CHAPTER I:
Introduction

the sections entitled 
“Insights into cyclin D1 overexpression and nuclear accumulation”
and
“Constitutively nuclear cyclin D1 drives genomic instability”
  have been published in part (including Figure 1.4/Table 1) in:

Pontano L.L. and Diehl J.A. (2008). 
Speeding through cell cycle roadblocks:
Nuclear cyclin D1-dependent kinase and neoplastic transformation.
Cell Division 3 (12).
				  (Pontano and Diehl, 2008)

	Cell cycle progression is intricately controlled by coordinated expression and ubiquitin-dependent degradation of key regulatory proteins. Temporally expressed mammalian cyclins serve as allosteric activators of cyclin dependent kinases (CDKs); active cyclin/CDK complexes catalyze phosphorylation events necessary for S-phase entry, DNA replication, and mitosis. CDK activity is regulated at each stage of the cell cycle, as labile cyclin proteins are targeted for proteasomal degradation by E3 ubiquitin ligases. The work presented herein focuses on mechanisms that control cell cycle-regulated accumulation of cyclin D1. Mitogen-dependent cyclin D1 expression and activation of CDK4 drives progression through G1 phase; critically, ubiquitin-mediated cyclin D1 proteolysis antagonizes nuclear cyclin D1/CDK4 activity following the G1/S transition. Cyclin D1 overexpression occurs frequently in human cancer and is thought to be a driving factor in neoplastic transformation. Recent evidence suggests that impaired protein degradation allows accumulation of active, nuclear cyclin D1/CDK4 kinase throughout the cell cycle; such aberrant nuclear cyclin D1/CDK4 activity in turn modulates neoplastic effects. Here, I will review cell cycle regulation by CDKs and ubiquitin ligases and discuss recent insights into cell cycle-dependent cyclin D1 regulation and mechanisms contributing to cyclin D1-driven tumorigenesis.

Temporal Cyclin/CDK activity drives cell proliferation. 
The concerted activity of multiple mammalian CDKs coordinates cell cycle progression, dependent on timely expression of corresponding activating cyclins and subsequent threonine phosphorylation within the CDK T-loop by the CDK activating-kinase (CAK), consisting of CDK7/Cyclin H (Russo et al., 1996). While CAK activity is thought to be constitutively active in eukaryotic cells, temporal CDK activation depends upon cyclin binding, which facilitates structural changes in the CDK allowing enhanced ATP catalysis (Jeffrey et al., 1995). The crystal structure of CDK2 alone, CDK2 in complex with cyclin A, and the cyclin A/CDK2/p27KIP1 ternary complex, provided valuable insight into the “molecular switch” function of CDK kinases in controlling cell cycle progression. For instance, mitogen-dependent cyclin induction in G1 phase coordinates growth signals with CDK activation. Alternatively, anti-proliferative signals such as senescence and DNA damage checkpoint activation lead to induction of cyclin dependent kinase inhibitors (CKIs) of the CIP/KIP and INK4 families, switching CDK activity off (Pavletich, 1999).
A large body of work established specific cyclin/CDK partnerships (Figure 1.1): G1-phase D-type cyclins and CDK4/6, S-phase cyclins E/A and CDK2, and G2/M phase cyclins A/B and CDK2/CDK1 (Aleem et al., 2005; Draetta et al., 1989; Sherr, 1994). Typically, cyclin D/CDK4/6 kinase activity promotes G1 progression by phosphorylating the retinoblastoma protein (Rb) and related pocket proteins p107/p130, relieving Rb-dependent E2F inhibition. Cyclin E/CDK2 also phosphorylates Rb in late G1; cyclin E mRNA levels rapidly increase at this time, and cyclin D/CDK4/6 complexes titrate the CIP/KIP class of CKIs from cyclin E/CDK2 complexes, facilitating activation. Rb hyperphosphorylation by site-specific, concerted CDK4 and CDK2 activity triggers complete Rb dissociation from E2F and commits the cell to S-phase entry (Connell-Crowley et al., 1997; Harbour et al., 1999; Weinberg, 1995). At this stage, Rb inactivation shifts from dependency upon mitogen signaling toward mitogen-independent cyclin E/CDK2 activity. Consequently, nuclear cyclin D/CDK4/6 activity is terminated by degradation of D-type cyclins (Figure 1.2). While distinct mechanisms governing cyclin D2 and D3 regulation are largely unknown, work over the past decade revealed that cyclin D1 is targeted for proteasomal degradation via phosphorylation-dependent nuclear export and cytoplasmic ubiquitylation (Diehl et al., 1998; Diehl et al., 1997).
As cells enter S-phase, CDK activity facilitates activation of replication origins and prevention of origin re-loading. This contradictory function is achieved through temporal CDK regulation; cyclin E/CDK2 activity in early S-phase triggers origin firing, while cyclin A/CDK2-dependent inactivation of pre-RCs occurs in late S- and G2-phases (Bell and Dutta, 2002). Early in S-phase, cyclin E is targeted for ubiquitin-mediated proteolysis (Clurman et al., 1996). Conversely, cyclin A is expressed in late G1, with maximal accumulation by late S/G2, where cyclin A/CDK2 activity regulates cell cycle progression in several ways. First, cyclin A/CDK2 activity terminates E2F-dependent gene transcription through phosphorylation of the DP-1 heterodimeric component of E2F transcription factors, precluding DNA binding (Sherr, 1996). Second, cyclin A/CDK2 terminates pre-RC assembly by phosphorylating CDC6, triggering its nuclear export in mammalian cells, and inhibiting MCM complex association with chromatin (Petersen et al., 1999; Petersen et al., 2000). Finally, cyclin A/CDK2 phosphorylates CDT1, targeting soluble CDT1 for SCFSKP2-dependent ubiquitylation. However, chromatin-bound CDT1 is degraded in a phosphorylation-independent, PCNA- and DNA-dependent manner by CUL4CDT2  (Havens and Walter, 2009; Nishitani et al., 2006; Vaziri et al., 2003).
In addition to S-phase functions, cyclin A/CDK2 regulates centrosome duplication and CDC25 phosphorylation in G2, and impaired cyclin A/CDK2 activity delays subsequent cyclin B/CDK1 activation (Baldin et al., 1997; De Boer et al., 2008; Meraldi et al., 1999). CDK1 activation requires dephosphorylation of T14 and Y15 at the G2/M boundary, when CDC25 dual specificity phosphatase activity exceeds inhibitory Wee1/Myt1-dependent CDK phosphorylation (Nigg, 2001). However, cyclin B/CDK1 is initially sequestered in the cytoplasm during S-phase and early G2, preventing pre-mature mitotic entry, followed by nuclear localization upon nuclear lamina breakdown at the G2/M transition (Pines and Hunter, 1991). Cyclin B phosphorylation within its cytoplasmic retention sequence is thought to facilitate nuclear accumulation. Although the distinct mechanism governing cyclin B/CDK1 nuclear entry remains elusive, recent work supports a role for coupling cyclin B1/CDK1 cytoplasmic activation to rapid nuclear import (Gavet and Pines; Yang et al., 1998). Cyclin B/CDK1 phosphorylates a plethora of substrates including the cytoskeleton, nuclear lamins, nucleolus, nuclear pore complexes, and the APC/C, leading to APC/C activation and cyclin destruction required for mitosis (Kraft et al., 2003). Clearly, cells have evolved intricate regulatory networks built upon post-translational modification of cyclins, CDKs, and inhibitory molecules necessary for temporal CDK activity and tight control of cell proliferation.

Functional redundancy among cell cycle-driving cyclin/CDK complexes.
A current model of CDK activity suggests that differential function of cyclin/CDK complexes is mainly controlled by spatiotemporal distribution during the cell cycle and less dependent upon specificity toward individual substrates (Moore et al., 2003). For instance, traditional models suggest that cyclin E/CDK2 functions in early S-phase, facilitating DNA replication, and cyclin A/CDK2 functions in late S-phase to inhibit origin relicensing. However, cyclin A/CDK2 can drive DNA replication in the absence of cyclin E, suggesting functional redundancy between these kinases (Rosenberg et al., 1995; Strausfeld et al., 1996). Furthermore, the idea that specific CDKs drive a single cell cycle event has been challenged by genetic studies, providing insight into the requirement for specific and redundant cyclin/CDK function during development. 
Genetic ablation of G1 CDKs revealed that mice develop beyond embryogenesis, with phenotypes observed in differentiated cells. CDK4-deficient mice are small and infertile; however, these mice develop diabetes, suggesting a requirement for CDK4 in -cell proliferation. Conversely, overexpression of a p16INK4A-refractory R24C CDK4 allele drives pancreatic hyperplasia, highlighting the importance for CDK regulation in this tissue (Rane et al., 1999). CDK4-deficient murine embryonic fibroblasts (MEFs) cycle normally under growth-stimulating conditions, yet quiescent cells exhibit a substantial S-phase entry delay, which is rescued by loss of p27. Compound deletion of CDK4 and CDK6 is embryonic lethal, as embryos exhibit severe anemia due to hematopoietic proliferative defects. These findings signify the importance of CDK4-dependent sequestration of p27, contributing to cyclin E activation (Malumbres et al., 2004; Tsutsui et al., 1999). Finally, CDK2 is required for meiotic cell division, but is dispensable in cycling mitotic cells, suggesting redundancy with CDK1 function (Ortega et al., 2003).
While CDK4 and CDK2 deletion independently results in normal embryogenesis, compound loss of both CDKs is embryonic lethal due to heart defects, and MEFs exhibit reduced Rb phosphorylation and E2F gene activation, consistent with impaired S-phase entry (Berthet et al., 2006). In contrast, mice lacking both CDK6 and CDK2 develop normally, with phenotypic abnormalities consistent with each individual knockout (Malumbres et al., 2004). Given these findings, it is quite interesting that exclusive expression of CDK1 is sufficient for mammalian cell cycle progression. Mice expressing CDK1 alone develop to mid-gestation, and MEFs proliferate in culture, despite prolonged cycle duration associated with inefficient Rb inactivation. CDK1 ablation results in embryonic lethality prior to the morula stage, suggesting that CDK1 can drive all cell cycle transition events required for cell division and early embryonic development (Santamaria et al., 2007). Further supporting this notion, knocking in CDK2 at the endogenous CDK1 locus failed to rescue CDK1 deficiency (Satyanarayana et al., 2008).
Similar to genetic CDK models, various individual and combinatorial cyclin knockouts revealed significant redundancy in cell cycle regulation. Knockout mouse models of each respective D-type cyclin demonstrated that mice develop normally, without severe proliferative defects. However, each model exhibited moderate, tissue specific defects, suggesting a non-redundant pattern of expression in various cell types. Tissue-specific phenotypes in cyclin D1-deficient mice include small size, retinal hypoplasia, neurological defects, and defective mammary gland development (Fantl et al., 1995; Sicinski et al., 1995). In contrast, cyclin D2-null animals exhibit defects in B-cell proliferation, -cell proliferation, and cerebellar development, and cyclin D3-null mice show impaired T-cell development (Huard et al., 1999; Kushner et al., 2005; Sicinska et al., 2003; Solvason et al., 2000). Together, these findings established a context-dependent role for the D-type cyclins in driving cell proliferation. 
Mice expressing a single D-type cyclin or lacking all three cyclins survive to mid-stages of embryogenesis. Mice lacking cyclin D1/D2 survive postnatally, exhibiting phenotypes consistent with each respective knockout. However, loss of cyclin D1/D3 and cyclin D2/D3 is lethal postnatally or in late embryogenesis due to neurological defects and severe anemia, respectively, suggesting that a single D-type cyclin is insufficient for tissue maturation and adult development (Ciemerych et al., 2002; Kozar et al., 2004). Consistent with the D-type cyclins, individual deletion of cyclin E1 or E2 does not impair viability or development, yet compound deletion results in embryonic lethality due to placental defects. Double knockout MEFs exhibit delayed S-phase progression, but both cyclin D/A can compensate for cyclin E function at the G1/S transition (Geng et al., 2003; Parisi et al., 2003). Additionally, loss of ubiquitously expressed cyclin A2 results in mortality after implantation (Murphy et al., 1997). Recent development of cyclin A2 conditional knockout mice revealed that this cyclin is essential for hematopoietic and embryonic stem cell proliferation (Kalaszczynska et al., 2009), supporting a non-redundant function that cannot be compensated by cyclin E/B. Finally, ablation of cyclin B1 results in embryonic lethality, while loss of cyclin B2 has no gross effects, suggesting redundancy among these cyclins, but a requisite role for cyclin B1 in mitotic function (Brandeis et al., 1998).
Genetic cyclin/CDK manipulation in vivo provided novel insight into how these kinases can be exploited to modulate cell cycle progression. Importantly, deregulation of G1 and S-phase CDKs and respective CKIs has been implicated in the genesis of human cancer. CDK4 overexpression due to gene amplification and less commonly mutation has been well documented, and loss or silencing of the INK4 locus occurs in a vast array of tumors (Sherr, 1996). Furthermore, cyclin D1 is frequently targeted in human cancer, driving tumorigenesis through activation of CDK4. For instance, knock-in mice harboring cyclin D1K112E (replacement of lysine with glutamic acid) alleles exhibit similar neurological defects and small size observed in cyclin D1-/- mice, and more importantly, these mice are refractory to Neu-mediated mammary tumorigenesis. Cyclin D1K112E binds CDK4 and p27KIP1 with normal affinity, but cannot facilitate CDK activation (Landis et al., 2006).  Interestingly, retinal and mammary gland development is unaltered in cyclin D1K112E mice, consistent with evidence suggesting that p27 is epistatic to cyclin D1 in these tissues (Geng et al., 2001; Landis et al., 2006). Additionally, overexpression of cyclin E occurs frequently in cancer, implicating the cyclin E/CDK2 kinase in cell cycle deregulation as well (Hwang and Clurman, 2005). Collectively, the large body of work describing CDK deregulation in cancer suggests that alteration of a single cyclin/CDK module is sufficient to promote aberrant cell growth, and subsequent loss of checkpoint constraints facilitates neoplastic transformation. 
Given the neoplastic effects of sustained CDK activity, it is not surprising that cells harbor multiple mechanisms to keep CDK activity in check. Intricate checkpoint mechanisms limit CDK activity following genotoxic insult and various other stresses, including induction of CKIs such as p21CIP1, degradation of cyclins, and inactivation of CDC25 phosphatases required for CDK activation. Following DNA damage, members of the PI(3)K (phosphatidylinositol-3-OH kinase)-like kinase (PIKK) family respond to double strand breaks (DSBs) and single strand DNA (ssDNA) and signal directly and through effector kinases, triggering p53 stabilization and p21CIP1 induction (Kastan and Bartek, 2004). These signals induce rapid cell cycle arrest in G1 phase. In S-phase and G2, signaling through PIKKs and downstream effectors also mediates phosphorylation of the CDC25 phosphatases; phosphorylated CDC25A is targeted for degradation by the SCF-TrCP E3 ubiquitin ligase, and CDC25B/C phosphorylation promotes 14-3-3 binding and nuclear exclusion. Additionally, the DNA damage effector kinase CHK1 activates WEE1, the kinase responsible for inhibitory phosphorylation on CDK2 and CDK1 (Reinhardt and Yaffe, 2009). Ultimately, tightly controlled CDK activity is achieved by coupling signaling events with E3 ubiquitin ligase function to maintain cell homeostasis.

E3 ubiquitin ligases control accumulation of cell cycle regulatory proteins.
Cell cycle progression is driven by alternating phases of cyclin accumulation and destruction via the ubiquitin-proteasome machinery, respectively (Evans et al., 1983; Glotzer et al., 1991). Ubiquitylation of target substrates requires the concerted activity of an ubiquitin activating enzyme (E1), ubiquitin conjugating enzyme (E2), and ubiquitin ligase (E3). Although certain E3 ligases, such as the HECT-domain family, directly transfer ubiquitin, many E3 ligases function as multi-subunit complexes that bridge the ubiquitin charged E2 enzyme with a given substrate protein (Ciechanover, 1994; Hershko and Ciechanover, 1998). Covalent, posttranslational ubiquitin linkages serve many cellular functions including proteasomal targeting, signaling, endocytosis, and protein interactions. Importantly, lysine-48 polyubiquitin conjugation targets substrate molecules for proteasomal degradation (Winget and Mayor).
Biphasic activity of two major E3 ligases, the SKP1-CUL1-F-Box (SCF) family and the anaphase promoting complex/cyclosome (APC/C), controls temporal cell cycle progression by targeting cyclins, CKIs, and other cell cycle regulators for proteasomal degradation. The SCF family of E3 ubiquitin ligases promotes polyubiquitylation of mainly phosphorylated substrates, including key regulators of G1 progression (Skowyra et al., 1997). The core cullin subunit, CUL1, functions as a scaffold for the RING-domain protein RBX1, required for E2 recruitment, and the adapter protein SKP1, required for substrate receptor recruitment. The SKP1 adapter associates with a given F-box protein, the specificity factor for substrate recognition (Figure 1.3A); over 70 F-boxes have been described in human cells, making the diversity of substrate recognition by SCF ligases quite vast (Petroski and Deshaies, 2005). SCF ligases function throughout the cell cycle and are regulated by cullin neddylation; this modification structurally alters the c-terminal RING binding domain of the cullin, increasing the processivity of ubiquitin transfer from the E2 to the substrate (Saha and Deshaies, 2008). The APC/C, on the other hand, is a modular ligase that promotes substrate degradation at the G2/M and M/G1 transitions. The APC/C consists of the APC11 RING domain protein, APC2 cullin homolog, two activating subunits, CDC20 and CDH1, and various scaffolding subunits (Harper et al., 2002; Petroski and Deshaies, 2005).
Ubiquitin-mediated destruction of inhibitory proteins ensures timely cell cycle progression; similarly, ubiquitylation and degradation of cyclins provides restrictive control necessary for DNA replication fidelity and mitosis. The APC/C coordinates mitotic progression and establishment of the next ensuing cell cycle by employing two respective substrate recognition adapters, CDC20 and CDH1. APC/CCDC20 controls the G2/M transition, degrading mitotic substrates such as cyclin A, cyclin B, and securin (Peters, 2006; Yu, 2007). APC/CCDH1 also assists APC/CCDC20 with mitotic exit, but functions mainly in G0/G1, targeting substrates including the CDK activator CDC25A, F-box protein Skp2, and components of the DNA replication machinery for degradation, thereby preventing premature cell cycle re-initiation (Manchado et al.). CDC20 and CDH1 recognize distinct domains within substrates called destruction box domains (D-box, amino acid sequence R-X-X-L-X-X-X-X-N), and CDH1 binds both D-box and KEN box domains (amino acid sequence K-E-N-X-X-X-E/D/N). This differential substrate recognition pattern is important as cells exit mitosis, since CDC20 levels decline due to recognition of its KEN box by APC/CCDH1 (Pfleger and Kirschner, 2000). While APC/CCDH1 regulates mitotic activity of APC/CCDC20, mitogen-dependent CDK activity in G1 promotes CDH1 phosphorylation and inactivation, and S-phase induction of the APC/C inhibitor EMI1 prevents premature degradation of APC/C substrates (Miller et al., 2006; Sorensen et al., 2001).
As APC/C activity decreases in G1, CDK activity increases, promoting replication licensing and E2F-dependent gene transcription (Figure 1.2). At this time, SCF E3 ligases come into play as cell cycle regulators, with F-boxes directing substrate recognition. F-box adapters are classified by conserved structural motifs that facilitate substrate binding. The “Fbw” class harbors WD40-repeat domains, the “Fbl” class harbors leucine-rich repeats, and the “Fbx” class lacks these aforementioned domains (Cenciarelli et al., 1999). Several SCF-like E3 ligases also control accumulation of cell cycle regulatory proteins. The CUL7-based ligase shares structural similarity with canonical SCFs, since it contains RBX1, SKP1, and Fbw8 (Figure 1.3B). This ligase has been implicated in insulin receptor substrate-1 degradation, and previous work suggested a role for SCFFbw8 and CUL7-Fbw8 ligases in cyclin D1 degradation (Okabe et al., 2006; Xu et al., 2008). A second SCF-like E3 ligase consists of the cullin scaffold CUL4, RBX1, DDB1 adapter, and various DDB1- and CUL4-associated factors (DCAFs), which serve as substrate-specific receptors (Figure 1.3B, C). Similar to the “FBW” class of F-box proteins, DCAF substrate recognition is achieved through substrate binding to WD40 repeats in the DCAF. The CUL4 ligase regulates multiple signaling pathways, including DNA replication, repair, and transcription; importantly, CUL4CDT2 prevents DNA re-replication by targeting CDT1 for degradation (O'Connell and Harper, 2007).
As cells proceed into G1, D-type cyclin expression drives CDK activation necessary for G1 progression. As described previously, cyclin D1/CDK4 kinase activity is not necessary following the G1/S transition; recent work revealed that SCFFbx4 targets phosphorylated cyclin D1 for proteasomal degradation. Cyclin D1 recognition requires Fbx4 in concert with the co-factor αB crystallin, a function analogous with the F-box protein Skp2 and co-factor Cks1, which target the CKI p27KIP1 for degradation (Figure 1.2) (Lin et al., 2006). Similar to Fbw7, an ubiquitin ligase for cyclin E, c-Myc, and Notch (Tang et al., 2007; Zhang and Koepp, 2006) and -TrCP, an E3 ligase for IB (Suzuki et al., 2000), Fbx4 undergoes dimerization; however, in contrast to either Fbw7 or -TrCP, Fbx4 dimerization occurs in a cell cycle-dependent manner, and requires GSK3-dependent phosphorylation at serine 12, which creates a 14-3-3 consensus site. 14-3-3 binding facilitates Fbx4 dimerizaton and ligase activation (Figure 1.2B) (Barbash et al.; Barbash et al., 2008). In contrast to other well-characterized F-box proteins, Fbx4 recognizes both phosphorylated and non-phosphorylated substrates, cyclin D1 and TRF1, respectively, and recent crystallographic evidence revealed that substrate binding occurs within a non-characteristic GTPase-like domain within its c-terminus (Zeng et al., 2010).
While SCF activity switches cyclin D1/CDK4 activity off following the G1/S transition, it turns on CDK2 and CDK1 activity in S- and G2-phase by regulating CKI accumulation. SCFSkp2 targets p27KIP1 and p21CIP1 for proteasomal degradation; p27 phosphorylation on T187 by CDKs facilitates recognition by this E3 ligase (Bornstein et al., 2003; Carrano et al., 1999; Tsvetkov et al., 1999). As cells begin DNA replication, cyclin E/CDK2 activity is no longer essential, and like cyclin D1, phosphorylated cyclin E is targeted for proteasomal degradation. Initial studies suggested that CDK2-dependent phosphorylation of T380 triggered cyclin E recognition by Fbw7, and subsequent work revealed that concerted GSK3-dependent phosphorylation of T380 and CDK2-dependent phosphorylation of S384 creates a phosphodegron motif that targets cyclin E for Fbw7-mediated degradation (Koepp et al., 2001; Welcker et al., 2003). More recently, biochemical analysis of the Fbw7 substrate-binding domain revealed critical arginine residues within the WD40 repeats that facilitate formation of the phosphodegron-binding pocket, and a dimerization domain proximal to the F-box facilitates Fbw7 dimerization, increasing the processivity of substrate ubiquitylation (Hao et al., 2007; Welcker et al.).
Although the remaining S- and G2/M phase cyclins are APC/C substrates, SCF ligases control CDK2 and CDK1 activity in response to various cellular cues. SCF-TrCP functions as an integrator of signaling pathways and tightly controls CDK function. -TrCP-dependent degradation of CDC25A following DNA damage prevents CDK2 activity until the checkpoint is resolved (Busino et al., 2003; Jin et al., 2003). -TrCP also negatively regulates CDK activity by targeting the APC/C inhibitor EMI1 for degradation in mitosis, leading to APC/C-mediated cyclin degradation (Guardavaccaro et al., 2003; Margottin-Goguet et al., 2003). In contrast, -TrCP facilitates mitotic CDK activation by targeting the CDK inhibitory kinase WEE1 for degradation, and -TrCP promotes CDK activation upon termination of checkpoint signaling by degrading the checkpoint mediator Claspin (Mailand et al., 2006; Peschiaroli et al., 2006; Watanabe et al., 2005; Watanabe et al., 2004). Like Fbw7, -TrCP recognizes conserved phosphodegron motifs (DpSGXXpS) containing two phosphorylated serine residues. -TrCP also undergoes dimerization, likely increasing ligase activity (Tang et al., 2007; Winston et al., 1999b). In addition to regulating CDK activity, -TrCP integrates growth and stress signaling with cell responses by targeting IB and -catenin for degradation, facilitating NFB-dependent activation of inflammatory and pro-survival genes and antagonizing pro-proliferative Wnt signaling, respectively (Winston et al., 1999b).
While a large number of F-box proteins have been identified, few have been well characterized (Fbw7, -TrCP, Skp2, and more recently Fbx4), highlighting the necessity to elucidate substrates and pathways regulated by novel SCF ligases. Furthermore, it is likely that most F-box proteins regulate multiple substrates, with recognition dictated by structural determinants within the F-box protein, and/or posttranslational substrate modifications. Given the complexity of SCF-mediated ubiquitylation in response to various cell stimuli, it is likely that activities of multiple ligases can perform synergistic or antagonistic functions regulating protein stability. This concept has been illustrated by recent work suggesting that -TrCP-mediated c-Myc N-terminal ubiquitylation antagonizes Fbw7-mediated c-Myc destruction (Popov et al., 2011). Furthermore, SCF ligase function is an attractive target for deregulation during neoplastic transformation, as accumulation of cell cycle drivers and survival proteins and downregulation of CKIs and pro-apoptotic proteins provide a growth and survival advantage for tumor cells.

