bonner, W. (2002). Histone H2A variants H2AX and H2AZ. Curr Opin Genet Dev 12, 162-169.
Ren, B., Cam, H., Takahashi, Y., Volkert, T., Terragni, J., Young, R. A., and Dynlacht, B. D. (2002). E2F integrates cell cycle progression with DNA repair, replication, and G(2)/M checkpoints. Genes Dev 16, 245-256.
138 Ren, B., and Dynlacht, B. D. (2004). Use of chromatin immunoprecipitation assays in genome-wide location analysis of mammalian transcription factors. Methods Enzymol 376, 304-315.
Rogoff, H. A., and Kowalik, T. F. (2004). Life, Death and E2F: Linking Proliferation Control and DNA Damage Signaling via E2F1. Cell Cycle 3, 845-846.
Rogoff, H. A., Pickering, M. T., Debatis, M. E., Jones, S., and Kowalik, T. F. (2002). E2F1 induces phosphorylation of p53 that is coincident with p53 accumulation and apoptosis. Mol Cell Biol 22, 5308-5318.
Rogoff, H. A., Pickering, M. T., Frame, F. M., Debatis, M. E., Sanchez, Y., Jones, S., and Kowalik, T. F. (2004). Apoptosis associated with deregulated E2F activity is dependent on E2F1 and Atm/Nbs1/Chk2. Mol Cell Biol 24, 2968-2977.
Rothkamm, K., Krüger, I., Thompson, L. H., and Löbrich, M. (2003). Pathways of DNA double-strand break repair during the mammalian cell cycle. Mol Cell Biol 23, 5706- 5715.
Saito, M., Helin, K., Valentine, M. B., Griffith, B. B., Willman, C. L., Harlow, E., and Look, A. T. (1995). Amplification of the E2F1 transcription factor gene in the HEL erythroleukemia cell line. Genomics 25, 130-138.
Sala, A., Nicolaides, N. C., Engelhard, A., Bellon, T., Lawe, D. C., Arnold, A., Grana, X., Giordano, A., and Calabretta, B. (1994). Correlation between E2F-1 requirement in the S phase and E2F-1 transactivation of cell cycle-related genes in human cells. Cancer Res 54, 1402-1406.
Sardet, C., Vidal, M., Cobrinik, D., Geng, Y., Onufryk, C., Chen, A., and Weinberg, R. A. (1995). E2F-4 and E2F-5, two members of the E2F family, are expressed in the early phases of the cell cycle. Proc Natl Acad Sci U S A 92, 2403-2407.
Sherr, C. J. (1996). Cancer cell cycles. Science 274, 1672-1677.
Smits, V. A., and Medema, R. H. (2001). Checking out the G(2)/M transition. Biochim Biophys Acta 1519, 1-12.
139 Souza, R. F., Yin, J., Smolinski, K. N., Zou, T. T., Wang, S., Shi, Y. Q., Rhyu, M. G., Cottrell, J., Abraham, J. M., Biden, K., et al. (1997). Frequent mutation of the E2F-4 cell cycle gene in primary human gastrointestinal tumors. Cancer Res 57, 2350-2353.
Stanelle, J., Stiewe, T., Theseling, C. C., Peter, M., and Putzer, B. M. (2002). Gene expression changes in response to E2F1 activation. Nucleic Acids Res 30, 1859-1867.
Stevaux, O., and Dyson, N. J. (2002). A revised picture of the E2F transcriptional network and RB function. Curr Opin Cell Biol 14, 684-691.
Stevens, C., Smith, L., and La Thangue, N. B. (2003). Chk2 activates E2F-1 in response to DNA damage. Nat Cell Biol 5, 401-409.
Szak, S. T., Mays, D., and Pietenpol, J. A. (2001). Kinetics of p53 binding to promoter sites in vivo. Mol Cell Biol 21, 3375-3386.
Takahashi, Y., Rayman, J. B., and Dynlacht, B. D. (2000). Analysis of promoter binding by the E2F and pRB families in vivo: distinct E2F proteins mediate activation and repression. Genes Dev 14, 804-816.
Taylor, W. R., Schonthal, A. H., Galante, J., and Stark, G. R. (2001). p130/E2F4 binds to and represses the cdc2 promoter in response to p53. J Biol Chem 276, 1998-2006.
Taylor, W. R., and Stark, G. R. (2001). Regulation of the G2/M transition by p53. Oncogene 20, 1803-1815.
Thornborrow, E. C., Patel, S., Mastropietro, A. E., Schwartzfarb, E. M., and Manfredi, J. J. (2002). A conserved intronic response element mediates direct p53-dependent transcriptional activation of both the human and murine bax genes. Oncogene 21, 990- 999.
Trimarchi, J. M., Fairchild, B., Wen, J., and Lees, J. A. (2001). The E2F6 transcription factor is a component of the mammalian Bmi1-containing polycomb complex. Proc Natl Acad Sci U S A 98, 1519-1524.
Trimarchi, J. M., and Lees, J. A. (2002). Sibling rivalry in the E2F family. Nat Rev Mol Cell Biol 3, 11-20.
