CYCLIN E PROVIDES A LINK BETWEEN CELL CYCLE,
DNA REPAIR AND APOPTOSIS
A dissertation submitted
to Kent State University in collaboration with the
Lerner Research Institute, Cleveland Clinic
in partial fulfillment of the requirements for the
degree of Doctor of Philosophy
by
Dragos Costin Plesca
May, 2008
Dissertation written by
Dragos Costin Plesca
Pharm.D., University of Medicine and Pharmacy “Carol Davila”, Romania, 2002
Ph.D., Kent State University, 2008
Approved by
______, Chair, Doctoral Dissertation Committee Alexandru Almasan, Ph.D
______, Member, Doctoral Dissertation Committee James Blank, Ph.D
______, Member, Doctoral Dissertation Committee Gail Fraizer, Ph.D
______, Member, Doctoral Dissertation Committee Olena Piontkivska, Ph.D
______, Graduate Faculty Representative Jennifer Marcinkiewicz, Ph.D
Accepted by
______, Director, School of Biomedical Sciences Robert V. Dorman, Ph.D
______, Dean, College of Arts and Sciences John R. D. Stalvey, Ph.D
ii
TABLE OF CONTENTS
List of Figures...... vi
List of Tables ...... ix
Acknowledgments ...... x
Chapter I. Introduction ...... 1
The Cell Cycle ...... 3
Cyclin E and cell cycle regulation ...... 8
Transcriptional regulation of Cyclin E ...... 14
Post-translational regulation of Cyclin E...... 20
Deregulated expression of Cyclin E in cancer...... 27
Therapeutic approaches involving cell cycle inhibitors...... 32
Apoptosis ...... 38
TRAIL, APO2L ...... 40
Adaptor proteins...... 41
Cytochrome c...... 42
Bcl-2 family members...... 42
Caspases...... 43
Other apoptosis-regulatory proteins...... 45
Apoptotic pathways ...... 46
iii DNA damage and repair mechanisms...... 51
Homologous recombination...... 55
Ku70/Ku80 heterodimer ...... 56
DNA-PKcs ...... 60
Artemis...... 62
XRCC4...... 62
Ligase IV...... 63
XLF...... 64
Non-Homologous End Joining ...... 66
Other DNA double-strand break repair pathways ...... 66
Generation of p18-Cyclin E and its role in modulating cellular
response to genotoxic stress...... 70
Chapter II. p18-Cyclin E regulates Non-Homologous End Joining by preventing
the recruitment of the XLF/XRCC4/Ligase IV heterocomplex to the
DNA repair complex ...... 73
Abstract...... 73
Introduction...... 74
Materials and Methods...... 78
Results...... 82
Discussion...... 95
iv Chapter III. Two new mechanisms for proteasome-mediated Cyclin E degradation
uncovered in hematopoietic tumor cells undergoing apoptosis ...... 102
Abstract...... 102
Introduction...... 103
Materials and Methods...... 106
Results...... 110
Discussion...... 137
Chapter IV. Summary and future directions ...... 145
Summary...... 145
Future directions ...... 148
References...... 157
Appendix. Abbreviations...... 193
v
LIST OF FIGURES
Chapter I
Fig 1.1 Cellular response to DNA damage ...... 2
Fig 1.2 Schematic representation of the cell cycle...... 4
Fig 1.3 Periodic oscillation in the expression levels of cyclins ...... 10
Fig 1.4 Schematic representation of the progression through the different cell cycle
phases...... 19
Fig 1.5 Schematic representation of the Cyclin E turnover mechanisms...... 23
Fig 1.6 Schematic representation of the cell cycle and the different therapeutic
approaches undertaken...... 37
Fig 1.7 Schematic representation of the extrinsic and intrinsic apoptotic pathways ...49
Fig 1.8 Three-dimensional structure of the Ku70/Ku80 heterodimer...... 59
Fig 1.9 Schematic representation of the XLF/XRCC4/Ligase IV heterocomplex...... 65
Fig 1.10 Schematic representation of the NHEJ pathway...... 69
Fig 1.11 Schematic representation of Cyclin E domains and cleavage sites ...... 71
vi Chapter II
Fig 2.1 Non-toxic levels of p18-Cyclin E sensitize cells to VP-16 and IR treatment..85
Fig 2.2 p18-Cyclin E inhibits in vitro DNA ligation and plasmid reactivation ...... 88
Fig 2.3 p18-Cyclin E prevents efficient repair of the DNA damage induced by IR and
CPT-11...... 91
Fig 2.4 p18-Cyclin E impairs the recruitment of XLF, XRCC4, and Ligase IV to the
DNA repair complex...... 94
Fig 2.5 Schematic representation of the NHEJ DNA repair pathway and the way in
which p18-Cyclin E affects the assembly of repair factors ...... 101
Chapter III
Fig 3.1 p18-Cyclin E is a short-lived protein that is stabilized by proteasome
inhibition...... 112
Fig 3.2 p18-Cyclin E is effectively ubiquitinated ...... 115
Fig 3.3 Fbw7 interacts with p18-Cyclin E independently of phosphorylation at the C-
terminal phosphodegron ...... 120
Fig 3.4 p18-Cyclin E is regulated by the SCFFbw7 complex and independently of Cdk2
binding ...... 125
Fig 3.5 Ku70 but not Cdk2 binding regulates p18-Cyclin E stability...... 130
Fig 3.6 p18-cyclin E sensitizes cells to Bortezomib treatment through enhanced
association with Ku70 and release of Bax from Ku70 ...... 135
Fig. 3.7 Chemical structure of the proteasome inhibitor, Bortezomib...... 141
vii Chapter IV
Fig 4.1 Dual regulation of apoptosis and DNA repair by p18-Cyclin E...... 147
Fig 4.2 Localization of p18-Cyclin E-GFP, Cyclin E-GFP, and Cyclin ER130A-GFP 151
Fig 4.3 Cyclin E sequence alignment across human, mouse, rat, rabbit, chicken, and
frog...... 153
viii
LIST OF TABLES
Table 1. Critical protein substrates of Caspase-3...... 45
ix
ACKNOWLEDGMENTS
None of the scientific results or achievements that I have had during my last five
years would have been possible without the sustained professional and moral support that
my adviser, Dr. Alex Almasan has given me. I thank him for his guidance, motivating
spirit and for nurturing my scientific creativity and independence. I feel very fortunate for
having studied in his lab as he provided me along the years with a unique Ph.D.
experience and great learning opportunities which will continue to serve me well in the future.
I would also like to thank the colleagues in my lab that throughout the years have supported and encouraged me both professionally and personally and who have shared with me the ups and downs of scientific research: Drs. Marcela Oancea, Meredith
Crosby, Erica DuPree, Damodar Gupta, Subrata Ray and John Hissong. I would like to thank in particular Dr. Suparna Mazumder for her never-ending patience, kindness and advice that she had for me at all times.
I would also like to extend my appreciation to my dissertation committee members Dr. Gail Fraizer, Dr. James Blank, Dr. Olena Piontkivska, and Dr. Bryan
Williams for their constructive comments and help at different stages during my studies.
Thank you also to Dr. Jennifer Marcinkiewicz for joining the committee as graduate
x faculty representative. Furthermore, I thank the School of Biomedical Sciences and the
Department of Cancer Biology for giving me the opportunity to pursue my Ph.D. degree.
Finally, I thank my mother and father, Gabriela and Gheorghe, and my dearest
Carolina whose unending love and emotional support have buffered the frustrations and gave me the strength to take critical decisions in the most difficult of times. They have been a constant source of energy and optimism and they have always had confidence in me whenever I seemed to lose mine. There is nothing that I have accomplished, or will achieve, that would be possible without them.
Time passes and when success or disappointment fades away, what matters the most is what we have influenced and changed with our presence and what we have been able to take with us in our hearts and minds to make us a better person. The pursuit of the
Ph.D. degree was a priceless and unique experience which has profoundly changed my perspectives and character.
xi
CHAPTER I
Introduction
The human DNA undergoes several thousand to a million damaging events each
day, generated by either external or internal metabolic processes. These damage events
can be caused by either gamma rays resulting from the decay of naturally occurring
radionuclides in the earth or by reactive oxygen species such as hydrogen peroxide or
superoxide anions (Chance et al., 1979). In response to DNA damage, a cell may arrest its progression through the cell cycle in order to repair the lesions (Kuerbitz et al., 1992).
Successful repair is essential for its survival and the cell resumes cycling. However, when the DNA damage overwhelms the DNA repair machinery’s capacity, an apoptotic signal is triggered leading to programmed cell death (Figure 1.1). This chapter describes the three cellular processes that are involved in the cellular response to DNA damage: cell cycle arrest, DNA repair, and apoptosis.
1 2
Figure 1.1 Cellular response to DNA damage. Upon infliction of DNA damage, the cell undergoes cell cycle arrest to allow the damage to be repaired. Depending on the severity of the lesion, the cell may be able to repair it, survive and resume cycling. On the other hand, when the damage is too extensive, the cell may choose to undergo apoptosis.
3
The Cell Cycle
The cell cycle is a fundamental cellular process characterized by a highly
conserved and ordered set of events, culminating in cell growth and division. It ensures
propagation of life from the unicellular to the multicellular organisms (Murray, 1993). In
order to generate a daughter cell, the parental cell has to undergo several changes among
which are, growth in mass, duplication of organelles, duplication and equal separation of
DNA in the two daughter cells (Mitchison, 1971). These processes take place in well
defined phases of the cell cycle. Generally the cell cycle can be broken down into
interphase and division or the mitotic (M) phase. During the interphase the cell goes
through elaborate preparation for its division including synthesis of necessary proteins
and DNA replication. The division consists of the breakdown of the nuclear envelope,
segregation of the chromosomes by the mitotic spindle and the actual separation of the
daughter cells or cytokinesis. Regulation of the cell cycle is critical for the normal
development of organisms, and loss of control could lead to cell death or onset of cancer
(Sherr, 1996).
The cell cycle can be divided into four stages: gap 1 (G1), synthesis (S), and gap 2
(G2), which form the interphase and the Mitosis (M) (Figure 1.2) (Baserga, 1985). The progression through these phases is governed by a precise series of events that allow the cell to grow and prepare for its division (Johnston, 1990).
4
Figure 1.2 Schematic representation of the cell cycle. During the G1 phase the cell grows is size preparing for the DNA synthesis that takes place in the S phase. After the sister chromatid has been synthesized, the cell passes through another gap phase, G2, ensuring that the environment is optimal for division. During the M phase, the chromatin condenses and the resulting chromosomes are evenly separated by the mitotic spindle into the two new daughter cells. The cells may choose to differentiate and thus stop cycling by entering the G0 phase.
5
During the G1 phase, the cell monitors its environment and size and faces the decision of either preparing for S phase or to differentiate and exit the cell cycle. Cells that leave the cell cycle in G1 in order to differentiate are found in G0. This quiescence may be permanent or temporary as they may re-enter the cell cycle at a later time. After the G1 phase, DNA replication occurs in S phase.
The G2 phase provides additional time for growth, with cells preparing for the M
phase. After the G2 phase, nuclear and cytoplasmic division occurs in the M phase. The
M phase is subdivided into prophase, metaphase, anaphase, and telophase. Each of these
cell cycle phases corresponds to specific events of the cell division. During prophase, the
chromatin is condensed and individual chromosomes become visible. In metaphase, sister
chromatids are paired and aligned at the center of the cell in what is called the metaphasic
plaque. In anaphase, the chromosomes are separated by being pulled to the opposite sides
of the cell by mitotic spindle fibers which are attached at their kinetochores. The mitotic
spindle disassembles and the chromosomes decondense during telophase. The physical
division of the cytoplasm containing the organelles takes place during cytokinesis
(Nasmyth, 2001).
During both G1 and G2 phases there are checkpoints whose role is to ensure the
DNA integrity (Andreassen et al., 2003). When challenged with DNA damage, a cell may decide to arrest its cycling to repair the lesion or to undergo other events that lead to apoptosis. Another role of the checkpoints is to ensure that the molecular events specific to a certain cell cycle phase have taken place before progressing to the next, downstream process (Hartwell and Weinert, 1989). Furthermore, they allow the system to integrate 6
and be controlled by signals from the environment (e.g. presence of mitogens). For example, the G1/S phase progression is controlled by the G1 checkpoint, also called the restriction point, whose role is to ensure that the cell is big enough and that the environment is favorable for DNA synthesis. Rapidly replicating human cells go through a round of division in approximately 24 hours: G1 lasting ~10 hours, S phase 10 hours,
G2 ~3.5 hours, and M ~ 30 minutes.
The tight control of the cell cycle is achieved through the temporal expression of cyclins which bind and activate their catalytic partners, the cyclin-dependent kinases
(CDK), and other accessory proteins important for their stability and activity, the cyclin- dependent kinase inhibitors (Murray and Kirschner, 1989; Murray and Kirschner, 1991).
The first molecular insights into understanding the molecular mechanisms governing the cell cycle emerged from studies of the M-phase Promoting Factor (MPF) or the Cyclin
B1/Cdk1 complex (Masui and Markert, 1971). Cyclins have originally been identified by studies of the cyclic synthesis and degradation of proteins in the eggs of the sea urchin
Lytechinus pictus (Evans et al., 1983).
The G1 cyclins, D and E were identified by homology screening, subtractive hybridization following stimulation with colony-stimulating factor-1 (Matsushime et al.,
1991) and yeast complementation assays (Koff et al., 1991; Lew et al., 1991; Xiong et al.,
1991). Cyclin D family of proteins consisting of Cyclin D1, D2, and D3 can interact with either Cdk4 or Cdk6. They regulate entry and progression through the cell cycle in response to mitogenic stimulation. Their main role is to phosphorylate and inactivate the cell-cycle inhibitory function of the retinoblastoma protein (pRB). The other G1 cyclin, 7
Cyclin E, interacts with Cdk2 and stimulates the G1/S phase transition. The activity of the G1 cyclins and CDKs may be blocked by the CDK inhibitors proteins (CIPs), p27Kip1, p57Kip2, p21CIP/Waf1, and the p16 family of proteins. Another regulatory protein, Cdc25A phosphatase, positively regulates activity of Cyclin A/Cdk2 and Cyclin E/Cdk2 complexes by removing the inhibitory phosphorylation on Cdk2 (Smits and Medema,
2001).
Cyclins A and B bind and regulate the kinase activity of Cdk1 (Minshull et al.,
1990) being the mammalian equivalent of MPF. Cyclin A expression starts at the beginning of the S phase and peaks during G2. As opposed to Cyclin B1, it also interacts with Cdk2 during the S phase. Cyclin B starts to be synthesized in late S phase, peaks during metaphase and decreases at the end of anaphase. The temporal destruction of
Cyclin B is performed by Anaphase-Promoting Complex (APC) ubiquitin ligase (Glotzer,
1995). The APC catalyzes the addition of polyubiquitin chains to Cyclin B, targeting it to proteasomal degradation.
The cell cycle progression is also regulated at the transcriptional level. Several genes important for cell division have been found to be controlled by the E2F family of transcription factors (Mudryj et al., 1991). For example, their transcriptional activity is important for the regulation of Cyclin E, dihydrofolate reductase (DHFR), Cdc6, and the minichromosome maintenance (MCM) proteins (Dyson, 1998). E2F’s cell cycle regulatory function resulted from studies in which E2F complexes were dissociated by
E1A (early region 1A product of adenovirus 5), which resulted in a loss of cellular proliferation control (Chellappan et al., 1991). Another cell cycle regulatory protein, the 8
retinoblastoma tumor suppressor protein (pRb), was found to interact with E2F
(Chellappan et al., 1991) and form a complex that acts as a growth suppressor and prevents progression through the cell cycle (Classon and Harlow, 2002). The pRb-
E2F/DP (dimerization protein) complex also attracts a histone deacetylase (HDAC) protein to the chromatin, further suppressing DNA synthesis. The E2F regulation in normal cells was first uncovered by studies using adenovirus E1A 12S product to dissociate E2F from pRb, leading to activation of E2F (Nevins, 1992).
Cyclin E and cell cycle regulation
The main families of regulatory proteins that play key roles in controlling the cell
cycle progression are the cyclins, the cyclin dependent kinases (CDKs) and, as their
critical downstream substrates, the tumor suppressor gene products of retinoblastoma
(pRb) and p53. The activity of CDKs oscillates, leading to the cyclical changes in the
activation of key proteins, which regulate the main cell cycle processes: DNA replication,
mitosis, and cytokinesis. G1/S-cyclins commit cells to DNA replication, which is driven
by the S-phase cyclins, whereas mitosis is governed by M-cyclins. Progression through
the early G1 phase is regulated by signal transduction cascades activated by polypeptide
growth factors and by extracellular matrix components.
The human G1 cyclins, the D- and E-type cyclins, were identified functionally by
screening of human cDNA libraries for sequences that could complement G1 cyclin
mutations in S. cerevisiae (Koff et al., 1991; Lew et al., 1991). This indicated that Cyclin
E plays an important role in the regulation of the cell cycle progression. In fission yeast 9
(Schizosaccharomyces pombe) cells have to reach a critical size before entering mitosis, which is achieved by associating Cdk1 activation with cell growth. Premature activation of the mitotic CDK (Cdk1) leads to mitosis entry at a reduced size. A similar phenomenon takes place in mammalian cells when cyclin E is overexpressed. The cells have a shorter G1 and enter mitosis at a reduced size (Ohtsubo and Roberts, 1993;
Resnitzky and Reed, 1995). Furthermore, Cyclin E may replace all functions of Cyclin
D1 indicating that it is one of its downstream targets as well as a key regulator of the S phase entry (Geng et al., 1999). Together with the importance of Cyclin E in G1/S transition, these data indicate that Cyclin E is playing an important role in critical limiting steps which are essential for S phase entry.
The human Cyclin E has several forms and splice variants, with the predominant species encoding a 395 amino-acid protein with a molecular weight of 50-55 kDa (Koff et al., 1991; Lew et al., 1991). The E-type cyclin family consists of two members, Cyclin
E1 (Cyclin E) and E2. Cyclin E2 also has a typical cyclin box motif and shares 47% similarity with Cyclin E1. The two proteins display a similar cycle-regulated expression pattern as well as biochemical properties. Cyclin E mRNA levels have a cyclic pattern of fluctuation, with synthesis starting during the G1 phase and reaching a maximum in late
G1 followed by a down-regulation in the S phase (Dulic et al., 1992; Koff et al., 1992;
Lew et al., 1991). The Cyclin E expression oscillates throughout the cell cycle reaching a peak at the G1/S transition and a minimum in the M phase (Dulic et al., 1992; Koff et al.,
1992) (Figure 1.3). 10
Figure1.3 Periodic oscillation in the expression levels of cyclins. Cyclin D partners
with Cdk4 and Cdk6 in early to mid-G1 phase to phosphorylate and inactivate the cell-
cycle inhibitory function of the retinoblastoma protein. Cyclin E levels peak at the G1/S
boundary where it exerts its roles in the G1/S phase transition. Cyclin A synthesis starts at the beginning of the S phase and reaches its peak at the beginning of the G2 phase while the mitotic Cyclin B levels attain a maximum during the M phase being required for the cell division.
11
Cyclin E is synthesized at the G1/S boundary under the influence of the E2F transcription factor family (Botz et al., 1996; Geng et al., 1996; Ohtani et al., 1995).
By interacting with and activating its catalytic partner Cdk2, Cyclin E is essential for regulating the G1/S transition (Ohtsubo et al., 1995; Resnitzky et al., 1994; Tsai et al.,
1993; van den Heuvel and Harlow, 1993) and the initiation of DNA replication (Jackson et al., 1995; Krude et al., 1997; Ohtsubo et al., 1995; Roberts and Sherr, 2003). Cyclin E2 was also found to interact with and activate Cdk2 forming a complex that can be inhibited by p27 (Gudas et al., 1999; Lauper et al., 1998; Zariwala et al., 1998). Both proteins were found to be redundant in development (Parisi et al., 2003). Cyclin E, unlike any other cyclin, can complement deficiencies of other cyclins in yeast (Koff et al., 1991;
Lew et al., 1991) and mouse (Geng et al., 1999) and it has unique and highly specialized functions. Cyclin E promotes events associated with genome replication (Reed, 1996;
Sauer and Lehner, 1995). These include regulation of key steps in the initiation of DNA replication through E2F regulation and thus of its critical S-phase target genes, CDC6 and loading of MCM and CDC45 to replication origins. Cyclin E/Cdk2 phosphorylates two of its own regulators: Cdc25A (Hoffmann et al., 1994) and p27Kip1 (Sheaff et al., 1997;
Vlach et al., 1997). Phosphorylation by Cyclin E/Cdk2 also regulates histone synthesis
(through p220/NPAT) (Ma et al., 2000; Zhao et al., 2000), centrosome duplication
(through nucleophosmin) (Okuda et al., 2000), chromatin modifications by p300/CBP
(Ait-Si-Ali et al., 1998) and SWI/SNF (Shanahan et al., 1999), pre-mRNA splicing
(through SAP155/SF3B1) (Seghezzi et al., 1998) and DNA replication (Ewen, 2000). 12
The other homologous protein, Cyclin E2, is believed to have similar functions
(Mazumder et al., 2004; Moroy and Geisen, 2004).
Cyclin/CDK complexes are negatively regulated by two families of CDK
inhibitors (Sherr and Roberts, 1995). The D-type cyclin/CDK complexes may be
inhibited by both Ink4 as well as Kip/Cip family of inhibitors, whereas the E-type cyclin/CDK complexes are inhibited exclusively by the latter one. p21Cip1 and p27Kip1 bind and inhibit Cyclin E/Cdk2 complexes by competing with their substrates but also by preventing the cyclin-activating kinase-mediated phosphorylation as well (Morgan, 1996;
Sherr and Roberts, 1995). Consequently, cells that lack these two proteins manifest a significant increase in the kinase activity of the Cyclin E/Cdk2 complex (Aleem et al.,
2005).
A number of biological functions have been attributed to Cyclin E, based mainly on its interaction with and phosphorylation of its substrates. Thus, Cyclin E/Cdk2 complexes have been shown to play an essential role in the initiation of DNA replication
(Jackson et al., 1995), in addition to the cell cycle transition (Ohtsubo et al., 1995;
Resnitzky et al., 1994). While Cyclin E complexes phosphorylate pRb, which is also phosphorylated by the D-type Cyclin-Cdk4/6 complexes (Dowdy et al., 1993) it is likely that other substrates exist during late G1. Unlike Cyclin D, Cyclin E remains essential in the absence of pRb, since: (i) its inducible expression in fibroblasts accelerates G1/S progression without affecting the kinetics of pRb phosphorylation (Resnitzky and Reed,
1995); (ii) unlike the D-type cyclins, Cyclin E is essential for cell-cycle progression in pRb-deficient cells (Ohtsubo et al., 1995); (iii) ectopic expression of cyclin E bypasses 13
pRb-mediated cell cycle arrest (Alevizopoulos et al., 1997; Lukas et al., 1997); and (iv)
Cyclin E is required for S phase entry in Drosophila (Duronio et al., 1996) and Xenopus
(Hua et al., 1997). These findings highlight a fundamental difference between the Cyclin
D and Cyclin E complexes, strongly suggesting that other key rate-limiting substrates
exist for Cyclin E/Cdk2 (Alevizopoulos et al., 1997; Lukas et al., 1997). Constitutive
activation of Cyclin E/Cdk2 results in uncoupling of the initiation of centrosome and
DNA duplication leading to unscheduled initiation of centrosome duplication prior to S-
phase entry (Mussman et al., 2000).
The Cyclin E/Cdk2 complex was thought to be a critical regulator of the somatic
cell cycle. Surprisingly, ablating these genes in the mouse had very limited consequences.
Embryonic fibroblasts deficient in Cdk2 had normal proliferation and become immortal
after continuous passage in culture. Cdk2-/- mice are viable and can survive for up to two years, suggesting that Cdk2 may not be necessary for proliferation and survival of most cell types. It was found, however, to be necessary for completion of prophase I during meiotic cell division in male and female germ cells (Berthet et al., 2003; Ortega et al.,
2003). Similarly, it was demonstrated that E-type cyclins are largely dispensable for mouse development (Geng et al., 2003). Nevertheless, in the absence of Cyclin E, endoreplication of trophoblast giant cells and megakaryocytes is severely impaired.