Deregulation of SCF E3 ligase activity promotes neoplastic transformation.
Considering that SCF ligases control accumulation of both oncogenes and tumor suppressor proteins, alterations in ligase activity can elicit detrimental consequences for the cell. Unrestricted SCFSkp2 activity, for example, is oncogenic, since this ligase targets tumor suppressor proteins such as p27KIP1 and FOXO1 for degradation (Carrano et al., 1999; Huang et al., 2005). Interestingly, it has been proposed that Skp2 mediates cyclin D1 degradation (Yu et al., 1998); however, direct ubiquitylation has not been demonstrated (Nakayama et al., 2000). A likely explanation for cyclin D1 accumulation in Skp2-/- cells is that stabilization of the Skp2 substrate p27 enhances cyclin D1 stability as ternary cyclin D1/CDK4/p27 complexes are formed. Additionally, Skp2-/-; p27-/- mice overcome phenotypes associated with Skp2 deficiency alone, suggesting that p27 is the critical substrate of this ligase (Kossatz et al., 2004). Consistent with this notion, mice lacking p27 display organ hyperplasia and develop pituitary tumors, and p27 is haploinsufficient for tumor suppression (Fero et al., 1998; Fero et al., 1996; Nakayama et al., 1996). Furthermore, other Skp2 substrates have alternative means of degradation; for example, p21 is degraded by APC/CCDC20, and chromatin-bound CDT1 is degraded by CUL4CDT2 (Amador et al., 2007; Nishitani et al., 2006).
Similar to SCFSkp2, SCF-TrCP targets tumor suppressor proteins for degradation, including IB and PDCD4, a protein translation inhibitory protein (Dorrello et al., 2006; Winston et al., 1999b). Aberrant IB degradation facilitates unrestricted NFB-dependent transcription of inflammatory proteins, pro-survival factors, signaling receptors, and adhesion molecules that promote tumor initiation and progression, and NFB activation is a hallmark of inflammation-associated tumors (Karin and Greten, 2005). Similarly, PDCD4 restricts cell growth and survival by inhibiting the translation factor eIF4A. PDCD4 is typically degraded in response to mTOR activation and S6 kinase function, and importantly, loss of PDCD4 is thought to promote cell transformation. PDCD4 protein levels are downregulated in hepatocellular carcinomas, and loss of PDCD4 is associated with poor prognosis in lung cancers (Chen et al., 2003; Dorrello et al., 2006; Zhang et al., 2006). Increased -TrCP expression has been documented in various cancers including colon cancer and melanoma, and many of these tumors exhibit positive NFB-dependent gene expression signatures, correlating with -TrCP-dependent inactivation of IB (Liu et al., 2007; Ougolkov et al., 2004). Furthermore, genetic studies revealed that loss of -TrCP confers a hypoplastic mammary gland phenotype, while transgenic -TrCP expression drives mammary epithelial cell proliferation, with a significant fraction of mice developing mammary, uterine, and ovarian carcinomas (Kudo et al., 2004). 
In contrast to Skp2 and -TrCP, Fbw7 functions as a tumor suppressor, as it controls accumulation of several cell cycle proteins and transcription factors deemed to be oncogenic. It has been established that Fbw7 targets cyclin E for degradation, and more recent work defined a large set of transcription factors as Fbw7 targets, including c-Myc, Notch, c-Jun, and SREBP (Inuzuka et al.; Nateri et al., 2004; Oberg et al., 2001; Sundqvist et al., 2005; Welcker and Clurman, 2008; Welcker et al., 2004). Mechanistic analysis of Fbw7 deregulation in cancer revealed mutation of critical arginine residues within the substrate phosphodegron binding pocket, as well as mutations in substrates that preclude Fbw7 binding. For example, the c-Myc oncogene is commonly mutated in the region surrounding T58 required for Fbw7 recognition in Burkitt lymphomas, leading to increased stability and tumor formation (Gregory and Hann, 2000). Fbw7 was initially thought to be a haploinsufficient tumor suppressor, given the frequency of hemizygous mutations in various human cancers including T-ALL; however, genetic studies revealed that Fbw7+/- mice develop normally, and deletion of one allele of Fbw7 in the lymphocyte compartment does not drive lymphomagenesis (Onoyama et al., 2007; Tetzlaff et al., 2004). Subsequent work investigating the effects of Fbw7 mutation and dimerization revealed that a majority of Fbw7 mutations confer dominant-negative character, thereby limiting ligase function (Welcker and Clurman, 2008). 
Similar to Fbw7, recent evidence suggests that Fbx4 is a tumor suppressor, as hemizygous Fbx4 mutations in human cancer impair ligase activity and lead to aberrant nuclear cyclin D1 accumulation (Barbash et al., 2008). Importantly, the frequent incidence of N-terminal mutations that impair ligase dimerization and full activity hints at the fact that Fbx4 could function as a haploinsufficient tumor suppressor. In the following section, I will elaborate on these findings and discuss recent work elucidating oncogenic properties of cyclin D1 when SCFFbx4-mediated degradation is disrupted. 

Insights into cyclin D1 overexpression and nuclear accumulation in cancer.
As described previously, nuclear cyclin D1 accumulation is antagonized by phosphorylation-dependent nuclear export and cytoplasmic degradation following the G1/S transition. Cyclin D1 is rapidly phosphorylated by GSK3 on T286, triggering CRM1-dependent nuclear export, and SCFFbx4-dependent proteasomal degradation (Diehl et al., 1998; Lin et al., 2006). Activation of SCFFbx4 ligase activity at this time also facilitates cyclin D1 destruction. Collectively, multiple levels of cyclin D1 regulation including nuclear export, increased ubiquitylation capacity, and degradation ensure that this cell cycle driver does not accumulate or re-enter the nucleus during S-phase, limiting its oncogenic potential (Figure 1.2).
Paradoxically, cyclin D1 overexpression is considered a causative factor in many tumor types, yet overexpression of wild type cyclin D1 is insufficient to drive neoplastic transformation. However, cyclin D1 mutants refractory to phosphorylation and subsequent proteasomal degradation are acutely transforming in vitro and in vivo, implying that compartmentalization of cyclin D1/CDK4 is essential for cell homeostasis (Alt et al., 2000; Gladden et al., 2006). Cyclin D1 overexpression occurs in carcinomas of the breast, esophagus, head and neck, and lung. In a majority of cases, alterations in gene expression cannot account for overexpression, suggesting that impaired degradation contributes to cyclin D1 stability. 
Mutations that inhibit cyclin D1 T286 phosphorylation are rare, occurring to date only in endometrial and esophageal cancer. Single-base substitutions changing P287 to a serine or threonine residue in endometrial carcinoma correlates with overexpression of cyclin D1 in the nucleus of neoplastic cells, and a 12-base pair in frame deletion of amino acids 289-292 was reported with overexpression of cyclin D1 (Moreno-Bueno et al., 2003). Subsequent analyses revealed that disruption of P287 abrogates GSK3β-dependent phosphorylation of T286, and deletion of residues 289-292 impairs cyclin D1 binding to CRM1, both resulting in nuclear accumulation (Benzeno and Diehl, 2004; Benzeno et al., 2006). In accordance with endometrial cancer, recently identified cyclin D1 mutations in esophageal cancer and tumor-derived cell lines also disrupt T286 phosphorylation. Cyclin D1 sequence analysis in a panel of 90 esophageal carcinomas revealed mutation of T286 to arginine and a deletion of carboxy-terminal residues 266-295, and screening of human tumor-derived esophageal carcinoma cell lines also identified a P287 to alanine mutation in three of these lines (Benzeno et al., 2006).
Given the low frequency of cyclin D1 mutations that directly impact its turnover, a natural prediction is the occurrence of inactivating mutations in its E3 ligase.  The first clue to cyclin D1 ligase involvement came from analysis of breast cancer cell lines exhibiting increased cyclin D1 half-life without mutations disrupting phosphorylation. These analyses revealed that MCF-7 and MDA-MB-231 cells lack αB crystallin expression as a consequence of chromosome 11 deletion, and tumor microarray analysis of esophageal carcinomas revealed a reduction in both αB crystallin and Fbx4 mRNA levels in tumor tissues (Lin et al., 2006). Significantly, recent assessment of the FBX4 sequence in 116 esophageal carcinoma samples identified hemizygous missense mutations in 14 percent of tumors. Many of these mutations reside within the FBX4 dimerization domain, while others target S12 in the N-terminus. Additionally, one mutation was identified within the F box, which could result in production of a dominant negative protein incapable of recruiting SKP1 and CUL1 (Barbash et al., 2008). Cyclin D1 and Fbx4 alterations in human cancer are summarized in Figure 1.4 and Table 1.
The Fbx4 residues targeted in cancer are consistent with data revealing that Fbx4 dimerization following S12 phosphorylation and 14-3-3 binding is important for ligase function, as described previously. Fbx4 phosphorylation correlates with low cyclin D1 expression during G2/M and early G1, with a marked decrease during cell cycle entry due to growth factor-dependent GSK3β inhibition (Cross et al., 1995). Fbx4 phosphorylation increases again at the G1/S boundary as GSK3β becomes active and temporally controls ligase activation and cyclin D1 phosphorylation. Consequently, cyclin D1 phosphorylation is intimately tied to mitogen signaling and ligase activation, with GSK3β at the apex of the regulatory pathway. Thus far, the tumorigenic potential of Fbx4 mutations has only been examined in vitro, revealing that such mutations are indeed transforming (Barbash et al., 2008). However, the role of Fbx4 as a tumor suppressor has not been elucidated in vivo. Work presented in chapter 3 of this thesis describes the first genetic study to evaluate the relevance of Fbx4 function and provides substantial evidence that Fbx4 is, in fact, a tumor suppressor in vivo. 

Constitutively nuclear cyclin D1 drives genomic instability.
Nuclear accumulation of cyclin D1 resulting from impaired degradation drives cellular transformation in vitro and B-cell lymphomagenesis in mice expressing cyclin D1T286A driven by the Eμ enhancer (Alt et al., 2000; Gladden et al., 2006). Furthermore, recent evidence linked nuclear retention of active cyclin D1/CDK4 complexes with genomic instability. Nuclear cyclin D1/CDK4 activity in S-phase promotes transcriptional repression of CUL4A and CUL4B, leading to stabilization of CDT1, the pre-RC component that promotes loading of the replicative helicase during G1, thereby facilitating maintenance of the MCM helicase on chromatin and subsequent DNA re-replication. DNA re-replication, in turn, promotes accumulation of DNA damage and activation of DNA damage checkpoint signaling. Checkpoint activation promotes p53-dependent apoptosis initially, but provides a selective pressure for p53 functional inactivation, resulting in genomic instability and neoplasia (Aggarwal et al., 2007). 
Strikingly, impaired cyclin D1 ligase function results in nuclear accumulation of active cyclin D1/CDK4 complexes, triggering cellular transformation analogous to cyclin D1T286A (Barbash et al., 2008). Taken together, data elucidating the role of nuclear cyclin D1 in neoplastic transformation support a model wherein disruption of cyclin D1 phosphorylation or SCFFbx4 function generate genomic instability, ultimately driving tumor formation (Figure 1.4). While it is clear that nuclear cyclin D1/CDK4 kinase promotes CUL4 repression in S-phase, the precise mechanism underlying this event remained elusive. Importantly, recent work identified a novel regulatory relationship between the protein arginine methyltransferase 5/Methylosome protein 50 (PRMT5/MEP50) methyltransferase complex and nuclear cyclin D1/CDK4. Briefly, nuclear cyclin D1/CDK4 kinase phosphorylates MEP50, triggering increased intrinsic methyltransferase activity of the PRMT5/MEP50 complex and subsequent histone methylation at the CUL4 promoters, leading to transcriptional repression, CDT1 stabilization, and DNA re-replication (Aggarwal et al.). A mechanistic description and biological implications of this process are presented in chapter 4 of this thesis.
Genomic instability is a hallmark of human cancer, implicated in initiation and promotion of tumorigenesis, and chronic DNA damage checkpoint activation provides selective pressure for deletion or mutation of tumor suppressors such as p53 (Bartkova J, 2005; Gorgoulis VG, 2005). DSB induction following genotoxic insults such as γ-irradiation (γIR) and stalled replication forks during normal replication activate the intra-S and DNA replication checkpoints, respectively, thereby maintaining replication fidelity (Figure 1.5A). PIKK kinases ATM (Ataxia Telangiectasia, Mutated) and ATR (ATM and Rad3- Related) respond to DSBs and ssDNA, respectively, and signal directly and through downstream effectors such as CHK1 and CHK2 to promote cell cycle arrest and DNA repair (Kastan and Bartek, 2004). 
Previous work implicated APC/C-mediated cyclin D1 destruction following DNA damage in promoting p53-independent G1-phase cell cycle arrest (Agami and Bernards, 2000). Additionally, recent work suggested that ATM-dependent stabilization of the F-box protein Fbxo31 promotes SCFFbxo31-mediated degradation of phosphorylated cyclin D1 in G1-phase. Interestingly, this work suggested that ERK-dependent cyclin D1 phosphorylation promotes cyclin D1 recognition by Fbxo31 in melanoma cells (Santra et al., 2009). Given the critical role for GSK3- and Fbx4-dependent cyclin D1 regulation following the G1/S transition and ensuing DNA damage that occurs when cyclin D1 is aberrantly maintained in the nucleus during S-phase, we proposed that cyclin D1 is regulated by DNA damage following the G1/S transition, dependent upon Fbx4 (Figure 1.5B), (Pontano et al., 2008). The work presented in chapter 2 of this thesis supports this model, demonstrating that rapid cyclin D1 destruction is essential to maintain genome integrity following S-phase DNA damage. 
Collectively, work over the past two decades established a model wherein temporal CDK activity drives cell cycle progression, and tight control of CDK activators and inhibitors by ubiquitin-mediated proteolysis keeps CDK activity in check. Critically, disrupting this balance promotes aberrant CDK activation, an underlying event in neoplastic transformation. This has been well demonstrated by cyclin D1/CDK4, as impaired cyclin D1 degradation facilitates nuclear accumulation of active cyclin D1/CDK4 kinase beyond the G1/S transition, thereby disrupting DNA replication fidelity and promoting genomic instability. Although previous studies described this oncogenic function of nuclear cyclin D1, the work presented in the next chapters further defines an intricate relationship where aberrant nuclear cyclin D1/CDK4 activity modulates genetic alterations necessary to promote genomic instability and neoplastic transformation.

Figure 1.1.

Figure 1.2.

Figure 1.3.

Figure 1.4.

Table 1

Figure 1.5.

CHAPTER II:
Genotoxic stress-induced Cyclin D1 phosphorylation and proteolysis are required for genomic stability 

this chapter has been published in:
Molecular and Cellular Biology 28 (23): 7245-7258. December 2008. 
Reprinted by permission from American Society of Microbiology
License Number: 2661040473670
Summary
While mitogenic induction of cyclin D1 contributes to cell cycle progression, ubiquitin-mediated proteolysis buffers this accumulation and prevents aberrant proliferation. Because failure to degrade cyclin D1 during S-phase triggers DNA re-replication, we investigated regulation of cyclin D1 following genotoxic stress. These data reveal that expression of cyclin D1 alleles refractory to phosphorylation- and ubiquitin mediated degradation increase the frequency of chromatid breaks following DNA damage. Double strand break-dependent cyclin D1 degradation requires ATM and GSK3, which in turn mediate cyclin D1 phosphorylation. Phosphorylated cyclin D1 is targeted for proteasomal degradation by SCFFbx4.  Loss of Fbx4-dependent degradation triggers radio-resistant DNA synthesis, thereby sensitizing cells to S-phase-specific chemotherapeutic intervention. These data suggest that failure to degrade cyclin D1 compromises the intra-S-phase checkpoint and suggest that cyclin D1 degradation is a vital cellular response necessary to prevent genomic instability following genotoxic insult.