140 Trouche, D., Le Chalony, C., Muchardt, C., Yaniv, M., and Kouzarides, T. (1997). RB and hbrm cooperate to repress the activation functions of E2F1. Proc Natl Acad Sci U S A 94, 11268-11273.
Tsantoulis, P. K., and Gorgoulis, V. G. (2005). Involvement of E2F transcription factor family in cancer. Eur J Cancer 41, 2403-2414.
Waghray, A., Schober, M., Feroze, F., Yao, F., Virgin, J., and Chen, Y. Q. (2001). Identification of differentially expressed genes by serial analysis of gene expression in human prostate cancer. Cancer Res 61, 4283-4286.
Wahl, G. M., and Carr, A. M. (2001). The evolution of diverse biological responses to DNA damage: insights from yeast and p53. Nat Cell Biol 3, E277-286.
Wang, H., Wang, M., Bocker, W., and Iliakis, G. (2005). Complex H2AX phosphorylation patterns by multiple kinases including ATM and DNA-PK in human cells exposed to ionizing radiation and treated with kinase inhibitors. J Cell Physiol 202, 492-502.
Wang, L., Wu, Q., Qiu, P., Mirza, A., McGuirk, M., Kirschmeier, P., Greene, J. R., Wang, Y., Pickett, C. B., and Liu, S. (2001). Analyses of p53 target genes in the human genome by bioinformatic and microarray approaches. J Biol Chem 276, 43604-43610.
Weinmann, A. S., Bartley, S. M., Zhang, T., Zhang, M. Q., and Farnham, P. J. (2001). Use of chromatin immunoprecipitation to clone novel E2F target promoters. Mol Cell Biol 21, 6820-6832.
Wells, J., Boyd, K. E., Fry, C. J., Bartley, S. M., and Farnham, P. J. (2000). Target gene specificity of E2F and pocket protein family members in living cells. Mol Cell Biol 20, 5797-5807.
Wu, L., Timmers, C., Maiti, B., Saavedra, H. I., Sang, L., Chong, G. T., Nuckolls, F., Giangrande, P., Wright, F. A., Field, S. J., et al. (2001). The E2F1-3 transcription factors are essential for cellular proliferation. Nature 414, 457-462.
Wu, X., and Levine, A. J. (1994). p53 and E2F-1 cooperate to mediate apoptosis. Proc Natl Acad Sci U S A 91, 3602-3606.
141 Xiong, Y., Connolly, T., Futcher, B., and Beach, D. (1991). Human D-type cyclin. Cell 65, 691-699.
Xu, B., Kim, S., and Kastan, M. B. (2001). Involvement of Brca1 in S-phase and G(2)- phase checkpoints after ionizing irradiation. Mol Cell Biol 21, 3445-3450.
Xu, B., Kim, S. T., Lim, D. S., and Kastan, M. B. (2002). Two molecularly distinct G(2)/M checkpoints are induced by ionizing irradiation. Mol Cell Biol 22, 1049-1059.
Xu, G., Livingston, D. M., and Krek, W. (1995). Multiple members of the E2F transcription factor family are the products of oncogenes. Proc Natl Acad Sci U S A 92, 1357-1361.
Yamasaki, L., Jacks, T., Bronson, R., Goillot, E., Harlow, E., and Dyson, N. J. (1996). Tumor induction and tissue atrophy in mice lacking E2F-1. Cell 85, 537-548.
Yan, T., Desai, A. B., Jacobberger, J. W., Sramkoski, R. M., Loh, T., and Kinsella, T. J. (2004). CHK1 and CHK2 are differentially involved in mismatch repair-mediated 6- thioguanine-induced cell cycle checkpoint responses. Mol Cancer Ther 3, 1147-1157.
Yoshitaka, T., Matsubara, N., Ikeda, M., Tanino, M., Hanafusa, H., Tanaka, N., and Shimizu, K. (1996). Mutations of E2F-4 trinucleotide repeats in colorectal cancer with microsatellite instability. Biochem Biophys Res Commun 227, 553-557.
Yu, J., Zhang, L., Hwang, P. M., Kinzler, K. W., and Vogelstein, B. (2001). PUMA induces the rapid apoptosis of colorectal cancer cells. Mol Cell 7, 673-682.
Yu, J., Zhang, L., Hwang, P. M., Rago, C., Kinzler, K. W., and Vogelstein, B. (1999). Identification and classification of p53-regulated genes. Proc Natl Acad Sci U S A 96, 14517-14522.
Zachos, G., Rainey, M. D., and Gillespie, D. A. (2003). Chk1-deficient tumour cells are viable but exhibit multiple checkpoint and survival defects. Embo J 22, 713-723.
Zhao, Y. L., Piao, C. Q., Wu, L. J., Suzuki, M., and Hei, T. K. (2000). Differentially expressed genes in asbestos-induced tumorigenic human bronchial epithelial cells: implication for mechanism. Carcinogenesis 21, 2005-2010.
142 Zhou, B. S., and Bartek, J. (2004). Targeting the checkpoint kinases: chemosensitization versus chemoprotection. Nature Reviews Cancer 4, 216-225.
Zhu, W., Giangrande, P. H., and Nevins, J. R. (2004). E2Fs link the control of G1/S and G2/M transcription. Embo J 23, 4615-4626.
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