Cyclin E-deficient cells proliferate normally under conditions of continuous cell cycling but are not able to re-enter the cell cycle from the quiescent G0 state. Molecular analyses revealed that cells lacking Cyclin E fail to normally incorporate MCM (minichromosome maintenance) proteins into DNA replication origins during G0 to S-phase progression. 14
These findings define a molecular function for E-type cyclins in cell cycle re-entry and
reveal a differential necessity for Cyclin E in normal versus oncogenic proliferation
(Geng et al., 2003).
In addition to the well characterized Cdk2-dependent roles in the cell cycle,
Cyclin E was also found to be involved in the G1/S progression in a kinase-independent
manner (Geng et al., 2007). This study showed that by interacting with CDT1 and MCM,
a kinase deficient Cyclin E mutant could restore the MCM replicative helicase loading
and cell cycling in Cyclin E-deficient mouse embryonic fibroblasts. Another kinase-
independent function of Cyclin E is at the G1/S transition by localizing at the
centrosomes in a Cdk2-independent manner (Matsumoto and Maller, 2004).
Transcriptional regulation of Cyclin E
Cyclin E is a regulator of proliferation of mammalian fibroblasts and thus, its
levels are tightly regulated (Koff et al., 1992; Lam and La Thangue, 1994; Nevins, 1992).
Cyclin E/Cdk2 activity is precisely regulated at multiple levels, including transcriptional
and post-transcriptional, by binding of Cdk inhibitors such as the members of the Cip/Kip family (p21 and p27) as well as through modification of the Cdk activity by inhibitory or activating phosphorylation (Hwang and Clurman, 2005). Normally, expression of Cyclin
E is tightly controlled so that its levels reach a peak at the G1/S phase transition and then decline to an eventual loss of expression during S phase (Dulic et al., 1992; Ekholm et al.,
2001; Koff et al., 1992). This expression pattern is achieved by regulation at both
transcriptional as well as post-transcriptional levels. There are a number of putative 15
binding sites for E2F in the cyclin E promoter region, suggesting an E2F-dependent
regulation (Le Cam et al., 1999). A variant E2F binding site is a cyclin E repressor
module responsible for the periodic down-regulation of the cyclin E promoter until the
growing cells have reached the late G1-phase (Le Cam et al., 1999). This site facilitates
transcriptional repression by binding to a large complex, containing E2F4, DP1, and a pocket protein, whose role is to delay the expression of Cyclin E until late G1 (Le Cam et al., 1999; Polanowska et al., 2001) (Figure 1.4). pRb forms a repressor complex with a histone deacetylase (HDAC) and the SNF2-like (BRG1 and hBRM) component of the mammalian hSW1/SNF nucleosome remodeling complex, starting from the end of S- phase until late G1 (Zhang et al., 2000). The SW1/SNF complexes have an important role in transcriptional regulation, altering the chromatin structure by relieving the transcription from the nucleosome-mediated repression, thereby, opening access to the activators of transcription. The phosphorylation of pRb by Cyclin D/Cdk4 abrogates its interaction with HDAC and transactivates Cyclin E and, thereby, overcomes the G1 arrest
(Zhang et al., 2000). Cyclin E/Cdk2 can phosphorylate pRb or the hSW1/SNF component after Cyclin E reaches a certain level, when the interaction of pRb-hSW1/SNF is disrupted.
Deregulation of any of the components of this transcriptional complex could lead
to the unscheduled expression of Cyclin E, which is very common in cancer cells. Our
laboratory and others have previously shown that constitutive levels of several E2F1
target genes are increased in Rb-deficient fibroblasts (Almasan et al., 1995b; Herrera et
al., 1996). Similarly, Cyclin E levels are increased constitutively when Rb has been 16
inactivated by the HPV16-E7 oncogene expression in human foreskin fibroblasts (Martin
et al., 1998).
The above reports suggest that the transcription factor responsible for Cyclin E
induction might be E2F. E2F1 has been reported to be upregulated in response to DNA
damage in a manner analogous to p53, with its levels being unchanged following DNA
damage of epithelial cells with mutated p53 (Blattner et al., 1999). In contrast, we found that Cyclin E was upregulated in cells with mutated p53 expressed either endogenously or following its stable transfection (Mazumder et al., 2000). Northern and run-on analyses have indicated that the Cyclin E levels induced by radiation result mostly from transcriptional regulation. Cyclin E induction is not restricted to radiation and the hematopoietic cell lines we have investigated. Cyclin E levels were also increased following treatment with chemotherapeutic agents, such as the topoisomerase II inhibitor
VP16 (Mazumder et al., 2000). In addition, we recently found that Cyclin E is also upregulated in other cell types, such as those of prostate and epithelial cell types.
Although further work is required to establish whether E2F plays a regulatory role in radiation-induced Cyclin E expression, it is quite possible that Cyclin E is regulated by different mechanism by mitogens and genotoxic stress such as radiation. Understanding how this regulation takes place may provide additional targets for therapy.
The oscillations in CDK activities are largely determined by the levels of their interacting partners, which in turn depend on their synthesis, through transcriptional regulation, and proteolytic degradation. Phosphorylation by Cyclin H/Cdk7 and
Wee1/Myt1 kinases and dephosphorylation by the KAP, Cdc25A, and PP2C phosphatase 17
families play another important role in the regulation of the cell cycle machinery (Cheng et al., 2000; Donzelli and Draetta, 2003; Lew and Kornbluth, 1996; Moroy and Geisen,
2004; Parker et al., 1995; Poon and Hunter, 1995). In normal cells, Cdc25A is negatively regulated by stress signals such as irradiation through checkpoint-mediated ubiquitination. In checkpoint-deficient cells, Cdc25A is overexpressed, leading to radioresistant DNA synthesis (Falck et al., 2001).
18
Figure 1.4 Schematic representation of the progression through the different cell
cycle phases. Cyclin D family of cyclins activate Cdk4 and Cdk6 to phosphorylate and
inactivate pRb and its related family members (p107 and p130). This phosphorylation event results in the release of the E2F family of transcription factors which can increase the levels of proteins essential for the S phase progression [Cyclin E, thymidylate synthase (TS), dihydrofolate reductase (DHFR)]. The INK4 family of inhibitors bind and inhibit the Cyclin D/Cdk4, 6 complexes while p21 and p27 cyclin-dependent kinase inhibitors (CKI) form inhibitory complexes with Cyclin E/Cdk2. Whenever the cell is challenged with genotoxic stress caused by DNA-damaging agents like ionizing radiation, a signaling pathway triggers the cell cycle arrest to allow the DNA repair to take place. The damage is sensed by several kinases including ATM (Ataxia telangiectasia mutated) and the signal is relayed either directly or indirectly by means of
Chk1 or Chk2 kinases to the p53 tumor suppressor protein. Induction of p53 leads to increased transcription of the CKIs, which trigger cell cycle arrest.
19
20
Post-translational regulation of Cyclin E
Cyclin E periodicity is not only determined at the transcriptional level, but it is also maintained by post-translational regulation mechanisms through ubiquitin-dependent proteolysis (Clurman et al., 1996; Won and Reed, 1996). Ubiquitin-mediated protein degradation is a regulatory mechanism of a wide variety of cellular and tissue processes
such as cell stress response, DNA repair, transcription, cell cycle regulation, and cellular transformation (Hershko and Ciechanover, 1998).
The programmed degradation of many proteins is performed by the 26S proteasome. This recognizes a degradation signal consisting of polyubiquitin chains attached to lysine residues of the substrate by a cascade of enzymes, generically named
E1, E2 and E3. This ubiquitination is carried out by the successive action of ubiquitin- activating (E1), ubiquitin-conjugating (E2) and ubiquitin-protein ligase (E3) enzymes
(Hershko and Ciechanover, 1998; Jentsch, 1992)}. The E1 activates the ubiquitin (an
ATP-dependent process) and transfers it to the E2 through the formation of a thioester bond between the E2 active-site cysteine and the ubiquitin carboxy terminus (Pickart and
Rose, 1985). E3s bind to both a cognate E2 and the substrate and in the case of the
RING-E3 family they catalyze the ubiquitination of the substrate lysines directly by the
E2. E3-substrate association is the primary determinant of substrate specificity and is
often regulated by phosphorylation or other post-translational modification of the
substrate’s E3 binding motif, termed degron (Ang and Wade Harper, 2005; Skowyra et
al., 1997). Finally, the 26S proteasome recognizes the polyubiquitin chain and degrades 21
the substrate into small peptides recycling the ubiquitin molecules at the same time
(Jentsch, 1992).
One major class of E3 ligases includes those that contain proteins called cullins.
The cullin component of these large complexes serves as a scaffold for the assembly of a substrate recognition subunit as well as an E2 ubiquitin-conjugating enzyme (Deshaies,
1999). For example, the Cul1-dependent E3 ligase, also known as the SCF complex
(Skp1-Cul1-F box), consists of an adapter protein called Skp1, a ring-finger protein
Rbx1/Roc1, and a substrate recognition protein that contains a motif called an F box
(named after a common sequence identified in Cyclin F (Bai et al., 1996)). The human F
box protein responsible for the turnover of Cyclin E is Fbw7 (also known as hCdc4,
hSEL-10, and hAgo) (Gupta-Rossi et al., 2001; Koepp et al., 2001; Moberg et al., 2001;
Strohmaier et al., 2001). Fbw7 is expressed as three splice-variant isoforms which differ from each other only by their amino terminus (Spruck et al., 2002). Each splice variant is comprised by ten common 3′ exons and a unique 5′ exon. The common exons encode two domains critical for hCdc4 function. The function of the first domain, known as the F box
(Bai et al., 1996), is to interact with Skp1, thus associating the F box protein with the
Cul1/Cdc53 scaffold. The second domain is comprised of eight repeats of the WD40 motif (van der Voorn and Ploegh, 1992). These fold adopting a structure termed β propeller, also confirmed by the crystal structure of the yeast homolog of hCdc4 (Orlicky et al., 2003). The role of the WD40 is to recognize specific substrates with which it interacts. The hallmark of substrate recognition by the SCF complex is that substrate phosphorylation is a requirement (Figure 1.5). 22
Figure 1.5 Schematic representation of the Cyclin E turnover mechanisms. Cdk2- bound Cyclin E requires at least two phosphorylation events by GSK3β (at Thr380) and by Cdk2 (at Ser384) in order to be recognized by Fbw7. The Skp1-Cul1-Rbx1-Fbw7 complex recognizes phosphorylated Cyclin E and polyubiquitinates it. Ubiquitin is provided by the E2 Ubiquitin-conjugating enzyme, Ubc. On the other hand, free Cyclin E does not require any phosphorylation and is ubiquitinated by a Cul3-based complex. The additional factors that form this complex have not been identified yet, but may involve
BTB proteins as specificity factors.
23
24
In the cell cycle, ubiquitination plays a central role in the cell-cycle transitions
and checkpoints by establishing the strict temporal control of proteins such as cyclins and
cyclin-dependent kinase inhibitors (Sherr and Roberts, 1999). Cyclin E accumulation is
caused in part by the inactivation of SCFFbw7 ubiquitin-protein ligase that mediates its ubiquitination (Koepp et al., 2001; Strohmaier et al., 2001). Fbw7 was found to be mutated in several tumors and tumor cell lines (Hubalek et al., 2004; Moberg et al., 2001;
Rajagopalan et al., 2004; Spruck et al., 2002; Strohmaier et al., 2001). The Fbw7 gene is located in the 4q32 chromosomal region, which has been reported to be deleted in as many as 30% of human tumors (Spruck et al., 2002). Its inactivation leads to chromosomal instability (Rajagopalan et al., 2004), while its loss in mice can lead to embryonic lethality (Tetzlaff et al., 2004; Tsunematsu et al., 2004).
Degradation of Cyclin E involves both Cul1 and Cul3 proteins. Cul1 knock-out
mice die early during development with a subset of cells containing high levels of Cyclin
E (Dealy et al., 1999; Wang et al., 1999). Cul3-/- mice also arrest early in development.
However, a different subset of cells show increased levels of Cyclin E. Cul3 has been shown to bind and ubiquitinate Cyclin E in a transient transfection system (Singer et al.,
1999). It was shown that Cul3 is responsible for a constitutive pathway of Cyclin E degradation, which did not require binding to Cdk2 (Clurman et al., 1996; Singer et al.,
1999) or phosphorylation. On the other hand, Cul1-dependent turnover of Cyclin E is dependent on its association with Cdk2 and on its GSK3β-mediated (Welcker et al.,
2003) and auto-phosphorylation of Cyclin E at a series of serines and threonines
(Strohmaier et al., 2001; Yeh et al., 2001). 25
The primary Cyclin E recognition site by SCFFbw7 is Thr380, which is phosphorylated in vivo by GSK3β resulting in a phosphodegron sequence (Clurman et al., 1996; Welcker et al., 2003; Won and Reed, 1996). The unbound Cyclin E is degraded by the proteasome, and binding to Cdk2 protects it from degradation. However, phosphorylation of Cyclin E on Ser384 by its Cdk2 subunit has also been shown to be important for Fbw7-mediated Cyclin E degradation (Welcker et al., 2003). Thus, Cyclin
E/Cdk2 activity reverses the stabilizing effect of complex assembly. Skp1 and
Fbw7/Cdc4 are responsible for ubiquitin-dependent proteolysis of Cyclin E. Skp2 was thought earlier to be the F-box protein responsible for Cyclin E degradation since Skp2 gene inactivation by homologous recombination results in p27 and Cyclin E accumulation (Bai et al., 1996; Winston et al., 1999). However, since p27 is the predominant substrate of Skp2, its accumulation leads indirectly to the inhibition of
Cyclin E/Cdk2 kinase activity and phosphorylation of Cyclin E on Thr380, thus preventing its recognition by the Cyclin E-specific F-box protein. Instead, Fbw7, specifically targets ubiquitin-mediated proteolysis of Cyclin E (Koepp et al., 2001;
Strohmaier et al., 2001). The interaction of Cyclin E with this F-box protein depends on
Cyclin E phosphorylation by GSK3β on Thr380 (Welcker et al., 2003) and Thr62 autophosphorylation (Won and Reed, 1996). Some tumor cell lines that have high levels of Cyclin E also have mutations in the Fbw7 gene or express low levels of its mRNA, suggesting that Fbw7 may function as a tumor suppressor (Ekholm-Reed et al., 2004;
Spruck et al., 2002; Welcker and Clurman, 2008). 26
Recent investigation of the Cyclin E turnover mechanism determined that only α
and γ Fbw7 isoforms are required for this process (van Drogen et al., 2006). Specifically,
Fbw7γ (localizing in the nucleoli) is directly involved in the ubiquitination of Cyclin E
while Fbw7α (localized in the nucleoplasm) plays a regulatory function facilitating this
event. Fbw7α serves as cofactor for the prolyl isomerization of the Cyclin E carboxy-
terminal phosphodegron performed by the peptidyl prolyl cis-trans isomerase Pin1 (van
Drogen et al., 2006). Apart from Cyclin E, the SCFFbw7 has also been shown to bind and ubiquitinate Notch1 (Wu et al., 2001), c-Myc (Welcker et al., 2004), and c-Jun (Wei et al., 2005), which further supports the role of Fbw7 as a tumor suppressor.
The cyclin E gene is under direct control by the E2F family of transcription factors, which places cyclin E transcription under the control of the retinoblastoma protein (pRb) and its upstream regulators (Botz et al., 1996; Duronio et al., 1996; Geng et al., 1996; Ohtani et al., 1995). Inactivating Rb as part of the normal cell cycle progression
or by viral oncoproteins leads to its elevated transcription (Weinberg, 1995). The large T
antigen (LT) of the simian virus 40 (SV40) disrupts the cellular checkpoint mechanisms
that regulate cell division, DNA transcription, replication, and repair. The key regulators
that are targeted by LT are p53 and Rb. This binds directly to pRb and its family member
pocket proteins (p107 and p130) via an N-terminal LxCxE motif. In the case of p53, LT
binds the DNA binding domain of p53 blocking the p53-dependent growth arrest and
apoptosis (Levine, 1997). Another way LT interferes with the normal activity of the cell
is apparently through binding Fbw7 and mislocalization of the Fbw7γ isoform from the
nucleoli to the nucleoplasm (Welcker and Clurman, 2005). This way LT may interfere 27
with the timely Cyclin E turnover in a substrate-like manner, thus leading to increased
Cyclin E-associated kinase activity. Unlike typical Fbw7 substrates, LT is not degraded
by the SCFFbw7, but rather acts as a competitive inhibitor.
Cross-talk between the regulation of cyclin E transcription and that of Cyclin E protein stability results in fine tuning of Cyclin E levels, emphasizing its crucial role in cell cycle regulation and predicting deleterious effects of a constitutively high level of expression of Cyclin E, as seen in many cancer cells.
Deregulated expression of Cyclin E in cancer
Normal cell proliferation is under strict regulation, governed by checkpoints
located at distinct positions in the cell cycle. Cell cycle progression is controlled by two
major checkpoints, one at the G1/S boundary, when cells commit to DNA replication,
with the other one at the G2/M boundary during the commitment of cells to mitotic
division (O'Connor et al., 2000). The G1/S transition is regulated by the Restriction point
(R point), which is a point of no return, such that, if the cell passes that point, it no longer
needs mitogens and is committed to completion of DNA replication and the cell cycle
(Pardee, 1989). Any type of deregulation of the G1/S transition, along with the
disappearance of the R-point, is a hallmark of cancer, leading to uncontrolled cell
proliferation. The periodic appearance of Cyclin E coincides precisely with the timing of
the R point. In contrast to normal cells, mitotic cyclins appear prior to G1 cyclins in
tumor cells (Keyomarsi and Pardee, 1993). Another view is that passage through the R
point is a prerequisite for Cyclin E accumulation (Ekholm et al., 2001). This study found 28
that the post-mitotic G1 cells that had not yet reached R were negative for Cyclin E
accumulation, while cells that had passed R accumulated Cyclin E at variable times (1 to
8 hours) after passage through R and 2 to 5 hours before entry into the S phase.
As these cyclins and CDKs are important components of cell cycle control and cell proliferation, alterations or mutational changes in the expression of the corresponding
genes leads to oncogenesis (Evan and Vousden, 2001; Hunter and Pines, 1994; Pardee,
1989; Sherr, 1996). Cyclin E has been shown to be deregulated and overexpressed in
several solid tumors, including breast, colon, and prostate carcinomas (Akli and
Keyomarsi, 2003; Erlandsson et al., 2003; Keyomarsi et al., 1995; Keyomarsi et al.,
1994; Keyomarsi et al., 2002; Moroy and Geisen, 2004; Sherr, 1996). High levels of
cyclin E expression have been associated with the progression of different other tumors,
such as leukemias and lymphomas. Loss of Cyclin E regulation has been associated with
aggressive disease and poor patient outcome (Donnellan and Chetty, 1999; Hwang and
Clurman, 2005; Sandhu and Slingerland, 2000). This was shown to take place either by
amplification of the gene or by its overexpression.
Overexpression of Cyclin E results in accelerated G1/S phase progression
(Ohtsubo and Roberts, 1993; Ohtsubo et al., 1995; Resnitzky et al., 1994). Transgenic
mouse models with constitutive overexpression of Cyclin E develop malignant diseases,
indicating that Cyclin E is a dominant oncoprotein (Karsunky et al., 1999; Moroy and
Geisen, 2004). For example, overexpression of Cyclin E in the mouse mammary
epithelium is associated with hyperplasia, which can ultimately lead to tumorigenesis
(Bortner and Rosenberg, 1997). Furthermore, cell lines derived from tumors 29
overexpressing Cyclin E rarely show amplification of the gene, suggesting that the deregulation is at the post-transcriptional level (Porter et al., 1997), specifically in the turnover process. Cul3-dependent degradation of Cyclin E is important for the maintenance of quiescence of hepatocytes (McEvoy et al., 2007). Moreover, Cyclin E- deficient mice show increased resistance to oncogenic transformation by c-Myc, H-Ras, and dominant-negative p53 (Geng et al., 2003). Deregulation of Cyclin E was also shown to promote chromosome instability, detected as both chromosome losses and gains
(Spruck et al., 1999). This may result in tumorigenesis by facilitating amplification of oncogenes or loss of heterozygosity at tumor suppressor loci.
Although the protein is both synthesized and degraded in the cytoplasm, it exerts its functions in the nucleus where it translocates. In the nucleus, the nascent Cyclin E associates with its Cdk2 partner, activating its serine/threonine kinase activity shortly before entry into S phase (D'Urso et al., 1990; Koff et al., 1992). Accumulation of Cyclin
E in the cytoplasm either reflects increased synthesis, decreased degradation, or failure of
transport to the nucleus. In some types of cancer, the cyclin E gene (CCNE1) was found
to be frequently amplified (Schraml et al., 2003). In others, however, the post-
translational regulatory mechanism is impaired as mutations have been identified in the
cellular machinery responsible for Cyclin E proteolysis (Calhoun et al., 2003; Ekholm-
Reed et al., 2004; McVey et al., 1989; Rajagopalan et al., 2004; Spruck et al., 2002;
Strohmaier et al., 2001). Loss or low p27 expression as well as overexpression of Cyclin
E or Cdk2 are significantly associated with malignancy in ovarian cancers (Porter et al.,
1997). 30
Levels of Cyclin E and its low molecular weight derivatives in these tumor tissues
correlate strongly with survival in these patients (Keyomarsi et al., 2002). There can be
significant changes of Cyclin E or Cdk2 levels during carcinogenesis. During
progression from the primary to the lymph node-metastatic foci, the levels of Cyclin E
protein remain the same, while Cdk2 levels increase significantly (Li et al., 2001a).
However, during a similar transition from the primary to the liver-metastatic foci, Cyclin
E levels are apparently reduced, and those of Cdk2 diminish almost completely.
Additionally, the decrease of Cyclin E is significantly associated with large tumor size
and lymph node metastasis in primary carcinomas, with large tumor size and hepatic
metastasis being strongly related to low Cdk2 levels. Induced Cyclin E protein is related
to increased Cdk2, which is further associated with Ki-67 staining. Thus, Cdk2
overexpression could facilitate lymph node metastasis and both Cyclin E and Cdk2
overexpression may trigger the progression of early cancer.
Cyclin E overexpression is apparent in 10-25% of breast cancers. It is also a
strong predictor of endocrine therapy failure (Span et al., 2003), and an accurate predictor of poor clinical outcome (Keyomarsi et al., 2002; Loden et al., 2002). Studies have found
Cyclin E to be processed into more active low molecular weight (LMW) forms (Porter et
al., 2001) that are resistant to p21 and p27 inhibition and therefore associated with higher
cyclin-dependent kinase activity (Akli et al., 2004). It was shown that the LMW isoforms
are more potent than the full length Cyclin E in enhancing angiogenesis and metastasis
(Bales et al., 2005). Different studies have found Cyclin E to be cleaved by elastase
(Porter et al., 2001) as well as calpain (Wang et al., 2003). Apparently Cyclin E is able to 31
transactivate calpain 2 transcription (Libertini et al., 2005) and thus regulates its own proteolysis as well of that of other calpain substrates such as focal adhesion kinase
(Cooray et al., 1996), α-spectrin (Stabach et al., 1997), talin (Franco et al., 2004), paxilin
(Yamaguchi et al., 1994), E-cadherin (Rios-Doria et al., 2003), and β-catenin (Rios-Doria et al., 2004).
In many human cancers Cyclin E is expressed at levels substantially higher than those in normal tissues and shows frequent amplification of the genomic locus at which the cyclin E gene is located (19q12-q13). In some tumors, cyclin E gene amplification and protein accumulation are late events, whereas in other neoplasms, the increase of
Cyclin E is observed during the early stages of malignancy. Increased protein expression does not necessarily reflect a consequence of mutations in the cyclin E gene. The question of whether Cyclin E is merely a link in the chain of events that leads to cell proliferation or it is the driving force for cell replication is difficult to ascertain, as it may be tumor dependent. Elevated transcript levels of cyclins E are found in breast, colorectal, lung, and ovary/uterus tumor samples as compared to levels in normal tissues. Its overexpression in breast carcinoma could pave the way for their genomic instability
(Carroll et al., 1999; Lingle et al., 1998). Expression levels of Cyclin E are most likely to be elevated in breast tumors that lacked the estrogen receptor as compared to breast tumors with the receptor and normal breast tissue (Keyomarsi et al., 1994; Nielsen et al.,
1996).