Introduction
Mitogenic signals coordinated by the small GTP binding protein Ras induce D-type cyclin expression during G1 phase of the cell cycle. Active cyclin D1/CDK4 complexes accumulate in the nucleus, triggering G1 progression via phosphorylation-dependent inactivation of the retinoblastoma protein (Rb) and related pocket proteins (Calbo et al., 2002; Harbour et al., 1999; Hatakeyama et al., 1994; Leng et al., 2002).  The cyclin D1/CDK4 kinase also facilitates activation of cyclin E/CDK2 through titration of the CDK inhibitors p21CIP1 and p27KIP1, contributing to Rb phosphorylation by the cyclin E/CDK2 kinase (Sherr, 1996).    
After the G1/S transition, cyclin D1 activation is terminated by GSK3β-dependent threonine-286 (T286) phosphorylation, which triggers CRM1-dependent nuclear export and ubiquitin-mediated proteolysis (Diehl et al., 1998). Proteolytic inactivation is essential as cyclin D1 synthesis does not decrease in the presence of continuous growth factor signaling (Matsushime et al., 1991).  Cytoplasmic T286 phosphorylated cyclin D1 is targeted for degradation via the SCFFbx4-αBcrystallin E3 ubiquitin ligase, and recognition of phosphorylated cyclin D1 is mediated by both Fbx4 and αB crystallin (Lin et al., 2006). 
    	 Of the numerous cell cycle regulatory proteins, cyclin D1 is one of the most frequently targeted in human cancer, with overexpression resulting from gene amplification, chromosomal translocation, and protein stabilization (Sherr, 1996). Cyclin D1 overexpression has been observed in various malignancies including carcinomas of the breast, esophagus, colon, and lung (Barnes and Gillett, 1998; Bartkova et al., 1994a; Bartkova et al., 1995; Bartkova et al., 1994b; Ikeguchi et al., 2001; Jin et al., 2001). While cyclin D1 overexpression occurs at a significant frequency, overexpression per se is insufficient to drive neoplastic transformation. In contrast, cyclin D1 mutants that are refractory to Fbx4-dependent proteolysis and accumulate in the nucleus are acutely transforming (Alt et al., 2000; Gladden et al., 2006; Lin et al., 2007). Consistent with disruption of cyclin D1 proteolysis functioning as an agonist of neoplastic growth, mutations that disrupt cyclin D1 nuclear export and thus, ubiquitin-mediated proteolysis, have been identified in human cancers (Benzeno and Diehl, 2004; Moreno-Bueno et al., 2003).  In addition, recent screening of human esophageal tumor samples revealed inactivating mutations in Fbx4, which will again drive nuclear cyclin D1 accumulation (Barbash et al., 2008). 
Genomic instability is a hallmark of human cancers implicated in initiation and promotion of tumorigenesis.  Double strand DNA breaks (DSBs) arise following γ-irradiation (γIR) or destabilization of stalled replication forks; checkpoint kinases respond to such lesions, promoting cell cycle arrest and DNA repair (Kastan and Bartek, 2004). Impaired detection or repair of DSBs results in chromosomal translocations and deletions, triggering genomic instability (Khanna and Jackson, 2001). The intra-S-phase checkpoint responds to DSB induction occurring at loci outside actively replicating DNA; failure to reduce the rate of DNA synthesis following such genotoxic insult results in radio-resistant DNA synthesis (RDS), a phenotype associated with abrogated checkpoint signaling through ATM (Ataxia Telangiectasia, mutated) and downstream effectors (Bartek et al., 2004). The primary mechanism of DSB-induced S-phase arrest involves inhibition of unfired replication origins (Bartek et al., 2004; Costanzo et al., 2000; Falck et al., 2002).  
Recent evidence revealed that ubiquitylation refractory cyclin D1 accumulates as an active D1/CDK4 kinase within the nucleus during S-phase, where it promotes DNA re-replication, thereby linking cyclin D1 deregulation with S-phase perturbation (Aggarwal et al., 2007).  Previous work implicated cyclin D1 destruction following DNA damage in promoting G1-phase cell cycle arrest (Agami and Bernards, 2000). Consequently, we have investigated the role of cyclin D1 degradation for genomic stability following genotoxic stress. The data provided reveal that cyclin D1 is rapidly degraded in a phosphorylation-dependent manner following DSB induction, and failure to degrade cyclin D1 compromises the intra-S-phase checkpoint response.  Furthermore, evidence is provided revealing that ATM signaling is critical for GSK3β-dependent cyclin D1 phosphorylation, highlighting the relevant signaling pathway that coordinates DSB induction with cyclin D1 regulation.  

Results
Expression of ubiquitylation-refractory cyclin D1T286A generates chromosomal instability following DNA damage. 	
	Transgenic mice expressing cyclin D1T286A driven by the Eμ enhancer develop B-cell lymphomas with significant aneuploidy and chromosomal translocations (Aggarwal et al., 2007).  Because cyclin D1T286A is refractory to ubiquitin-mediated proteolysis (Diehl et al., 1997), we determined whether its expression also compromised genomic integrity in response to transient exposure to DNA damaging agents. Non-malignant splenocytes derived from Eμ-D1T286A mice that were exposed to mitomycin-C (MMC) exhibited a significant increase in chromatid breaks relative to non-transgenic controls (Figure 2.1A). Similarly, exposure to hydroxyurea (HU) increased the incidence of chromatid breaks in D1T286A-expressing cells compared to non-transgenic controls (Figure 2.1B). Importantly, untreated non-transgenic and Eμ-D1T286A splenocytes exhibited infrequent chromatid breaks (Figure 2.1C), indicating synergy between cyclin D1T286A and DNA damage in promoting genomic instability. 

DSBs trigger phosphorylation-dependent cyclin D1 degradation.	
The enhanced frequency of chromatid breaks in cells expressing cyclin D1T286A might reflect the refractory nature of this allele to proteasome-dependent destruction (Diehl et al., 1997). In fact, genotoxic stress normally triggers the rapid loss of cyclin D1, which has been attributed to ubiquitin-mediated destruction during G1 phase (Agami and Bernards, 2000). Based on these findings, we investigated the mechanism governing cyclin D1 loss following S-phase DNA damage. NIH3T3 cells were treated with HU, a compound that stalls DNA replication by depleting nucleotide pools and generates double breaks as a consequence of replication fork collapse (Bianchi et al., 1986; Lundin et al., 2002). HU treatment promoted a rapid decline in both endogenous cyclin D1 protein and exogenously expressed, Flag-tagged cyclin D1 (Figure 2.2A; data not shown). Cyclin D1 mRNA levels remained constant, consistent with post-translational regulation (data not shown). We subsequently assessed cyclin D1 loss at varied HU concentrations.  Cyclin D1 loss was readily apparent by 0.5mM HU and was maximal at 1-2mM HU (Figure 2.2A); an intermediate dose of 2mM was selected for further analyses.  
We next determined whether proteasome-resistant, non-phosphorylatable D1T286A was refractory to DNA damage-dependent downregulation. NIH3T3 cells stably expressing equivalent levels of either wild type Flag-D1 (3T3-D1) or Flag-D1T286A (3T3-D1T286A) were utilized (Diehl et al., 1997). Treatment of cells synchronously proliferating through S-phase with 2mM HU triggered loss of wild type cyclin D1 protein; loss was inhibited by concurrent application of MG132, revealing proteasome-dependence (Figure 2.2B). In contrast, D1T286A levels were constant following HU treatment (Figure 2.2 B, C).  If accelerated loss of cyclin D1 is determined by increased proteolysis, the cyclin D1 half-life should decrease following DNA damage. S-phase NIH3T3 cells were left untreated or treated with 2mM HU for 30 minutes, followed by addition of cycloheximide (CHX). Western analysis revealed that the cyclin D1 half-life was reduced following HU compared to untreated cells (Figure 2.2D). These results support a model wherein DNA damage induced by HU triggers phosphorylation- and proteasome-dependent degradation of cyclin D1.
	We then ascertained whether cyclin D1 proteolysis required the generation of DSBs. Asynchronously proliferating 3T3-D1 or 3T3-D1T286A cells were subjected to 10Gy gIR and cyclin D1 levels were assessed by immunoblot.  IR, like HU, triggered the rapid loss of wild type cyclin D1 but not D1T286A (Figure 2.2E). Similar to IR, bleomycin treatment also induced rapid, phosphorylation-dependent destruction of cyclin D1 (Figure 2.2F), implying that DSBs promote cyclin D1 degradation. 
To directly address whether stalled DNA replication without DSB induction can trigger cyclin D1 destruction, cyclin D1 stability was compared for aphidicolin and HU treatment. NIH3T3 or 3T3-D1 cells were synchronized by serum starvation for 24 hours and serum stimulated for 15 hours to enter S-phase; cells were then subjected to short-term, low dose aphidicolin treatment to promote the accumulation of single strand DNA without collapsing replication forks (Liu et al., 2003) or 2mM HU, previously shown to induce cyclin D1 degradation (Figure 2.2A-D). A significant percentage (>80%) of cells were in S-phase throughout the duration of treatment with DNA damaging agents as determined by flow cytometry (data not shown). In contrast to HU or γIR, cyclin D1 levels remained stable following aphidicolin treatment (Figure 2.2G, H). Significantly, cyclin D1 is only phosphorylated in a damage-dependent manner following HU treatment (Figure 2.2G), consistent with activation of Chk2 signaling as an indicator of DSB induction. Chk1 is activated following both aphidicolin and HU treatment, consistent with stalled replication; however, a mobility shift in Chk2 due to ATM-dependent phosphorylation was not observed in aphidicolin-treated cells, indicating that this short-term treatment fails to activate the ATM pathway (Figure 2.2G). Collectively, these data demonstrate that stalled replication is not sufficient to induce cyclin D1 loss despite activation of the ATR (ATM and Rad3-related)-Chk1 DNA damage response pathway. 
To ensure that the stability of D1T286A was not a reflection of ectopic expression, we also examined cyclin D1 response to DSBs in human tumor-derived cells harboring either wild type D1 (KYSE520) or an endogenous cyclin D1P287A allele (TE3/TE7) (Benzeno et al., 2006). KYSE520 cells exhibited rapid cyclin D1 degradation following γIR, while cyclin D1P287A, expressed in TE3 and TE7 cells, was refractory to degradation following moderate or high γIR dose (Figure 2.3A-B). Cyclin D2 and cyclin E degradation was not triggered by irradiation, consistent with previous work indicating that DNA damage-induced regulation is restricted to cyclin D1 (Agami and Bernards, 2000); Figure 2.3C). Collectively, our data suggest that bona fide DNA breaks are a prerequisite for cyclin D1-specific proteolysis in response to DNA damage.

GSK3β catalyzes cyclin D1 T286 phosphorylation following genotoxic stress.	
The stability of D1T286A to DSB-mediated destruction suggests that phosphorylation of this residue is essential for genotoxic stress-induced cyclin D1 loss. If so, genotoxic stress should trigger increased T286 phosphorylation (p-T286). Indeed, both irradiation (Figure 2.4A) and bleomycin (Figure 2.2F) triggered an increase in p-T286 prior to cyclin D1 proteolysis. Because the cyclin D1 E3 ligase is cytoplasmic, ubiquitin-mediated proteolysis requires both p-T286 and cytoplasmic redistribution (Alt et al., 2000; Diehl et al., 1997; Lin et al., 2006). If genotoxic stress utilizes the same machinery to eliminate cyclin D1, then nuclear export-deficient cyclin D1 mutants should accumulate as p-T286 proteins in the nucleus.  To test this notion, we examined the regulation of cyclin D1-290/95A, a cyclin D1 mutant harboring an inactive nuclear export sequence, but retaining the GSK3 phosphorylation site (Benzeno and Diehl, 2004).  Consistent with our hypothesis, D1-290/95A was resistant to DSB-dependent proteolysis and accumulated as a phosphorylated protein (Figure 2.4A, right panel).	
GSK3 catalyzes phosphorylation of T286 at the G1/S boundary to promote cyclin D1 loss (Diehl et al., 1998). To determine whether GSK3 mediates DSB-dependent T286 phosphorylation, we initially utilized a kinase-dead GSK3β (kdGSK3β) allele. Cyclin D1 phosphorylation was assessed 20 minutes after γIR in order to visualize this highly labile protein in the absence of proteasome inhibition; importantly, significant cyclin D1 degradation is not evident at this early time following DNA damage in NIH3T3 cells (Figure 2.2G, H). Expression of kdGSK3 attenuated T286 phosphorylation following γIR (Figure 2.4B). As independent assessment of GSK3β dependence, NIH3T3 cells were treated with LiCl (Stambolic et al., 1996) or a small molecule inhibitor of GSK3β, SB216763. Both inhibitors attenuated cyclin D1 phosphorylation following γIR (Figure 2.4C). As a third method for assessing GSK3-dependent T286 phosphorylation, shRNA technology was utilized to attenuate GSK3 expression. 293T cells were transfected with vectors encoding cyclin D1 and CDK4 along with control or GSK3-specific shRNA. Immunoblotting confirmed a reduction in GSK3 protein levels (Figure 2.4D). Attenuation of GSK3 expression significantly reduced cyclin D1 phosphorylation compared to control shRNA-transfected cells (Figure 2.4D). Collectively, three independent methods of GSK3 inhibition reveal that this kinase regulates T286 phosphorylation following DNA damage.
Since cyclin D1 is rapidly phosphorylated following DNA damage, we determined whether GSK3 kinase activity was increased following IR. Previously published work revealed an increase in GSK3 kinase activity following UV irradiation (Lee et al., 2007). IP-kinase reactions from irradiated NIH 3T3 cells revealed increased GSK3 kinase activity following IR (Figure 2.4E). GSK3 can also phosphorylate cyclin D2 on threonine 280 (T280) and thereby trigger its proteasome-dependent loss (Kida et al., 2007). Consequently, we determined whether cyclin D2 phosphorylation was regulated by IR.  Cyclin D2 T280 phosphorylation was not induced above basal levels following IR (data not shown). This finding suggests that while GSK3 kinase activity is increased following DNA damage, additional regulatory mechanisms exist to allow GSK3-dependent cyclin D1 phosphorylation specifically. A second kinase, p38SAPK, has been implicated in stress-induced cyclin D1 phosphorylation (Casanovas et al., 2000), and DNA damage increases p38SAPK activity (Raman et al., 2007). We therefore determined whether p38SAPK contributes to DNA damage-dependent cyclin D1 destruction. Significantly, p38 inhibition did not stabilize cyclin D1 protein, suggesting that p38 is not a key regulator of DSB-induced cyclin D1 loss (Figure 2.4F).  Collectively, these results indicate that GSK3β is the primary kinase responsible for DSB-dependent cyclin D1 phosphorylation.

ATM signaling coordinates DSB induction with cyclin D1 phosphorylation. 
	Because the ATM kinase is the primary signal transducer following DSB induction, we determined whether either ATM or the related kinase ATR (which responds to stalled replication) is necessary for cyclin D1 loss following genotoxic stress. We utilized ATM+/+ or ATM-/- murine fibroblasts for this analysis.  To facilitate detection, we expressed Flag-tagged cyclin D1 in these cells as our analysis has revealed that ectopic protein is regulated as effectively as endogenous (see Figure 2.2).  In ATM-/- cells, cyclin D1 levels remained elevated, while cyclin D1 was degraded in matched passage ATM+/+ cells following stress (Figure 2.5A). Consistent with ATM coordinating cyclin D1 loss, cyclin D1 phosphorylation was only induced in ATM+/+ cells following γIR (Figure 2.5B). Furthermore, pharmacological inhibition of ATM with KU55933 also inhibited cyclin D1 phosphorylation (Figure 2.5C).  
Since ATM and Mre11-dependent DNA resection following DSBs will also trigger activation of ATR (Jazayeri et al., 2006), we determined whether ATR signaling contributed to DSB-induced cyclin D1 loss. Stalled DNA replication induced by aphidicolin treatment was insufficient to induce cyclin D1 phosphorylation or degradation despite activation of Chk1, a checkpoint kinase downstream of ATR (Figure 2.2G, H) suggesting that the ATR pathway does not regulate cyclin D1. The role of ATR was directly assessed using fibroblasts harboring a tamoxifen inducible ATR allele, ATRflox/- (Brown and Baltimore, 2003).  Cells were treated with DMSO or 200nM 4-OH tamoxifen (TAM) for 24 hours to induce ATR deletion, followed by 12-hour incubation to allow for ATR depletion. Cyclin D1 levels were equivalent in the presence or absence of ATR following IR (Figure 2.6A) or HU treatment (Figure 2.6B). Additionally, attenuation of endogenous ATR expression via ATR-specific siRNA did not alter cyclin D1 phosphorylation or degradation following IR, despite efficient knockdown (Figure 2.6C). Taken together, these data reveal that cyclin D1 regulation following DNA damage occurs downstream of ATM activation.
Because Chk2 is a primary effector of ATM (Kastan and Bartek, 2004), we assessed whether activation of Chk2 is required for cyclin D1 loss.  Knockdown of Chk2 via siRNA modestly attenuated, but did not abrogate DSB-dependent cyclin D1 destruction (Figure 2.7A). We also assessed whether inhibition of Chk1, the downstream ATR effector, affects cyclin D1 phosphorylation and loss following DSB induction (Figure 2.7B, C).  Significantly, use of the Chk1-specific inhibitor SB218078 or attenuation of Chk1 expression via shRNA did not alter cyclin D1 phosphorylation or degradation in S-phase cells treated with HU (Figure 2.7B and C, respectively). Finally, we determined whether there is synergy between Chk1 and Chk2 in regulating cyclin D1 following DSB induction. siRNA-mediated knockdown of Chk2 modestly attenuates cyclin D1 degradation at early time points in U20S cells; in contrast, cyclin D1 levels are equivalent in control and Chk2 siRNA conditions at later time points (Figure 2.7D). Significantly, addition of a Chk1 inhibitor, SB218078, did not further stabilize cyclin D1 levels, suggesting that Chk1 and Chk2 activity is not synergistic in regulating cyclin D1 (Figure 2.7D). These results suggest that Chk2 may play a minor or indirect role in cyclin D1 regulation; however, additional ATM-dependent effectors also function in DNA damage-induced cyclin D1 degradation. Collectively, our data support a model wherein ATM activation following DSB induction is necessary for cyclin D1 phosphorylation and subsequent degradation. However, future investigation of regulators downstream of ATM, including Chk2, is important to completely delineate the DNA damage-specific signaling network that coordinates cyclin D1 destruction.

SCFFbx4-αB crystallin is required for cyclin D1 degradation following DNA damage.	SCFFbx4-αBcrystallin was recently shown to direct p-T286-dependent ubiquitylation of cyclin D1 in both fibroblasts and epithelial cells (Lin et al., 2006). Fbx4 and αB crystallin serve as substrate-specific adaptors that recognize phosphorylated cyclin D1. To directly assess the role of Fbx4 and αB crystallin in DNA damage-induced cyclin D1 proteolysis, we utilized NIH3T3 cells wherein either Fbx4 or αB crystallin have been stably knocked down by shRNA (Lin et al., 2006).  Knockdown of either Fbx4 or αB crystallin significantly attenuated cyclin D1 proteolysis following DNA damage (Figure 2.8A, B). 
Previous work revealed disruption of the B crystallin locus in several breast cancer-derived cell lines (Lin et al., 2006).  We utilized one such cell line, MDA-MB 231 cells, to independently determine whether αB crystallin is required for cyclin D1 degradation following DSB induction. Consistent with the results obtained utilizing NIH3T3 cells expressing αB crystallin shRNA, irradiation of MDA-MB 231 cells did not trigger cyclin D1 loss, while re-introduction of exogenous αB crystallin restored DNA damage-induced cyclin D1 degradation (Figure 2.8C, compare 1h and 3h treatment in left panel to corresponding treatments in right panel).  
While our data reveal cyclin D1 destruction following DSB initiation to be dependent upon p-T286 and the SCFFbx4-αBcrystallin ligase, the anaphase promoting complex/cyclosome (APC/C) has also been implicated in the regulation of cyclin D1 degradation during G1 phase following γIR (Agami and Bernards, 2000).  Importantly, the cell line utilized for these studies, MCF7, lacks endogenous αB crystallin expression, thereby rendering the Fbx4 ligase inactive toward cyclin D1 in these cells (Lin et al., 2006).  Since cyclin D1 turns over, albeit with slower kinetics in this cell line, an alternative degradation pathway is likely.  To determine if APC/C contributes to cyclin D1 degradation in cells expressing a functional Fbx4-dependent ligase, we mutated L32 of cyclin D1 within the putative destruction box (Agami and Bernards, 2000) to alanine and examined cyclin D1 stability following DNA damage.  The D1-L32A mutant is degraded similarly to wild type protein (Figure 2.8D); we also demonstrate that this mutant has impaired CDK4 binding potential, consistent with disruption of residues proximal to the cyclin box (Figure 2.8D).  Additionally, D1-L32A is degraded in MCF7 cells reconstituted with exogenous αB crystallin (data not shown), suggesting that restored SCFFbx4-αBcrystallin ligase activity shifts cyclin D1 destruction from alternative mechanisms to the phosphorylation-dependent destruction pathway. 
Recent work revealed that disruption of SCFFbx4-αB crystallin allows accumulation of cyclin D1 in the nucleus, and impaired ligase function results in neoplastic transformation (Barbash et al., 2008). We therefore determined whether knockdown of Fbx4 sensitizes cells to chromatid breaks following DNA damage, analogous to cyclin D1T286A (Figure 2.1B). NIH3T3 cells stably expressing vector control, Fbx4 shRNA, or cyclin D1T286A were synchronized and treated with HU during S-phase. Metaphase spreads were then prepared and analyzed for chromatid breaks. In agreement with splenocytes derived from Eμ-D1T286A transgenic mice, NIH3T3 cells expressing cyclin D1T286A exhibited a significant increase in chromatid breaks following HU treatment. Analogous with cyclin D1T286A expression, knockdown of Fbx4 also results in increased chromatid breaks following HU compared to the vector control (Figure 2.8E, F). 

Cyclin D1 stabilization compromises the intra-S-phase checkpoint.
	Cyclin D1 mutants refractory to ubiquitin-mediated proteolysis accumulate in the nucleus as active cyclin D1/CDK4 complexes and drive DNA re-replication. This deregulated replication results from stabilization of the pre-RC licensing factor CDT1, which promotes loading of the MCM helicase (Aggarwal et al., 2007). Cdt1 degradation in response to DNA damage is necessary to inhibit replication of damaged DNA (Higa et al., 2003; Hu et al., 2004; Nishitani et al., 2006). Since cyclin D1 is rapidly degraded following DSB induction, we considered whether failure to degrade cyclin D1 would impede CDT1 destruction. To test this notion, we assessed CDT1 stability following DNA damage; HeLa cells were transiently transfected with wild type or cyclin D1T286A constructs along with CDK4 and treated with bleomycin. Cdt1 levels were attenuated in the presence of wild type cyclin D1, while D1T286A expressing cells exhibited CDT1 stabilization up to 2 hours of bleomycin treatment (Figure 2.9A), consistent with published results (Aggarwal et al., 2007). 
Maintenance of CDT1 should have a direct impact on MCM chromatin loading following DNA damage. We assessed chromatin retention of MCM complexes in human cancer-derived cells that express endogenous wild type cyclin D1 or P287A. MCM3 (a surrogate marker of MCM loading) dissociates from chromatin with a concomitant increase in soluble MCM3 protein following DNA damage in the presence of wild type cyclin D1; however, MCM3 was stabilized on chromatin in P287A-expressing cells (Figure 2.9B).  PCNA serves as a representative chromatin bound protein, and loading of the chromatin fraction was also examined by Ponceau S stain. These data suggest that improper maintenance of pre-RC components on chromatin contributes to cyclin D1T286A-driven perturbations in DNA replication, a finding consistent with previously published reports (Aggarwal et al., 2007).	
To directly determine whether cyclin D1 stabilization overrides the intra-S-phase checkpoint, we assessed RDS in cells expressing equivalent steady-state levels (prior to DNA damage) of either wild type cyclin D1 or D1T286A (Figure 2.2E).  DNA synthesis decreases significantly following γIR in naïve 3T3 and 3T3-D1 cells (p<0.05); however, DNA synthesis was not significantly decreased in D1T286A expressing cells (Figure 2.9C). We also measured RDS in NIH3T3 cells where Fbx4 levels are reduced through shRNA technology. Analogous with degradation-refractory D1T286A, knockdown of Fbx4 or B crystallin promoted RDS (Figure 2.9D).  In contrast, knockdown of Fbx4 in cyclin D1-/- fibroblasts failed to trigger RDS (Figure 2.9E, F), which is consistent with cyclin D1 as the primary regulatory target of Fbx4 pertaining to cell cycle progression. Although RDS is often associated with a failure of cells to inhibit CDK2, our analysis revealed effective inhibition of CDK2 kinase in D1T286A expressing cells (data not shown) suggesting that maintenance of cyclin D1/CDK4 activity directly contributes to the RDS phenotype.
	Since cyclin D1T286A expression promotes RDS, we hypothesized that cells expressing stabilized cyclin D1 would be sensitized to the S-phase-specific chemotherapeutic agent camptothecin (CPT).  To address this hypothesis, 3T3-D1 or D1T286A cells were treated with 25 or 50M CPT for 24 or 48 hours; cells were harvested, stained with propidium iodide, and subjected to FACS analysis to determine cell cycle distribution. A significant increase in sub-G1 cells was observed following CPT treatment for 48h (p<0.05), providing evidence that cyclin D1T286A-expressing cells are sensitized to CPT (Figure 2.9G). Taken together, the RDS phenotype and S-phase chemotherapy-sensitivity observed in D1T286A-expressing cells highlights a potential therapeutic strategy for tumors exhibiting stabilized cyclin D1.

Discussion
Recent work revealed that alterations in post-translational control of cyclin D1 contribute to neoplastic growth in model systems and in human cancers (Barbash et al., 2008; Benzeno et al., 2006; Gladden et al., 2006; Lin et al., 2006).   Cyclin D1 accumulation in the nuclear compartment, where it functions as an allosteric activator of CDK4, contributes directly to G1 progression and cell proliferation. Importantly, accumulation is limited by ubiquitin-mediated proteolysis catalyzed by SCFFbx4-αBcrystallin, and loss of proteolytic control contributes to neoplastic growth (Diehl et al., 1997; Lin et al., 2006). The findings presented here reveal that nuclear stabilization of cyclin D1 compromises the intra-S-phase checkpoint following DNA damage.  While cyclin D1 is an inherently unstable protein, genotoxic stress further accelerates its proteolysis in a manner that depends upon GSK3β-mediated p-T286 and ubiquitylation by the SCFFbx4-αBcrystallin E3 ligase.  Mutations that impair cyclin D1 phosphorylation or inactivate SCFFbx4-αBcrystallin result in the accumulation of active cyclin D1/CDK4 in the nucleus, which in turn, impairs checkpoint-dependent inhibition of DNA replication.  Ultimately, the ensuing RDS results in the accumulation of chromatid breaks.  
T286 phosphorylation is required for both cell cycle- and DNA damage-mediated destruction of cyclin D1.  While additional kinases have been implicated as regulators of p-T286 (Casanovas et al., 2000; Kwak et al., 2005; Okabe et al., 2006), our data provide strong evidence for GSK3β as the T286 kinase.  GSK3β inhibition via LiCl treatment enriches nuclear cyclin D1 (Alt et al., 2000); LiCl, a small molecule GSK3β inhibitor (SB216763), kdGSK3β, and shRNA all significantly attenuate DNA damage-induced T286 phosphorylation.  Importantly, inhibition of the two additional kinases implicated as regulators of cyclin D1, IKK (data not shown) and p38 (Figure 2.4F), had no effect on cyclin D1 phosphorylation or degradation in response to DSB induction.  
GSK3β contributes directly to cyclin D1 regulation in multiple ways.  First, phosphorylation of T286 is required for CRM1-mediated nuclear export (Alt et al., 2000).  Second, phosphorylated cyclin D1 is recognized by Fbx4, the specificity factor for the SCF E3 ligase, once re-localized to the cytoplasm (Lin et al., 2006).  Our data reveal that shRNA-mediated reduction of either Fbx4 or αB crystallin stabilizes cyclin D1 in the presence of DSBs. Consistently, cyclin D1 is also refractory to degradation in breast cancer cells lacking endogenous αB crystallin expression, while reintroduction of exogenous αB crystallin restores cyclin D1 turnover.  Collectively, these data support a model wherein upregulation of GSK3β-dependent cyclin D1 phosphorylation following genotoxic stress targets the protein for SCFFbx4-mediated destruction.  
Previous work implicated the anaphase promoting complex/cyclosome (APC/C) in DNA damage-induced cyclin D1 degradation (Agami and Bernards, 2000).  A putative RxxL destruction box in the N-terminus of cyclin D1 was identified as being responsible for directing APC/C-dependent degradation. Our work reveals that mutation of T286, loss of Fbx4, or inhibition of GSK3β impairs DSB-mediated cyclin D1 loss; we also assessed the potential contribution of the RxxL motif. Leucine 32 to alanine (L32A) mutation did not attenuate DSB-induced loss. Importantly, this mutation profoundly impairs CDK4 binding, most likely resulting from disruption of protein structure just proximal to the cyclin box. Thus, alterations in folding of this mutant could contribute to its stabilization under some conditions. 
While mutations near the cyclin box clearly alter structural interactions with CDK4, this may not be the only reason accounting for discrepancies regarding the role of different E3 ligases in cyclin destruction.  An additional possibility is that this previous work utilized MCF7 cells, a cell line lacking αB crystallin (Lin et al., 2006), thereby inactivating the SCFFbx4-αB crystallin ubiquitin ligase.  While cyclin D1 proteolysis is significantly attenuated in these cells, restoration of αB crystallin rescues both mitogen and DSB-dependent cyclin D1 proteolysis (data not shown). A third E3 ligase utilizing Fbw8 as a specificity co-factor has been implicated in regulating cyclin D1 (Okabe et al., 2006).  We have not assessed Fbw8 directly due to a lack of suitable antibodies and cannot rule out its potential role in regulating D1. However, Cul7, a core component of the Fbw8 ligase, is undetectable in NIH3T3 cells (Kasper et al., 2005), and our data revealing that knockdown of either Fbx4 or B crystallin is sufficient to inhibit cyclin D1 proteolysis emphasizes their requisite role in this regulatory process. 
During S-phase, DNA must be replicated with high fidelity to ensure error-free propagation of genetic material. DSBs resulting from various stresses invoke a checkpoint that attenuates cell cycle progression until DNA is repaired (Bartek et al., 2004). The intra-S-phase checkpoint is characterized by replication fork stalling, inhibition of origin firing, and accumulation of single strand DNA, promoting ATR activation; in contrast, the accumulation of DSBs, regardless of replication status, triggers ATM activation (Bartek et al., 2004; Kastan and Bartek, 2004).  Strikingly, inhibition of cyclin D1 degradation impairs checkpoint response and cells exhibit an RDS phenotype, implying checkpoint failure. The RDS phenotype observed in cyclin D1T286A-expressing cells is accompanied by CDT1 stabilization, which is dependent upon CDK4 activation by cyclin D1.  Ultimately, the continued presence of CDT1 contributes to retention of MCM helicase on chromatin, suggesting that the presence of this mutant within the nucleus perturbs pre-RC disassembly following DNA damage. The significance of the cyclin D1T286A RDS phenotype is highlighted by increased presence of chromatid breaks in primary splenocytes treated with DNA damaging agents. 	
While the ability of constitutively nuclear D1T286A to promote CDT1 stabilization is clear, how does inactivation of Fbx4 ligase in cells harboring wild type cyclin D1 equate with the same biochemical function?  Importantly, published work reveals that inactivation of SCFFbx4-αB crystallin results in the nuclear accumulation of endogenous cyclin D1/CDK4 kinase leading to CDT1 stabilization and function in both cycling cells and following DSB generation (Aggarwal et al., 2007; Barbash et al., 2008).  Accordingly, our data are consistent with these findings and further reveal the retention of MCM on chromatin following DSBs. Additionally, shRNA-mediated attenuation of Fbx4 expression promotes an RDS phenotype that correlates with increased genomic instability following treatment with DNA damaging agents. 
Previous work suggested that inhibition of cyclin D1/CDK4 activity is critical for genotoxic stress induced cell cycle arrest (Agami and Bernards, 2000). Significantly, cyclin D1-driven DNA re-replication requires CDK4 kinase activity (Aggarwal et al., 2007).  Our findings here not only reveal the molecular nature of the ligase that regulates cyclin D1 but also importantly provide mechanistic insight with regard to the consequences of the accumulation of stabilized cyclin D1 and thus, cyclin D1/CDK4 kinase.  In the absence of CDK4 kinase, pre-malignant cells could successfully halt DNA replication, thereby reducing genomic instability that may promote transformation. 
Given the RDS phenotype associated with maintenance of cyclin D1 protein following genotoxic stress, it is not surprising that ATM signaling is required to coordinate cyclin D1 destruction. Consistent with our model of DSB induction as a prerequisite for cyclin D1 phosphorylation and destruction, accumulation of single strand DNA and subsequent ATR activation is insufficient to direct cyclin D1 loss. Significantly, damage-induced cyclin D1 phosphorylation and destruction are abrogated in ATM null cells and use of an ATM-specific inhibitor dramatically attenuates cyclin D1 phosphorylation. However, it is possible that ATR could contribute to ATM activation following fork stalling (Stiff et al., 2006). The fact that ATR depletion did not attenuate cyclin D1 phosphorylation/degradation and that cyclin D1 is not phosphorylated following stress that does not trigger DSB implies that cyclin D1 loss is ATM-dependent and ATR-independent.
While ATM is required for DNA damage-induced cyclin D1 phosphorylation, the precise effector-signaling network remains unclear. Chk2 appears to function in the regulation of cyclin D1 degradation at early time points following γIR; however, Chk2 knockdown/inhibition fails to rescue cyclin D1 degradation at later time points after damage.  As loss of ATM and GSK3 function to stabilize cyclin D1 at all time points, this result suggests an alternative ATM effector in the mediation of GSK3-dependent cyclin D1 degradation. Delineating the molecular signaling network connecting ATM with cyclin D1 T286 phosphorylation is an exciting area for future study.

Figure 2.1.

Figure 2.2.

Figure 2.3.

Figure 2.4.

Figure 2.5.

Figure 2.6.

Figure 2.7.

Figure 2.8.

Figure 2.9.

CHAPTER III:
The fbx4 tumor suppressor regulates cyclin D1 accumulation and prevents neoplastic transformation

SUMMARY
SCF E3 ubiquitin ligase complexes modulate the accumulation of key cell cycle regulatory proteins. Following the G1/S transition, SCFFbx4 targets cyclin D1 for proteasomal degradation, a critical event necessary for DNA replication fidelity.  Deregulated cyclin D1 drives tumorigenesis, and inactivating mutations in Fbx4 have been identified in human cancer, suggesting that Fbx4 may function as a tumor suppressor. Fbx4+/- and Fbx4-/- mice succumb to multiple tumor phenotypes including lymphomas, histiocytic sarcomas, and less frequently, mammary tumors and hepatocellular carcinomas. Tumors and pre-malignant tissue from Fbx4+/- and Fbx4-/- mice exhibit elevated cyclin D1, consistent with cyclin D1 as a target of Fbx4. Molecular dissection of the Fbx4 regulatory network in murine embryonic fibroblasts (MEFs) revealed that loss of Fbx4 results in cyclin D1 stabilization and nuclear accumulation throughout cell division. Increased proliferation in early passage primary MEFs is antagonized by DNA damage checkpoint activation, consistent with nuclear cyclin D1-driven genomic instability. Furthermore, Fbx4-/- MEFs exhibited increased susceptibility to Ras-dependent transformation in vitro, analogous to tumorigenesis observed in mice. Collectively, these data reveal a requisite role for the SCFFbx4 E3 ubiquitin ligase in regulating cyclin D1 accumulation, consistent with tumor suppressive function in vivo.

Introduction
Cell cycle progression is intricately controlled by coordinated expression and ubiquitin (Ub)-dependent degradation of key regulatory proteins. Ubiquitylation of target substrates requires the concerted activity of an Ub-activating enzyme (E1), recruitment of an Ub-conjugating enzyme (E2), and an E3 ubiquitin ligase. Although some E3 ubiquitin ligases directly mediate transfer of ubiquitin, for example the HECT-domain family, many E3 ligases function as multi-subunit complexes that bridge the charged E2 enzyme with a given substrate (Hershko and Ciechanover, 1998). The SKP1-CUL1-F-box (SCF) family of E3 ubiquitin ligases promotes polyubiquitylation of mainly phosphorylated substrates, including mediators of the G1/S phase transition (Deshaies, 1999; Skowyra et al., 1997). Substrate specificity of SCF E3 ligases is conferred by the F-box protein, which serves as a molecular adapter bridging substrate molecules with the core ligase machinery and E2 enzyme. F-box proteins contain a conserved F-box domain within the N-terminal region, required for specific SKP1 binding and recruitment of core ligase components CUL1, RBX1, and E2, as well as a substrate-recognition domain within the C-terminus (Nakayama and Nakayama, 2005; Winston et al., 1999a).  
The F-box protein Fbx4 facilitates ubiquitylation of cyclin D1 following the G1/S transition (Lin et al., 2006). Similar to Fbw7, an ubiquitin ligase for substrates including cyclin E, c-Myc, and Notch (Tang et al., 2007; Zhang and Koepp, 2006), and -TrCP, an IB E3 ligase (Suzuki et al., 2000), Fbx4 undergoes dimerization; however, in contrast to Fbw7, Fbx4 dimerization occurs in a cell cycle-dependent manner. Fbx4 homodimerization occurs following GSK3-dependent phosphorylation on serine 12, resulting in ligase activation at the G1/S transition and subsequent ubiquitylation of phosphorylated cyclin D1 (Barbash et al., 2008; Lin et al., 2006). Critically, failure to degrade cyclin D1 following the G1/S transition permits active cyclin D1/CDK4 nuclear accumulation and drives cell transformation (Aggarwal et al., 2007; Aggarwal et al.; Alt et al., 2000). While cyclin D1 overexpression occurs in multiple human malignancies (Bani-Hani et al., 2000; Bartkova et al., 1994a; Bartkova et al., 1995; Bartkova et al., 1994b; Gillett et al., 1994; Herman et al., 1995; Hibberts et al., 1999; Hosokawa and Arnold, 1998; Hosokawa et al., 1999; Jin et al., 2001), overexpression of wild type cyclin D1 in an Fbx4-proficient system is insufficient to drive spontaneous transformation (Alt et al., 2000; Barbash et al., 2008), supporting a model wherein cytoplasmic recognition of phosphorylated cyclin D1 by Fbx4 is sufficient to maintain cellular integrity. 
Cyclin D1 mutations that impede its timely degradation and trigger constitutively nuclear accumulation promote neoplastic growth (Aggarwal et al., 2007; Alt et al., 2000; Barbash et al., 2008; Benzeno et al., 2006; Lu et al., 2003); however such mutations are rare in human cancer. Recent work identified inactivating Fbx4 mutations in human cancers that impair ligase phosphorylation and dimerization; such tumors exhibit marked cyclin D1 nuclear accumulation, highlighting a novel mechanism for cyclin D1 deregulation in cancer (Barbash et al., 2008). Consistent with these findings, in vitro studies revealed that shRNA-mediated knockdown of Fbx4 promotes anchorage-independent growth, a phenotype that is rescued by loss of cyclin D1 (Barbash et al., 2008).  Taken together, these findings suggest that the Fbx4 protein functions as a tumor suppressor, preventing aberrant cyclin D1 accumulation during S-phase.
To investigate the role of Fbx4 as a putative tumor suppressor in vivo, we ablated the murine Fbx4 gene. Fbx4-/- mice are viable and lack major developmental defects; however, both Fbx4+/- and Fbx4-/- mice succumb to multiple tumor phenotypes, with a marked increase in cyclin D1 levels in pre-malignant and tumor tissue. Likewise, Fbx4-/- MEFs exhibit cyclin D1 stabilization and subsequent nuclear localization, with concomitant induction of DNA damage-associated nuclear foci, consistent with a model wherein nuclear cyclin D1/CDK4 kinase drives genomic instability. Finally, Fbx4 deficiency facilitates cellular transformation in vitro, supporting the notion that Fbx4 is a bona fide tumor suppressor. 