The status of cyclin E expression is also important for other cancer types, such as sarcomas, NSCLC, leukemias, and lymphomas for prognosis and staging of tumors 32
(Akama et al., 1995; Dong et al., 2000; Erlanson and Landberg, 2001). The finding that
cyclin E is expressed at high levels in several types of leukemias, chronic lymphocytic leukemia, Hodgkin’s and Non-Hodgkin’s lymphoma (Erlanson and Landberg, 2001;
Erlanson et al., 1998), point to a role of Cyclin E in the development of these neoplastic diseases. Cyclin E and HPV expression seem to also correlate very well, thus making
Cyclin E a good biomarker for infection and early stage of preneoplastic lesions (Martin et al., 1998). Cervical, prostate, breast (Erlandsson et al., 2003), and renal cell carcinoma
(65%) (Hedberg et al., 2002) as well as leiomyosarcomas (Noguchi et al., 2000) exhibit
high expression of Cyclin E. Cyclin E may be also used as an independent marker in the
prognosis of metastasis-free survival of lymph node-negative breast cancer (Kuhling et
al., 2003) as well as in hepatocellular carcinomas (Ohashi et al., 2001) and testicular
germ cell tumors (Sugimoto et al., 2002).
Therapeutic approaches involving cell cycle inhibitors
The deregulation of cell cycle checkpoints and the molecules associated with
them may transform a normal cell into a cancer cell. This most often occurs by interfering
with the basic cell cycle regulatory machinery to activate cell cycle entry that leads to
deregulation of the cell cycle in dividing somatic cells. Finding anticancer drug treatment
strategies tailored to modulate specific cell cycle components in particular tumors is of
great interest. Candidate targets for such strategies include critical cell cycle molecules
involved in the G1 to S phase or G2 to M phase transitions (Boonstra, 2003; Gali-
Muhtasib and Bakkar, 2002; Lee and Yang, 2003). S phase sensitization through G1/S 33
cyclin or E2F deregulation could be an effective therapeutic approach. Most tumor cells
are characterized by aberrations in the cell cycle control primarily incurred through
inactivation of pRb by the phosphorylation-induced hyperactive CDKs or inactivating
viral proteins (Senderowicz, 2003). Clearly, Rb deficiency allows cells to progress into the S phase inappropriately, conditions under which they are vulnerable to chemo and radiotherapy (Almasan et al., 1995a). Rb deficiency leads to increased expression of E2F target genes that include cyclin E. Increasing levels of cyclins using proteasome inhibitors and thus sensitizing target cells has been also used effectively to target S phase
cyclin complexes as an effective therapeutic approach to induce apoptosis (Bunn, 2004).
Uncontrolled CDK activity is often the cause of human cancer. The function of
CDKs is tightly regulated by cell cycle inhibitors such as p21Cip1/Waf1 and p27Kip1.
Following DNA damage or anti-mitogenic signals, p21 and p27 associate with
Cyclin/CDK complexes to inhibit their catalytic activity and induce cell cycle arrest.
Loss of function of these genes is often observed in different types of human cancers
(Facchinetti et al., 2004; Porter et al., 1997). Molecular targeting of the p21 and p27 inhibitors has been reported in epithelial cells. Thus, cleavage of p21Cip1/Waf1 and p27Kip1 releases Cyclin A/Cdk2 from inhibition raising its expression levels and induces apoptosis (Jin et al., 2000; Levkau et al., 1998; Poon and Hunter, 1998).
Deregulation of the cell cycle components is one of the hallmarks of neoplastic cells (Sherr and McCormick, 2002). However, the most common chemotherapeutic agents used to prevent the growth of cancer cells are also toxic to normal cells. In order to identify a novel cell cycle target that would be selective against cancer cells, 34
determination of the expression pattern as well as the mechanism of the target in normal
versus tumor cells will have to be pursued so that therapeutic application of that target could be achieved. Genetic abnormalities of CDKs (e.g.1, 2, 7), cyclins (e.g. A and E) and CKIs (p21 and p27) have been reported in human cancers. Enforced perturbations of
CDKs and their interacting partners can enhance neoplastic transformation and, thereby, making them extremely attractive targets for cancer therapy (Figure 1.6).
Structural studies of cyclins and CDKs have revealed that CDKs provide suitable drug targets given their more rigid structure and identifiable domains, such as that responsible for ATP-binding. In contrast, cyclins have a highly flexible structure that is not amenable for target development (Davies et al., 2002a; Davies et al., 2002b).
Inhibitors targeting CDKs have become of great interest in cancer therapy with more than
50 compounds being evaluated for antitumor activity, some of which are already in preclinical trials (Dai and Grant, 2003; Senderowicz, 2003). These inhibitors either target the CDKs themselves, through their ATP-binding site, or their upstream regulators. The first group includes flavopiridol and roscovitine, the second UCN-01, perifostine, and lovastatin (Senderowicz, 2003). Indolinones, UCN-01, and paullones, are currently tested in clinical trials (Dai and Grant, 2003; Senderowicz, 2003). While demonstrating clinical activity, neither acts specifically against Cdk2. Other more specific Cdk2 inhibitors are currently in preclinical development (Lane et al., 2001; Monaco et al., 2004; Yu et al.,
2003a). These include purine based CDK inhibitors, such as olomoucine, purvalanols, and roscovitine, which display greater selectivity for Cdk1, 2, and probably 5. Novel derivatives, such as R-roscovitine (CYC202) show great promise (Whittaker et al., 2004). 35
The efficacy of these inhibitors may not be the result of their CDK-inhibitory activity alone but also of their effect on transcription of CDK complex components, such as
Cyclin D1 as well as apoptotic regulators (Dai and Grant, 2003).
36
Figure 1.6 Schematic representation of the cell cycle and the different therapeutic
approaches undertaken. Several pharmacologic inhibitors have been generated to target the cyclins/CDKs at different phases of the cell cycle and several of them are in preclinical trials. Most of the inhibitors prevent the activity of more than one Cyclin/CDK complex. The effect of Cyclin E can be blocked by the expression of a dominant negative
(dn) Cdk2 which may constitute another therapeutic approach to target Cyclin E.
37
38
A novel class of therapeutics comprises inhibitors of the proteasome (bortezomib) which have been shown for example to cause the accumulation of Cyclin A and B, G2/M arrest and induce apoptosis in lung cancer cell lines (Bunn, 2004). Another new emerging compound with antitumoral activity is a fusion protein made up of a Cyclin A/Cdk2 binding peptide and an F-box protein, which targets the complex to ubiquitination and proteasomal degradation. This leads to a decrease of Cyclin A and Cdk2 levels followed by apoptosis both in vivo and in vitro with no significant toxicity to normal cells.
Apparently, the down-regulation of Cyclin A is essential for the cytotoxic activity.
However, decreasing both Cyclin A and Cdk2 expression causes the highest cytotoxic effect (Chen et al., 2004). Previous studies have shown that such Cyclin A/Cdk2-binding peptides induced apoptosis by further deregulating E2F expression (Chen et al., 1999).
Thus, an S-phase checkpoint seems to be activated and this involves E2F whose DNA binding function is in turn negatively regulated by Cyclin A (Krek et al., 1995; Pagano et al., 1992; Pruschy et al., 1999). A different type of Cdk2-Cyclin A antagonists (which do not block the ATP site) bearing the RXL peptidic motif were found to selectively induce apoptosis in cells in which pRb and Cyclin D were deregulated (Mendoza et al., 2003).
Apoptosis
Apoptosis is a universal genetic program of cell death in higher eukaryotes that is
an essential component of cellular development and differentiation, as well as to the 39
responses to various types of stress (Danial and Korsmeyer, 2004). Programmed cell death was shown to be critical for morphogenesis (Zuzarte-Luis and Hurle, 2005), differentiation (Hutchins and Barger, 1998), maintenance of homeostasis (Vaux and
Strasser, 1996), as well as regulation of the immune system (Rathmell and Thompson,
2002). Loss of control of apoptosis may lead to a series of pathological effects such as cancer, neurodegenerative and autoimmune disorders, viral infections, and ischemic diseases (Fadeel and Orrenius, 2005).
Apoptosis is defined by the typical morphology of the dying cell (Wyllie et al.,
1980), specifically by the chromatin condensation which is often accompanied by nucleus fragmentation. The cell membrane blebs and appears budding, resulting in membrane- enclosed structures called apoptotic bodies (Arends and Wyllie, 1991). These apoptotic bodies are then rapidly recognized and cleared by phagocytic cells without mounting an inflammatory response. On the other hand, necrosis results in rupture of the plasma membrane, followed by the spilling of the inner cellular contents, resulting in a strong inflammatory response (Leist and Jaattela, 2001).
Many apoptotic biochemical markers have been identified and are currently used to monitor it in experimental settings: DNA fragmentation by TUNEL assay (terminal deoxynucleotidyl transferase-mediated dUTP-digoxigenin nick end labelling) or subdiploid DNA content assays, activation of caspases by western blotting or immunofluorescence, cell surface externalization of phosphatidylserine by binding of
Annexin V (Hengartner, 2000). 40
Apoptosis can be triggered by a variety of factors: DNA damage induced by
ultraviolet light, ionizing radiation or chemotherapeutic drugs, elevated concentration of
oxidants within the cell or cell surface stimuli such as death activators, and growth factor
withdrawal such as serum starvation. Genotoxic agents, such as ionizing radiation, induce
apoptosis in many cell types, most potently in those of hematopoietic origin. Apoptosis is
frequently associated with proliferating cells, implying the existence of molecules in late
G1 and S whose activities facilitate execution of the apoptotic process.
Apoptosis is mediated through two major pathways, the death receptor pathway
and the mitochondrial pathway (Hengartner, 2000). Many proteins regulate or actively take part in the execution of the apoptotic process, but the main four types of molecules that are involved in this process are the caspases, the members of the Bcl-2 family, members of the tumor necrosis factor family and its receptors, and adaptor proteins.
Apoptosis is orchestrated by the function of a set of proteases termed caspases (cysteine aspartate proteases) which are critical regulators of the execution phase of apoptosis, triggered by many factors, including genotoxic agents (e.g. γ-irradiation or treatment with anti-cancer agents) (Chen et al., 2000b; Gong and Almasan, 2000; Gong et al., 1999).
These enzymes cleave specific substrates causing the characteristic cellular morphology.
TRAIL, APO2L
TRAIL (TNF-Related Apoptosis-Inducing Ligand) is a member of the Tumor
Necrosis Factor (TNF) family of death ligands. It is a transmembrane protein which can 41
be cleaved to generate a soluble ligand. There are two types of TRAIL receptors: TRAIL-
R1 and TRAIL-R2 which are agonistic receptors and TRAIL-R3 and TRAIL-R4 which
compete with the first two for binding of the ligands. Binding of TRAIL to its agonistic
receptors results in receptor trimerization, recruitment of adaptor molecules (FADD), and
Caspase-8 to their cytosolic tails to form the Death-Inducing Signaling Complex (DISC).
Adaptor proteins
The adaptor proteins are molecules that relay the apoptotic signal from the
transmembrane TRAIL receptors to the cell death regulators and effectors, such as
caspases. One of these is the Fas-Associated Death Domain (FADD or MORT-1). Upon
engagement of the death receptors by their specific ligands or agonistic antibodies,
FADD is recruited to the cytosolic tail of these receptors by homotypic interactions with
their Death Domains (DD) (Boldin et al., 1995; Kischkel et al., 1995; Micheau and
Tschopp, 2003). In turn, FADD, recruits pro-caspase-8 by interacting with its Death
Effector Domain (DED). The complex comprised of the TRAIL-R, FADD and pro-
caspase-8 is known as the DISC.
Apaf-1 is another adaptor protein which acts as a scaffold for the formation of the
apoptosome complex. Its caspase-recruitment (CARD) domain acts as a docking station
for pro-caspase-9. With the contribution of cytochrome c and dATP, Apaf-1 oligomerizes
to form the apoptosome complex (Cain et al., 2002; Rodriguez and Lazebnik, 1999; Zou
et al., 1997). This facilitates the activation of Caspase-9 which is crucial for the
subsequent activation of the executioner Caspase-3. 42
Cytochrome c
Once the cells are committed to cell death, apoptogenic factors, the best known of which is cytochrome c, are released from mitochondria to initiate the caspase cascade
(Chen et al., 2000b). Cytochrome c acts as a cofactor to stimulate the association of
Apaf-1 with Caspase-9 (Li et al., 1997), which then initiates activation of the caspase cascade.
Bcl-2 family members
The Bcl-2 family members are the main regulators of the mitochondrial death pathway, The first discovered member of this family, Bcl-2, was identified in human B cell lymphoma while studying the chromosomal translocation t(14;18) (Tsujimoto et al.,
1984). Many Bcl-2 family members have been uncovered to date and their main role is to regulate the release of cytochrome c from the mitochondria (Kaufmann and Hengartner,
2001). Based on their structure, they may either manifest anti-apoptotic properties such as
Bcl-2, Bcl-xL, Mcl-1, Bok or promote apoptosis such as Bax, Bcl-xS, Bak, Bad, Bim,
Puma, Noxa. These proteins have the ability to homo- or hetero-dimerize and the balance in the ratio between the pro- versus anti-apoptotic determines the susceptibility of the cell to the apoptotic signal (Oltvai et al., 1993).
The cell death antagonist proteins are characterized by the presence of all Bcl-2 homology domains (BH1-BH4). As most of these have a hydrophobic transmembrane domain, they can localize to different organelles, in particular the mitochondria (Hsu et al., 1997; Janiak et al., 1994; Wilson-Annan et al., 2003). These proteins block cell death 43
by preventing the disruption of the mitochondrial membrane. On the other hand, the
death promoting proteins have all the BH domains except for the BH4 domain. Following
the apoptotic signal, Bax which is found in the cytosol, translocates to the outer mitochondrial membrane (Griffiths et al., 1999; Hsu and Youle, 1998; Wolter et al.,
1997) where it oligomerizes to form a pore-like structure, promoting the release of
cytochrome c (Antonsson et al., 2000; Wei et al., 2000). This translocation can be
promoted by another type of Bcl-2 members, those that have only the BH3 domain, such
as tBid [resulted from cleavage of Bid by caspase-8 (Desagher et al., 1999)], Bim and
Bmf [microtubule-associated proteins (Puthalakath et al., 1999; Puthalakath et al., 2001)],
Noxa and Puma [induced by p53 (Nakano and Vousden, 2001; Oda et al., 2000)]. These
hetero-dimerize with the pro-apoptotic, Bax and Bak to induce their activation. This can
be prevented by Bcl-2 or Bcl-xL by sequestering the BH-3 only proteins or by directly
interfering with channel formation, thus inhibiting the release of mitochondrial
apoptogenic factors, such as cytochrome c, AIF, Smac/Diablo and HtrA/Omi.
Caspases
Caspases are the most important proteins in the apoptotic pathway. They are
members of a cysteine protease family that can hydrolyze peptidic bonds after an aspartic
acid (Asp) residue. Fourteen members of this family have been identified to date with
eleven of them being present in humans (Earnshaw et al., 1999). Caspases are involved in
apoptosis signaling (caspases 2, 3, 6, 7, 8, 9, and 10) as well as cytokine processing
(Earnshaw et al., 1999; Strasser et al., 2000). They are synthesized as inactive zymogens 44
containing a prodomain, a large, and a small unit. They are activated by proteolytic cleavage between the large and the small subunits followed by cleavage between the large subunit and the prodomain. Upon maturation, caspases form a tetramer of two small and two large subunits. Caspases are able to activate each other resulting in an amplification of activity through a protease cascade (Thornberry and Lazebnik, 1998).
The caspases involved in apoptosis fall within two categories: the initiators
(caspase 2, 8, 9, and 10) and the effectors (caspase 3, 6, and 7). The initiator caspases have long prodomains containing protein-protein interaction domains such as DED or
CARD while the effector caspases have short domains. Caspase-8 and -10 are the initiator caspases in the death receptor pathway (Denault and Salvesen, 2002). Binding of
Fas or TRAIL to their receptors results in receptor aggregation and recruitment of FADD and Caspase-8 through their DED domains to form the DISC (Muzio et al., 1996; Peter and Krammer, 2003). Once recruited, Caspase-8 is activated and initiates apoptosis by direct cleavage of effector caspases (Alnemri et al., 1996). Caspase-8 may also be cleaved and activated by Caspase-6. On the other hand, Caspase-9 is activated following its recruitment to Apaf-1 through its CARD domain. The effector caspases -3, -6, and -7 exist in the cytosol as inactive pro-caspase dimers and are activated through cleavage by the initiator caspases to generate the fully active enzymes (Boatright et al., 2003). They target various cytosolic or nuclear substrates whose degradation leads to the characteristic apoptotic cell morphology. Among these, there are proteins crucial for the maintenance of cellular cytoskeleton, DNA repair, transcription and translation, signal transduction, and cell cycle control (Hengartner, 2000). 45
Type / pathway Caspase-3 substrates
Kinases PKCδ, MEKK-1, FAK, PAK-2, PITSLRE
Transcription / SREBPs, hnRNPs C1/C2, U1-70 kD, smRNP Translation
DNA repair / DNA-PKcs, PITSLRE, pRb, D4-GDP-DI, NuMa, Mdm-2, PARP Cell Cycle
Structural lamin A,B, G-actin, fodrin, Gas2, presenilins, gelsolin, keratin1β
Table 1. Critical protein substrates of Caspase-3
Other apoptosis-regulatory proteins
The inhibitor of apoptosis proteins (IAPs) play an important regulatory role in the
apoptotic signaling pathway by tightly controlling the activation of caspases. c-IAP1, c-
IAP2, and XIAP interact and directly inhibit Caspase-3, -7, and -9 through their BIR
domains (Salvesen and Duckett, 2002; Takahashi et al., 1998).
The activation of caspase-8 in the DISC may be inhibited by proteins belonging to the FLICE-inhibitory protein (FLIP) family. There are several viral FLIPs named v-
FLIPs while the human homolog is termed c-FLIP. This may be present in two splice variants, the short c-FLIPs and the long c-FLIPL. These proteins are characterized by the presence of two DED domains. They act by preventing the activation of Caspase-8 and -
10 (Donepudi et al., 2003; Krueger et al., 2001). 46
Another protein with an important role in the execution of apoptosis is the
Caspase-Activated DNAase (CAD). Its function is to degrade the chromosomal DNA.
CAD is kept inactive bound to ICAD (inhibitor of CAD) (Enari et al., 1998; Nagata et al., 2003). When ICAD is degraded by caspase-3, CAD translocates to the nucleus and cleaves the DNA in small fragments which is a hallmark of apoptosis.
Apoptosis Inducing Factor (AIF) is a mitochondrial protein that bears both mitochondrial and nuclear localization sequences. During apoptosis, AIF is released from the mitochondria and translocates to the nucleus where it is responsible for caspase- independent induction of condensation of chromatin and DNA fragmentation (Cande et al., 2002; Susin et al., 1999).
Apoptotic pathways
Apoptosis may be initiated and executed through two well described pathways: the death receptor pathway (the extrinsic pathway) or the mitochondrial pathway (the intrinsic pathway). The death receptor pathway is initiated when ligands belonging to the
TNF family such as FasL (Apo1L), TNF-α, and TRAIL (Apo2L) engage their receptors
(Fas or CD95, DR1, DR2) on the plasma membrane. This interaction leads to the recruitment to their cytosolic tails of the adaptor protein FADD and pro-caspase-8 or -10 to form the DISC. The formation of this complex leads to the activation of Caspase-8
(Kischkel et al., 1995; Kischkel et al., 2000; Sprick et al., 2000). In turn, activated
Caspase-8 or -10 cleaves and activates the effector Caspases -3, -6, and -7 (Chen et al.,
2001) resulting in cleavage of cellular substrates (Figure 1.7). 47
Alternatively, Caspase-8 may cleave the BH3-only protein Bid generated tBid
(truncated Bid) which activates Bax, thus initiating the mitochondrial apoptotic pathway
(Wei et al., 2000). This pathway may also be activated by virus infection or DNA damage. In this pathway, the pro-apoptotic protein, Bax, translocates from the cytosol to the outer mitochondrial membrane (Nomura et al., 1999). Here, it oligomerizes following a change in conformation, and forms pores in the membrane through which cytochrome c is being released. Together with it, many other apoptosis regulatory factors such as
Smac/DIABLO (Du et al., 2000; Srinivasula et al., 2001; Verhagen et al., 2000),
HtrA/Omi (Hegde et al., 2002; Martins et al., 2002), ICAD, AIF (Joza et al., 2001), and endonuclease G (Li et al., 2001b) are being released into the cytoplasm. Cytochrome c and pro-caspase-9 are being complexes by Apaf-1 in presence of dATP to form a large complex named the apoptosome. This leads to the activation of caspase-9 (Strasser et al.,
2000), which in turn cleaves and activates the effector caspase-3.
48
Figure 1.7 Schematic representation of the extrinsic and intrinsic apoptotic
pathways. The extrinsic apoptotic pathway consists of Apo1 and Apo2 ligands interacting with their receptors on the cell membrane leading to the formation of DISC comprised of the cyotosolic part of the receptor, FADD, and pro-caspase-8. This causes the activation of caspase-8 which cleaves activating caspase-3. The intrinsic apoptotic pathway consists of cleavage of Bid by caspase-8 generating tBid which activates Bax.
Activated Bax translocates to the mitochondria where forms pores through which cytochrome c is being released into the cytosol. Cytochrome c complexes with Apaf-1 and pro-caspase-9 forming the apoptosome. This leads to activation of caspase-9 which, in turn, cleaves and activates caspase-3. Caspase-3 cleaves many cellular protein
substrates including ICAD thus releasing the nuclease CAD. In the nucleus, p53 up-
regulates the transcription of Bax, Puma and Noxa. Together with cytochrome c, other
pro-apoptotic factors are released from the mitochondria, such as, AIF and Smac/Diablo.
49
50
Cells that are able to undergo apoptosis through the DR pathway are termed type I
cells, whereas cells in which caspase-8 cleaves Bid, which induces apoptosis through the
mitochondrial pathway are called type II cells (Scaffidi et al., 1998).
Apoptosis represents the most severe cellular response to genotoxic stress, which is mounted when the cellular damage, primarily to the DNA, is beyond the capacity of the DNA repair proteins. Experimental evidence suggests that the cytotoxic effects of radiation and many forms of chemotherapy are mediated through a final common pathway that involves the activation of apoptosis. Radiation leads to cell death mainly through apoptosis in numerous normal tissues. Lymphoid cells, immature hematopoietic cells, and epithelial cells of the small intestine are among the most radiosensitive in the
body. Many cancer therapies are designed to target the apoptotic pathways within a cell, and the failure of cytotoxic therapies in many cancer cells have been attributed to apoptosis impairment. Two key early cellular events associated with radiation-injured apoptosis are the exposure of the phosphatidylserine on the plasma membrane and activation of caspases. Once cells are committed to cell death, apoptogenic factors, the best known of which is cytochrome c, are released form mitochondria to initiate a caspase cascade. Caspase activation is a critical regulator of the execution phase of apoptosis induced by many factors, including ionizing radiation. Upon activation through proteolytic cleavage, caspase may further cleave other caspases or cellular target proteins that include Bcl-2, Cyclin E, and poly (ADP-ribose) polymerase-1 (PARP-1). While other forms of cell death have been recently described (e.g. autophagy, necrosis with 51
different variations), it is believed that the apoptotic response is the predominant
mechanism of cell death.
DNA damage and repair mechanisms
Cellular DNA damage can be responsible for mutagenesis and the development of
cancer (van Gent et al., 2001). The human DNA may be subjected to several thousand to a million damaging events per day, generated by either external or internal metabolic processes. Changes in the genome can lead to errors in transcription and subsequently translation of the proteins necessary for cellular function. Certain DNA mutations can also be carried over into daughter cells if they are not repaired prior to mitosis.
When cells lose their ability to effectively repair damaged DNA, they can have
three possible fates. One possibility is to undergo senescence and become dormant. It has been reported that senescence could occur in cancer cells in vivo as well as in vitro, by stopping mitosis and preventing the cell from evolving further (Braig et al., 2005; Chen et al., 2005b; Collado et al., 2005; Michaloglou et al., 2005). Another outcome is cell death.
Sufficient DNA damage may trigger an apoptotic signaling cascade, forcing the cell into programmed cell death (O'Driscoll and Jeggo, 2006). Finally, the cells may undergo malignant transformation, become immortal, and divide uncontrollably (Mills et al.,
2003). To be able to cope with the different types of DNA damage that may occur, cells have developed several repair mechanisms such as mismatch (MMR), base excision 52
(BER), and nucleotide excision repair (NER). Double-strand breaks are resolved by the non-homologous end joining (NHEJ) (Lieber et al., 2003) or homologous recombination
(HR) (Haber, 1995) pathways. Cells may choose to proceed into apoptosis or senescence if overwhelming damage has taken place instead of expending energy to repair the DNA damage. The rate at which a cell is able to perform DNA repair depends on the cell type and its age.