Results
Generation of Fbx4-/- mice. 
To directly assess the tumor suppressor function of Fbx4 we generated a targeting vector to facilitate deletion of murine Fbx4 by homologous recombination in embryonic stem (ES) cells. LoxP sites were inserted flanking exon 2, with a downstream neomycin (Lan et al.) cassette flanked by a third LoxP site. Cre-mediated recombination and deletion of exon 2 removes sequences critical for assembly of a functional ligase F-box and results in a frameshift and premature termination (Figure 3.1A). The targeting construct was electroporated into ES cells, and correctly targeted clones were identified by Southern hybridization (Figure 3.1B). Four ES clones out of 132 exhibited a correctly targeted allele, with complete excision of the Neo cassette and exon 2. ES cell clones harboring a correctly targeted allele with a normal karyotype were injected in C57BL/6 blastocysts, and two clones resulted in germline transmission. Chimeric mice were then intercrossed with EIIa-Cre transgenic mice to generate Fbx4 heterozygous progeny (genotype confirmation, Figure 3.1C). Both Fbx4+/- and Fbx4-/- animals are viable and fertile, with no apparent developmental defects.

Fbx4+/- and Fbx4-/- mice are tumor prone. 
Multiple tissues harvested from young (1-2 month old) animals were assessed for cyclin D1 and Fbx4 expression. Importantly, targeting exon 2 results in complete loss of Fbx4 protein expression. Among the tissues examined, cyclin D1 protein accumulation was noted in the mammary gland, lung, thymus and spleen (Figure 3.2A-D). Increased cyclin D1 accumulation was also observed in both Fbx4+/- and Fbx4-/- primary MEFs (Figure 3.2E). RT-PCR analysis revealed equivalent levels of cyclin D1 message in Fbx4+/+, Fbx4+/-, and Fbx4-/- MEFs, suggesting that protein accumulation occurs at the post-translational level (Figure 3.2F).
Given the observed cyclin D1 accumulation in normal tissues derived from Fbx4+/- and Fbx4-/- mice, we interrogated whether Fbx4+/- and Fbx4-/- mice develop spontaneous tumors with increasing age. We monitored Fbx4+/+, Fbx4+/-, and Fbx4-/- mice for up to 24 months and observed significant spontaneous tumor formation in both Fbx4+/- and Fbx4-/- mice (Figure 3.3A). Fbx4+/- and Fbx4-/- mice exhibit tumors of lymphoblastic lineage, frequent intense myeloid proliferation resulting from severe extramedullary hematopoiesis, and early myeloid malignancy (Table 2). These mice develop large cell lymphomas, as confirmed by flow cytometric and immunohistochemical analysis. Additionally, dendritic cell tumors/histiocytic sarcomas of the spleen, thymus and intestinal wall/mesentery, liver hemangiomas and hepatocellular carcinoma, and tumors of the mammary and uterine epithelium were observed (Table 2). Fbx4+/- and Fbx4-/- mice succumb to significant tumor burden with a mean survival of 23 months (Figure 3.3A). Importantly, 60.6 percent and 64.3 percent of Fbx4+/- and Fbx4-/- mice, respectively exhibit significant pathology before 24 months of age, highlighting the robust penetrance of these phenotypes. 
To examine whether tumorigenesis in Fbx4+/- mice was due to haploinsufficiency or loss of heterozygosity (LOH), Fbx4 protein expression was analyzed in representative tumor samples. Immunoblot analyses revealed that Fbx4+/- tumors and pre-malignant tissues retain Fbx4 expression, albeit an approximate fifty percent reduction in expression compared to wild type, suggesting that Fbx4 dosage is an important determinant in malignant transformation (Figure 3.3B). Consistent with this observation, one wild type age matched control mouse exhibited severe extramedullary hematopoiesis, a phenotype typically observed in Fbx4+/- and Fbx4-/- mice; strikingly, immunoblot analysis revealed significant Fbx4 downregulation in this spleen.
Since normal and pre-malignant Fbx4+/- and Fbx4-/- tissues exhibit cyclin D1 protein accumulation, we ascertained whether cyclin D1 protein expression is elevated in the various tumors. Consistently, elevated cyclin D1 protein was apparent in a majority of lymphomas originating in the mesenteric node of both Fbx4+/- and Fbx4-/- mice and in livers exhibiting lymphoma infiltrates or hepatocellular carcinoma (Figure 3.4A, B). Furthermore, intestinal histiocytic sarcomas and mesenteric node lymphomas exhibit nuclear cyclin D1 accumulation, correlating with increased proliferation as evidenced by Ki-67 staining (Figure 3.4C).

Fbx4 deletion promotes cyclin D1 stabilization and nuclear accumulation.
To dissect the molecular mechanism underlying neoplastic transformation in Fbx4+/- and Fbx4-/- mice, we prepared MEFs from Fbx4+/+, Fbx4+/-, and Fbx4-/- embryos and examined cell proliferation and cyclin D1 accumulation. Primary Fbx4+/- and Fbx4-/- MEFs exhibit cyclin D1 stabilization in S-phase, with significantly increased cyclin D1 half-life (Figure 3.5A). Importantly, cyclin D1 stabilization correlates with its robust nuclear accumulation in primary MEFs (Figure 3.5B). Consistent with cyclin D1 deregulation in the absence of Fbx4, early passage primary MEFs also exhibit an increased rate of proliferation compared to wild type MEFs (Figure 3.5C). 
Subsequently, we determined whether cyclin D1 protein is stabilized throughout S-phase in Fbx4-/- cells. Primary early-passage MEFs were immortalized with p19Arf shRNA, allowing extended culture and effective synchronization, and cyclin D1 levels were assessed across late G1 through mid S-phase. In Fbx4+/+ cells, cyclin D1 is rapidly degraded following the G1/S transition; conversely, cyclin D1 protein is elevated, reflecting increased stability, through S-phase in Fbx4-/- cells (Figure 3.5D). Furthermore, threonine 286 phosphorylated (pT286) cyclin D1 accumulates in Fbx4-/- cells, indicating defective turnover of this short-lived cyclin D1 modification (Figure 3.5E). We also determined whether cyclin D1 S-phase ubiquitylation requires Fbx4. Wild type and null cells were synchronized and stimulated to enter S-phase with serum-derived growth factors; cells were treated with the proteasome inhibitor MG-132 to allow accumulation of poly-ubiquitylated proteins, and cyclin D1 was immunopurified. Immunoblot analysis revealed markedly reduced cyclin D1 polyubiquitylation in Fbx4-/- cells compared to Fbx4+/+ controls, suggesting that impaired cyclin D1 ubiquitylation/degradation drives nuclear cyclin D1 accumulation in Fbx4-/- cells (Figure 3.5F).

Enhanced proliferation is antagonized by DNA damage in Fbx4-/- MEFs. 
Primary MEFs were maintained on a 3T9 passage protocol to evaluate the onset of cellular senescence. While early passage Fbx4-/- MEFs initially accumulate to a higher density, proliferation is dramatically reduced between passage 6-7, compared to the gradual proliferative decline observed in Fbx4+/+ MEFs (Figure 3.6A). Importantly, both wild type and null MEFs exhibit similar patterns of senescence-associated p19Arf and p16Ink4a induction, and p19Arf shRNA facilitates immortalization both wild type and null cells with similar kinetics, suggesting that reduced proliferation in primary null cells may result from alternative mechanisms such as DNA damage (Figure 3.6B, C) Significantly, Fbx4-/- MEFs accumulate H2AX, as well as p53 and its downstream target, p21Cip1, consistent with induction of the DNA damage checkpoint in later passages (Figure 3.6 D-E, Figure 3.5D). Fbx4-/- cells also accumulate DSB-associated nuclear foci (Figure 3.6F). Previous studies established that primary MEF proliferation under atmospheric oxygen is limited by oxidative DNA damage, despite maintenance of long telomeres (Parrinello et al., 2003). Importantly, Fbx4-/- MEFs display markers of DNA damage earlier than Fbx4+/+ counterparts, and DNA damage in Fbx4-/- MEFs cannot be rescued by culture under physiological oxygen (3% O2, Figure 3.6F), providing evidence that oxidative stress alone does not account for DNA damage accumulation in Fbx4-/- MEFs. 
Knockdown of p19Arf bypasses cellular senescence, allowing immortalization of Fbx4+/+ and Fbx4-/- MEFs; such immortalized cells do not exhibit proliferative defects (Figure 3.6C). Next, we assessed whether immortalized Fbx4+/+ and Fbx4-/- cells are selectively sensitive to S-phase DNA damage. Fbx4+/+ and Fbx4-/- cells were synchronized in S-phase, pulsed with 2mM HU for 2 hours, followed by HU washout, incubation in complete media, and treatment with colcemid to arrest cells in metaphase. Metaphase spreads prepared from Fbx4-/- cells exhibited a significant increase in chromosome alterations compared to wild type controls (Figure 3.6G). 
Previous work established that nuclear cyclin D1 accumulation promotes DNA re-replication during S-phase and continued DNA replication in the presence of S-phase DNA damage, thereby facilitating genomic instability; these phenotypes are driven by nuclear cyclin D1/CDK4-mediated CUL4 repression and CDT1 stabilization (Aggarwal et al., 2007; Pontano et al., 2008). Consistent with these findings, CDT1 protein is stabilized throughout S-phase in Fbx4-/- cells, with a concomitant reduction in CUL4A expression (Figure 3.7A). Collectively, these data reveal that loss of Fbx4 triggers nuclear cyclin D1 accumulation, DNA damage checkpoint activation, and ultimately genomic instability.
Our data revealed attenuated cyclin D1 turnover in primary S-phase Fbx4-/- MEFs (Figure 3.5A); furthermore, cyclin D1 degradation in response to S-phase DNA damage is impaired in Fbx4-/- MEFs (Figure 3.7B), suggesting that Fbx4 is a critical mediator of cyclin D1 destruction in unperturbed S-phase progression and following S-phase DNA damage. Since GSK3 phosphorylates Fbx4, triggering ligase dimerization and activation (Barbash et al., 2008), and GSK3 activity increases following S-phase DNA damage (Lee et al., 2007; Pontano et al., 2008), we next ascertained whether Fbx4 phosphorylation and ligase activity are regulated by S-phase DNA damage. Treatment of S-phase cells with ionizing radiation enhanced Fbx4 phosphorylation, concomitant with elevated cyclin D1 phosphorylation (Figure 3.7C). Moreover, SCFFbx4 E3 ubiquitin ligase activity toward cyclin D1 substrate in vitro is markedly increased when SCFFbx4 complexes are purified from -irradiated S-phase NIH 3T3 cells (Figure 3.7D). Together, these data indicate that both phosphorylation-dependent cyclin D1 nuclear export, as well as phosphorylation-dependent Fbx4 ligase activation synergize to promote cyclin D1 ubiquitylation following S-phase DNA damage, thereby preserving genome integrity.

Fbx4 loss cooperates with RasV12 in vitro.
	The data provided demonstrate that Fbx4+/- and Fbx4-/- mice are tumor prone and this correlates with deregulation of cyclin D1 accumulation.  Because cyclin D1 overexpression cooperates with Ras to induce cellular transformation, we considered whether Fbx4+/- or Fbx4-/- primary MEFs were susceptible to transformation induced by oncogenic Ha-Ras(V12). Infection of early passage MEFs with retroviruses harboring empty vector or oncogenic Ha-Ras(V12) resulted in foci formation in both Fbx4+/- and Fbx4-/- MEFs, but not Fbx4+/+ MEFs (Figure 3.8A). While overexpression of wild type cyclin D1 alone does not promote cell transformation or tumorigenesis in immune compromised mice (Quelle et al., 1993), expression of cyclin D1 mutants that are stably retained in the nucleus is oncogenic (Alt et al., 2000). To determine whether Fbx4 loss and subsequent nuclear accumulation of endogenous cyclin D1 promotes cellular transformation, we examined the capacity of multiple p19ArfshRNA-immortalized cell lines to form foci in vitro. While immortalized Fbx4+/+ cells were refractory to spontaneous transformation, Fbx4+/-, and to a greater extent, Fbx4-/- cells readily exhibited spontaneous transformation (Figure 3.8B), suggesting that Fbx4 plays a requisite role in suppressing tumorigenesis in vivo and cellular transformation in vitro.

Discussion
The SCFFbx4 E3 ubiquitin ligase has been previously implicated in regulating cyclin D1 turnover in cell culture models. Further work revealed a significant role for Fbx4 function in maintaining cell integrity, highlighted by the relative frequency of mutations that impair its biochemical activity in human tumors. Such mutations occur in a hemizygous manner, consistent with the notion that reduced Fbx4 expression is sufficient for cyclin D1 deregulation and neoplastic transformation (Barbash et al., 2008). While these lines of evidence suggest that Fbx4 may function as a tumor suppressor, genetic studies in vivo have not been performed to directly address this question. We have interrogated Fbx4-dependent tumor suppression in vivo, through targeted disruption of murine Fbx4. Our data reveal that loss of Fbx4 in mice does not impact viability or development; however, a significant increase in age-associated malignancy was observed in Fbx4+/- and Fbx4-/- mice compared to wild type. Furthermore, Fbx4 loss in MEFs triggered cyclin D1 stabilization and nuclear accumulation, and aberrant cyclin D1 regulation in the absence of Fbx4 drives the accumulation of DNA damage and subsequent genomic instability, a hallmark of neoplastic transformation. Consistently, loss of Fbx4 facilitates cell transformation in vitro, further emphasizing the tumor suppressive properties of Fbx4. 

Fbx4 is a tumor suppressor.
Both Fbx4+/- and Fbx4-/- aged mice commonly develop lymphoid malignancies involving the mesenteric node, spleen, liver, and intestine, as well as histiocytic sarcomas of the intestinal wall and mesentery, dendritic cell tumors of the spleen and thymus, early myeloid malignancies, and mammary, hepatocellular, and uterine carcinomas occurring at lesser frequencies. Fbx4+/- tumors retain Fbx4 expression, consistent with human cancer data revealing hemizygous Fbx4 mutations (Barbash et al., 2008). In addition, Fbx4+/- and Fbx4-/- MEFs are susceptible to single step, Ras-mediated transformation, supporting the notion that Fbx4 functions as a haploinsufficient tumor suppressor. While an elevation in cyclin D1 protein in Fbx4+/- and Fbx4-/- in epithelial tissue might be anticipated, the increased cyclin D1 expression is in lymphocytes, which typically do not express cyclin D1, and lymphoid tumors were indeed enlightening.  These results support a requisite role for posttranslational degradation as a means to limit cyclin D1 accumulation in normal lymphoid tissue, highlighting the tumor suppressive function of Fbx4 in this tissue. Interestingly, not all tissues exhibiting elevated cyclin D1 develop neoplastic lesions; for example cyclin D1 is elevated in normal Fbx4+/- and Fbx4-/- lung tissue; however, spontaneous lung adenocarcinomas were not observed. This raises the possibility that development in the absence of Fbx4 results in compensatory pathways for cyclin D1/CDK4 regulation, ultimately antagonizing nuclear function when Fbx4-mediated degradation is impaired. 
Fbx4+/- and Fbx4-/- mice develop tumors with a protracted latency that is comparable to pituitary tumors observed in p18Ink4c-/- mice, sarcomas in p15Ink4b-/- mice, and B-cell lymphomas, lung adenocarcinomas, and hepatocellular carcinomas in U19/Eaf2-/- mice (Latres et al., 2000; Xiao et al., 2008). Although tumors arise in aged mice, it is important to note that this is a remarkably penetrant phenotype, with 38 percent of Fbx4+/- and Fbx4-/- mice exhibiting substantial lymphoma burden and 23.4 percent exhibiting EMH before 24 months of age. Furthermore, the frequent incidence of EMH and associated early myeloid malignancy suggests bone marrow failure, and it is of interest to assess the role of Fbx4 in hematopoietic development and bone marrow function in future work. 

Loss of Fbx4 promotes cyclin D1 nuclear accumulation and DNA damage.
	Previous work has linked nuclear cyclin D1 accumulation with tumorigenesis (Gladden et al., 2006; Lin et al., 2007). However, the rare incidence of cyclin D1 mutations that impair nuclear export and cytoplasmic degradation suggests that other components of the cyclin D1 regulatory machinery, such as Fbx4, are targeted during neoplastic transformation. Consistent with this notion, we observed a substantial extension of cyclin D1 half-life in S-phase Fbx4-/- MEFs, with a moderate, yet significant, increase in cellular proliferation and substantial nuclear cyclin D1 accumulation. Importantly, accumulating evidence suggests that cyclin D1 nuclear localization during S-phase, not simply overexpression, underlies its oncogenic function (Aggarwal et al., 2007; Aggarwal et al., 2010; Gladden et al., 2006; Lin et al., 2007). 
Consistent with the aforementioned nuclear cyclin D1 phenotype, the initial proliferative advantage observed in Fbx4-/- MEFs dramatically decreases in later passages, suggesting that cells either activate a senescence program prematurely, or decrease proliferation as a consequence of DNA damage and/or additional stress responses. Our data support the generation of DNA damage, as DNA damage checkpoint activation is clearly evident in passage 3-5 MEFs. Importantly, DNA damage observed in Fbx4-/- MEFs is not simply due to oxidative stress, as growth under physiological (3%) O2 tension does not reduce DNA damage. In addition, DNA damage is not likely to result from stabilization of the other known Fbx4 substrate, Trf1, given that telomere uncapping effects of Trf1 stabilization would take many generations, due to the extremely long telomeres observed in most laboratory mouse lines (Lee et al., 2006). Nuclear cyclin D1 retention during S-phase promotes DNA re-replication and subsequent DNA damage checkpoint activation; mechanistically, nuclear cyclin D1/CDK4 kinase activity is necessary for elevated histone arginine methylation at the CUL4 promoters, CUL4 repression, and consequent stabilization of the replication factor CDT1 (Aggarwal et al., 2007; Aggarwal et al.). In fact, tumors arising in Fbx4+/- and Fbx4-/- mice exhibit a dramatic increase in histone 4 arginine 3 symmetric di-methylation (H4R3, Figure 3.4A), and Fbx4-/- MEFs exhibit CDT1 stabilization in S-phase, with a concomitant reduction in CUL4 levels, supporting the notion that CDT1 stabilization downstream of nuclear cyclin D1/CDK4 activity drives DNA damage accumulation.
Our previous and current data reveal a tightly regulated mechanistic program for cyclin D1 control in S-phase and following DNA damage. Fbx4 serine 12 phosphorylation, an event preceding ligase dimerization and activation, ensures an increased rate of cyclin D1 ubiquitylation following the G1/S transition. Intriguingly, both nuclear cyclin D1 T286 and cytoplasmic Fbx4 S12 are phosphorylated by GSK3 at this phase of the cell cycle, highlighting the dual regulatory role posttranslational modifications play in coordinating timely cyclin D1 destruction (Barbash et al., 2008; Diehl et al., 1998). Therefore, it is not surprising that both cyclin D1 T286 and Fbx4 S12 phosphorylation events are induced above basal levels following S-phase DNA damage, thereby facilitating increased ubiquitin ligase activity toward cyclin D1 substrate (Figure 3.7C, D). Considering that nuclear cyclin D1 accumulation promotes DNA re-replication and checkpoint activation, and that impaired cyclin D1 degradation in response to S-phase DNA damage drives genomic instability, loss or functional inactivation of Fbx4 provides oncogenic insults necessary for neoplastic transformation. 
Collectively, our current work supports the notion that Fbx4 expression is essential for cell homeostasis in somatic cells. Fbx4 ablation predisposes mice to multiple tumor phenotypes, with concomitant cyclin D1 deregulation and nuclear accumulation. Furthermore, our data suggest that Fbx4 status is an important determinant of neoplastic potential in cells harboring wild type cyclin D1 overexpression, functioning as a haploinsufficient tumor suppressor. Identification of novel Fbx4 substrates will elucidate additional signaling pathways contributing to the neoplastic phenotype in Fbx4-/- animals functioning in concert with cyclin D1, and future development of Fbx4 conditionally-deficient mice will enable Fbx4 ablation in various adult tissues, mirroring the somatic inactivating mutations observed in human cancers. 

Figure 3.1.

Figure 3.2.

Figure 3.3.

Table 1.

Figure 3.4.

Figure 3.5.

Figure 3.6.

Figure 3.7.

Figure 3.8.