For many years, it was believed that the main cause of DNA mutations leading to cancer was DNA damage induced by exogenous sources. However, more recently it was suggested that malignancy could also result from DNA damage caused by endogenous sources (Jackson and Loeb, 2001). There are two types of DNA damaging agents, physical and chemical. Physical mutagens are mainly sources of radiation, including UV
(200-300 nm) and ionizing radiation (X-rays). UV radiation is responsible for the production of covalent bonds that crosslink adjacent pyrimidine (cytosine and thymine) bases in DNA. On the other hand, IR causes DNA damage by generating free radicals within the cell that create reactive oxygen species (ROS). These, if not neutralized, may cause single-strand and double-strand breaks in the double helix. Chemical agents may alter the DNA structure by covalently attaching alkyl groups to DNA bases. Others are found as chemical inert precursors, which have to be converted metabolically to highly reactive carcinogens. These can further react with the DNA forming DNA adducts. For example, benzopyrene has to be activated to benzopyrenediol epoxide by the cellular oxidases to be able to form covalent DNA adducts. 53
DNA damage may also result from different other endogenous biochemical processes (De Bont and van Larebeke, 2004). Depurination (loss of adenine or guanine from the deoxyribosyl phosphate chain) is the most common event (Lindahl and Nyberg,
1972). Depyrimidation (loss of thymine or cytosine) also takes place but it is a lot less common than depurination. Another type of damage is the deamination of adenine, guanine, and cytosine rings to form the unnatural bases hypoxanthine, xanthine, and uracil. If the DNA repair enzymes do not recognize and remove these bases, an uncorrected uracil may be misread as a thymine during DNA replication and result in a C to T point mutation.
Another type of DNA modification, which could lead to misreading of DNA is
DNA methylation. The molecule responsible for transferring the methyl group onto DNA is S-adenosylmethionine (SAM). SAM is an intracellular metabolic intermediate that contains a highly reactive methyl group. The addition of a methyl group results in the formation of a DNA CpG sequence as this process occurs at the 5-position of the cytosine ring of a cytidine base that is 5’ to a guanosine base. The 5-methyl-cytosine product is very prone to spontaneous deamination, which represents a significant source of mutation error. Loss of the amine group results in a thymine base. Thus, the whole process leads to the creation of a C to T point mutation.
Reactive oxygen species (ROS) resulting from cellular metabolic processes can modify both purine and pyrimidine bases by oxidation. The most common change is guanine oxidized to 8-oxo-7,8-dihydroguanine, which results in the nucleotide 8-oxo- deoxyguanosine (8-oxo-dG). However, 8-oxo-dG instead of pairing with deoxycytidine, 54
can base-pair with deoxyadenosine. Enzymes belonging to the mismatch repair pathway
are responsible for detection and correction of this error. However, if the error is not
repaired, the DNA subsequently replicated will contain a C to A point mutation. ROS
may also cause depurination, depyrimidation and single-strand or double-strand breaks in
the DNA.
The most deleterious DNA damage consists of the single strand (SSB) and double
strand breaks (DSB) in the DNA (Khanna and Jackson, 2001). SSBs may result from
damage to the deoxyribose moiety of the DNA chain. SSBs may also be caused by the
base excision repair pathway after removal of the deoxyribose phosphate by AP-1
endonuclease. DSBs occur most often during the S phase, when the DNA may be more
susceptible to breakage as it is unwinding to serve as a template for replication (Brem and
Hall, 2005). Another source of DSBs is the V(D)J recombination process (Raghavan et
al., 2001). The extraordinary variability of the immunoglobulins and the T-cell receptors
is achieved through recombination of the V, D, and J gene segments to create mature
exons encoding the final protein. The process of recombination is mediated by the RAG1,
RAG2 (recombination activating genes 1 and 2) endonucleases and HMG1 (high
mobility group 1). RAG1 and 2 bind to specific signal sequences flanking the coding
regions and create a double-strand cleavage (Fugmann et al., 2000; Gellert, 2002). The
coding sequences and the two signal ends will be joined using the NHEJ pathways
(Lieber, 1999; Lieber et al., 2003).
The danger posed by the presence of DNA DSBs caused by DNA-damaging agents, such as ionizing radiation or chemotherapeutic drugs arises from their ability to 55
cause genomic instability through loss of heterozygosity and chromosomal translocations
(van Gent et al., 2001). These will eventually lead to cancer or apoptosis. As few as one or two unrepaired breaks would be sufficient to trigger the death of a cell. Throughout the
mammalian cell cycle, it is believed that DSBs are being repaired by HR during late S
and G2 phases while NHEJ is responsible for the repair during the rest of the cell cycle
(Takata et al., 1998). While repair by HR is an error-free process (West, 2003), NHEJ can
be very imprecise and thus lead to potential gene alterations. During the repair performed by NHEJ, the broken DNA ends get resected or nucleotides are filled in to generate regions of microhomology. In general, DNA ends that do not share significant homology will be repaired and joined by the NHEJ following resection to generate sites of microhomology (Roth and Wilson, 1986). However, when the NHEJ machinery is not entirely functional due to genetic mutation or absence of one the proteins involved in it, other pathways or enzymes may be employed for the repair of the lesion (Ferguson and
Alt, 2001).
Homologous recombination
The most common type of HR employed in repair of DNA DSBs is gene
conversion. This process takes advantage of a donor sequence with homology to both
sides of the DSB (the sister chromatid) which serves as template for the fill in process
(Elliott et al., 1998; Taghian and Nickoloff, 1997). The information is faithfully
incorporated resulting in an error-free repair process which is very important for the
maintenance of genomic integrity (Haber, 1995). The DSBs repaired by this mechanism 56
are usually caused in the S phase during attempts of replicating the DNA across a single-
strand break or an unrepaired lesion which results in the collapse of the replication fork.
Even though there is evidence of preference of HR or NHEJ repair for certain
types of DNA DSBs [V(D)J recombination is performed by NHEJ while DNA breaks
caused during meiosis by Spo1 require HR (Keeney, 2001)] and despite a cell cycle
phase preference, the two pathways are still coupled for DSB repair (Richardson et al.,
1998). This indicates that the choice regarding which pathway will be used for a particular DNA repair pathway, is influenced by many other factors. Studies in Ku70-/- cells have shown increased HR activity which suggests that upon binding of Ku70 to the ends of a DSB it prevents the access of HR repair machinery (Pierce et al., 2001). It appears that NHEJ and HR may compete for the repair of the same DNA DSB.
Ku70/Ku80 heterodimer
The Ku70/Ku80 heterodimer is a critical component in the NHEJ pathway (Gullo et al., 2006; Lees-Miller and Meek, 2003). Other important roles of Ku70 are in V(D)J recombination of the antigen receptor genes (Gellert, 2002), telomere maintenance, and in the transcriptional activation or repression of several genes (Shi et al., 2007).
Furthermore, both Ku70 and Ku80 are autoantigens that have been associated with connective tissues diseases such as systemic lupus erythematosus and scleroderma. Ku70 was found to be regulated through several mechanisms: heterodimerization with Ku80, intracellular localization (Koike and Koike, 2005), post-translational modifications such 57
as phosphorylation, acetylation, ubiquitination (Gama et al., 2006), and sumoylation
(Yurchenko et al., 2008).
In the nucleus, Ku70 is found in a very tight heterocomplex with Ku80. Ku70 is
very abundant and manifests high affinity for free DNA ends (Lieber et al., 2003). The
three-dimensional structure of the Ku70/Ku80 heterodimer has the shape of a doughnut
with a bulky bottom and a narrow bridge which can encircle and bind the DNA in a
sequence-independent manner (Walker et al., 2001). Even if they share minimal sequence
homology, the Ku70 and Ku80 subunits adopt a very similar three-dimensional structure, each of them contributing to the base, pillars, and bridge (Figure 1.8). One of the main roles in the NHEJ is to recruit DNA-PKcs. However, to accommodate the large 460-kDa
DNA-PKcs, Ku70/Ku80 must slide inward on the DNA in an ATP-dependent manner
(Turchi et al., 2000; Yoo and Dynan, 1999). The recruited DNA-PKcs then phosphorylates several substrates including Artemis (Poinsignon et al., 2004), the
Mre11/Rad50/Nbs1 complex (Uziel et al., 2003), and XRCC4/Ligase IV (Wang et al.,
2001b) which ultimately performs the repair of the DNA lesion.
Studies in Ku70-/- and Ku80-/- have shown that the absence of Ku leads to
dramatic radiation hypersensitivity, deficiency in DNA DSB repair, V(D)J
recombination, and impaired lymphocyte development. Apart form the radiosensitivity
imparted by the defects in the NHEJ process, these also lead to severe combined
immunodeficiency due to impaired V(D)J recombination (O'Driscoll et al., 2004; Rooney
et al., 2004; Schwarz et al., 2003). Even though most of its functions are nuclear, Ku70 58
also plays a cytoprotective role in the cytosol by interacting with and sequestering the pro-apoptotic protein Bax (Gomez et al., 2007; Yoshida et al., 2004).
59
Figure 1.8 Three-dimensional structure of the Ku70/Ku80 heterodimer. Ku70 (pink) and Ku80 (blue) interact very tightly through a very large interface forming a heterocomplex. At the carboxy terminus of Ku70 lies the Bax binding site (yellow).
(PDB structure 1JEQ(Walker et al., 2001) visualized with Cn3D)
60
DNA-PKcs
DNA-PKcs is a 460 kDa protein and a member of the phosphoinositol-3-OH
kinase (PI3K)-related kinase (PIKK) family which contains other proteins involved in
DNA repair such as ATM and ATR (Abraham, 2004). It is recruited by the Ku70/Ku80
heterodimer to the site of the DNA DSB. Together with Ku70 and Ku80 it promotes
synapsis of the broken DNA ends. The sheer size of DNA-PKcs warrants the belief that it
acts as a molecular scaffold for the recruitment for other NHEJ proteins with which it has
been shown to interact: XRCC4/Ligase IV (Calsou et al., 2003; Hsu et al., 2002; Leber et al., 1998), Artemis (Ma et al., 2005b; Ma et al., 2002), PNK (Koch et al., 2004), and the
X family polymerases (Ma et al., 2004) which are employed for making the free ends compatible for ligation. However, DNA-PKcs was also shown to be able to bind DNA directly, independently of Ku, which led to stimulation of its kinase activity (Yaneva et al., 1997). When activated, DNA-PKcs can phosphorylate itself as well as a series of substrates involved in DNA repair, such as Artemis and H1 histone, which is released from nucleosomes upon phosphorylation (Kysela et al., 2005). Studies have shown that
DNA-PKcs kinase activity is required for the Artemis endonuclease activity (Goodarzi et al., 2006). Furthermore, it is required for the conversion of 5’-OH ends to 5’-phosphate by polynucleotide kinase (Chappell et al., 2002). When assembled on double-stranded
DNA in vitro, the DNA-PK holoenzyme phosphorylates transcription factors and other proteins, including Sp1, Oct-1 (Schild-Poulter et al., 2007), c-fos, c-jun, p53, and the 34- kDa subunit of replication protein A (Anderson, 1993; Barnes et al., 1998). 61
DNA-PKcs was also found to regulate cell cycle arrest through down-regulation
of H2B and U2 RNA levels by phosphorylating the Oct-1 transcription factor (Schild-
Poulter et al., 2003). DNA-PKcs is also involved in the signaling pathway following
DNA damage by regulating p53-induced apoptosis (Wang et al., 2000; Woo et al., 2002).
Furthermore, it has been implicated in the regulation of NF-kB (Basu et al., 1998) and
H2AX (Burma et al., 2001) following IR. Moreover, DNA-PKcs was shown to undergo
proteolytic cleavage in cells undergoing apoptosis as well as during infection with the
poliovirus (Casciola-Rosen et al., 1995; Graham et al., 2004). This appeared to be
independent of caspase activity but directly dependent on picornaviral 2A protease.
Upon binding to DNA, DNA-PKcs undergoes autophosphorylation, which results
in loss of its kinase activity and disassembly from the DNA repair complex (Chan and
Lees-Miller, 1996). When DNA-PKcs cannot undergo autophosphorylation due to
mutation of specific amino-acid residues residing between position 2609 and 2647, cells
manifest marked radiosensitivity and V(D)J coding joints show little end processing. This is confirmed by several findings which demonstrate that efficient ligation by
XRCC4/Ligase IV requires previous DNA-PKcs autophosphorylation (Block et al., 2004;
Reddy et al., 2004; Weterings et al., 2003). Several reports made a link between the
NHEJ pathway and cellular redox stress. It was shown that cellular oxidative stress would
result in decreased DNA-PKcs activity (Boldogh et al., 2003). This impacts the
autophosphorylation capability of DNA-PKcs (Bacsi et al., 2005), which is required for
its activation and role in NHEJ (Chen et al., 2005a; Reddy et al., 2004). Interestingly,
cellular deficiency in DNA-PKcs autophosphorylation leads to an even higher 62
radiosensitivity than the absence of DNA-PKcs altogether. Apparently, this is due to the
fact that DNA-PKcs restricts the access of HR proteins to the DNA damage site (Convery et al., 2005). This suggests that the kinase disassembly from the DSB site would be as important for the completion of DNA repair as for initiating the repair process (Chan and
Lees-Miller, 1996; Douglas et al., 2001; Merkle et al., 2002). Furthermore, consistent with the belief that NHEJ is active mainly in the G0 and G1 phases of the cell cycle, while HR predominates in S and G2, several studies have shown that DNA-PKcs autophosphorylation occurs primarily in G0 and G1, thus limiting the involvement of HR when the sister chromatid is not present (Chen et al., 2005a).
Artemis
DNA DSBs caused by ionizing radiation gene have “dirty”, unligatable ends.
These aberrant structures, such as the 3’-phosphate or 3’-phosphoglycolate groups must be removed prior to the ligation event mediated by Ligase IV. This resection is performed by Artemis, a protein with endonuclease activity (Ma et al., 2005b). Its activity was shown to be stimulated by phosphorylation by DNA-PKcs (Goodarzi et al., 2006).
XRCC4
XRCC4 is another important protein in the NHEJ pathway. This protein was found in complex with Ligase IV (Grawunder et al., 1998; Modesti et al., 1999) as well as forming homodimers (Modesti et al., 1999). XRCC4 is constitutively phosphorylated.
However, following treatment with ionizing radiation is further phosphorylated by DNA- 63
PKcs (Matsumoto et al., 2000; Yu et al., 2003b). A recent study showed that XRCC4 is also subject of sumoylation, a process that regulates its nuclear localization (Yurchenko et al., 2006). Furthermore, XRCC4 was shown to be mono-ubiquitinated (Foster et al.,
2006), but the exact role of this process has not yet been defined. Even though not completely understood, it appears that it plays a regulatory role on the repair complex by stabilizing Ligase IV (Bryans et al., 1999) as well as by stimulating its activity
(Grawunder et al., 1997).
Ligase IV
Ligase IV is the essential enzyme that performs the last step in the NHEJ pathway. The ligation takes place in three steps involving an adenylate intermediary
(Tomkinson et al., 2006). Ligase IV and XRCC4 knock-out mice are embryonic lethal and this has been associated with massive neuronal cell death (Frank et al., 2000; Gao et al., 1998; Pan et al., 1994). Ligase IV function is specifically required for the NHEJ pathway of DSB repair and is completely dependent on Ku. In mice deficient in Ligase
IV or XRCC4, cells harboring Ku-bound DSBs eventually die, because these lesions cannot be repaired by HR. In contrast, in the absence of Ku, HR can partially substitute for the lost NHEJ function, resulting in the less severe phenotype observed in the Ku deficient mice (Adachi et al., 2001). Of the DNA repair complex proteins, Ku70, Ku80,
XRCC4, and Ligase IV are conserved throughout evolution in all eukaryotic species known, whereas DNA-PKcs and Artemis have not been found in lower eukaryotes such as yeast (Critchlow and Jackson, 1998). 64
XLF
A new factor with an important role in the NHEJ has recently been identified
(Ahnesorg et al., 2006; Buck et al., 2006). It was named Cernunnos or XLF (XRCC4-like factor) as it showed very high structural similarity with XRCC4, despite a low sequence homology. Several mutants of XLF have been shown to cause immunodeficiency with microcephaly (Buck et al., 2006). XLF was found to form homodimers (Ahnesorg et al.,
2006) which interact with XRCC4 through their globular head domains (Deshpande and
Wilson, 2007). Many reports showed a complex comprised of XLF, XRCC4 and Ligase
IV, but apart from the role of Ligase IV in the final step of repair, much needs to be done to ascertain the biochemical functions of these proteins. It appears that XLF stimulates the ability of XRCC4/Ligase IV to join incompatible ends (Gu et al., 2007; Tsai et al.,
2007). Its newly discovered crystal structure (Li et al., 2008) confirms that XLF forms homodimers through a coiled-coil region in a similar manner to that of XRRC4. This confirms the similarity in general structure with XRCC4 but points out important structural differences indicating distinct functions in the DNA repair process. It further casts doubt upon the interaction between XLF and Ligase IV and suggests that this might take place indirectly through XRCC4, hypothesizing that the stoichiometry of the
XRCC4/XLF/Ligase IV complex is 2:2:1 (Figure 1.9).
65
Figure 1.9 Schematic representation of the XLF/XRCC4/Ligase IV heterocomplex
66
Non-Homologous End Joining
As soon as DSBs are generated, the free DNA ends are bound by Ku70/Ku80
heterodimers (Lieber et al., 2003) (Figure 1.10). This interaction is facilitated by the
doughnut-like structure of the complex (Figure 1.8). The DNA is encircled by the
Ku70/Ku80 complex ensuring a tight binding. This structure functions as a scaffold for
the subsequent recruitment of other DNA repair proteins (Ma et al., 2005a; Ma et al.,
2004). DNA-PKcs is being recruited by the Ku heterocomplex and is activated upon
binding of the DNA end (West et al., 1998). This 460-kDa serine/threonine kinase
phosphorylates and recruits other important proteins for the repair of the lesion. One of
its substrates is the endonuclease Artemis. Another protein recruited at the DSB site is a polymerase, specifically µ or λ (Ma et al., 2004; Mahajan et al., 2002) via its BRCT domains. This may perform some of the fill-in synthesis (Wilson and Lieber, 1999). The ligation of the free DNA ends is performed by Ligase IV, which is recruited as part of the
XLF/XRCC4/Ligase IV heterocomplex (Chen et al., 2000a; Nick McElhinny et al.,
2000). XLF/XRCC4/Ligase IV was shown to be recruited directly by the Ku70/Ku80 heterodimer. However, this interaction is facilitated in the presence of DNA-PKcs
(Costantini et al., 2007). The exact physiological role of XLF in this complex still needs to be elucidated.
Other DNA double-strand break repair pathways
Interestingly, cells that harbor mutations in the NHEJ pathway even though they
show a marked reduction in the DNA repair ability, still rejoin the majority of their DSBs 67
albeit by a slow operating process (DiBiase et al., 2000; Nevaldine et al., 1997; Wang et al., 2001a). Additionally, this process is completely independent of HR (Wang et al.,
2001a). This acts as an alternative back-up NHEJ pathway to the classical one involving
Ku, DNA-PKcs, Artemis, XRCC4, and Ligase IV. The essential proteins taking part in this alternate pathway are PARP-1 and DNA Ligase III (Audebert et al., 2004; Wang et al., 2006), however much remains to be characterized in this pathway.
68
Figure 1.10 Schematic representation of the NHEJ pathway. DNA DSBs caused by
IR or chemotherapeutic agents are repaired preferentially by NHEJ. The first step in the repair pathway is the binding of the Ku70/Ku80 heterodimer to the free DNA ends. This is followed by the recruitment of the serine/threonine kinase DNA-PKcs. This, in turn, phosphorylates and recruits Artemis, an endonuclease, and µ or λ polymerases, whose
roles are to trim or fill in the DNA overhangs in order to make them compatible for
ligation. The final step in the pathway is the recruitment of the XLF/XRCC4/Ligase IV
heterocomplex. Ligase IV is responsible for the ligation of the DNA ends while XLF and
XRCC4 appear to have regulatory functions.
69
70
Generation of p18-Cyclin E and its role in modulating cellular response to genotoxic
stress
Another Cdk2-independent role of Cyclin E was identified in the regulation of
programmed cell death. We have previously shown that Cyclin E is regulated by genotoxic stress caused by ionizing radiation or other chemotherapeutic drugs and that
this plays a functional role in the apoptosis of hematopoietic cells (Mazumder et al.,
2000). We observed a surge in Cyclin E levels during genotoxic stress, which is followed
by caspase activation and membrane display of phosphatidylserine, all hallmarks of
apoptosis. We have further reported the cleavage of the full length 50 kDa Cyclin E to
generate an 18 kDa fragment which was identified as the carboxy terminus of the Cyclin
E (Mazumder et al., 2002). This fragment, termed p18-Cyclin E, lacks the Cdk2 binding
domain and therefore, fails to activate Cdk2 (Figure 1.11).
Overexpression of p18-Cyclin E leads to onset of apoptosis in a caspase-
dependent fashion. We identified caspase-3 as the responsible caspase for the cleavage of
Cyclin E and generation of p18-Cyclin E. Overexpression of the p50-Cyclin E as well as
of a cleavage resistant mutant of Cyclin E, which was generated by mutating the cleavage
site, did not result in the activation of apoptosis. However, overexpression of the cleavage
resistant mutant of Cyclin E not only prevented the induction of apoptosis, but also the
generation of p18-Cyclin E, thus acting in a dominant negative manner (Mazumder et al.,
2002). Taken together, these data suggest that in addition to the cell cycle regulatory
function, Cyclin E plays another important role in the mechanism of apoptosis. 71
Figure 1.11 Schematic representation of Cyclin E domains and cleavage sites. Cyclin
E was found to be cleaved by calpain resulting in low molecular weight (LMW)
hyperactive forms. On the other hand, Cyclin E was also found to be cleaved by elastase
resulting in a truncated form (81-363) whose three-dimensional structure has been solved.
Based on this structure, the interface between Cyclin E and Cdk2 was defined as amino-
acids 94-112, 182-256, and 340-344. Through analogy with the interaction between
Cyclin A and p27, it was suggested that 129-136 and 170-173 are the residues responsible for the binding and inhibition of Cyclin E by p27. The 129-215 sequence illustrated in blue represents the Cyclin box, a conserved domain among cyclins. p18-
Cyclin E is generated following a Caspase-3 mediated cleavage at the Asp275 residue and thus, lacks the Cdk2 binding domain failing to bind and activate it. Human Cyclin
E1, accession number AAM54043 (NM_057182), was used as a reference.
72
Later on, using a yeast-two-hybrid assay, we identified several potential p18-
Cyclin E-interacting proteins (Mazumder et al., 2007b). Among all candidates, Ku70 stood out through its roles in Non-Homologous End-Joining DNA repair (Lieber et al.,
2003) as well as its cytoprotective function in the cytosol (Cohen et al., 2004;
Subramanian et al., 2005). As a result of this interaction, we witnessed a release of Bax from its cytoprotective sequestration to Ku70 in the cytosol and its activation, which could be responsible for the observed onset of apoptosis following generation of p18-
Cyclin E (Mazumder et al., 2007a; Mazumder et al., 2007b).
In this dissertation I will discuss a novel, important role of p18-Cyclin E in the regulation of NHEJ DNA repair pathway as well as newly uncovered regulatory mechanisms of p18-Cyclin E turnover and their implications in clinical therapeutics.
CHAPTER II
p18-Cyclin E regulates Non-Homologous End Joining by preventing the recruitment
of the XLF/XRCC4/Ligase IV heterocomplex to the DNA repair complex
Abstract
Cyclin E/Cdk2 is a critical regulator of cell cycle progression from G1 to S phase in mammalian cells that is deregulated in many types of neoplasms. We found that genotoxic stress leads to a dramatic decrease in Cyclin E levels in hematopoietic tumor cells, coinciding with the timely appearance of its proteolytic fragment, p18-Cyclin E.
Overexpression of p18-Cyclin E in a variety of cell types induces apoptosis. We have identified Ku70, a critical component of non-homologous end joining (NHEJ) DNA repair as a new interacting partner of p18-Cyclin E. Cells stably expressing p18-Cyclin E at non-toxic levels are more sensitive to etoposide or ionizing radiation treatment. Neutral comet assays used to determine residual double-strand breaks (DSBs) indicative of an ineffective NHEJ, showed significant increase in both tail length and tail moment in the presence of p18-Cyclin E following irradiation. Moreover, a plasmid reactivation assay indicated that p18-Cyclin E reduced the colony formation ability known to be associated with NHEJ activity as compared to cell lysates containing wild-type Cyclin E. Gel electrophoretic analyses indicated an impairment of end-ligation dependent on the expression levels of p18-Cyclin E. DNA pull-down assays showed that the assembly of
73 74
Ku70/Ku80 and DNA-PKcs is not affected by the presence of p18-Cyclin E. However, the recruitment of XRCC4, Ligase IV, and the recently identified accessory factor XLF was greatly impaired. These data indicate a profound effect of p18-Cyclin E on cellular survival and on NHEJ that is dependent on its interaction with Ku70 and probably caused by interference with the recruitment of the XLF/XRCC4/Ligase IV heterocomplex to the sites of DSBs. Interaction of Ku70 with p18-Cyclin E and Bax may be unique in providing the molecular switch between cell cycle control, DNA repair, and apoptosis activation for the cellular decisions made following genotoxic stress.
Introduction
The integrity of the genome is a critical requirement for propagation of life.
Damage to cellular DNA is involved in mutagenesis and development of cancer. Among the different types of DNA damage encountered daily by a human cell, the DSBs are the most dangerous for the survival of the cell, making their repair a necessity (Hoeijmakers,
2001; Jackson, 2001; Shiloh, 2003; Valerie and Povirk, 2003). Double strand breaks are naturally produced as a consequence of ionizing radiation (IR), DNA replication over a nicked template or as a result of enzymatic cleavage such as by RAG1 and RAG2 endonucleases during V(D)J recombination. The emergence of DNA DSBs caused by IR or other chemotherapeutic drugs leads to activation of a signal transduction cascade resulting in cell cycle arrest, assembly of a DNA repair complex, and changes in gene transcription. Inability to efficiently repair these DSBs causes genomic instability as shown by the presence of chromosome fusions (Khanna and Jackson, 2001). DSBs are 75
potentially catastrophic lesions which, if not repaired, would lead to loss of genetic
information, genomic instability or cell death.
Depending on the cell cycle phase, this type of lesion may be repaired by
homologous recombination (HR) or NHEJ. The general belief is that HR is employed in
late S and G2 cell cycle phases when the sister chromatid is available and may be used as
a template while NHEJ is used during the rest of the cell cycle. For this reason, NHEJ is considered as the most commonly used repair mechanism for the repair of DSBs. HR requires extensive sequence homology so that one DNA strand could invade a homologous sequence and initiate the repair (Paques and Haber, 1999). On the other hand, NHEJ rejoins the broken DNA with little or no sequence homology but it requires processing of the ends prior to ligation, a process during which some nucleotides are
deleted or inserted (Jeggo, 1998). While HR takes place with high accuracy as one
chromatid serves as template, NHEJ is largely an imprecise DNA repair process. Mice
that are deficient in components of the NHEJ machinery show radio hypersensitivity,
immunodeficiency, and are prone to tumor formation (Ferguson and Alt, 2001; van Gent
et al., 2001).
The NHEJ machinery relies on the formation of a large DNA repair complex
encompassing the Ku70/Ku80 heterodimer, DNA-PKcs, and XLF/XRCC4/Ligase IV.
The absence of any of these proteins confers increased radiation hypersensitivity as well
as immune deficiency due to impaired V(D)J recombination (Weterings and van Gent,
2004). Ku70/Ku80 shows very high affinity for free DNA ends and binds them in order
to keep them in close proximity (DeFazio et al., 2002). Next, the Ku70/Ku80 heterodimer 76
recruits DNA-PKcs, a serine/threonine kinase which phosphorylates many substrates
including itself (Smith and Jackson, 1999). These substrates include the nuclease Artemis
(Ma et al., 2002), that is responsible for the resection of 5’ and 3’ overhangs and DNA polymerases µ and λ (Bebenek et al., 2003; Capp et al., 2006; Mahajan et al., 2002;
Zhang et al., 2001), which perform some of the fill-in synthesis. The final step of ligation is performed by Ligase IV, whose activity is regulated by two proteins with which it is found in complex: XRCC4 (Critchlow et al., 1997; Grawunder et al., 1997) and
XLF/Cernnunos (Ahnesorg et al., 2006; Buck et al., 2006). Given the important role of
NHEJ in DNA DSB repair and cell survival, inhibiting this pathway represents an attractive opportunity to increase cell sensitivity to IR or chemotherapy (He et al., 2007;
Li et al., 2003; Marangoni et al., 2000).
Cyclin-dependent kinase 2 (Cdk2) and its regulatory factor Cyclin E form the key regulatory complex of the G1 to S phase cell cycle progression. Cyclin E has been shown to be deregulated and overexpressed in several solid tumors, including breast, colon, and prostate carcinomas (Akli and Keyomarsi, 2003; Erlandsson et al., 2003; Keyomarsi et al., 1995; Keyomarsi et al., 1994; Keyomarsi et al., 2002; Moroy and Geisen, 2004;
Sherr, 1996). High levels of Cyclin E expression have been associated with the progression of different other tumors, such as leukemias and lymphomas (Erlanson and
Landberg, 2001; Erlanson et al., 1998).
We have previously shown that Cyclin E is critical for genotoxic stress-induced apoptosis of tumor cells of hematopoietic origin (Mazumder et al., 2000). An 18-kDa
Cyclin E fragment (p18-Cyclin E) is generated through caspase-mediated proteolytic 77
cleavage of Cyclin E (Mazumder et al., 2002) in hematopoietic cells undergoing
apoptosis. Recently, we have identified Ku70 as a specific p18-Cyclin E-interacting
protein which mediates p18-Cyclin E-induced apoptosis (Mazumder et al., 2007a;
Mazumder et al., 2007b). Apart from its role in the NHEJ, Ku70 also plays a
cytoprotective role by binding and sequestering Bax in the cytosol (Cohen et al., 2004;
Subramanian et al., 2005). Mapping the p18-Cyclin E interaction domain of Ku70 using a
series of deletion mutants of Ku70, revealed that their interface resides at the N-terminus
of Ku70, which is different from the Bax binding site located at the C-terminus.
Furthermore, we have shown that binding of p18-Cyclin E to Ku70 results in Bax release
leading to amplification of the apoptotic signal. However, the presence of p18-Cyclin
both in the cytosol and in the nucleus, taken together with the important role of Ku70 in
NHEJ DNA repair, led to the hypothesis that p18-Cyclin E may also be involved in
regulation of DNA repair during cell death induced by genotoxic stress.
In our study we observed that cells that stably express p18-Cyclin E at non-toxic levels are more sensitive to DNA damaging agents, such as ionizing radiation and etoposide. Furthermore, p18-Cyclin E impairs DNA repair in vitro as shown by the failure to re-ligate linear plasmid DNA. Moreover, a plasmid reactivation assay showed that the presence of p18-Cyclin E led to a dramatic reduction in colony formation. Tail length and tail moment of unrepaired DSBs determined by neutral comet assays, indicated that the DNA DSBs repair is less efficient in cells expressing p18-Cyclin E as compared to control cells. Mechanistically, DNA pull-down assays which recapitulate the
DNA repair complex assembly showed that the DNA binding of Ku70/Ku80 and the 78
subsequent DNA-PKcs association are not affected by p18-Cyclin E. Conversely, the recruitment of XLF/XRCC4/Ligase IV to the DNA repair complex is severely reduced in the presence of p18-Cyclin E.
Here we report a novel role of the C-terminal fragment of Cyclin E (p18-Cyclin
E) in regulating the NHEJ DNA repair pathway in cells undergoing genotoxic stress. This may be critical in hematopoietic tumor cells that are known to have high NHEJ activity that may be activated by the DSBs produced by apoptotic nucleases. Blocking NHEJ could prevent, for example translocations leading to further tumor progression in these cells. Thus, p18-Cyclin E upon being generated may on one hand trigger the release of
Bax from its complex with Ku70 in the cytosol and on the other hand prevent DNA repair by NHEJ in the nucleus. This way, p18-Cyclin E may play an important role in the cellular decision making process between cellular survival and apoptosis.
Materials and methods
Cell lines and treatments. HEK293T human kidney cells were maintained in Dulbecco modified Eagle medium (Gibco-BRL) containing 10% fetal bovine serum (Hyclone,
Logan, UT) with L-glutamine and 100 units/ml penicillin/streptomycin at 37°C and in an atmosphere containing 5% CO2. Cells ~75% confluent were either treated with etoposide
(VP16) or irinotecan (CPT-11) from Sigma (St. Louis, MO), or irradiated with 10 Gy
(137Cs source; fixed dose rate of 2.0 Gy/min). HA-tagged p18-Cyclin E stably expressing 79
293T cells were generated by infecting cells with a lentivirus carrying HA-p18-Cyclin E
and EGFP separated by an IRES2 sequence and under the control of the EF1α promoter.
Antibodies. The following primary antibodies were used: anti-cyclin E (HE-12), anti-
Ku70 (E-5), anti-Ku80 (B-1) from Santa Cruz Biotechnology (Santa Cruz, CA), anti-β- actin from Sigma, anti-HA (HA.11) from Covance (Berkeley, CA), anti-PARP-1 from
Cell Signaling (Beverly, MA), anti-DNA-PKcs (Ab-1, 18-2) from Lab Vision (Fremont,
CA), anti-XLF (A300-730A) from Bethyl Laboratories (Montgomery, TX), anti-XRCC4
(AHP387) and anti-Ligase IV (AHP554) from AbD Serotec (Raleigh, NC).
Immunoblot analyses. Following irradiation or treatment with VP16, cells were lysed at
4°C for 30 min in buffer A (20 mM HEPES pH 7.5, 150 mM NaCl, 1 mM EDTA, 1%
NP-40, containing 1 mM phenylmethylsulfonyl fluoride (PMSF), 1 µg/mL pepstatin A, aprotinin, and leupeptin. Proteins (10-75 µg/lane) were resolved by SDS-PAGE, followed
by transfer to 0.1 µm nitrocellulose membrane and blocking in 5% nonfat dry milk for 1 h
at room temperature. The blots were then incubated with the appropriate primary
antibody in phosphate-buffered saline with 0.1% Tween 20 (PBST) containing 5% nonfat
dry milk for 16 h at 4°C followed by horseradish peroxidase (HRP)-conjugated
secondary antibodies.
Caspase-3 activation assay. Treated cells were lysed in lysis buffer containing 1% NP-
40, 20 mM HEPES (pH 7.5), 4 mM EDTA and aprotinin (10 µg/ml), leupeptin (10 80
µg/ml), pepstatin (10 µg/ml), and PMSF (1 mM). Equal amounts of protein lysates were taken in a microtiter plate to 200 µL with reaction buffer containing 100 mM HEPES (pH
7.5), 20% v/v glycerol, 5 mM dithiothreitol (DTT), 0.5 mM EDTA and caspase-3 substrate (Ac-DEVD-pNA; Calbiochem EMD, Gibbstown, NJ) was added to 100 µM.
The samples were incubated at 37°C for 1-2 h and the enzyme-catalyzed release of p-NA was determined by measuring the absorbance spectrophotometrically at 405 nM.
Comet assay (single-cell gel electrophoresis). HEK-293T cells were transiently transfected with p18-Cyclin E-HA using Lipofectamine 2000 (Invitrogen, Carlsbad, CA) according to the manufacturer’s indications. 24 h post-transfection cells were irradiated with 10 Gy and collected after 16 h. To evaluate the degree of DNA damage, we used
CometAssay Silver (Trevigen, Gaithersburg, MD), which combines a neutral lysis in low point melting agarose with silver staining or SYBR Green for visualization of the DNA.
Lysis and electrophoresis were performed according to the manufacturer’s protocol.
Image analysis and quantification have been performed with the software package NIH
ImageJ. Tail Moment (TM) and Tail Length (TL) were used to quantify the DNA damage. TM = % of DNA in the tail x TL; where % of DNA in the tail = tail area (TA) x tail average intensity (TAI) x 100/(TA x TAI) + [head area (HA) x head area intensity
(HAI)].
DNA-agarose pull-down. Nuclear extracts of HEK293T stably expressing HA-tagged p18-Cyclin E were prepared by hypotonic swelling lysis in 20 mM Tris-HCl pH 8.0, and 81
1 mM DTT. After incubation on ice for 30 min, nuclei were collected by centrifugation at
8000xg for 5 min and extracted in 20 mM Tris-HCl, pH 8.0, 500 mM NaCl, 5 mM
o MgCl2, 10% glycerol, and 1 mM DTT for 30 min, gently rocking at 4 C. All buffers contained 1x HALT phosphatase inhibitor cocktail (Pierce, Rockford, IL), 1µg/mL aprotinin, leupeptin, pepstatin, and 1 mM phenylmethylsulfonyl fluoride. Extracts were clarified by centrifugation at 16.000xg for 10 min and diluted (1:3) with 20 mM Tris-HCl pH 8.0, 5 mM MgCl2, 10% glycerol, 1 mM DTT. A slurry (50 µL) of denatured calf
thymus DNA-agarose beads (Amersham, Piscataway, NJ), were washed three times, added to the extracts and incubated at 4oC with rotation overnight. Beads were washed 3 times with 1 mL of buffer and the proteins eluted with 500 mM KCl and separated by
SDS-PAGE.
DNA end-ligation and plasmid reactivation assay. 1 µg of EcoR1-digested pUC18
DNA (DSB) was incubated with nuclear extracts of p18-Cyclin E and Cyclin E transfected 293T cells or control (GFP-expressing) and p18-Cyclin E stably expressing
293T cells, in reaction buffer X (40 mM Tris-HCl (pH 7.5), 10 mM MgCl2, 50 µM dNTPs, 2 mM ATP, 1 mM DTT and 100 µg/ml BSA). The end ligation mixture was incubated at 37oC for 1 h. The product was separated and analyzed by electrophoresis on
0.6% agarose gels together with monomer pUC18, used as a negative control and monomer pUC18 ligated with T4 DNA ligase, used as a positive control. For the plasmid reactivation assay, EcoRI-linearized pUC18 DNA was incubated with nuclear extract of transfected 293T cells in reaction buffer X at 37oC for 1 h. DNA was purified, 82
transformed in DH5α E.coli and transformed cells were plated on LB with Amp, X-gal, and IPTG.
Results
Non-toxic levels of p18-Cyclin E sensitize cells to VP-16 and IR treatment
Our previous experiments have shown that in hematopoietic cells subjected to
genotoxic stress, Cyclin E undergoes a caspase-3-mediated cleavage to generate an 18-
kDa carboxy-terminal fragment termed p18-Cyclin E (Mazumder et al., 2002). We have
also demonstrated that the pro-apoptotic effect of this protein is taking place in the
cytosol and it is mediated by its interaction with Ku70 through release and activation of
Bax (Mazumder et al., 2007b). To study the effect p18-Cyclin E may have on NHEJ in
the nucleus in the absence of its action in the cytosol, we have generated an HA-p18-
Cyclin E stably expressing 293T cell line using a lentiviral approach (Figure 2.1A).
These cells express p18-Cyclin E at moderate non-toxic levels and are viable and
thriving. Furthermore, as Cyclin E cleavage does not take place, therefore these cells are
ideally suited to study the regulation and role of p18-Cyclin E.
To determine the effect of p18-Cyclin E on cell viability under these experimental
conditions, we treated control cells (GFP-expressing) or p18-Cyclin E stably expressing
293T cells with etoposide at increasing concentrations. Increased PARP-1 cleavage in
p18-Cyclin E-expressing cells suggested that p18-Cyclin E may be a mediator of
etoposide-induced cell death (Figure 2.1B). Moreover, E cells stably expressing p18- 83
Cyclin were more sensitive than control (GFP-expressing only) cells to irradiation as shown by a caspase-3 activation assay. Taken together, these data suggest that p18-
Cyclin E is able to sensitize cells to DNA damaging agents and we hypothesize that this
effect could result from its interaction with Ku70 in the nucleus and subsequent
interference with NHEJ repair.
84
Figure 2.1 Non-toxic levels of p18-Cyclin E sensitize cells to VP-16 and IR treatment. A) p18-Cyclin E lentivirus expression vector. B) HEK293T cells stably expressing p18-Cyclin E or EGFP were treated with increasing doses of etoposide (VP-
16) and cell death was determined by PARP-1 cleavage [full length (fl), cleaved (cl)] .
Levels of PARP-1, p18-Cyclin E, and β-actin, used as a loading control were determined by immunoblotting. C) 293T cells stably expressing p18-Cyclin E or EGFP were irradiated with 10 Gy ionizing radiation and cell death was assessed by activation of caspase-3. (Data are representative of three independent experiments)
85
A
B
C
86
p18-Cyclin E inhibits in vitro DNA ligation and plasmid reactivation
To determine whether p18-Cyclin E is capable of interfering with DNA repair we tested this hypothesis in an in vitro DNA end-ligation study. In the presence of p18-
Cyclin E the plasmid DNA was observed only in its monomeric form. In contrast, in the presence of Cyclin E or T4 DNA ligase used as positive control, the plasmid DNA was efficiently re-ligated, as shown by the higher molecular weight concatemeric forms
(Figure 2.2A left and middle panel). The expression of p18-Cyclin E and Cyclin E was
confirmed by immunoblotting using anti-HA antibodies (Figure 2.2A right panel). This
suggests that p18-Cyclin E was able to inhibit the plasmid re-ligation as observed by the
absence of the concatemeric DNA forms. Furthermore, when we examined the ability of
the re-ligated plasmid to form colonies following transformation in E. coli, we observed a
dramatic reduction in the number of colonies resulted in the presence of p18-Cyclin E to
~10% as compared to ~60% for Cyclin E or T4 DNA ligase considered to be 100%
(Figure 2.2B). These data suggest that p18-Cyclin E is able to interfere and prevent the
DNA repair in an in vitro assay.
87
Figure 2.2 p18-Cyclin E inhibits in vitro DNA ligation and plasmid reactivation. A)
Nuclear cellular extracts from control (GFP-expressing), 293T stably expressing p18-
Cyclin E (left panel) or transfected with increasing DNA amounts of p18-Cyclin E and
Cyclin E (middle panel) were incubated with a DNA plasmid previously linearized with
the EcoRI restriction enzyme. The reaction was incubated at 37oC to allow the plasmid to be re-ligated. T4 DNA ligase was used as a positive control. The DNA ligation reaction was separated by gel electrophoresis (left and middle panel). Expression of the transfected constructs was determined by immunoblotting with anti-HA antibody (right panel). The second and third lanes of the left panel represent identical reactions. B) The
DNA ligation reaction mix was transformed in E. coli that were then plated on agar
containing X-Gal and IPTG. The colonies were counted and the number was expressed as
percentage of the T4 DNA ligase treated colonies, that was considered to be 100%.
88
89
p18-Cyclin E prevents efficient repair of the DNA damage induced by IR or CPT-11
To determine whether p18-Cyclin E affects DNA repair in vivo, we examined the
ability of human cells to repair the DNA damage induced by either IR or irinotecan
(CPT-11, inhibitor of topoisomerase I) in the presence or absence of p18-Cyclin E. The
treated cells were subjected to neutral comet assay (single-cell gel electrophoresis). The residual, unrepaired DSBs were determined by quantification of the tail lengths and
moments of the resulting comets (Figures 2.3A and 2.3B upper panels). We found that p18-Cyclin E impairs the repair of DNA damage induced by IR (Figure 2.3A lower panel) or CPT-11 (Figure 2.3B lower panel). Taken together, these data indicate that p18-
Cyclin E interferes with the DNA repair in both in vitro and in vivo experiments.
90
Figure 2.3 p18-Cyclin E prevents efficient repair of the DNA damage induced by IR
and CPT-11. A) 293T cells were transfected with HA-p18-Cyclin E (p18). At 24 h post-
transfection cells were irradiated (10 Gy) and collected after 16 h. Lysis and
electrophoresis were performed according to the manufacturer’s indications (upper
panel). After the slides were silver stained, the image analysis and quantification was
performed with NIH ImageJ. Tail length (lower left panel) and tail moment (lower right panel) were determined and calculated according to the following formula: Tail moment
= % of DNA in the tail x Tail length; where % of DNA in the tail = tail area (TA) x tail average intensity (TAI) x 100/(TA x TAI) + [head area (HA) x head area intensity
(HAI)]. B) 293T cells were transfected with HA-p18-Cyclin E. At 24 h post-transfection,
cells were treated with 100 ng/mL CPT-11 and collected after 16 h. Neutral comet assay
image analysis was performed as described above. DNA was stained in this case with
SYBR Green. (Data are representative of three independent experiments)
91
92
93
p18-Cyclin E impairs the recruitment of XLF/XRCC4/Ligase IV heterocomplex to
the DNA repair complex
We have shown previously that p18-Cyclin E interacts specifically with Ku70, a protein very well studied for its role in the NHEJ DNA repair pathway (Mazumder et al.,
2007b). Upon appearance of DNA DSBs, Ku70/Ku80 heterodimers bind the free DNA ends to keep them in close proximity and recruit other important DNA repair factors such as DNA-PKcs, Artemis, several polymerases, XRCC4, XLF, and Ligase IV. The interaction between p18-Cyclin E and Ku70, together with the demonstrated ability of p18-Cyclin E to impair the DNA repair, prompted the question whether p18-Cyclin E inhibits the NHEJ machinery by interfering with the Ku70-DNA binding step or with the subsequent recruitment of DNA repair factors. To determine at which step in the NHEJ pathway p18-Cyclin E acts we reproduced the NHEJ repair in nuclear extracts from cells expressing p18-Cyclin E or only GFP as control. The DNA DSBs were mimicked by denatured calf thymus DNA fragments crosslinked to agarose beads. The DNA repair
complexes were allowed to form at the ends of the DNA fragments, then eluted with high
saline buffer and separated by SDS-PAGE. Interestingly, the binding of the Ku70/Ku80
heterodimer to the DNA fragments was not affected at all. Similarly, p18-Cyclin E did
not impact the recruitment of DNA-PKcs to DSBs. In contrast, the recruitment of the
XLF/XRCC4/Ligase IV heterocomplex was dramatically reduced in presence of p18-
Cyclin E (Figure 2.4 upper panel) suggesting its interference with the last step in NHEJ,
the ligation of the DNA ends. Moreover, p18-Cyclin E was present at the sites of DSBs
(Figure 2.4 lower panel). 94
Figure 2.4 p18-Cyclin E impairs the recruitment of XLF, XRCC4, and Ligase IV to the DNA repair complex. Nuclear extracts of HEK293T cells stably expressing p18-
Cyclin E or EGFP were prepared by hypotonic swelling lysis. Extracts were incubated with denatured calf thymus DNA-agarose beads overnight. Beads were washed and the proteins eluted with a high salt buffer and separated by SDS-PAGE. The levels of the recruited proteins were determined with the respective antibodies: DNA-PKcs, Ligase IV,
Ku80, Ku70, XRCC4, and XLF (upper panel). A higher percentage gel was used to determine the presence of p18-Cyclin E, as well as that of Ku70 and Ku80 as controls
(lower panel). (Data are representative of three independent experiments)
95
Discussion
Ionizing radiation and chemotherapeutic treatment are responsible for the induction of several types of DNA damage including base damage, single strand breaks and DSBs (Collis and DeWeese, 2004; Hoeijmakers, 2001; Petrini and Stracker, 2003).
Among these, the DSBs are the most toxic ones, their presence being able to lead to either cell death or chromosomal translocation, which is a major cause for malignant transformation (van Gent et al., 2001). Therefore, their accurate repair is essential for cell survival. In eukaryotic cells, DNA DSBs can be repaired by two major pathways, homologous recombination (HR) (Haber, 1995) or non-homologous end joining (NHEJ)
(Lieber et al., 2003). HR performs the repair with very high accuracy since it employs the sister chromatid as template for the DNA synthesis. For this reason, HR takes place mainly in late S and G2 phases of the cell cycle. On the other hand, NHEJ which is the most common pathway for DSB repair in mammalian cells is very imprecise and may lead to mutations in genome. The purpose of treating tumor cells with DNA-damaging agents such as IR or chemotherapeutics is to induce enough genotoxic stress that would lead to cell death. Attempts to replicate damaged DNA would cause increased cell death, making tumor cells with high replication rates more sensitive to this type of therapy.