CHAPTER IV:
nuclear cyclin D1/CDK4 kinase regulates cul4 expression and triggers neoplastic growth via activation of the prmt5 methyltransferase

this chapter has been published as a co-first author article with Dr. Priya Aggarwal in:
Cancer Cell 18 (4): 329-340. October 19, 2010. 
Reprinted by permission from Elsevier
License Number: 2661040720840

SUMMARY
Cyclin D1 elicits transcriptional effects through inactivation of the retinoblastoma protein and direct association with transcriptional regulators. The current work reveals a molecular relationship between cyclin D1/CDK4 kinase and protein arginine methyltransferase 5 (PRMT5), an enzyme associated with histone methylation and transcriptional repression. Primary tumors of a mouse lymphoma model exhibit increased PRMT5 methyltransferase activity and histone arginine methylation. Analyses demonstrate that MEP50, a PRMT5 co-regulatory factor, is a CDK4 substrate, and phosphorylation increases PRMT5/MEP50 activity. Increased PRMT5 activity mediates key events associated with cyclin D1-dependent neoplastic growth, including CUL4 repression, CDT1 overexpression, and DNA re-replication. Importantly, human cancers harboring mutations in Fbx4, the cyclin D1 E3 ligase, exhibit nuclear cyclin D1 accumulation and increased PRMT5 activity.

INTRODUCTION
Cyclin D1, the allosteric regulator of CDK4 and CDK6, is an integral mediator of growth factor-dependent G1-phase progression. Growth factor stimulation induces cyclin D1 expression, association with CDK4 and nuclear accumulation during mid-G1; active cyclin D1/CDK4 kinase catalyzes phosphorylation-dependent inactivation of the retinoblastoma (RB) family proteins (Diehl, 2002).  Following the G1/S transition, cyclin D1 accumulation is opposed by Pro287-directed phosphorylation of threonine 286 (T286) by glycogen synthase kinase 3β (GSK3β), which promotes its nuclear export (Alt et al., 2000). Cytoplasmic, phosphorylated cyclin D1 is polyubiquitylated by SCFFbx4 and degraded by the 26S proteasome (Lin et al., 2006).
Overexpression of cyclin D1 occurs in numerous human malignancies including carcinomas of the breast, esophagus, colon, and lung (Bani-Hani et al., 2000; Bartkova et al., 1994a; Bartkova et al., 1995; Bartkova et al., 1994b; Gillett et al., 1994; Herman et al., 1995; Hibberts et al., 1999; Hosokawa and Arnold, 1998; Hosokawa et al., 1999; Jin et al., 2001). While overexpression of cyclin D1 is often observed, overexpression per se is insufficient to drive spontaneous transformation.  The primary cellular mechanism that restricts cyclin D1/CDK4 activity is cytoplasmic, ubiquitin-mediated degradation of cyclin D1 during S-phase.  Mutations that disrupt this event directly contribute to neoplastic growth (Aggarwal et al., 2007; Alt et al., 2000; Barbash et al., 2008; Benzeno et al., 2006; Lu et al., 2003).  Specifically, inhibition of cyclin D1 proteolysis during S-phase via mutations within the cyclin D1 degron or inactivation of the cyclin D1 E3 ligase, Fbx4, triggers constitutive nuclear accumulation of active cyclin D1/CDK4 complexes which, in turn, disrupt temporal regulation of DNA replication, thereby contributing to neoplastic growth.  The disruption of S-phase fidelity reflects accumulation of the replication licensing protein CDT1. Inappropriate CDT1 stabilization is a result of nuclear cyclin D1/CDK4-dependent repression of CUL4A and CUL4B, encoding scaffolding proteins for the E3 ligase that directs CDT1 degradation during S-phase (Aggarwal et al., 2007).  
Numerous previous studies have linked cyclin D1 with transcriptional repression.  Current models focus on an intrinsic capacity of cyclin D1 to associate directly with transcriptional regulators and either interfere with the recruitment of co-factors that direct gene repression or conversely, facilitate co-activator recruitment as observed in the collaboration between cAMP signaling and cyclin D1 in the activation of estrogen receptor (ER)-mediated transcription in mammary epithelial cells (Lamb et al., 2000).  The strongest evidence for cyclin D1 directly contributing to the regulation of gene expression stems from recent work utilizing genome-wide chromatin immune precipitation to identify promoters occupied by cyclin D1 (Bienvenu et al., 2010).  
A common theme throughout this previous work is that cyclin D1 modulates gene expression in a kinase-independent manner.  However, transcriptional repression of CUL4A and CUL4B by cyclin D1 requires S-phase accumulation of catalytically active cyclin D1/CDK4 (Aggarwal et al., 2007). The current study dissects the molecular mechanism of CUL4A/B transcriptional regulation by cyclin D1 and its contribution to nuclear cyclin D1-driven neoplastic growth. 

RESULTS
Association of the PRMT5/MEP50 methyltransferase with cyclin D1T286A/CDK4.
To decipher mechanisms of nuclear cyclin D1-driven neoplasia, we immunopurified cyclin D1T286A complexes from tumors derived from E-D1T286A transgenic mice (Gladden et al., 2006) and identified co-purifying proteins by mass spectrometry (data not shown).  Given previous reports linking cyclin D1 with transcriptional repression, we were intrigued by co-purification of PRMT5, a Type II methyltransferase, along with MEP50, a WD40 repeat containing protein that contributes to PRMT5 activity and substrate recruitment (Krause et al., 2007).  Co-precipitation of cyclin D1T286A, CDK4, MEP50 and PRMT5 from B-cell tumors confirmed the interaction (Figure 4.1A-B). To ascertain the existence of this complex in human cancer cells, we used TE3 and TE7 cell lines that harbor cyclin D1P287A (Benzeno et al., 2006). Cyclin D1P287A is refractory to GSK3β-dependent phosphorylation and is stabilized in the nucleus, analogous to D1T286A. Consistent with the above results, we observed co-precipitation of cyclin D1P287A along with MEP50, PRMT5, and CDK4 (Figure 4.1C). 
Association of cyclin D1T286A with PRMT5 in concert with previous work demonstrating that cyclin D1T286A reduces CUL4A/B expression in a CDK4-dependent manner (Aggarwal et al., 2007) suggested a potential regulatory relationship involving cyclin D1T286A and chromatin modifying proteins. To interrogate this putative relationship, we co-expressed Myc-PRMT5 together with CDK4 and either wild type cyclin D1 or D1T286A in HeLa cells. Following synchronization at the G1/S boundary and release into S-phase, complexes were immune precipitated and associated proteins identified by immunoblot.  Cyclin D1T286A was enriched in PRMT5 complexes at the G1/S boundary and declined as cells progressed through S-phase (Figures 4.1D). In contrast, low levels of wild type cyclin D1 were associated with PRMT5, with no obvious enrichment.  The reduced binding of wild type cyclin D1 reflects both its low abundance, due to proteolysis during S-phase, and cytoplasmic localization of the remaining protein (Alt et al., 2000; Barbash et al., 2008).  
To determine whether known components of chromatin remodeling complexes were also present in the cyclin D1T286A-PRMT5 complex, we performed a two-step affinity purification of this complex.  Initially, Flag-D1T286A-containing complexes were collected from pooled tumor lysates by M2-affinity chromatography, which was used to isolate flag-tagged cyclin D1. Complexes were eluted with flag peptide then re-precipitated with PRMT5 antibodies.  In addition to the expected components (cyclin D1T286A, CDK4, PRMT5, MEP50) we also noted Brg1 (Figure 4.1E). Importantly, these complexes do not contain RB, PRMT5-related PRMT1 and PRMT7, mSin3a (a component of large multi-subunit histone deacetylase (HDAC) co-repressor complexes), or Mi2 (a nucleosome remodeling and HDAC complex component, Figure 4.1E). This result reveals the existence of a cyclin D1T286A, CDK4, MEP50, PRMT5 and Brg1 complex. Consistently, we also noted co-precipitation of Brg1 with endogenous cyclin D1P287A in TE3 and TE7 cells (Figure 4.1C). Consistently, size exclusion chromatography revealed co-fractionation of cyclin D1T286A, CDK4, PRMT5, MEP50, and Brg1 in murine lymphomas (data not shown). 

PRMT5/MEP50 mediates cyclin D1T286A/CDK4-dependent CUL4A/B loss and CDT1 stabilization. 
Because PRMT5-dependent di-methylation of histone H3 arginine 8 (H3R8) and histone H4 arginine 3 (H4R3) is associated with transcriptional repression (Pal et al., 2004), we considered whether PRMT5 would mediate cyclin D1T286A/CDK4-dependent repression of CUL4A/B. We first used the general methyltransferase inhibitor 5’ Deoxy-5”-methyl-thioadenosine (MTA) (Hou et al., 2008; Iwasaki and Yada, 2007), following identification of a concentration that inhibited H4R3 methylation but not H3K4 methylation (Figure 4.2A). As previously observed, expression of catalytically active cyclin D1T286A/CDK4 resulted in the maintenance of CDT1 and loss of CUL4A/B proteins and mRNAs specifically during S-phase (Figures 4.2B-D). Treatment with 100mM MTA restored CUL4A/B proteins and mRNAs and triggered CDT1 degradation during S-phase (Figure 4.2B). A similar restoration of CUL4A/B was observed with another PRMT inhibitor, AMI-1 (Cheng et al., 2004) (data not shown). 
As an independent assessment of PRMT5 function, we utilized siRNA directed at PRMT5 or MEP50. Knockdown of PRMT5 (Figures 4.2E; G-H) or MEP50 (data not shown) restored expression of CUL4A/B and CDT1 loss during S-phase. Knockdown of PRMT5 also reduced H4R3 and H3R8 methylation but did not change levels of PRMT1, 2, and 7 (Figure 4.2E). We confirmed previous observations that regulation of CUL4 and CDT1 requires the kinase activity of CDK4 as expression of a kinase defective mutant, CDK4(K35M), does not support cyclin D1T286A-dependent CUL4 loss and eliminates the impact of PRMT5 knockdown (Figure 4.2F).
Since attenuation of PRMT5 relieves CUL4 repression, we questioned whether cyclin D1T286A regulates HDAC activity, contributing to CUL4 repression. Synchronized HeLa cells were treated with the HDAC inhibitors sodium butyrate, trichostatin A and nicotinamide; each inhibitor increased CUL4 expression independent of cyclin D1T286A.  Furthermore, acetylation of histone H3 and H4 within the CUL4A promoter and CUL4B promoter (data not shown) was not influenced by cyclin D1T286A, suggesting that HDAC activity is not coordinately regulated with histone arginine methylation by cyclin D1T286A/CDK4 at the CUL4 promoters. 

Cyclin D1T286A/CDK4 kinase directs increased PRMT5 activity and PRMT5-dependent methylation of CUL4 promoters.  
If PRMT5 mediates the action of cyclin D1T286A/CDK4 in vivo, we reasoned that increased H3R8 di-methylation, a PRMT5-specific mark (Pal et al., 2004), should be apparent in tumors derived from E-D1T286A mice. Methylated H3R8 was elevated in tumors compared to non-transgenic controls (Figure 4.3A-B). Given the increase in total H4R3/H3R8 methylation, we determined whether there was an increase in H4R3/H3R8 methylation on the proximal CUL4A/B promoter regions by ChIP using antibodies detecting di-methylated H4R3 and di-methylated H3R8 (Primer design, Figure 4.3C).  We observed an ~2-3 fold increase in methylation of both H4R3 and H3R8 at the proximal promoter regions (~500bp upstream of the first coding exon) of CUL4B (Figure 4.3D) in the presence of cyclin D1T286A/CDK4 as compared to cyclin D1/CDK4 or untransfected controls in S-phase HeLa cells.  No enrichment was detected with primers directed to regions corresponding to 1000bp upstream (data not shown) or ~5000bp upstream of first coding exon of CUL4B (control/blue bars Figure 4.3D).  Using PRMT5 specific antibodies for ChIP, we noted PRMT5 occupancy of the same region of the CUL4 promoters (Figure 4.3E).  Critically, siRNA-mediated knockdown of PRMT5 resulted in significant attenuation of cyclin D1T286A/CDK4 dependent H4R3/H3R8 methylation at CUL4A/B promoter regions during S-phase (Figure 4.3F). In agreement with cyclin D1T286A directly contributing to increased PRMT5 activity, cyclin D1T286A occupancy of both Cul4A/B promoters was observed in cyclin D1T286A expressing primary murine lymphoma cells (Figure 4.3G, data not shown). 
To determine whether increased methylation of histones reflected increased PRMT5 activity, we assessed methyltransferase activity of PRMT5 immune-purified from control or tumor-burden spleens and found that four of six tumors exhibited an increase in PRMT5 activity (Figure 4.4A).  While PRMT5 levels are increased in cyclin D1T286A tumors, it does not appear sufficient to account for the dramatic increase in PRMT5 catalysis observed. To more directly determine whether this increase reflects the action of cyclin D1T286A/CDK4, we reconstituted this pathway in HeLa cells.  Consistently, PRMT5 isolated from S-phase cells expressing cyclin D1T286A exhibited enhanced methyltransferase activity; a small but significant increase was also observed in cells overexpressing wild type cyclin D1/CDK4 (Figure 4.4B). Furthermore, we observed increased methylation of recombinant H3R8 in vitro (Figure 4.4C).

Phosphorylation of MEP50 by cyclin D1T286A/CDK4 in vitro and in vivo.
We next considered whether either MEP50 or PRMT5 might be a cyclin D1/CDK4 substrate. PRMT5/MEP50 complexes were precipitated from HeLa cells expressing cyclin D1 or cyclin D1T286A along with either CDK4 or CDK4(K35M) with a MEP50 antibody. Upon immunoblotting with an antibody detecting phosphorylated serine or threonine when either residue is followed by a proline, we noted a band in cells expressing cyclin D1T286A/CDK4 that migrated with the expected mobility of MEP50 (Figure 4.5A, noted by arrow).  No such band was apparent in cyclin D1 expressing cells or in cyclin D1T286A/CDK4(K35M) expressing cells.  In addition, we did not detect any apparent p-SP/TP signal corresponding to PRMT5.
MEP50 contains four putative CDK phosphorylation sites: T5, S176, S264, and S306 (Figure 4.5B). Consistent with MEP50 being a direct substrate, recombinant MEP50 was phosphorylated by cyclin D1/CDK4 and D1T286A/CDK4 in vitro (Figure 4.5C). To identify phosphorylated residues, recombinant MEP50 was phosphorylated in vitro with purified cyclin D1T286A/CDK4 and subjected to mass spectrometry; we also affinity purified MEP50 from cells co-expressing cyclin D1T286A/CDK4 for the same analysis. Both approaches revealed phosphorylation of MEP50 at threonine 5 (T5, data not shown). We generated recombinant MEP50 proteins harboring alanine substitutions at all four putative CDK phosphorylation sites and used these as substrates.  Mutation of T5 dramatically reduced phosphate incorporation; mutation of S264 and S306 also resulted in a moderate reduction while S176A had no apparent effect (Figure 4.5D). 
If CUL4 repression and CDT1 stabilization depends upon MEP50 phosphorylation by cyclin D1T286A/CDK4, we reasoned that overexpression of key non-phosphorylatable MEP50 mutants should be inhibitory. Indeed, expression of myc-tagged MEP50-T5A and to a lesser degree MEP50-S264A interfered with cyclin D1T286A-dependent CDT1 stabilization and CUL4 loss during S-phase (Figure 4.5E). Collectively, these results reveal that MEP50 is a CDK4 substrate and that phosphorylation of MEP50 predominantly on T5 dominantly mediates downstream function. 

Cyclin D1T286A/CDK4 regulates PRMT5 methyltransferase activity via MEP50 phosphorylation.
To determine whether phosphorylation of MEP50 can directly regulate PRMT5 methyltransferase activity independent of association with high molecular weight chromatin remodeling complexes, coupled in vitro kinase/methyltransferase reactions were performed with purified recombinant PRMT5/MEP50 produced in Sf9 cells. PRMT5-dependent methyltransferase activity was increased following a kinase reaction with purified cyclin D1T286A/CDK4 (Figure 4.6A). Additionally, increased PRMT5/MEP50 SAM-dependent methyltransferase activity directed towards histone H4 was readily apparent following incubation of PRMT5/MEP50 complexes purified from HeLa cells with cyclin D1T286A/CDK4 and to a lesser degree with cyclin D1/CDK4 (Figure 4.6B).  To evaluate the role of MEP50 phosphorylation, we expressed either wild type MEP50 or mutant MEP50 alleles in cells treated with MEP50 siRNA. Expression of MEP50-T5A maintained basal methyltransferase activity in vitro, but was refractory to the CDK4-dependent increase of PRMT5 activation (Figure 4.6C).  Serine-264 to alanine substitution also attenuated the response to cyclin D1T286A/CDK4 whereas alanine substitution at 176/306 was without influence.  These results demonstrate that cyclin D1/CDK4 can increase histone methyltransferase activity of PRMT5/MEP50 through phosphorylation of T5 and perhaps through S264. 

PRMT5/MEP50 mediates cyclin D1T286A-dependent re-replication, transformation and survival of cyclin D1T286A expressing lymphoma cells. 
Cyclin D1T286A/CDK4 expression induces re-replication in a CDT1-dependent fashion (Aggarwal et al., 2007). If PRMT5/MEP50 mediates re-replication, knockdown of PRMT5 should be inhibitory. Consistent with this hypothesis, we observed a dramatic reduction of >4N population from 26.8% to 2.28% with PRMT5 knockdown in T286A/CDK4/CDT1 transfected HeLa cells (Figure 4.7A-B). 
We next determined whether PRMT5 mediates cyclin D1T286A-dependent transformation by knockdown of PRMT5 in murine fibroblasts, concurrent with expression of oncogenic RasV12 and cyclin D1T286A. Indeed, while co-transfection of both RasV12 and cyclin D1T286A induced focus formation (Figure 4.7C), foci number was reduced by 50% following knockdown of PRMT5. Importantly, PRMT5 knockdown was not accompanied by cell cycle arrest, suggesting that foci reduction is not a consequence of cell cycle arrest (data not shown). In addition, PRMT5 knockdown did not significantly attenuate transformation mediated by Ras plus c-Myc, suggesting mechanistic specificity between PRMT5-dependent methyltransferase activity and constitutively nuclear cyclin D1 (Figure 4.7C).  
To investigate therapeutic potential of targeting PRMT5, we determined whether inhibition of PRMT5 would compromise survival of primary Em-D1T286A lymphoma cells ex vivo.  Because we were unable to efficiently introduce siRNA into these primary tumor cells, we utilized AMI-1. AMI-1 treatment increased death of tumor cells relative to normal controls (Figure 4.7D). In addition, treatment of primary lymphoma cells with MTA also triggered increased cell death relative to untreated tumor cells or MTA treated control lymphocytes (data not shown). Together, these observations suggest that PRMT5 plays an important role in promoting cyclin D1T286A/CDK4-dependent DNA re-replication, cell transformation and survival of cells harboring constitutively active cyclin D1/CDK4.
To examine the role of MEP50 phosphorylation in cyclin D1T286A-dependent transformation, wild type or phosphorylation-deficient T5A or S264A mutant MEP50 was co-expressed with RasV12 and cyclin D1T286A in NIH3T3 cells. Compared to wild type MEP50, expression of the T5A or S264A MEP50 mutant significantly reduced foci number, suggesting that phosphorylation of MEP50 and subsequent increase in PRMT5 activity is a driving force for cellular transformation in the presence of cyclin D1T286A (Figure 4.7E).

Fbx4 inactivation promotes increased PRMT5-dependent histone methylation.
While mutations directly targeting cyclin D1 do occur in human cancer (Benzeno et al., 2006), they occur infrequently relative to inactivation of the cyclin D1 E3 ligase (Barbash et al., 2008).  Because inactivation of Fbx4 also drives nuclear accumulation of cyclin D1 kinase during S-phase, we postulated that Fbx4 loss should also impact PRMT5 function.  We initially assessed this hypothesis in NIH 3T3 cells harboring Fbx4 shRNA (Lin et al., 2006). Fbx4 knockdown increased cyclin D1 and CDT1 and decreased CUL4A/B; critically, the level of H4R3 methylation was dramatically increased (Figure 4.8A). Previous work identified esophageal tumors harboring inactivating mutations in Fbx4 that resulted in strong cyclin D1 overexpression (Barbash et al., 2008).  We chose these same tumors to ascertain whether loss of Fbx4 function and overexpression of wild type cyclin D1 kinase also correlated with increased H4R3 methylation.  Immunohistochemical staining on tumor sections revealed a marked increase in H4R3 methylation relative to an esophageal tumor harboring wild type Fbx4 or normal esophageal epithelium (Figure 4.8B). Because our analysis was restricted to tumors with known Fbx4 mutations and tissue availability for IHC (n=4), our analysis did not achieve statistical significance. To support this analysis, we analyzed an esophageal tumor cell line harboring a mutant Fbx4, Fbx4(S8R), TE10 cell line) (Barbash et al., 2008) and another expressing both wild type Fbx4 and cyclin D1 (TE15)  and found that inactive Fbx4 correlates with increased CDT1 and methylated H4R3 (Figure 4.8C).  Together, these observations demonstrate that stabilized nuclear cyclin D1 resulting from either cyclin D1 mutation or inactivation of Fbx4 results in increased PRMT5-dependent histone methylation.