However, the toxicity of the DNA-damaging drugs may be reduced in cells that manifest very high levels of DNA repair activity, thus leading to inefficacy of the treatment and survival of the tumor cells. Furthermore, as some of the DNA repair pathways are inactivated in many types of cancers, the DNA repair mechanisms are an attractive target for cancer therapy. 96
Cyclins in association with their catalytic subunits, the cyclin-dependent kinases play an important regulatory role in the cell cycle progression (Murray, 2004). Cyclin
E/Cdk2 complex has an essential role at the rate-limiting step in the G1 to S phase
transition (Dulic et al., 1992; Koff et al., 1991). Loss of its regulation has been shown to lead to aberrant cellular division and malignant transformation (Ohtsubo and Roberts,
1993; Ohtsubo et al., 1995; Resnitzky et al., 1994). We have reported that in cells of hematopoietic origin, genotoxic stress induced by IR leads to caspase-3-mediated proteolytical cleavage of Cyclin E resulting in generation of its C-terminal proteolytic fragment, p18-Cyclin E (Mazumder et al., 2002). Unlike Cyclin E, p18-Cyclin E lacks the Cdk2 binding domain and therefore cannot interact with Cdk2 suggesting that its functions are independent of Cdk2 and different from those of Cyclin E. Furthermore, we have recently demonstrated that by binding to Ku70 in the cytosol, p18-Cyclin E causes the dissociation of Bax from Ku70 and its activation, thus leading to apoptosis
(Mazumder et al., 2007a; Mazumder et al., 2007b).
The nuclear abundance of Ku70 as well as the cytosolic and nuclear presence of
p18-Cyclin E warranted the investigation of its involvement in NHEJ DNA repair. This
hypothesis was also supported by the finding that stable, non-toxic levels of p18-Cyclin E
were able to sensitize cells to different DNA-damaging agents such as IR or etoposide.
The fact that no apoptosis was observed in untreated control cells suggests that p18-
Cyclin E was expressed at levels below those that would trigger release of Bax in the
cytosol. As p18-Cyclin E and Bax expression levels are quite low compared to those of
Ku70, stoichiometrically, it is less likely that a p18-Cyclin E molecule would meet and 97
interact with a Ku70/Bax complex to result in the release of Bax. This stable cell line allowed us to study the effect of p18-Cyclin E strictly on DNA repair in the nucleus. The observed sensitivity of p18-Cyclin E-expressing cells to genotoxic stress suggested that this may be a result of unrepaired DNA damage which caused the cells to undergo apoptosis.
To test this hypothesis, we next examined the effect of p18-Cyclin E on in vitro plasmid DNA ligation. Increasing amounts of p18-Cyclin E in nuclear extracts effectively inhibited the ligation reaction as shown by the presence of DNA only in linear, monomeric form. On the other hand, Cyclin E did not interfere with the end ligation and allowed the reaction to take place to a similar extent as that catalyzed by T4 DNA ligase.
This indicated that the presence of p18-Cyclin E could account for the inability of DNA repair enzymes in the nuclear extracts to perform the ligation. This conclusion was further confirmed by the extensive reduction in number of colonies resulted from transformation of the DNA in E.coli. The capability of p18-Cyclin E to prevent the repair of DNA damage induced by IR or CPT-11 was also indicated by the increased tail length and moment measured in a neutral comet assay. Taken together, these data clearly prove the involvement of p18-Cyclin E in the regulation of DNA repair and presumably of
NHEJ as suggested by its interaction with Ku70.
The NHEJ DNA repair pathway relies on the assembly of a large protein complex at the DNA DSB. This process starts with the binding of the Ku70/Ku80 heterodimer to the free DNA ends (DeFazio et al., 2002) followed by the recruitment of the large molecular weight DNA-PKcs (Smith and Jackson, 1999). This kinase, in turn, 98
phosphorylates and recruits other important repair enzymes such as Artemis and several
polymerases (Koch et al., 2004; Ma et al., 2002). The final step of the repair process, the
ligation, is performed by the XLF/XRCC4/Ligase IV heterocomplex (Critchlow et al.,
1997; Grawunder et al., 1997). The interaction of Ku70 with p18-Cyclin E would suggest that the most likely step in the pathway that p18-Cyclin E may interfere with, would be the association of Ku70/Ku80 with the DNA ends, a destabilization of the Ku70/Ku80 complex, or the recruitment of DNA-PKcs. However, a DNA pull-down assay which reconstitutes the DNA repair complex assembly did not show any difference in the synapsis formation by the Ku70, Ku80, and DNA-PKcs proteins. Interestingly, the recruitment of the XLF/XRCC4/Ligase IV heterocomplex was severely impaired, suggesting that this may be prevented by the binding of p18-Cyclin E to Ku70.
Several studies have shown that XRCC4/Ligase IV can be recruited to the DSB repair site even in the absence of DNA-PKcs (Costantini et al., 2007; Mari et al., 2006) and that it is Ku70/Ku80 that is indispensable for recruiting Ligase IV. Furthermore, it has been reported that Ligase IV interacts directly with Ku and that Ku70 and Ku80 have to be present in the heterodimeric form. But, even though DNA-PKcs may not be required for attracting Ligase IV to the DNA repair complex, it appears that this has a stabilizing effect on its recruitment (Calsou et al., 2003). This provides an explanation for the mechanism of how p18-Cyclin E regulates NHEJ. Binding of p18-Cyclin E to Ku70 could prevent the interaction between Ku and the XLF/XRCC4/Ligase IV complex resulting in inhibition of the ligation step (Figure 2.5). 99
The fact that Ku stimulates Ligase IV activity and that XRCC4 stabilizes the
NHEJ complex by interacting with DNA-PKcs (Calsou et al., 2003; Hsu et al., 2002;
Kysela et al., 2003) explains how diminished recruitment of XLF, XRCC4, and Ligase
IV caused by p18-Cyclin E could lead to impairment of the DNA ligation.
The ability of p18-Cyclin E to prevent the DNA repair in the nucleus and to trigger the release and activation of Bax in the cytosol are indicative of the important role played by p18-Cyclin E in the amplification of the apoptotic signal, thus committing the cells to apoptosis. In conclusion, the recently discovered interaction between p18-Cyclin
E and Ku70 uncovers a new role of Cyclin E in the regulation of cellular physiology which may be unique in linking cell cycle control, DNA repair, and activation of apoptosis following genotoxic stress as well as provide mechanistic insights into the choice between cell death and cell survival depending on the cell type and nature of the genotoxic challenge.
100
Figure 2.5 Schematic representation of the NHEJ DNA repair pathway and the way
in which p18-Cyclin E affects the assembly of repair factors. The free DNA ends of a
DSB are bound very rapidly and tightly by the Ku70/Ku80 heterocomplexes. These complexes slide inwards on the DNA and recruit the large serine/threonine kinase DNA-
PKcs which together with Ku70/Ku80 forms the synaptic complex keeping the DNA
ends in close proximity. DNA-PKcs phosphorylates and recruits Artemis (overhang
resection) and several polymerases (nucleotide fill-in) necessary for the processing of the
DNA ends in order to make them compatible for ligation. Finally, XLF/XRCC4/Ligase
IV is recruited to the DNA repair complex to perform the final step of the repair pathway,
the ligation of DNA ends. However, in the presence of p18-Cyclin E, the recruitment of
this heterocomplex is impaired and the repair of the DNA damage fails to take place,
resulting in cell death. 101
CHAPTER III
Two new mechanisms for proteasome-mediated Cyclin E degradation uncovered in
hematopoietic tumor cells undergoing apoptosis
Abstract
We have shown earlier that in hematopoietic tumor cells, caspase-mediated
cleavage of Cyclin E generates p18-Cyclin E. Its expression can induce apoptosis or
sensitize to apoptotic stimuli in many cell types by associating with Ku70, leading to dissociation of Bax from Ku70 and its activation. However, it has a much shorter half-life than Cyclin E. Here, we show that p18-Cyclin E is more effectively ubiquitinated and degraded by the 26S proteasome. Interestingly, recognition by the Skp1-Cul1-Fbw7
(SCF) complex and the interaction with the Fbw7 family of protein isoforms can take place independently of phosphorylation of p18-Cyclin E or its association with Cdk2.
However, the SCFFbw7 complex is implicated in this ubiquitination process only to a limited extent, with a novel pathway involving Ku70 and Hdm2 being more effective in the degradation of p18-Cyclin E. In addition, blocking its degradation using specific proteasome inhibitors, such as Bortezomib and MG132 can stabilize p18-Cyclin E as well as prolong its association with Ku70 and thereby, lead to enhanced apoptosis.
Moreover, cells expressing p18-Cyclin E are more sensitive to Bortezomib treatment. By preventing its proteasomal degradation, p18-Cyclin E may become an effective
102 103
therapeutic target for Bortezomib and apoptotic effectors in various hematopoietic
malignancies.
Introduction
Deregulation of cellular proliferation together with defects in apoptosis control are two critical processes responsible for tumor formation (Green and Evan, 2002).
Cyclins, in association with their catalytic subunits, the cyclin-dependent kinases
(CDKs), control cell cycle progression by regulating events that drive the transition between cell cycle phases (Elledge, 1996; Murray, 2004; Reed, 2003). The Cyclin
E/Cdk2 complex plays an essential and rate-limiting role in the transition between the G1 and S phases of the cell cycle (Ohtsubo et al., 1995; Resnitzky and Reed, 1995) and in the initiation of DNA replication (Jackson et al., 1995; Ohtsubo et al., 1995). Regulation of
Cyclin E takes places at both transcriptional and post-transcriptional level.
Transcriptionally, Cyclin E is a target of the E2F transcription factor family whereas post-transcriptionally, Cyclin E is targeted by the ubiquitin-proteasomal pathway to 26S proteasomal degradation (Clurman et al., 1996; Won and Reed, 1996). The required ubiquitination is achieved following recognition by and interaction with the E3 ubiquitin ligase, the F-box tumor suppressor protein, Fbw7 (also known as hCdc4). Fbw7 is a specificity factor for the Skp1-Cul1-F-box protein ubiquitin ligase complex that targets a variety of proto-oncogene products for ubiquitin-mediated degradation, including Cyclin
E (Zhang and Koepp, 2006). Cullin-3 (Cul3) on the other hand is responsible for the free 104
Cyclin E degradation (McEvoy et al., 2007; Singer et al., 1999). The role of Cyclin E expression in apoptosis, particularly independent of its interaction with Cdk2 is not clear.
The ubiquitin-proteasome pathway is essential for the degradation of the majority of intracellular proteins (Hershko and Ciechanover, 1998). A number of key regulatory proteins involved in cell proliferation, differentiation, survival, as well as cell death are regulated by proteasome-mediated proteolysis, that results in the activation or inhibition of specific cell signaling pathways (Adams, 2004). Apoptosis is regulated by the ubiquitin/proteasome system at different levels and importantly, inhibition of proteasome function may induce cell death (Herrmann et al., 1998). Recently, inhibition of the proteasomal function with specific inhibitors has evolved as a novel approach for the treatment of various cancers (Aghajanian et al., 2002; Chauhan et al., 2005; Papandreou et al., 2004). Highly specific inhibitors of the 26S proteasome, such as the reversible inhibitor MG132 have been shown to induce apoptosis in many tumor cell lines (Adams and Kauffman, 2004; Zhang et al., 1999). Bortezomib (PS-341, Velcade), a modified dipeptydil boronic acid is a potent and selective inhibitor of the 26S proteasome, a multisubunit protein complex responsible for the regulation of the turnover of cellular proteins, thus maintaining cellular homeostasis. It controls protein expression and function by degradation of ubiquitinated proteins as well as cleanses the cell of abnormal or misfolded proteins. Bortezomib inhibits the proteasome by binding reversibly to the chymotrypsin-like site in the 20S core. The proteolytic inhibition disrupts the homeostasis leading to cell death. 105
We have shown that Cyclin E is critical for genotoxic stress-induced apoptosis of
tumor cells of hematopoietic origin (Mazumder et al., 2000). An 18-kDa Cyclin E
fragment (p18-Cyclin E) is generated through caspase-mediated proteolytic cleavage of
Cyclin E (Mazumder et al., 2002). Cleavage of Cyclin E results in the generation of p18-
Cyclin E, which lacks the Cdk2-interaction domain as well as the N-terminal
phosphodegron known to be necessary for the optimal ubiquitination and degradation of
Cyclin E. However, the presence of the main phosphodegron comprised of the Ser372,
Thr380, and Ser384 residues would suggest a similar regulatory mechanism of p18-
Cyclin E as Cyclin E by the SCF complex. Here we show that, indeed, the turnover of
p18-Cyclin E can be regulated by SCFFbw7. Strikingly, the recognition of p18-Cyclin E by the SCFFbw7 can be independent of its phosphorylation at the Cdc4 phosphodegron (CPD)
(Nash et al., 2001) or binding to Cdk2. Moreover, SCF has a minor contribution to the
degradation of p18-Cyclin E suggesting a parallel mechanism responsible for its turnover
that is independent of Cul1 or Cul3 complexes or the C-terminus of Hsp70 interacting
protein (CHIP). Here we propose that a novel pathway involving Ku70, known as a
critical component of Non Homologous End-joining DNA repair, that we have recently
identified as a specific p18-Cyclin E-interacting protein, plays a significant role in the
proteasomal degradation of p18-Cyclin E. We are reporting here the interaction between
Ku70 and the E3 ubiquitin ligase Hdm2 (Human homologue of Murine Double Minute).
Hdm2 is known for negatively regulating and ubiquitinating the p53 tumor suppressor
protein (Haupt et al., 1997; Kubbutat et al., 1997). Furthermore, Hdm2 has been shown to 106
associate with Ku70 in human umbilical vein endothelial cells (HUVEC) (Gama et al.,
2008, unpublished).
While attractive, the short half-life of p18-Cyclin E limits its clinical use. Since extensive ubiquitination after its generation leads to rapid proteasomal degradation, we tested whether enhancing its stability using proteasome inhibitors may potentiate its effect. Indeed, blocking its degradation caused a massive elevation of its levels as well as its association with Ku70 leading to Bax activation and enhanced apoptosis. Cells expressing p18-Cyclin E were more sensitive to cell death induction upon treatment with
Bortezomib, as compared to the parental cells. By preventing its proteasomal degradation, p18-Cyclin E may become an effective therapeutic target for Bortezomib when it is used in the therapy of hematopoietic or solid cancers.
Materials and Methods
Cell lines and treatments. HEK293T human kidney and NCI-H1299 non-small cell lung
carcinoma cells were maintained in DMEM (Gibco-BRL) containing 10% fetal bovine
serum (Hyclone, Logan, UT) with L-glutamine, and 100 units/ml penicillin/streptomycin.
Cells ~75% confluent were irradiated with 5 Gy (137Cs source; fixed dose rate of 2.0
Gy/min) (Gong et al., 1999). Bortezomib (Velcade) was obtained from Millenium
Pharmaceuticals (Cambridge, MA), MG132 and cycloheximide from Sigma (St. Louis,
MO). For the half-life study, cells were treated with 10 µM cycloheximide and collected 107
at the indicated times after treatment. HA-p18-Cyclin E stably expressing 293T and
H1299 cells were generated by infecting cells with a lentivirus carrying HA-p18-Cyclin E as well as EGFP separated by an IRES2 sequence and under the control of an EF1α
promoter.
Western blotting and immunoprecipitation analyses. For immunoblots, cells were
lysed in a buffer consisting in 20 mM HEPES, pH 7.5, 1 mM EDTA, 150 mM NaCl, 1%
NP-40, 1 mM DTT and protease inhibitors (1 mM PMSF, 2 µg/ml aprotinin, leupeptin,
and pepstatin) and immunoblotting was performed with anti-HA (Covance; Berkeley,
CA), Cyclin E1 (HE-12), Cdk2 (D-12), p53 (DO1), Ku70 (E-5 and H-308; Santa Cruz
Biotechnology; Santa Cruz, CA), PARP-1 (1:1000; Cell Signaling; Beverly, MA), and β-
actin (Sigma) antibodies. For immunoprecipitation, the lysis buffer composition was
similar to that employed for Westerns except that instead of 1% NP-40, for the
interactions between Cdk2 and p18-Cyclin E or Cyclin E, 0.1% NP-40 was used, while
for the interactions between p18-Cyclin E and Fbw7α, β, or γ, 0.2% NP-40 was used. For the interaction between Bax and Ku70 cells were lysed in 1% CHAPS.
Immunoprecipitation was performed with Bax antibody (N-20, Santa Cruz
Biotechnologies). Cell lysates (500-1000 µg) were incubated with 0.5-2 µg antibody for 2 h at 4oC, followed by incubation with Protein A plus G agarose beads for 1 h at 4oC.
Beads were washed with lysis buffer and boiled with 2x sample buffer. The eluted
proteins were resolved by SDS-PAGE and Western blotting was performed using the
appropriate antibodies, as described above. For determining the interaction between 108
Hdm2 and Ku70 in HEK293T cells, untreated or 2h etoposide (10 µM) treated cells were lysed in 1% Triton-X buffer in PBS, containing 1x protease inhibitor cocktail (Sigma), and 1 mM PMSF. Immunoprecipitation was performed with 2 µg anti-Hdm2 antibody
(2A10; Calbiochem: San Diego, CA) for 2 h at 4°C. 2 µg normal mouse IgG were used as
a negative control. For the determination of Ku70 and Hdm2 interaction in IM-9 cells,
these were irradiated with 4 Gy and collected after 8h.
For determining the ubiquitination status, cells were treated with 25 µM MG132
prior to harvesting and lysed in the same 1% NP-40 lysis buffer as described above also
containing 1x HALT phosphatase inhibitor cocktail (Pierce, Rockford, IL), 5 µM MG132 and 5 µM N-ethyl-maleimide (Sigma). Immunoprecipitation was performed with HA or
FLAG antibody (Sigma) for tagged p18-Cyclin E, with (HE-111, Santa Cruz
Biotechnology) for Cyclin E for 3 h at 4oC followed by incubation with Protein A plus G agarose beads for 1 h. Immunoblotting was performed with either HA antibody, anti-
Ubiquitin mouse monoclonal antibody (Chemicon, Temecula, CA) or rabbit polyclonal antibody (Dako, Carpinteria, CA).
Transfections. H1299 cells were transfected with 10 µg p18-Cyclin E-HA and 3 µg
FLAG-Fbw7α, β, or γ. At 24 h post-transfection, cells were collected and lysed.
Immunoprecipitation was performed with anti-HA antibody followed by immunoblotting
with anti-FLAG antibody. For the overexpression of FLAG-tagged Fbw7α, β, γ and dominant negative Cul1, dominant negative Cul3, 3 µg of construct and Lipofectamine
2000 (Invitrogen, Carlsbad, CA) was used according to the protocol provided by the 109
manufacturer. Knock-down of Cdk2 and Fbw7 was performed using 75-100 nM of
siRNA from Dharmacon (Lafayette, CO). As a negative control we used siCONTROL
non-targeting siRNA pool #1 from Dharmacon. In both the overexpression and knock-
down experiments cells were collected after 48 h. To effectively knock-down Ku70,
293T cells were transfected with 100 nM siKu70 or siCONTROL (Dharmacon). At 24 h
post transfection, cells were collected and plated followed by a second transfection the
second day with cells being finally collected the next day (72 h after the initial
transfection). Overexpression of CHIP (kind gift form Dr. E. Ficker, Metrohealth,
Cleveland) was achieved by transfection of an expression construct followed by collection after 24h. CHIP was knocked-down using siRNA from Dharmacon. The transfected cells were collected after 24 and 48 h respectively.
Construction of expression vectors and p18-Cyclin E phosphomutants. FLAG-tagged
p18-Cyclin E was constructed by subcloning p18-Cyclin E into the HindIII/BamHI
restriction sites of the p3xFLAG-myc-CMV-24 vector (Sigma) using the forward primer
5’-CGACAAGCTTGCCATGGACTGCCTTG-3’ and the reverse primer 5’-
TAGATGGATCCTCTCACGCCATTTCCGG-3’ resulting in an expression construct
that has the 3xFLAG at the N-terminus, but no myc tag. S384A-p18-Cyclin E was created
by mutating the Ser384 to Ala in the HA-tagged p18-Cyclin E using the ExSite PCR-
based site-directed mutagenesis kit from Stratagene (La Jolla, CA) and the primers
S384A-b: 5’-CTGTGGCGGGGTGAGGAGC-3’ and S384A-P: 5’-phospho-
GCCGGTAAGAAGCAGAGCAGCG-3’. C-terminal HA-tagged T380A-p18-Cyclin E 110
was created by subcloning T380A-p18-Cyclin E from the T380A-Cyclin E-GFP
construct obtained from Dr. Geisen (Burnham Institute) (Geisen and Moroy, 2002) into
the C-terminal HA-containing pcDNA3.1 vector. The resulting construct was used for the
generation of S372A/T380A-p18-Cyclin E by mutating the Ser372 to Ala using the
primers S372A-b: 5’-AGCCCTATTTTGTTCAGACAACATG-3’ and S372A-P: 5’-
phospho-GCTCCTCTCCCCAGTGGGCTCCT-3’. The resulting construct was further
used for the generation of S372A/T380A/S384A-p18-Cyclin E using the primers S384A-
T380A: 5’-CTGTGGCGGCGCCAGGAGC-3’ and S384A-P. The T380A/S384A-p18-
Cyclin E was generated by mutating Ser384 to Ala in T380A-p18-Cyclin E using the primers S384A-T380A and S384A-P. All mutations were verified by DNA sequencing.
Results
p18-Cyclin E is a short-lived protein that is stabilized by proteasome inhibition
Our previous experiments have shown that Cyclin E is cleaved during genotoxic
stress induced by IR generating an 18-kDa C-terminal fragment, p18-Cyclin E
(Mazumder et al., 2007b). To examine the stability of p18-Cyclin E compared to full
length Cyclin E (Cyclin E), the half-life of these two proteins was determined following
protein translation inhibition by Cycloheximide treatment. Results of these experiments
indicate that p18-Cyclin E had a remarkably short half life of less than 0.5 h. In contrast,
the half-life of Cyclin E was longer than 8 h in HEK293T cells stably expressing p18-
Cyclin E (Fig 3.1A). These cells were generated in order to overcome the drawbacks 111
related to transient expression of p18-Cyclin E. Furthermore, these cells express p18-
Cyclin E at low, non-toxic levels and Cyclin E cleavage does not take place.
Strikingly, proteasome inhibition achieved by treatment with either MG132 or the new, clinically approved compound, Bortezomib (Velcade) led to a dramatic stabilization of p18-Cyclin E, indicating that this protein is very rapidly turned over by its proteasome- mediated degradation. Protein stabilization took place in both HEK293T and NCI-H1299 cells stably expressing p18-Cyclin E (Fig 3.1B). Interestingly, the stabilizing effect of proteasome inhibition on Cyclin E was much less robust than that on p18-Cyclin E.
112
Fig 3.1 p18-Cyclin E is a short-lived protein that is stabilized by proteasome
inhibition. A) 293T cells stably expressing HA-tagged p18-Cyclin E were treated with
10 µM Cycloheximide for the indicated times. The levels of p18-Cyclin E, Cyclin E, and
β-actin used as a loading control were determined by Western blotting using the respective primary antibodies. B) 293T and H1299 cells stably expressing HA-p18-
Cyclin E were either untreated (Control, C) or treated with 1 µM Bortezomib
(Velcade:Vel) or 10 µM MG132 (MG) and collected after 3 h. The levels of p18-Cyclin
E, Cyclin E, and β-actin were determined by Western blotting. (Data are representative of three independent experiments)
A
0.25 .5124824Time(h) - Cyclin E
- p18-Cyclin E
- β-actin
B
293T H1299 C Vel MG C Vel MG -CyclinE - p18-Cyclin E - β-actin 113
p18-Cyclin E is effectively ubiquitinated
The above experiments revealed that proteasome inhibition led to a massive
increase in the levels of p18-Cyclin E. Poly-ubiquitination of the target protein is the
most common signal for proteasome-mediated degradation. The higher molecular weight,
modified forms of p18-Cyclin E were readily visible following a long exposure upon
MG132 treatment in the presence of N-ethylmaleimide (NEM), an inhibitor of the
ubiquitin isopeptidases, indicating that p18-Cyclin E is degraded through the addition of
multiple ubiquitin or ubiquitin-like residues that target it for proteasomal degradation
(Fig 3.2A). To determine whether p18-Cyclin E is modified by addition of Ubiquitin,
Flag-tagged p18-Cyclin E was co-transfected with an HA-Ubiquitin expression construct
in 293T cells. Immunoprecipitation with anti-FLAG antibody for p18-Cyclin E followed by immunoblotting with anti-HA antibody for Ubiquitin shows the characteristic smear
(Fig 3.2B left panel) as well as the characteristic laddering (Fig 3.2B right panel) of poly-
ubiquitinated p18-Cyclin E. To compare the extent of ubiquitination between p18-Cyclin
E and Cyclin E, the latter was immunoprecipitated followed by immunoblotting with an
anti-HA antibody for the ubiquitinated protein forms (Fig 3.2B left panel). This
experiment revealed that the extent of Cyclin E ubiquitination was less than that of p18-
Cyclin E, a result which confirms the finding above that Cyclin E has a much longer half-
life than its carboxy terminal fragment, p18-Cyclin E (Fig 3.1A). Addition of poly-
ubiquitin chains was also observed when HA-p18-Cyclin E was immunoprecipitated from stably expressing 293T cells followed by immunoblotting with two different anti-
Ubiquitin antibodies (Fig 3.2C). 114
Fig 3.2 p18-Cyclin E is effectively ubiquitinated. A) 293T cells were transfected with
HA-tagged p18-Cyclin E or empty vector (C) and treated 24 h later with 2 µM or 10 µM
MG132 for an additional 16 h and 4 h, respectively. The levels of p18-Cyclin E and β- actin were determined by Western blotting. B) 293T cells were co-transfected with
FLAG-tagged p18-Cyclin and HA-tagged Ubiquitin. At 24 h post-transfection, cells were treated with 25 µM MG132 for 3 h. p18-Cyclin E and Cyclin E were immunoprecipitated from whole-cell lysates (WCL) with anti-FLAG (1st and 2nd lane) or anti-Cyclin E (3rd lane) antibodies and separated by SDS-PAGE. Ubiquitination of p18-Cyclin E and Cyclin
E (left top panel), levels of p18-Cyclin E and β-actin in WCL (right panel) were
determined by Western blotting. C) 293T cells stably expressing HA-p18-Cyclin E or
only GFP as control were treated with 25 µM MG132 for 3 h. p18-Cyclin E was immunoprecipitated using anti-HA antibody. Ubiquitination of p18-Cyclin E (left panel rabbit anti-Ubiquitin, middle panel mouse anti-Ubiquitin), levels of p18-Cyclin E (right panel HA antibody), and β-actin were determined by Western blotting.
115
A
C p18-Cyclin E --2µM10µM MG132 16h 4h
ubiquitinated forms p18-Cyclin E
- β-actin
B
Ub-HA + + + + + + + + + FLAG-p18-Cyclin E + - - + - - + - - poly-Ubiquitinated - protein poly-Ub- p18-Cyclin E
- p18-Cyclin E IgGκ - - β-actin WB: HA
IgGλ - p18-Cyclin E - IP: FLAG CycE WB: HA WB: HA
116
C
GFP p18 GFP p18 GFP p18
poly-Ub poly-Ub p18-Cyclin E p18-Cyclin E poly-Ub- p18-Cyclin E -IgGκ -IgGκ -IgGλ IP: HA -IgGλ WB: pAb Ubiquitin - β-actin IP: HA WB: mAbUbiquitin
117
Fbw7 interacts with p18-Cyclin E independently of phosphorylation at the C-
terminal phosphodegron
Cyclin E is known to be degraded through the ubiquitin-proteasomal pathway that
uses Fbw7 as a component of the Skp1-Cul1-F-box (SCF) complex (Koepp et al., 2001;
Moberg et al., 2001; Strohmaier et al., 2001). There are three splice-derived isoforms of
Fbw7 designated as α, β, and γ that localize to different cellular compartments (Spruck et al., 2002). To determine whether p18-Cyclin E is also targeted for proteasomal degradation by the SCFFbw7 complex, we examined the interaction between p18-Cyclin E and all three Fbw7 isoforms. To determine the interactions of p18-Cyclin E with the different isoforms of Fbw7, HEK293T cells were transiently transfected with HA-p18-
Cyclin E and FLAG-Fbw7α, β, γ expression constructs and immunoprecipitated with
anti-HA to pull-down p18-Cyclin E followed by Western blot performed with an anti-
FLAG antibody. p18-Cyclin E associated with all three isoforms of Fbw7, which indicates that it is a ubiquitination target of Fbw7, which regulates its degradation (Fig
3.3A).
Phosphorylation plays an important role in the interaction between the Fbw7 isoforms and Cyclin E (Koepp et al., 2001; Moberg et al., 2001; Strohmaier et al., 2001).
To examine the influence of phosphorylation upon the expression levels of p18-Cyclin E, amino-acid residues known to be important for the phosphorylation-dependent turnover
of Cyclin E (Ye et al., 2004) were mutated alone or in combination. We generated the
following phosphorylation mutants: T380A, S384A, T380A/S384A, S372A/T380A, and
S372A/T380A/S384A. Unlike the effect that is known for these point mutations on 118
Cyclin E (Ye et al., 2004), transient transfection of these mutant p18-Cyclin E expression
constructs has shown very striking differences. The Thr380 residue, which has been
reported to play a crucial role in the turnover of Cyclin E (Clurman et al., 1996; Won and
Reed, 1996), does not affect the stability of p18-Cyclin E. Moreover, the mutation of
Ser384 instead of increasing the levels of p18-Cyclin E leads to its destabilization. Only
mutation of Ser372 showed the expected increase in levels of the protein (Fig 3.3B), with
the triple mutant having less effect, most likely because of the destabilizing effect of the
Ser384 mutation. Furthermore, co-transfection of the HA-tagged triple phosphorylation
mutant S372A-T380A-S384A-p18-Cyclin E with the Fbw7α, β, and γ isoforms in 293T cells still showed their interaction (Fig 3.3C). These data indicate that p18-Cyclin E may
interact with Fbw7α, β, and γ independently of phosphorylation at S327, T380 or S384 that constitute the C-terminal phosphodegron of Cyclin E.
119
Fig 3.3 Fbw7 interacts with p18-Cyclin E independently of phosphorylation at the
C-terminal phosphodegron. A) H1299 cells were co-transfected with HA-tagged p18-
Cyclin E and FLAG-tagged Fbw7α, β, or γ. p18-Cyclin E was immunoprecipitated from whole cell lysates with anti-HA antibody and separated by SDS-PAGE. Levels of immunoprecipitated Fbw7α, β or γ were determined by Western blotting with anti-Flag antibody. B) 293T cells were transfected with empty pcDNA3.1 vector or containing HA- p18-Cyclin E and its derivative mutants: T380A, S384A, T380A/S384A,
S372A/T380A/S384A, and S372A/T380A. Their levels were determined by Western blotting using the anti-HA antibody. C) 293T cells were co-transfected with HA-
S372A/T380A/S384A-p18-Cyclin E (triple mutant - TM) and FLAG-Fbw7α, β, or γ.
S372A/T380A/S384A-p18-Cyclin E was immunoprecipitated from whole cell lysates with anti-HA antibody and separated by SDS-PAGE. Levels of immunoprecipitated
Fbw7α, β, or γ were determined by Western blotting with anti-Flag antibody. (* - cross- reacting protein) D) Schematic representation of Cyclin E phosphodegrons showing the relative positions of the nuclear localization sequence (NLS) and its two groups of phosphorylation sites at the N-terminus (S58, T62, S75, S88) and C-terminus (S372,
T380, S384) necessary for its optimal recognition by SCFFbw7 and ubiquitination.
Caspase-3 dependent cleavage at Asp275 generates an 18-kDa carboxy terminal fragment, which carries only the C-terminus phosphodegron. (Data are representative of three independent experiments)
120
A
Fbw7 C p18 Lys -Fbw7α
-IgG -IgG
-Fbw7β
-Fbw7γ
-IgG IP: HA WB: Flag B
- p18-Cyclin E - β-actin
121
C
IP TM Lys ++++TM +--+ Fbw7 -Fbw7β -IgG -Fbw7γ -IgG IP TM Lys + + + + TM - + - + Fbw7 -Fbw7α *
D
122
p18-Cyclin E is regulated by the SCFFbw7 complex and independently of Cdk2 binding
To further determine the role of Fbw7 in p18-Cyclin E regulation, p18-Cyclin E- expressing 293T cells were transfected with the α, β, and γ isoforms of Fbw7. Fbw7
overexpression enhanced the turnover of p18-Cyclin E as well as that of Cyclin E.
Conversely, the ectopic expression of dominant negative (dn) Cul1 led to a decrease in
their degradation (Fig 3.4A). The effect on p18-Cyclin E levels was limited while that on
Cyclin E was dramatic leading to a massive accumulation of the protein. The striking
difference in the increase in levels of Cyclin E as compared to p18-Cyclin E would
suggest a different mechanism employed for their regulation by the SCFCul1 complex.
Cyclin E is known to be regulated through both the Cul1 and Cul3 pathways
(Koepp et al., 2001; McEvoy et al., 2007; Singer et al., 1999). To determine the influence
of Cul1 and Cul3 upon the degradation of p18-Cyclin E, stably expressing 293T-p18-
Cyclin E cells were transfected with dnCul1 (Jin et al., 2003) or dnCul3 (Cullinan et al.,
2004) expression constructs. Even though this led to a minor stabilization of p18-Cyclin
E (Fig 3.4B), this increase was far less compared to the robust increase in levels of Cyclin
E (Fig 3.4A). This suggests that even if SCFCul1-Fbw7 plays a role in the degradation of
p18-Cyclin E, this mechanism represents only a minor component and that another
regulatory pathway plays a predominant role in its turnover. Moreover, these data
indicate that Cul3, known to be important for degradation of free Cyclin E (McEvoy et
al., 2007; Singer et al., 1999) has no role in p18-Cyclin E regulation. Finally, to clearly
establish the role of Fbw7 in the turnover of p18-Cyclin E, RNA interference (RNAi) was 123
used to diminish Fbw7 levels in 293T cells stably expressing p18-Cyclin E.
Immunoblotting with anti-HA antibody for p18-Cyclin E and anti-Cyclin E (HE-12)
antibody for Cyclin E (Fig 3.4B) at 48 h post-transfection, indicated that their levels
increased, indicating a regulatory role for Fbw7.
Cdk2 is known to be responsible for the phosphorylation of Ser384 on Cyclin E
when it is present in complex with Cdk2 (Welcker et al., 2003). To assess whether Cdk2
affects the levels of p18-Cyclin E, even in the absence of their interaction (Fig 3.5A and
3.5B), Cdk2 was knocked-down by siRNA (Fig 3.4C). However, despite a robust
depletion of Cdk2 levels, there was only a minimal increase in p18-Cyclin E levels
despite the lack of Cdk2 binding domain and its implicit interaction with Cdk2. Cyclin E
increased to some extent in the absence of Cdk2 but much less compared to when Fbw7
was knocked down. The C-terminus of Hsp70 interacting protein (CHIP) is an E3 ligase
that partners with chaperones Hsp70, Hsc70, and Hsp90 targeting the degradation of misfolded proteins. Furthermore, it has been shown to participate in the proteasome- mediated degradation of base excision repair proteins such as XRCC1 and Pol β, which have not been recruited to a DNA repair complex (Sobol, 2008). To determine whether
CHIP contributes to the turnover of p18-Cyclin E, it was overexpressed in 293T cells stably expressing HA-p18-Cyclin and at 48 h post-transfection the levels of p18-Cyclin E were determined by Western blotting. It appears that CHIP does not potentiate the degradation of p18-Cyclin E. At most, it may show a slightly opposite effect (Fig 3.4D).
Moreover, knocking-down CHIP by siRNA also showed no significant effect on the levels of p18-Cyclin E (Figure 3.4E). 124
Fig 3.4 p18-Cyclin E is regulated by the SCFFbw7 complex and independently of
Cdk2 binding. A) 293T cells stably expressing HA-tagged p18-Cyclin E were transfected with a combination of FLAG-tagged Fbw7α, β and γ or dominant negative
Cul1 (dnCul1). The levels of Fbw7α, β, γ, dnCul1, Cyclin E, p18-Cyclin E, and β-actin were determined by Western blotting. B) 293T cells stably expressing HA-p18-Cyclin E were transfected with dominant negative Cul1 (dnCul1) and dominant negative Cul3
(dnCul3) expression constructs. The levels of dnCul1, dnCul3, p18-Cyclin E, and β-actin, as a loading control, were determined by Western blotting. C) 293T cells stably
expressing HA-p18-Cyclin E were transfected with 100 nM siFbw7, siCdk2, and
siControl. The levels of Cyclin E, Cdk2, p18-Cyclin E, and β-actin were determined by
Western blotting. D) 293T cells stably expressing HA-p18-Cyclin E were transfected with CHIP or empty vector (C) and cells were collected after 48 h. The levels of CHIP, p18-Cyclin E and β-actin were determined by Western blotting. E) 293T stably
expressing HA-p18-Cyclin E were transfected with 100 nM siCHIP or siCONTROL and
cells were collected after 24 or 48h. The levels of CHIP, p18-Cyclin E, and β-actin were determined by Western blotting. (Data are representative of three independent experiments)
125
A
C -Fbw7α -Fbw7β,γ -dnCul1 - Cyclin E
- p18-Cyclin E - β-actin
B
C -dnCul1 -dnCul3 - p18-Cyclin E -dnCul1 - β-actin C
- Cyclin E -Cdk2
- p18-Cyclin E
- β-actin 126
D
C CHIP -CHIP - p18-Cyclin E - β-actin E
-CHIP -p18-CyclinE - β-actin
127
Ku70 but not Cdk2 binding regulates p18-Cyclin E stability
We have previously identified p18-Cyclin E as the carboxy terminal fragment of
Cyclin E, generated following a caspase-3-mediated cleavage (Mazumder et al., 2002).
p18-Cyclin E encompasses the 276-395 amino-acid domain of Cyclin E and thus, is
lacking the Cdk2 binding domain (Mazumder et al., 2007a). To confirm that indeed
Cyclin E binds Cdk2 while p18-Cyclin E has lost this ability, HA-p18-Cylin E was
immunoprecipitated from 293T cells stably expressing it followed by immunoblotting for
Cdk2 (Fig 3.5A). Conversely, Cdk2 was immunoprecipitated from 293T expressing HA-
Cyclin E or HA-p18-Cyclin E respectively, followed by immunoblotting with HA
antibody (Fig 3.5B). These experiments, together with our previous report (Mazumder et
al., 2002) clearly indicate that p18-Cyclin E does not interact with Cdk2 while Cyclin E,
as expected, does. Taken together these data suggest that even though p18-Cyclin E does
not interact with Cdk2, Fbw7 can recruit it to the SCF complex and promote its
ubiquitination.
To examine the possible role of Ku70 binding in the degradation of p18-Cyclin E,
293T cells were transfected with HA-p18-Cyclin E alone or in the presence of FLAG-
Ku70. This led to a dramatic reduction in the levels of p18-Cyclin E (Fig 3.5C). To compare this change with the effect of Fbw7, HA-p18-Cyclin E was co-transfected with
Fbw7α, β, and γ. Fbw7 overexpression led to a much more limited increase in the turnover of p18-Cyclin E as compared to that induced by Ku70. To clearly establish the role of Ku70, its levels were substantially depleted by specific siRNA oligonucleotides.
Down-regulation of Ku70 by siRNA led to an increase in p18-Cyclin E levels, a change 128
that could not be observed for Cyclin E (Fig 3.5D). Hdm2 is an ubiquitin E3 ligase
known to be transcriptionally induced by p53 following DNA damage (Haupt et al.,
1997; Kubbutat et al., 1997). Its role is to polyubiquitinate p53 targeting it to proteasomal degradation, thus limiting its effect. The newly discovered interaction between Hdm2 and
Ku70 (Gama et al., 2008, unpublished) raised the question of whether Hdm2 interacts with Ku70 in our experimental system. In IM-9 cells this association was clearly induced following irradiation, that corresponded to increased p53 levels, known to lead to increased levels of Hdm2 (Figure 3.5E). Hdm2 was also immunoprecipitated from control or etoposide (VP16)-treated HEK293T cells and the presence of Ku70 was determined by immunoblotting. Ku70 and Hdm2 interacted in both control and etoposide-treated cells (Figure 3.5F), as p53 is inactive in these cells and therefore levels of Hdm2 are not expected to be induced. Taken together, these data strongly implicate
Ku70 and Hdm2 in a degradation pathway of p18-Cyclin E, the exact role of which and its mechanism still need to be investigated.
129
Fig 3.5 Ku70 but not Cdk2 binding regulates p18-Cyclin E stability. A) 293T cells
stably expressing HA-p18-Cyclin E or GFP as control were lysed with 0.1% NP-40 lysis
buffer. p18-Cyclin E was immunoprecipitated with anti-HA antibody. Levels of
immunoprecipitated Cdk2 and of p18-Cyclin E present in whole cell lysates were
determined by Western blotting. B) Cdk2 was immunoprecipitated from 293T cells transfected with HA-Cyclin E and p18-Cyclin E. Immunoprecipitated Cyclin E and p18-
Cyclin E as well as their levels from whole cell lysates were determined by Western blotting. C) 293T cells were transfected with FLAG-Ku70, or HA-p18-Cyclin E alone, or in combination with Fbw7α, β, γ or Ku70 in presence of HA-p18-Cyclin E. The levels of
Ku70, Fbw7α, β, γ, p18-Cyclin E, and β-actin were determined after 24 h post-
transfection by Western blotting. D) 293T cells stably expressing HA-p18-Cyclin E were transfected with 100 nM siKu70 or siCONTROL. At 48 h post transfection, a second
round of transfection was performed to achieve a satisfactory down-regulation of Ku70.
The levels of Ku70, p18-Cyclin E, and β-actin used as a loading control were determined by Western blotting at 72 h after the initial transfection. E) IM-9 cells were irradiated
with 4 Gy and Hdm2 was immunoprecipitation after 8h. Levels of Ku70, Hdm2, and p53
were determined by immunoblotting. F) Hdm2 was immunoprecipitated from HEK293T
cells and the levels of associated Ku70 were determined by immunoblotting.
130
A
WCL +-HA-p18-CyclinE -Cdk2 -IgGλ -IgGλ - HA-p18-CyclinE
IP: HA B
- + HA-Cyclin E WCL + - HA-p18-Cyclin E - Cyclin E
- p18-Cyclin E
IP: Cdk2 C
---+Fbw7α,β,γ +-+-Ku70 -+++p18-Cyclin E -Fbw7α -Ku70
- p18-Cyclin E
- p18-Cyclin E (long exp.) - β-actin
131
D
-Ku70 - Cyclin E - β-actin - p18-Cyclin E - β-actin E
IP Hdm2 Lysate CIR C IR -Hdm2 - Ku70 -p53 -IgGκ
F
Lys IP Hdm2 CEIgG C E -Ku70
132
p18-cyclin E sensitizes cells to Bortezomib treatment through enhanced association
with Ku70 and release of Bax from Ku70
Of the commonly used proteasome inhibitors tested Bortezomib was more
effective than MG132 in stabilizing p18-Cyclin E (Fig 3.6A). To examine the dynamics
of proteasome inhibition and onset of apoptosis, stably expressing 293T-p18-Cyclin E
cells were treated with 100 nM Bortezomib for 0.5, 1, 2, 4, 8, 24, and 48 h. Western blot
analysis was performed for PARP-1 to determine the extent of cell death, HA for the
expression of p18-Cyclin E, HE-12 for the expression of Cyclin E and β-actin as a
loading control. Apoptosis was detectable as early as 8 h, with ~10% of PARP-1 being
cleaved by 24 h and 50% by 48 h (Fig 3.6B). The level of p18-Cyclin E was increased in
the presence of Bortezomib, due to the inhibition of its proteasomal degradation. p18-
Cyclin E levels reached a maximum as early as 2 h, while Cyclin E levels did not vary
significantly during the same time interval.
While attractive, the perspective of using p18-Cyclin E for cancer treatment is limited by its very short half-life (Fig 3.1A). Therefore, enhancing its stability may
potentiate its effect. To determine whether the levels of p18-Cyclin E can enhance the
response of cells to Bortezomib treatment, control (GFP-expressing) or p18-Cyclin E-
stably expressing cells were examined at 24 h following treatment with increasing
concentrations of Bortezomib. Increased PARP-1 cleavage in p18-Cyclin E-expressing
cells suggested that p18-Cyclin E may be a mediator of Bortezomib-induced cell death
(Fig 3.6C). 133
To determine the mechanism responsible for the observed cell sensitivity, we
examined the interaction between p18-Cyclin E and Ku70 before and after treatment with
50 nM Bortezomib or irradiation. At 16 h after treatment, HA-p18-Cyclin E was
immunoprecipitated with anti-HA antibody followed by immunoblotting for Ku70.
Treatment with Bortezomib, but not IR caused an elevation of the Cyclin E and to an
even larger extent that of p18-Cyclin E levels (Fig 3.6D). Increased p18-Cyclin E levels led to an increase in its association with Ku70 (Fig 3.6E). To determine whether this
association leads to release of Bax from Ku70, Bax is immunoprecipitated followed by immunoblotting for Ku70. As expected, elevated levels of p18-Cyclin E bound more
Ku70, resulting in release of Bax (Fig 3.6F), which correlated with induction of cell death as indicated by cleavage of PARP-1 (Fig 3.6D).
134
Fig. 3.6 p18-cyclin E sensitizes cells to Bortezomib treatment through enhanced association with Ku70 and release of Bax from Ku70. A) 293T cells stably expressing
HA-p18-Cyclin E were treated with 100 nM of Bortezomib (Velcade:Vel) or MG132 and collected at the indicated intervals. The levels of PARP-1, Cyclin E, p18-Cyclin E and β-
actin were determined by Western blotting. B) 293T cells stably expressing HA-p18-
Cyclin E were treated with 100 nM Bortezomib and collected at the indicated times. The
levels of PARP-1, p18-Cyclin E, Cyclin E and β-actin as a loading control were detected by Western blotting. C) 293T cells stably expressing GFP as control or HA-p18-Cyclin E
were treated with the indicated concentrations of Bortezomib and collected after 24 h.
The levels of PARP-1, p18-Cyclin E and β-actin were determined by Western blotting.
D) 293T cells stably expressing HA-p18-Cyclin E were treated with 5 Gy IR or 50 nM
Velcade (Vel) and collected after 16 h. The levels of PARP-1, Cyclin E, p18-Cyclin E and β-actin were determined by Western blotting. E) 293T cells stably expressing HA- p18-Cyclin E were treated with 5 Gy IR or 50 nM Bortezomib and collected after 16 h.
p18-Cyclin E was immunoprecipitated with anti-HA antibody. Levels of
immunoprecipitated Ku70 as well as present in WCL, were determined by Western blotting. F) 293T cells stably expressing HA-p18-Cyclin E were treated with 5 Gy IR or
50 nM Bortezomib and collected after 16 h. Bax was immunoprecipitated with anti-Bax
(N-20) antibody and the levels of associated Ku70 were determined by Western blotting.
135
A
Bortezomib MG132 0 8 24 48 0 8 24 48 Time (h) -PARP
- Cyclin E
- p18-Cyclin E
- β-actin B
0.512482448Time (h) -PARP116kD -PARP85kD p18-Cyclin E - (long exp.) - p18-Cyclin E (short exp.) - Cyclin E - β-actin - β-actin C
293T 293T p18 0 25 100 200 0 25 100 200 Bortezomib (nM) -PARP
- p18-CyclinE
- β-actin
136
D
C IR Bort -PARP -p85 -CyclinE
- p18 Cyclin E
- β-actin
E
C IR Bort -Ku70(shortexp) - Ku70 (long exp)
IP: HA WB: Ku70 F
C IR Bort -Ku70 -IgGκ IP: Bax WB: Ku70
137
Discussion
The expression of Cyclin E, the master regulatory protein of the cell cycle
transition from the G1 to S phase, is controlled at the mRNA level as well as post-
transcriptionally., Cyclin E is regulated post-transcriptionally by the 26S proteasome
following its ubiquitination by the SCF complex. While the Cdk2-bound Cyclin E
requires phosphorylation at a conserved phosphodegron in order to be recognized and
efficiently ubiquitinated by the Skp1-Cul1-Fbw7 complex (Zhang and Koepp, 2006), the
free Cyclin E is degraded following direct interaction with Cul3 (McEvoy et al., 2007;
Singer et al., 1999).