DISCUSSION
Cyclin D1 is subject to intricate post-translational control to ensure its temporal regulation. Cyclin D1 nuclear export followed by Fbx4-dependent ubiquitylation and degradation during S-phase are key features that serve to restrain nuclear cyclin D1/CDK4 activity and ensure normal cell division (Alt et al., 2000; Benzeno et al., 2006; Lin et al., 2006). Previous work revealed that aberrant nuclear accumulation of cyclin D1 during S-phase promotes transformation in vitro (Alt et al., 2000) and drives both B-cell lymphomas and mammary carcinomas in mice (Gladden et al., 2006; Lin et al., 2008).  Molecular analysis of these tumors revealed a potential mechanism wherein the cyclin D1-dependent kinase interferes with CUL4-dependent CDT1 proteolysis during S-phase (Higa et al., 2006; Sansam et al., 2006), due to transcriptional repression of CUL4A and CUL4B; the resulting overexpression of CDT1 during S-phase triggers DNA re-replication, enhancing genomic instability and the acquisition of fortuitous mutations (Aggarwal et al., 2007).  A significant feature of the regulatory pathway leading to loss of CUL4 expression is the noted dependence upon cyclin D1/CDK4 activity. The current work revealing that phosphorylation of MEP50 on T5 by cyclin D1T286A/CDK4 mediates PRMT5-dependent transcriptional repression provides key mechanistic insights into the neoplastic activities of the cyclin D1/CDK4 enzyme.
	While PRMT5/MEP50 was identified through affinity purification of D1T286A complexes, it is the demonstration that MEP50 is a direct substrate for D-type cyclin dependent kinase that contributes a key advance in our understanding of cyclin D1-driven tumorigenesis.  Phosphorylation of MEP50 on T5 by D1T286A/CDK4 is necessary and sufficient to increase the intrinsic methyltransferase activity of PRMT5, resulting in increased H4R3/H3R8 methylation and repression of the CUL4A/B expression.  Notably, although we did not detect phosphorylation of S264 in vivo, mutation of S264 to alanine in MEP50 reduced cyclin D1-dependent induction of PRMT5 activity, suggesting that phosphorylation of this site may also contribute.  The fact that inhibition of PRMT5/MEP50 activity by siRNA or overexpression of non-phosphorylatable MEP50 attenuates CUL4 repression is consistent with a model where PRMT5/MEP50 represents a key molecular target of nuclear cyclin D1/CDK4 during S-phase (Figure 4.9).

Regulation of PRMT5 by the cyclin D1-dependent kinase
PRMT5 methylates arginines in multiple proteins including myelin basic protein (Ghosh et al., 1988), SM  proteins (Friesen et al., 2001) and p53 (Jansson et al., 2008).  However, it is PRMT5-dependent histone di-methylation that is associated with transcriptional repression. Of the eleven PRMTs, PRMT5 is closely associated with transcriptional repression (Pal and Sif, 2007; Pal et al., 2004); its repressive function is attributed to symmetric di-methylation of histone 4 (arginine 3) and histone 3 (arginine 8) (Krause et al., 2007).  These histone modifications were recently linked with the subsequent recruitment of DNMT3, linking histone methylation with DNA methylation (Zhao et al., 2009).  
Regulation of PRMT5 function remains poorly understood and modification of PRMT5 by phosphorylation, acetylation or ubiquitylation has not been reported. In contrast, association of PRMT5 with MEP50 is an activating event (Krause et al., 2007). In addition, association of COPR5 with PRMT5 changes the balance of activity from H3R8 towards H4R3 (Lacroix et al., 2008). While PRMT5 is not detectably modified in a CDK4-dependent manner, a multiplicity of approaches revealed MEP50 to be a direct substrate.  The data provided demonstrate that phosphorylation of T5, and perhaps S264, of MEP50 contributes to increased PRMT5/MEP50 methyltransferase activity in cells harboring nuclear cyclin D1/CDK4. As addition of purified MEP50 to recombinant PRMT5 is sufficient to activate PRMT5 methyltransferase function (Krause et al., 2007), the most direct model is that cyclin D1-dependent phosphorylation functions as a switch to facilitate MEP50-dependent, structural alterations in PRMT5 that contribute to enhanced catalysis or perhaps increased affinity for substrates which translates into more efficient methylation.  

Transcriptional regulation of CUL4A/B by cyclin D1
	Previous work demonstrated that nuclear, active cyclin D1/CDK4 during S-phase leads to reduced CUL4A/B expression (Aggarwal et al., 2007).  While the underlying mechanism remained elusive, a large body of work associating cyclin D1 with transcriptional regulation suggested that regulation could be direct. For example, cyclin D1 was first noted to associate with the estrogen receptor (ER) to coordinate ligand-independent expression of ER targets independent of CDK activation (Lamb et al., 2000; McMahon et al., 1999; Zwijsen et al., 1998; Zwijsen et al., 1997).  Subsequently, cyclin D1 was shown to bind and regulate the activities of androgen receptor (Knudsen et al., 1999), DMP1 (Hirai and Sherr, 1996), and C/EBPb (Lamb et al., 2003).  Our current work demonstrates that regulation reflects CDK4-dependent phosphorylation of MEP50, which in turn increases PRMT5-dependent histone methylation of target genes, reducing their expression. While previous work supports a model where cyclin D1 functions as a molecular bridge between DNA-bound transcription factors and co-activators/repressors, our data reveal not only that cyclin D1 occupies promoter regions of a target gene, but also that regulation reflects CDK-dependent phosphorylation of a recruited epigenetic regulatory enzyme.  Consistently, recent work utilizing genome-wide ChIP analysis demonstrated that cyclin D1 is enriched in regions near the transcriptional start site of a large number of genes (Bienvenu et al., 2010),  providing additional evidence for cyclin D1 directly regulating gene expression.  It is intriguing to note that the strongest H4R3 methylation occurs near the predicted transcription start site of both CUL4A and CUL4B. 
The role of CDK4, while perhaps unexpected in the context of transcription, is not totally unexpected in the context of tumorigenesis given that CDK activation is central to the function of cyclin D1 during neoplastic transformation. That MEP50 is a substrate for cyclin D1/CDK4 makes the PRMT5/MEP50 complex one of a limited number of cyclin D1-dependent substrates. Previously, only the RB family members (Kato et al., 1993) and Smad3, a key mediator for TGF- anti-proliferative responses (Liu and Matsuura, 2005; Matsuura et al., 2004), have been established as substrates in vitro and in vivo. 
While increased cyclin D1T286A/CDK4-dependent PRMT5/MEP50 function is required for CUL4 loss and CDT1 overexpression, we cannot rule out the contribution of additional mechanisms that contribute to CUL4 regulation.  For example, neither cyclin D1 nor PRMT5/MEP50 has intrinsic DNA binding properties.  Thus, it remains unclear how cyclin D1T286A/CDK4 promotes the targeting to this locus. In addition, we questioned the possibility that cyclin D1T286A might result in the recruitment of histone deacetylases and thereby impact CUL4 expression. Our results indicated that HDAC inhibition resulted in increased CUL4 expression regardless of cyclin D1T286A/CDK4 suggesting a non-specific effect (data not shown).  
	
 Signaling through PRMT5/MEP50 is not limited to lymphoid tumors harboring mutant cyclin D1 alleles.  
Cyclin D1T286A/CDK4 increases PRMT5 methyltransferase activity in vivo through MEP50, revealing a pathway where PRMT5 facilitates cyclin D1T286A/CDK4 dependent re-replication and survival of murine tumors harboring a lymphoid-specific D1T286A transgene. However, it is important to note that this mechanism is not restricted to this model system or this particular cyclin D1 allele for the following reasons.  First, knockdown of Fbx4, the specificity component of the cyclin D1 E3 ligase, stabilizes nuclear cyclin D1/CDK4 complexes in the nucleus during S-phase and thereby increases PRMT5-dependent histone methylation resulting in CUL4 loss.  Second, human tumors harboring inactivating mutations in Fbx4 and subsequent nuclear accumulation of endogenous cyclin D1-dependent kinase, exhibit a dramatic increase in H4R3 methylation. Finally, cyclin D1/CDK4-MEP50/PRMT5 complexes are observed in esophageal carcinoma cell lines harboring cyclin D1P287A. These independent findings implicate aberrant nuclear cyclin D1 in transcriptional repression of CUL4. 
Collectively, our current work supports a model wherein accumulation of nuclear cyclin D1/CDK4 during S-phase triggers increased PRMT5/MEP50 activity, thereby specifically reducing CUL4 expression (and triggering downstream stabilization of CUL4 ligase targets such as CDT1, Figure 4.9).  Our functional analysis revealed that PRMT5 is necessary for cyclin D1-mediated cell transformation and further that tumors harboring deregulated cyclin D1 exhibit increased PRMT5-dependent histone methylation.  Future efforts to determine the global gene expression pattern altered through cyclin D1-dependent regulation of PRMT5 and which of these specifically contribute to neoplastic growth will be of significant importance.

Figure 4.1.

Figure 4.2.

Figure 4.3.

Figure 4.4.

Figure 4.5.

Figure 4.6.

Figure 4.7.

Figure 4.8.

Figure 4.9.

 

CHAPTER V: 
DISCUSSION, CONCLUSIONS, AND FUTURE DIRECTIONS

the section entitled
“Cyclin D1 regulation following the G1/S transition and genotoxic stress” and
“Checkpoint signaling directs cyclin D1 degradation”	
 have been published in part (including Figure 5.1) in:

Pontano L.L. and Diehl J.A. (2009). 
DNA damage-dependent cyclin D1 proteolysis: 
GSK3 holds the smoking gun. Cell Cycle 8 (6): 824-827.
(Pontano and Diehl, 2009)

	Temporal CDK regulation is achieved through alternating phases of cyclin expression, followed by cyclin destruction via ubiquitin-mediated proteolysis. Disruption in this balance promotes aberrant CDK activity, thereby creating a cellular environment conducive to neoplastic transformation. This concept has been well illustrated by cyclin D1/CDK4; alterations in cyclin D1 phosphorylation, nuclear export, and Fbx4-mediated degradation facilitate accumulation of active cyclin D1/CDK4 kinase in the nucleus following the G1/S transition, driving DNA re-replication and genomic instability. While previous work defined these molecular events, mechanistic details of nuclear cyclin D1/CDK4-dependent genomic instability remained elusive. The work presented in this thesis described a role for SCFFbx4 in regulating cyclin D1 following genotoxic stress, and genetic analysis revealed that Fbx4 functions as a haploinsufficient tumor suppressor in vivo by controlling cyclin D1 accumulation. Critically, this work elucidated a novel mechanism wherein nuclear cyclin D1/CDK4 kinase elicits oncogenic effects by modulating epigenetic regulation. This chapter provides a broad perspective of how deregulated cyclin D1/CDK4 drives cell cycle progression and genomic instability necessary for tumor initiation, highlighting the impact of aberrant cyclin D1/CDK4 activity on checkpoint responses and gene expression. Finally, I will discuss how these mechanistic studies may influence potential therapeutic strategies for cyclin D1-overexpressing tumors.

Cyclin D1 regulation following the G1/S transition and genotoxic stress.
Ubiquitin-mediated cyclin D1 proteolysis following the G1/S transition counters its mitogen-dependent accumulation during G1 phase. Previous work established that cyclin D1 accumulation is antagonized by GSK3-dependent T286 phosphorylation, triggering nuclear export, and SCFFbx4-mediated ubiquitylation within the cytoplasm, targeting cyclin D1 for proteasomal degradation (Alt et al., 2000; Diehl et al., 1998; Diehl et al., 1997; Lin et al., 2006). Although the cellular machinery responsible for this process was previously identified, how this regulatory pathway interfaces to cellular stress responses remained to be established. Here, I identified a phosphorylation- and Fbx4-dependent cyclin D1 degradation mechanism in response to genotoxic stress. Following DSB induction, cyclin D1 is rapidly phosphorylated by GSK3, facilitating nuclear export and subsequent recognition by Fbx4. Critically, nuclear cyclin D1/CDK4 retention following DNA damage compromises the intra-S-checkpoint response, resulting in chromosomal instability. 
Previous work suggested that phosphorylation may be superfluous for DNA damage mediated cyclin D1 proteolysis, as phosphorylation-independent cyclin D1 degradation mediated by APC/C was shown to elicit G1 arrest (Agami and Bernards, 2000). Consistent with this notion, cyclin D1 is not phosphorylated in G1 phase following genotoxic stress; however, rapid cyclin D1 turnover was not observed at this time (unpublished data not shown). Furthermore, phosphorylation-refractory cyclin D1T286A is not degraded in response to DNA damage. Several kinases have been implicated in cyclin D1 T286 phosphorylation during normal cell cycle progression and various stress conditions, including IB kinase alpha (IKK) and p38SAPK (Casanovas et al., 2000; Kwak et al., 2005). While manipulation of these kinases did not impact DNA damage-dependent cyclin D1 phosphorylation, our work revealed that GSK3 activity increases in response to genotoxic stress, and inhibition of GSK3 via shRNA, small molecule inhibitors or dominant negative alleles all effectively inhibit cyclin D1 phosphorylation, highlighting the significance of this kinase for proper control of cyclin D1 accumulation. 
The prerequisite for cyclin D1 phosphorylation in DNA damage-driven degradation suggested a role for the SCFFbx4 ligase in this process.  Consistent with this notion, shRNA-mediated knockdown of both Fbx4 and B crystallin attenuated cyclin D1 degradation; of significance, cyclin D1 regulation could be restored in B crystallin-deficient cell lines following introduction of ectopic B crystallin, suggesting that this SCFFbx4 co-factor is essential for cyclin D1 recognition. A question that arises, then, is whether other G1/S cyclins are also subject to accelerated degradation. Cyclin D2 and cyclin E, for instance, are phosphorylated by GSK3, but are not regulated by Fbx4. Importantly, cyclin D2 and cyclin E accumulation is unaffected by DNA damage, highlighting the specific nature of cyclin D1 regulation in the DNA damage response. 
My work supports a model where genotoxic stress-mediated cyclin D1 degradation occurs in an Fbx4-dependent manner at the G1/S transition, in contrast to phosphorylation-independent degradation via APC/C (Agami and Bernards, 2000) and recent work suggesting Fbxo31-mediated degradation during G1 (Santra et al., 2009). What accounts for these differences? Work describing APC/C-dependent cyclin D1 regulation was performed in MCF-7 cells, a cell line deficient for B crystallin and thus, SCFFbx4 ligase function toward cyclin D1 (Lin et al., 2006), implying that these cells use alternative degradation pathways to terminate cyclin D1 protein expression. Consistent with this notion, phosphorylation-deficient cyclin D1 proteins are degraded, albeit with dramatically extended kinetics. Furthermore, GSK3 is inactive in G1 and cannot facilitate Fbx4 ligase activation or cyclin D1 phosphorylation. The Fbxo31 model suggests that direct, ATM-dependent phosphorylation facilitates Fbxo31 stabilization, permitting recognition of ERK-phosphorylated cyclin D1 by this SCF ligase in G1-phase melanoma cells. Again, it is possible that these cells have adopted alternative signaling solutions for regulating cyclin D1, as our work using fibroblasts and various epithelial cell lines indicate that cyclin D1 is not phosphorylated following G1-phase DNA damage. Furthermore, Fbxo31 knockdown in fibroblasts does not influence genotoxic stress-mediated cyclin D1 destruction (unpublished data not shown). While these alternative models predict that G1-phase cyclin D1 degradation is important for cell cycle arrest prior to S-phase, our current model suggests that perhaps a more critical time to ensure cyclin D1 destruction is following the G1/S transition, as nuclear retention of cyclin D1/CDK4 kinase perturbs S-phase fidelity and DNA damage checkpoint integrity. 

Checkpoint signaling orchestrates cyclin D1 degradation.	
	Rapid GSK3- and Fbx4-dependent cyclin D1 destruction suggests that the canonical cyclin D1 proteolysis pathway is activated by DNA damage signaling effectors. Replication stress associated-ATR activation has been proposed to regulate cyclin D1 phosphorylation and degradation during S-phase (Hitomi et al., 2008). In contrast, we found that DSB-inducing treatments including HU, IR, and camptothecin (CPT) induced rapid cyclin D1 phosphorylation and degradation downstream of ATM activation, whereas activation of ATR via aphidicolin treatment was not sufficient to trigger cyclin D1 loss. Furthermore, ATM inhibition or ablation abrogates cyclin D1 phosphorylation following genotoxic stress; however, ATR deletion or shRNA-mediated knockdown does not influence cyclin D1 regulation. 
Although ATM signaling is required for GSK3-dependent cyclin D1 phosphorylation following genotoxic stress, the precise molecular underpinnings that connect ATM, GSK3, and cyclin D1 remain to be established. Several different models of ATM-dependent cyclin D1 phosphorylation can be considered. First, activation of ATM and its downstream effectors, such as Chk2, could directly activate GSK3 (Lee et al., 2007). Consistent with this notion, an increase in GSK3 kinase activity following DNA damage was noted. Second, ATM-dependent regulation of GSK3 interacting proteins could bring the kinase in proximity to cyclin D1 in the nucleus, increasing the efficiency of phosphorylation. Finally, ATM-dependent modification of a cyclin D1-interacting protein could induce a conformational change, making cyclin D1 a better GSK3 substrate. Our initial work to parse out this ATM-dependent signaling pathway reveal that Chk2 only modestly stabilizes cyclin D1 at early time points following DNA damage, suggesting that ATM itself or alternative downstream effector(s) regulate this DNA damage response. Consequently, further characterizing the connection between ATM and cyclin D1 regulation is an exciting question and a future area for investigation.
Considering the above models for increased GSK3 activity, the question arises whether SCFFbx4 ligase activity is also regulated by DNA damage. GSK3β catalyzes both nuclear cyclin D1 phosphorylation, promoting nuclear export, and cytoplasmic Fbx4 phosphorylation, triggering its dimerization and activation (Barbash et al., 2008; Diehl et al., 1998). Are these events linked following DNA damage? Our data suggest that cyclin D1 phosphorylation is the critical event for degradation, as phosphorylation-deficient mutants never reach the cytoplasm. However, the possibility of both increased cyclin D1 phosphorylation and increased Fbx4-dependent ligase activity contributing to rapid kinetics of cyclin D1 loss was quite intriguing. Work presented in chapter 3 began to address this question, as I demonstrated that GSK3β-dependent Fbx4 phosphorylation increases following S-phase DNA damage. Moreover, my work revealed that enhanced phosphorylation correlates with a marked increase in SCFFbx4 ubiquitin ligase activity following S-phase DNA damage. While these data indicate that both cyclin D1 and Fbx4 are targeted by GSK3βthe distinct mechanisms that control both nuclear/cytoplasmic activation and localization of this kinase remain to be established.
Cyclin D1 proteolysis downstream of ATM activation is a cellular checkpoint response to maintain genome integrity, supporting a model wherein DSB-dependent ATM activation triggers rapid, Fbx4-dependent cyclin D1 degradation (Figure 5.1A). In contrast, expression of phosphorylation-deficient cyclin D1 or inactivation of Fbx4 promotes an RDS phenotype associated with CDT1 stabilization and maintenance of MCM proteins on chromatin. Sustained replication in the face of DNA damage potentiates genomic instability. While genomic instability can drive genetic alterations that provide a growth advantage to proliferating cells, severe DNA damage invokes cell death through apoptosis or mitotic failure. Significantly, our recent work demonstrated that nuclear accumulation of cyclin D1 sensitizes cells to the DNA damaging, S-phase-specific chemotherapeutic agent CPT. Ultimately, these findings imply that loss of Fbx4 control compromises the cellular checkpoint response to DNA-damaging chemotherapy, thereby driving substantial genomic instability and cell death (Figure 5.1B). 