We have previously shown that Cyclin E is regulated by genotoxic stress caused by ionizing radiation (IR) or other chemotherapeutic drugs, and which plays a functional role in apoptosis of hematopoietic cells (Mazumder et al., 2000). Moreover, we have demonstrated that caspase-3 mediates cleavage of Cyclin E that results in generation of its C-terminal proteolytic fragment, p18-Cyclin E. Protein synthesis inhibition studies with cycloheximide in HEK293T cells stably expressing p18-Cyclin E have now determined a half-life of less than 0.5 h for p18-Cyclin E as opposed to more than 8 h for
Cyclin E. This finding indicates that after being generated, p18-Cyclin E is degraded rapidly through the proteasomal degradation pathway. Indeed, the addition of proteasome inhibitors such as MG132 and Bortezomib significantly inhibited the degradation of p18-
Cyclin E, indicating that p18-Cyclin E is a target of the ubiquitin-proteasomal degradation pathway. The higher molecular weight ubiquitinated forms of p18-Cyclin E are detectable using proteasome inhibitors, suggesting the susceptibility of these 138
ubiquitinated forms to degradation. Unlike Cyclin E, p18-Cyclin E cannot interact with
Cdk-2 as it is missing the Cdk-2 binding domain. It could, alternatively be bound and
regulated by the Cul3 degradation pathway similar to free Cyclin E (McEvoy et al., 2007;
Singer et al., 1999). However, we show here that Cul3 does not have a role in the
degradation of p18-Cyclin E.
Degradation of p18-Cyclin E is dependent on ubiquitination by the Skp1-Cul1-
Fbw7 (SCF) indicated by its interaction with the different isoforms of Fbw7, an
interaction directly responsible for the ubiquitin-dependent proteolysis of a number of
proteins including proto-oncogenes such as Cyclin E and c-Myc (Yada et al., 2004).
Strikingly, we find that the interaction between Fbw7 and p18-Cyclin E takes place even in the absence of p18-Cyclin E binding to Cdk2 and independently of its phosphorylation at three key residues at the well-characterized C-terminal phosphodegron of Cyclin E.
We found that p18-Cyclin E interacts stronger with the α and γ isoforms of Fbw7, which has also been reported for Cyclin E (van Drogen et al., 2006). It should be noted that there is a possibility that when p18-Cyclin E is generated in cells following cleavage of
Cyclin E during apoptosis, it may be phosphorylated at this C-terminal phosphodegron, whereas these phosphorylation events do not take place in cells stably expressing p18-
Cyclin E. The observed difference in turnover efficiency by Cul1 suggests a different regulatory mechanism for p18-Cyclin E than that for full length Cyclin E, which may involve a parallel pathway independent of Cul1 and Cul3.
We have recently identified a specific p18-Cyclin E-interacting protein, Ku70, known as a critical component of the Non Homologous End-joining (NHEJ) DNA repair 139
pathway (Mazumder et al., 2007b). This unexpected interaction raised the possibility of
p18-Cyclin E being targeted to degradation by a pathway involving Ku70. Indeed,
overexpression of Ku70 led to a decrease in p18-Cyclin E levels while knock-down of
Ku70 by siRNA stabilizes the protein. Furthermore, we have identified Ku70 in a
complex with the ubiquitin ligase Hdm2. Taken together, these data would suggest a
novel degradation mechanism for p18-Cyclin E involving its binding to Ku70 and Hdm2,
the exact role of which and its characterization still need to be determined. It is possible that Ku70 acts as a scaffold bringing Hdm2 and p18-Cyclin E together, thus facilitating its ubiquitination and degradation. This may suggest a two-step degradation process of
Cyclin E to achieve its accelerated turnover. These observations provide new insights into the regulation of Cyclin E as well as on proteasome-mediated degradation pathways that may affect other cellular proteins.
The ubiquitin-proteasome pathway is the major non-lysosomal proteolytic system
in the cytosol and nucleus of all eukaryotic cells. As most of the cell death-regulatory
proteins are targets of the ubiquitin-proteasomal pathway, proteasome inhibitors have
emerged as an attractive target for human cancer therapy (Aghajanian et al., 2002;
Chauhan et al., 2005). Bortezomib (PS-341) and MG132 are highly specific proteasome
inhibitors that induce apoptosis in a variety of tumor cell lines. Proteasome inhibitors have been demonstrated to overcome chemoresistance of tumor cells by increasing chemosensitivity and also by working in synergy with other agents to induce apoptotic cell death of tumor cells (Hayden et al., 2007; Russo et al., 2007). However, their specific targets and the intimate mechanism responsible for inducing cell death are unclear. 140
Bortezomib was the first small molecule proteasome inhibitor (Fig 3.7) that has
shown antitumoral activity in a number of cell types and is currently undergoing clinical trials for hematopoietic as well as solid tumors (Bross et al., 2004; Hideshima et al.,
2003; Papandreou et al., 2004; Richardson et al., 2003). It has already been approved by
the FDA for the treatment of multiple myeloma and mantle cell lymphoma in patients
who have received at least one prior therapy and who indicated disease progression.
Bortezomib alone or in combination with other chemotherapeutic agents represents a new attractive therapeutic approach and is currently investigated in clinical trials for various malignancies, such as multiple myeloma, chronic lymphocytic leukemia (CLL) (Pahler et al., 2003), acute lymphocytic leukemia, follicular lymphoma, non-Hodgkin’s lymphoma, myelodysplastic syndrome, non-small cell lung carcinoma, and head and neck cancer.
MG132 is a reversible specific and potent inhibitor of the 26S proteasome that has been
shown to induce cell death in many malignancies (Drexler, 1997).
141
Fig 3.7 Chemical structure of the proteasome inhibitor, bortezomib
HO OH B
N NH
H N O N
O
142
We have demonstrated recently that by binding to Ku70, p18-Cyclin E causes the
dissociation of Bax from Ku70 and its activation, thus leading to apoptosis (Mazumder et
al., 2007b). Therefore, p18-Cyclin E may be an attractive target for cancer therapy.
However, the scope of its clinical use is limited due to its labile nature and therefore
enhancing its stability may potentiate its effect. In this study we show that as a
consequence of proteasome inhibition with Bortezomib, p18-Cyclin E is stabilized and its
association with Ku70 increased leading to enhanced apoptosis. Previously, we have
reported that p18-Cyclin E-induced cell death is due to the interaction of Ku70 with p18-
Cyclin E, which influences the interaction of Bax with Ku70 and thereby, activation of
Bax. In this context, the association of p18-Cyclin E with Ku70 is also enhanced by
Bortezomib treatment, most probably due to the increased levels of p18-Cyclin E and
counteracting the destabilizing effect of Ku70 on p18-Cyclin E. Importantly, this
interaction leads to release of Bax followed by its activation, suggesting that increased
cell death is directly related to the binding of p18-Cyclin E to Ku70. We have determined
that cells expressing p18-Cyclin E are more sensitive to cell death induction when treated
with Bortezomib, as compared to the parental cells, which is most probably due to
inhibition of proteasomal degradation of p18-Cyclin E. Moreover, the combination of
Bortezomib with a DNA-damaging agent such as VP-16 is much more effective in
stabilizing p18-Cyclin E as well as sensitization of the cells (data not shown).
We have reported earlier that following treatment with DNA-damaging agents such as irradiation and VP16, p18-Cyclin E is generated through proteolytic cleavage of
Cyclin E (Mazumder et al., 2002; Mazumder et al., 2007b). Interestingly, Bortezomib 143
treatment also generates endogenous p18-Cyclin E in hematopoietic IM-9 cells at a
significantly higher level compared to treatment with irradiation alone (data not shown).
Furthermore, Bortezomib treatment sensitized cells much more compared to the radiation treatment. Most likely, the increased cell death in case of Bortezomib treated cells is due to stabilization leading to its increased p18-Cyclin E levels. Although p18-Cyclin E is not generated in all cell types, once expressed, it can induce apoptosis in a wide variety of cellular systems that we have tested. These findings support the development of p18-
Cyclin E as a novel therapeutical approach for the treatment of cancer.
Although the exact mechanism of how proteasome inhibition causes apoptosis is
not fully understood, the fact that malignant cells are much more sensitive to the death-
promoting aspects of proteasome inhibition than normal cells (Hayden et al., 2007)
makes the ubiquitin/26S proteasome system a target for cancer therapy. Bortezomib is the
first proteasome inhibitor that has become available for clinical use. The excellent results
achieved in patients with multiple myeloma suggest the use of Bortezomib in solid
cancers, especially in combination with chemotherapy or radiotherapy. Several phase I/II
studies have used Bortezomib as a mono therapy in advanced chemotherapy refractory
solid cancers but, knowledge of in-vivo effects of combined modality approaches using
Bortezomib is limited. In this study, we demonstrate that treatment of cancer cells with
the highly potent and selective proteasome inhibitors Bortezomib and MG132 triggers
activation of caspase and apoptosis by significant induction and stabilization of p18-
Cyclin E. Furthermore, by preventing its proteasomal degradation, p18-Cyclin E may 144
become an important Bortezomib target while an adjuvant for Bortezomib treatment of solid tumors and hematopoietic malignancies.
CHAPTER IV
Summary and future directions
Summary
Conventional therapies including ionizing radiation, alkylating agents, anti- metabolites, and different targeted inhibitors have tried to take advantage of the particular biology of the tumor cells. However, many patients develop resistance over time or show no favorable response at all. Therefore, a clear understanding of the biochemical processes specific to cancer cells or which lead to malignant transformation in order to exploit them for therapeutical purposes is of utmost importance.
Our studies have uncovered two new important roles of Cyclin E in the cellular response to genotoxic stress caused by DNA-damaging agents. We have initially reported that in hematopoietic cells undergoing apoptosis induced by DNA-damaging agents,
Cyclin E generates an 18-kDa carboxy-terminal fragment, p18-Cyclin E (Mazumder et al., 2002). Then we have shown that p18-Cyclin E plays an important role in regulating the apoptotic pathway in the cytosol by binding Ku70 and releasing Bax (Mazumder et al., 2007b). Apart from its role in modulating the apoptotic response, p18-Cyclin E impairs the non-homologous end joining (NHEJ) DNA double-strand break repair pathway (Chapter II). Therefore, we have shown that Cyclin E, traditionally believed to be involved only in cell cycle regulation, has two other non-canonical roles, in regulating
145 146
apoptosis and DNA repair (Figure 4.1). Furthermore, we have characterized the
biochemical processes responsible for the turnover of p18-Cyclin E and how these can be
taken advantage of to sensitize tumor cells to therapy (Chapter III). These novel findings
extend the role of Cyclin E beyond the cell cycle control to include regulation of
apoptosis as well as that of NHEJ DNA repair.
As therapeutic resistance continues to be an important obstacle in cancer
treatment, one must consider the diverse pathways that tumor cells may utilize in
acquiring such survival capacity. Future therapeutical approach strategies may rely on
using p18-Cyclin E or a peptidic derivative to sensitize tumor cells to conventional
ionizing radiation or drug therapies.
Understanding the regulation of these three cellular processes, cell cycle, DNA
repair, and apoptosis in cancer cells would facilitate the development of targeted therapies which would result in tumor cell killing while protecting the normal cells.
147
Figure 4.1 Dual regulation of apoptosis and DNA repair by p18-Cyclin E. Genotoxic stress caused by DNA-damaging agents leads to a caspase-3-mediated cleavage of Cyclin
E generating its C-terminal fragment, p18-Cyclin E. This fragment binds Ku70 in the cytosol, probably inducing a change in its conformation which results in release and activation of Bax from its inhibitory association with Ku70. Bax translocates to the mitochondria, thus amplifying the apoptotic signal. On the other hand, p18-Cyclin E interacts with Ku70 in the nucleus impairing the NHEJ DNA repair. Both effects of p18-
Cyclin E lead to an amplification of the cell death signal committing the cell to apoptotic death.
148
Future directions
p18-Cyclin E affects DNA damage signaling and slows down the kinetic of DNA
repair
Formation of foci of different nuclear proteins is a widely used method for
assessing DNA double-strand breaks (DSB) formation and its repair. Among these are
53BP1, Nbs1, Rad51, and BRCA1 (Paull et al., 2000), with γ-H2AX being the most frequently used. Histone H2AX is phosphorylated in response to DNA damage on Ser139
(Rogakou et al., 1998) forming a nuclear focus. γ-H2AX foci serve as flags on the
chromatin for further DNA repair. This molecule links global chromatin changes and
accessory proteins with DNA repair (Redon et al., 2002). Quantification of γ-H2AX foci
has become a standard in the detection and quantification of the extent of DNA damage
following IR (Wang et al., 2005).
A very conclusive evidence of the influence of p18-Cyclin E on DNA repair
would be provided by showing that p18-Cyclin E slows down the kinetics of DNA repair
in vivo. This would be monitored by observing the appearance and disappearance of γ-
H2AX foci, as a measure of DNA damage and its repair. p18-Cyclin E stably expressing
293T cells would be irradiated or treated with etoposide and the presence of γ-H2AX would be determined by immunofluorescence or flow cytometry. If p18-Cyclin prevents the DNA repair, a persistence of the γ-H2AX foci in these cells as compared to control cells would be expected. A drawback in using the 293T cell system for this type of experiment could be the fact that the DNA damage signaling is not completely functional 149
in these cells due to the inactivation of p53 by the large T antigen. A different cellular system (e.g. normal diploid fibroblasts, epithelial cells or PBMC) in which the DNA damage signaling pathway is intact and which would also stably express p18-Cyclin E might be more indicated.
p18-Cyclin E impairs the recruitment of the XLF/XRCC4/Ligase IV heterocomplex to the Ku70/Ku80 heterodimer
In Chapter II I have shown that p18-Cyclin E impairs NHEJ DNA repair by preventing the recruitment of XLF/XRCC4/Ligase IV to the DNA repair complex at the damage site. However, recent studies have shown that XRCC4/Ligase IV can be recruited directly to Ku70/Ku80 in the absence DNA-PKcs (Costantini et al., 2007). Determining the ability of p18-Cyclin E to prevent the recruitment of XLF/XRCC4/Ligase IV using the experimental conditions used by Costantini et al. would provide conclusive mechanistic evidence regarding the influence of p18-Cyclin E in the final step of NHEJ. p18-Cyclin E stably expressing cells would be lysed according to the description in
Costantini et al. and Ku70/Ku80 would be immunoprecipitated followed by determination of Ligase IV and XRCC4 levels by immunoblotting. A reduction in the association between Ku70/Ku80 and XLF/XRCC4/Ligase IV would be expected. This experiment would determine unequivocally the mechanism by which p18-Cyclin E prevents NHEJ DNA repair.
150
Determination of p18-Cyclin E subcellular localization and its implications
Immunofluorescence studies showed that while Cyclin E localization is mainly in
the nucleus, where it exercises its well known role in cell cycle control and DNA
replication, p18-Cyclin E is both nuclear and cytosolic. Intriguingly, although present throughout the cytosol, preliminary data indicate that p18-Cyclin E is concentrated in peri-nuclear clusters resembling the Golgi apparatus (Figure 4.2). In contrast, a Cyclin
ER130A mutant that does not bind Cdk2 due to intramolecular destabilization localizes
mainly to the cytosol.
To determine whether, indeed, p18-Cyclin E localizes to the Golgi apparatus,
confocal microscopy will be used to examine the co-localization of p18-Cyclin E with
Golgin 97 (a unique protein specific to the Golgi apparatus) or a transfected Yellow
Fluorescence Protein bearing a Golgi localization signal. Furthermore, dynamics of the
p18-Cyclin E subcellular localization will be examined by time lapse confocal
microscopy using a p18-Cyclin E-GFP construct.
151
Figure 4.2 Localization of p18-Cyclin E-GFP, Cyclin E-GFP, and Cyclin ER130A-
GFP. Nuclei are stained with DAPI (blue). Nuclear and cytosolic localization of Cyclin E and Cyclin ER130A respectively, versus the Golgi-like formation localization of p18-
Cyclin E (green). Arrows indicate p18-Cyclin E clusters.
152
Characterize the mechanism of generation of p18-Cyclin E
The mechanism by which p18-Cyclin E is generated would be of particular interest as we have witnessed its appearance following treatment with DNA-damaging agents only in cells of hematopoietic origin. We examined the generation of p18-Cyclin E from Cyclin E in the NCI-H1299 non-small cell lung carcinoma cell line. A HA-Cyclin E stably expressing H1299 cell line was generated by transfection followed by selection with neomycin. These cells were either irradiated or treated with etoposide.
Unfortunately, in either case, the appearance of HA-p18-Cyclin E was not observed. The same experiment should be repeated in a hematopoietic cell line such as IM-9, Molt-4, or
Nalm-6, as it appears that due to yet unexplained mechanisms, p18-Cyclin E is generated only in cells of hematopoietic origin.
Another alternative of studying the circumstances in which p18-Cyclin E is being generated would be in a murine system. However, ionizing radiation and etoposide treatment failed to result in the generation of p18-Cyclin E. A careful analysis of the
Cyclin E amino-acid sequence led to the conclusion that the amino-acid residue where caspase-3-mediated cleavage is supposed to take place is mutated (Figure 4.3). A Cyclin
E sequence alignment across human, mouse, rat, rabbit, chicken, and frog indicated that only in human the LDVD sequence with the Asp275 (or 292 according to a longer isoform) residue is present. In all other species, the Asp275 residue is changed to Gly or
Cys. This explains why the generation of p18-Cyclin E was observed only in human cell lines. 153
Figure 4.3 Cyclin E sequence alignment across human, mouse, rat, rabbit, chicken,
and frog. The blue wire box identifies the Cyclin E caspase-3 cleavage site, LDVD. Only in humans, Cyclin E maintains the Asp275 (NM_05172 short isoform) or Asp292
(NM_001238 long isoform) residue, other species having this residues mutated to glycine. The sequences Homo sapiens (NM_001238), Mus musculus (NM_007633),
Rattus norvegicus (NM_001100821), Oryctolagus cunniculus (EU137106, partial sequence), Gallus gallus (NM_001031358), Xenopus laevis (NM_001087976) were aligned using ClustalW.
154
155
Furthermore, to test whether p18-Cyclin E is generated from the free or Cdk2-
bound Cyclin E, three expression constructs would be used: wt Cyclin E-HA, Cyclin
ED275A-HA (cleavage resistant mutant), and Cyclin ER130A-HA that does not bind Cdk2 due to intramolecular destabilization. Even though Cyclin E is known to shuttle from the
nucleus to the cytosol in a Crm1-independent manner (Jackman et al., 2002), its main
localization is in the nucleus, where it is imported after being synthesized and assembled with Cdk2 in the cytoplasm. Consequently, Cyclin ER130A is not expected to be retained in the nucleus and will be found mainly in the cytoplasm as also shown by immunocytochemistry (Figure 4.2). To determine from which pool of Cyclin E p18-
Cyclin E is generated, IM-9 cells stably expressing HA-Cyclin E or HA-Cyclin ER130A
will be irradiated and the generated fragments will be identified with anti-HA antibody. If
following IR, Cyclin E generates p18-Cyclin E and Cyclin ER130A does not generate it, p18-Cyclin E is most likely generated from cleavage of Cdk2-bound Cyclin E. If both
Cyclin E and Cyclin ER130A are able to generate p18-Cyclin E then the Cdk2-free pool of
Cyclin E is responsible for generating p18-Cyclin E. As negative control, the Cyclin E cleavage resistant mutant that is effective in blocking apoptosis (Mazumder et al., 2002) will be used.
The fact that the role of p18-Cyclin E is to commit the cell for apoptotic death by amplifying the apoptotic signal as well as by impairing DNA repair, warrants its use as a biomarker in determining the sensitivity of hematopoietic tumor cells to different chemotherapeutic approaches.
156
Study the Ku70/Hdm2-mediated p18-Cyclin E degradation
The ubiquitin E3 ligase activity of Hdm2, its recent shown interaction with Ku70
(Gama et al., 2008, unpublished) corroborated with the association between p18-Cyclin E and Ku70 raised the question of whether Hdm2 contributes to the degradation of p18-
Cyclin E. To unequivocally prove that Hdm2 is responsible for the ubiquitination of p18-
Cyclin E, Hdm2 will be either overexpressed or down-regulated using the siRNA technology and levels of p18-Cyclin E will be determined by immunoblotting.
Furthermore, nutlin, an inhibitor of Hdm2 will be used to block its activity in order to assess its effect on p18-Cyclin E (Wang and El-Deiry, 2008).
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APPENDIX
Abbreviations
53BP1 p53 binding protein 1
AIF apoptosis inducing factor
Apaf-1 apoptotic protease activating factor 1
APC anaphase promoting complex
ATM ataxia-telangiectasia mutated
ATP adenosine triphosphate
ATR ataxia-telangiectasia and rad3 related
Bad Bcl-2 antagonist of cell death
Bak Bcl-2 antagonist killer
Bax Bcl-2-associated X protein
Bcl-2 B-cell CLL/lymphoma 2
Bcl-xL, Bcl-2 like 1, long form
BH Bcl-2 homology domain
BH3 Bcl-2 homology domain 3
Bik Bcl-2 interacting killer
Bim Bcl-2-interacting protein
BIR baculoviral IAP repeats
193 194
bp base pair
BRCA1 breast cancer 1 gene
BRCT BRCA1 C-termini
BSA bovine serum albumin
CAD caspase-activated DNase
CARD caspase recruitment domain
Caspase cysteine aspartate protease
Caspase-3 apoptosis-related cysteine protease 3
CDK cyclin-dependent kinase
Cdk2 cyclin-dependent kinase 2
cDNA complimentary deoxyribonucleic acid
Cernnunos XLF, XRCC4-like factor
CKI cyclin-dependent kinase inhibitors
CIP CDK inhibitory protein
Cyt C cytochrome c
DED death effector domain
DD death domain
DHFR dihydrofolate reductase
DN dominant negative
DNA deoxyribonucleic acid
DNA-PKcs DNA protein kinase catalytic subunit
DP1/DP2 dimerization protein 1/2 195
DR death receptor
DSB double-strand break
E1A early region 1A (E1A) products of adenovirus 5 (Ad5)
E2F E2 factor, cellular factor required for E2 viral promoter activation
EDTA ethylenediaminotetraacetic acid
EF1α elongation factor 1 α
EGFP enhanced green fluorescence protein
FADD Fas associated protein with death domain
FLIP FLICE-inhibitory protein
G1 phase gap 1 phase of the cell cycle
G2 phase gap 2 phase of the cell cycle
GFP green fluorescence protein
GSK3β glycogen synthase kinase 3β
Gy gray (dose unit of IR)
HDAC histone deacetylase
Hdm2 human homologue of mouse double minute 2
HMG1 high mobility group 1
HR homologous recombination
IAP inhibitor of apoptosis protein
ICAD inhibitor of CAD
ICE interleukin-1beta converting enzyme
IgGλ immunoglobulin G light chain 196
IR ionizing radiation
kDa kilodalton
LMW low molecular weight
LT large T antigen
LV lentivirus
M phase mitosis phase of the cell cycle
Mcl-1 myeloid cell leukemia sequence 1
MCM minichromosome maintenance protein(s)
MEF mouse embryonic fibroblast
MEK mitogen-activated protein/ERK kinase
MMR mismatch repair
MPF maturation promoting factor or M-phase promoting factor
mRNA messenger ribonucleic acid
Nbs1 Nijmegen breakage syndrome, nibrin
NER nucleotide excision repair
NF-κB nuclear factor κB
NHEJ non-homologous end-joining repair
NOXA phorbol-12-myristate-13-acetate-induced protein 1
NP-40 Nonidet P-40, non-ionic detergent p21 cyclin-dependent kinase inhibitor-1A (CDKN1A) p27/Kip1 cyclin-dependent kinase inhibitor
p53 tumor suppressor protein p53 197
p107 retinoblastoma-like 1; RBL1 p130 retinoblastoma-like 2; RBL2
PAGE polyacrylamide gel electrophoresis
PARP-1 poly (ADP-ribose) polymerase-1
PBS phosphate buffered saline
PCR polymerase chain reaction
PMSF phenylmethylsulphonylfluoride
PUMA p53-upregulated modulator of apoptosis
PI propidium iodide
PI3K phosphatidylinositol 3-kinase
PS phosphatidylserine
R point restriction point
Rb retinoblastoma
RNA ribonucleic acid
ROS reactive oxygen species
S phase synthesis phase of the cell cycle
SCF Skp1-Cul1-F box or Skp1-Cul1-Fbw7
SD standard deviation
SDS-PAGE sodium dodecyl sulfate polyacrylamide gel electrophoresis siRNA small interference RNA
SSB single-strand break tBid truncated Bid 198
TK thymidine kinase
TM transmembrane region
TNF tumor necrosis factor
TNFR tumor necrosis factor receptor
TRAIL tumor necrosis factor-related apoptosis-inducing ligand tRNA transfer RNA
TS thymidilate synthase
VEGF vascular endothelial growth factor
XRCC4 X-ray repair complementing defective repair in Chinese hamster cells 4
XLF XRCC4-like factor, Cernnunos