Fbx4 is a bona fide tumor suppressor.
	Identification of Fbx4 and B crystallin as key cyclin D1 regulatory components ultimately suggested that these proteins could function as potential tumor suppressors. Indeed, this notion was first validated when a screen of human tumors revealed downregulation of Fbx4 and B crystallin at the protein and mRNA levels (Lin et al., 2006). Additionally, the B crystallin gene is located on chromosome 11q22.3-q23.1, a region commonly deleted in cancer (Dohner et al., 1999). Subsequent analyses revealed that Fbx4 is frequently mutated in esophageal carcinoma, with a majority of mutations disrupting residues necessary for ligase dimerization/activation (Barbash et al., 2008). Such sequencing efforts support a role for Fbx4 in tumor suppression, and future analysis of many different tumor subsets harboring cyclin D1 protein overexpression will likely reveal additional alterations. For example, a prime suspect for Fbx4 mutation or inactivation is mantle cell lymphoma (MCL). A genetic hallmark in MCL is the t(11;14) chromosomal translocation, resulting in ectopic cyclin D1 expression (Marzec et al., 2006). While overexpression of wild type cyclin D1 is typically insufficient for neoplastic transformation, mutation or loss of Fbx4 could facilitate nuclear cyclin D1 accumulation necessary for this process. Furthermore, a subset of 11q22-23 deletions in MCL affect ATM expression; loss of ATM could, in turn, prevent cyclin D1 degradation in response to DNA damage during the transformation process, thereby providing a “second hit” through mechanisms described in chapter 2 of this thesis.
	In addition to tumor sequencing studies, cell culture and orthotopic transplant models suggested that Fbx4 attenuation or inactivation permits cell transformation. Knockdown of Fbx4 promotes anchorage-independent growth; critically, Fbx4 knockdown in cyclin D1-/- MEFs fails to recapitulate this phenotype, suggesting that transformation depends on cyclin D1 stabilization. Furthermore, inhibition of Fbx4 function cooperates with cyclin D1 in driving tumor growth, as MMTV-D1 transgenic mammary epithelial cells infected with F-box Fbx4 (dominant negative) readily formed tumors in fat pads of NOD/SCID mice (Barbash et al., 2008). Finally, Fbx4 attenuation facilitates nuclear cyclin D1 accumulation and chromosomal instability following genotoxic stress. While these findings indirectly implicate Fbx4 as a tumor suppressor, experiments that directly address this question have not been documented to date. Consequently, in chapter 3 of thesis, Fbx4-dependent tumor suppression was directly interrogated through targeted disruption of the murine Fbx4 locus. These data reveal that Fbx4 loss does not impact development; however, a significant percentage of Fbx4+/- and Fbx4-/- mice succumb to age-associated malignancy.
	The most common tumor phenotype in Fbx4+/- and Fbx4-/- aged mice are large cell lymphomas of B- and T-cell origin initiating in the spleen and mesenteric node; lymphoma infiltrates were also observed in the liver and intestine. Less frequent malignancies including histiocytic sarcoma/dendritic cell tumors, hemangioma, early myeloid tumors, and mammary, uterine, and hepatocellular carcinomas were observed (Table 2). While tumors of epithelial cell origin are expected based on the requirement for cyclin D1 in proliferation of these tissues, the frequent occurrence of lymphoid tumors is quite intriguing, since a large body of evidence suggested that cyclin D2/D3 drives proliferation of the lymphocyte compartment, while cyclin D1 protein is not expressed at significant levels. It was initially thought that control of cyclin D1 expression in lymphocytes occurred at the transcriptional level; however, our data revealing cyclin D1 accumulation in Fbx4-deficient normal lymphoid tissue and tumors support a requisite role for posttranslational degradation in limiting cyclin D1 accumulation. Additionally, frequent incidence of extramedullary hematopoiesis and early myeloid malignancy suggests bone marrow failure; assessing the role of Fbx4 in hematopoietic development and bone marrow homeostasis is an area for future work. 
Fbx4+/- lymphomas retain Fbx4 expression, consistent with human cancer data revealing hemizygous Fbx4 mutations (Barbash et al., 2008), suggesting that Fbx4 functions as a haploinsufficient tumor suppressor in vivo. Furthermore, Fbx4+/- and Fbx4-/- MEFs are susceptible to single step, Ras-mediated transformation, also supporting haploinsufficiency in vitro. Another well-studied F-box protein, Fbw7, was initially thought to be a haploinsufficient tumor suppressor based on frequent hemizygous mutations in cancer and rare incidence of loss or deletion; however, biochemical characterization of such mutations within the substrate binding domain revealed dominant-negative character toward the remaining wild type protein, and Fbw7+/- mice are refractory to spontaneous tumorigenesis (Onoyama et al., 2007; Tetzlaff et al., 2004; Welcker and Clurman, 2008). In contrast to Fbw7, our work demonstrates that expression of a single Fbx4 allele is not sufficient to maintain cell homeostasis. 
While pre-malignant or normal Fbx4+/- and Fbx4-/- tissues including the lung exhibit cyclin D1 protein accumulation, lung adenocarcinomas and other epithelial-lineage tumors are not frequently observed. It is possible that conventional Fbx4 ablation allows developmental re-wiring of cyclin D1 regulatory circuits in these tissues, resulting in compensatory pathways for cyclin D1/CDK4 regulation. This concept is also illustrated in MEFs. Cyclin D1 stability increases from 15 minutes in primary wild type MEFs to approximately 35 minutes in Fbx4-/- MEFs. Although Fbx4 loss permits cyclin D1 stabilization and nuclear accumulation, cyclin D1 is still relatively unstable compared to constitutively nuclear cyclin D1T286A. Furthermore, the protracted tumor latency in Fbx4+/- and Fbx4-/- mice suggests that alternative mechanisms may initially repress neoplastic effects of cyclin D1/CDK4 or a second hit, such as p53 inactivation, may be required for neoplastic transformation. Therefore, development of a conditional Fbx4 allele is required for tissue-specific Fbx4 ablation in the adult mouse; this genetic manipulation will mirror the somatic inactivating mutations observed in human cancers. Additionally, crossing conventional Fbx4-/- mice onto a p53 heterozygous background could potentially address whether p53 inactivation is required for tumorigenesis; one would suspect that p53 LOH would accelerate tumor onset in conventional Fbx4-/- mice. Similarly, crossing Fbx4-/- mice with an oncogenic driver such as K-Ras (G12D) could answer the question as to whether loss of Fbx4 accelerates tumor progression.
In addition to tumor analysis, molecular characterization of Fbx4+/- and Fbx4-/- MEFs revealed significant nuclear cyclin D1 accumulation, with concomitant DNA damage checkpoint activation, as evidenced by the presence of H2AX foci and p53 accumulation. Fbx4+/- and Fbx4-/- MEFs exhibit similar onset of senescence, and culture under physiological oxygen conditions did not alleviate the DNA damage phenotype, suggesting that an alternative, cell intrinsic mechanism underlies the DNA damage. In addition, DNA damage is not likely to result from stabilization of the other known Fbx4 substrate, Trf1, given that telomere uncapping effects of Trf1 stabilization would take many generations due to the extremely long telomeres observed in laboratory mouse lines (Lee et al., 2006). Consistent with this notion, nuclear cyclin D1 accumulation correlated with CDT1 stabilization in S-phase MEFs, suggesting that DNA re-replication promotes DNA damage and subsequent chromosomal instability when cells are challenged with extrinsic DNA damaging agents. Taken together, our data support a model wherein cyclin D1 nuclear accumulation in the absence of Fbx4 promotes CDT1 stabilization, DNA re-replication, and genomic instability, thereby charging the cell for potential transformation. p53 induction in response to DNA damage initially limits proliferation; however, subsequent alterations in the p53 pathway then facilitate neoplastic growth and tumor formation (Figure 5.2).
The studies presented in chapter 3 support the notion that Fbx4 expression is essential for homeostasis in somatic cells. However, several questions regarding Fbx4 regulation and function remain to be answered. For example, it is unclear whether accumulation of novel Fbx4 substrates contributes to neoplastic transformation in concert with cyclin D1. To address this in a timely manner, Fbx4-/-; cyclin D1-/- MEFs should be generated and utilized in cell transformation assays with constitutively active Ras in vitro. Furthermore, the Fbx4 crystal structure in complex with its other known substrate, Trf1 was recently solved, revealing that the hydrophobic interface between Fbx4 and Trf1 is necessary for substrate-Fbx4 association (Li and Hao, 2010; Zeng et al., 2010). It is of interest to determine whether critical residues within this region are altered in cancer, thereby precluding cyclin D1 binding. Interestingly, the Fbx4 structure revealed that its C-terminal substrate-binding region consists of an atypical small GTPase domain. While this domain lacks nucleotide-binding capacity, it likely functions as scaffold for protein interactions. This region may be important for B crystallin binding and subsequent cyclin D1 recognition. In contrast, Trf1 recognition occurs in a phosphorylation- and B crystallin-independent manner. Elucidation of these different mechanisms for substrate recognition is required to fully understand the molecular architecture and function of SCFFbx4. Additionally, F-box independent functions of Fbx4, such as serving as a scaffold for protein-protein interactions through its GTPase-like domain, remain to be established.

Additional cyclin D1 ubiquitin ligases?
As mentioned previously, both APC/C and SCFFbxo31 have been implicated in cyclin D1 regulation following genotoxic stress. The work presented in chapter 2 suggests that the APC/C does not play a major role in cyclin D1 regulation following G1 or S-phase DNA damage in fibroblasts and several epithelial cell lines, since cyclin D1 levels are unaltered in G1 phase and phosphorylation-dependent degradation mediated by Fbx4 regulates cyclin D1 accumulation following the G1/S transition. Interestingly, recent work revealed that C/EBP induces expression of the APC/C component CDC27, leading to degradation of cyclin D1, along with well established APC/C targets Skp2 and cyclin B (Pawar et al., 2010). In agreement with previous work describing APC/C mediated cyclin D1 degradation, this group utilized MCF-7 cells lacking B crystallin, and CDC27-APC/C mediated cyclin D1 ubiquitylation remains to be demonstrated in vitro. It is possible that APC/C activity may serve as an additional mechanism to limit cyclin D1 accumulation in G1 phase when GSK3 is inactive; however, precise signaling events that potentially facilitate specific recognition by CDC27 or APC/C itself are unclear.
Additional evidence suggested that CUL7/Fbw8 or SCFFbw8 complexes regulate polyubiquitylation of ERK-phosphorylated cyclin D1 at the G1/S transition (Okabe et al., 2006). However, loss of Fbw8 or CUL7 in vivo results in reduced body size and severe proliferative defects in MEFs (Tsutsumi et al., 2008). These results are inconsistent with the expected phenotype of cyclin D1 accumulation, given that overexpression of cyclin D1 facilitates cell growth. ERK-mediated cyclin D1 phosphorylation is also proposed to facilitate Fbxo31-dependent cyclin D1 ubiquitylation in G1 phase. Importantly, in vivo evidence for direct ERK-mediated T286 phosphorylation is weak, and ERK has not been convincingly demonstrated as a cyclin D1 T286 kinase in vitro. Furthermore, Fbxo31 is an inherently unstable protein that is stabilized following ATM-dependent phosphorylation; Fbxo31 stabilization is thought to allow its association with cyclin D1 in response to G1 phase genotoxic stress (Santra et al., 2009). Contrary to these findings, knockdown of Fbxo31 does not affect DNA damage-dependent cyclin D1 degradation, and cyclin D1 is not phosphorylated in G1 phase fibroblasts or epithelial cell lines (data not shown). These results suggest that the ATM-Fbxo31-cyclin D1 axis may be specific to the particular melanoma cell line utilized in that study. 
Correlative evidence linking Fbx4 or B crystallin loss or inactivation in tumor-derived cells with cyclin D1 overexpression suggests that SCFFbx4 function keeps cyclin D1 in check. Consistently, studies directly testing this hypothesis using Fbx4- or B crystallin-specific shRNAs or genetic Fbx4 ablation also reveal significant cyclin D1 accumulation. Although these data support a requisite role for Fbx4 in regulating cyclin D1 accumulation, it is important to note that cyclin D1 is still degraded, despite dramatically reduced kinetics, suggesting that additional ligases may play a secondary role in limiting cyclin D1 protein levels. Nevertheless, Fbx4 ablation or shRNA-mediated knockdown is sufficient to elicit cyclin D1 nuclear accumulation, unmasking cyclin D1’s oncogenic potential. The Fbx4-cyclin D1 axis, therefore, is a classic tumor suppressor-oncogene relationship that must remain balanced in cycling cells.

Mechanism of nuclear cyclin D1-driven genomic instability.
Cyclin D1 is subject to strict posttranslational regulation to ensure its timely destruction at the G1/S transition. Phosphorylation-dependent nuclear export and Fbx4-dependent ubiquitylation/degradation during S-phase constrain nuclear cyclin D1/CDK4 kinase activity (Alt et al., 2000; Benzeno et al., 2006; Lin et al., 2006). This regulation is essential since aberrant nuclear cyclin D1/CDK4 activity during S-phase promotes DNA re-replication (Aggarwal et al., 2007), transformation in vitro (Alt et al., 2000), and tumorigenesis in vivo (Gladden et al., 2006; Lin et al., 2008).  Initial molecular dissection of this process in E-D1T286A-driven lymphomas and cell culture models revealed downregulation of CUL4 expression in a cyclin D1/CDK4-dependent manner; CUL4  loss facilitates CDT1 stabilization, triggering DNA re-replication, genomic instability and acquisition of fortuitous mutations necessary for tumor initiation (Aggarwal et al., 2007).  
This previous work highlighted dependence upon CDK4 activity, suggesting that cyclin D1/CDK4 may phosphorylate novel nuclear substrates during S-phase, leading to transcriptional repression at key target loci. The collaborative work presented in chapter 4 identified a critical regulatory pathway wherein nuclear cyclin D1/CDK4 kinase elicits neoplastic effects through modulation of the PRMT5/MEP50 methyltransferase; MEP50 phosphorylation on T5 mediates PRMT5-dependent histone methylation and subsequent transcriptional repression, providing a novel mechanism for cyclin D1/CDK4-driven cell transformation. Importantly, inhibition of PRMT5/MEP50 activity via siRNA, arginine methyltransferase inhibitors, or overexpression of non-phosphorylatable MEP50 T5A or kinase-defective CDK4 (K35M) restores CUL4 expression and CDT1 degradation, consistent with a model where PRMT5/MEP50 represents a key molecular target of nuclear cyclin D1/CDK4 during S-phase (Figure 4.9).
Identification of a PRMT5-cyclin D1 relationship was especially intriguing, considering that PRMT5 catalyzes arginine methylation on a variety of substrates, including myelin basic protein (Ghosh et al., 1988), SM  proteins (Friesen et al., 2001), p53 (Jansson et al., 2008), and most importantly, histone H3 and H4. Furthermore, there are 11 different PRMTs expressed in mammalian cells, yet only PRMT5 is associated with transcriptional repression (Pal and Sif, 2007; Pal et al., 2004). Additionally, H4R3 and H3R8 methylation marks were recently linked with the recruitment of DNMT3 to DNA, suggesting that coordinated histone methylation with DNA methylation mediates transcriptional repression (Zhao et al., 2009).  
Implication of cyclin D1 in transcriptional regulation is not a new concept, as various studies suggested that cyclin D1 directly associates with transcription factors and transcriptional regulatory complexes. For instance, early work suggested that cyclin D1 association with estrogen receptor (ER) coordinates ligand-independent induction of ER targets (Lamb et al., 2000; McMahon et al., 1999; Zwijsen et al., 1998; Zwijsen et al., 1997). Subsequent work also supported a role for cyclin D1 in regulating androgen receptor activity (Knudsen et al., 1999), DMP1 (Hirai and Sherr, 1996), and C/EBP (Lamb et al., 2003). Therefore, a model for cyclin D1 in transcriptional regulation was that cyclin D1 functions as a molecular bridge between transcription factors and co-activator or repressor complexes. However, all of these studies suggested that cyclin D1 elicits transcriptional effects independent of CDK4 activation. 
Critically, the results presented in chapter 4 indicate that cyclin D1 modulates PRMT5 activity through CDK4-dependent phosphorylation of the activating co-factor MEP50, challenging the previous model of cyclin D1-dependent, CDK4 independent transcriptional regulation. This is the first documented example of cyclin D1/CDK4-dependent phosphorylation of an epigenetic regulatory enzyme, demonstrated both in vivo and in vitro. The exact mechanism underlying methyltransferase activation upon MEP50 phosphorylation is yet to be defined; however, the most direct model is that CDK4-dependent phosphorylation functions as a switch to facilitate MEP50-dependent structural alterations in PRMT5 that contribute to enhanced catalysis or increased affinity for substrates, leading to more efficient substrate methylation. Furthermore, cyclin D1/CDK4 and PRMT5/MEP50 occupy the promoter regions of CUL4A and CUL4B; whether cyclin D1/CDK4 is directly associated with DNA or binding through PRMT5/MEP50 or other chromatin modifying enzymes remains to be established. Consistent with our findings, recent work utilizing genome-wide ChIP revealed that cyclin D1 is enriched in regions near transcriptional start sites of many genes, providing additional evidence for cyclin D1 directly regulating gene expression (Bienvenu et al.).
While it is clear that cyclin D1/CDK4 and PRMT5/MEP50 occupy the CUL4 promoters, future work is required to define the precise mode of cyclin D1/CDK4 recruitment to chromatin. Cyclin D1 and PRMT5/MEP50 lack defined intrinsic DNA binding properties, suggesting that association with additional DNA binding proteins is important for recruitment. Furthermore, the cyclin D1/CDK4/PRMT5/MEP50 ternary complex is likely associated with large chromatin remodeling complexes, since the SWI/SNF component Brg1 is present in cyclin D1/CDK4-containing PRMT5/MEP50 complexes. Recent evidence suggests that the Kruppel-like associated box-zinc finger protein ZNF224 interacts with PRMT5, facilitating PRMT5 recruitment to the L-type aldolase A promoter and subsequent transcriptional repression (Cesaro et al., 2009). Identification of additional PRMT5/MEP50/cyclin D1/CDK4 complex components, such as DNA binding zinc-finger proteins, will assist in elucidating how this ternary complex is assembled and recruited to chromatin.
CDK4-dependent transcriptional regulation is a novel concept; however, this is not unexpected in the context of tumorigenesis given that CDK activation is central to the function of cyclin D1 during cell transformation. Furthermore, identification of MEP50 as a novel cyclin D1/CDK4 target is of interest, given the small repertoire of previously characterized CDK4 substrates, including the Rb family members (Kato et al., 1993; Liu and Matsuura, 2005; Matsuura et al., 2004). Establishment of MEP50 as a CDK4 substrate that contributes to epigenetic regulation of target genes associated with cancer establishes a paradigm whereby cyclin D1 regulation of gene expression occurs independently of Rb yet remains kinase dependent. This raises the question of whether cyclin D2/D3/CDK4 and cyclin E/CDK2 complexes are capable of phosphorylating MEP50. Little is known with regard to tumorigenic activity of constitutively nuclear cyclin D2 and D3, and cyclin E is typically nuclear at the G1/S transition, making a regulatory relationship between these molecules and PRMT5 unlikely. A more attractive model is one wherein exclusive sequences within cyclin D1 or association of a novel bridging adapter protein facilitate specific cyclin D1/CDK4-PRMT5/MEP50 ternary complex assembly and subsequent regulation.
Collectively, the work presented in chapter 4 supports a model wherein accumulation of active, nuclear cyclin D1/CDK4 during S-phase triggers increased PRMT5/MEP50 activity, thereby specifically reducing CUL4 expression (and triggering downstream stabilization of CUL4 ligase targets such as CDT1). Functional analysis revealed that PRMT5 is necessary for cyclin D1-mediated DNA re-replication and cell transformation, and tumors harboring deregulated cyclin D1 exhibit increased PRMT5-dependent histone methylation. Critically, inhibition of arginine methyltransferase activity in cyclin D1T286A expressing transformed splenocytes impairs survival of these tumor cells in vitro, making PRMT5 an attractive target for therapeutic development. Finally, future efforts to determine the global gene expression pattern altered through cyclin D1-dependent regulation of PRMT5 and which genes specifically contribute to neoplastic growth will be of significant importance.
	
Loss of checkpoint integrity and nuclear cyclin D1-driven tumorigenesis.
A current model for cyclin D1-mediated genomic instability and neoplastic transformation is as follows: nuclear cyclin D1/CDK4 accumulation in S-phase activates the PRMT5 methyltransferase through MEP50 phosphorylation. PRMT5/ MEP50 then methylates H3 and H4 at key target promoters, including CUL4A and CUL4B, triggering transcriptional repression. Loss of CUL4 promotes CDT1 stabilization, facilitating DNA re-replication and subsequent genomic instability (Figure 5.3A). DNA damage (DSBs) accrued due to re-replication/replication fork collision activates the ATM-dependent checkpoint response, leading to cell cycle arrest, DNA repair, or p53-dependent apoptosis if damage is irreparable; this checkpoint function is critical to suppress nuclear cyclin D1-driven transformation. For example, pre-malignant spleens expressing the E-D1T286A transgene exhibit markers of DNA damage checkpoint activation such as phosphorylated ATM, Chk2 and H2AX, with a concomitant induction of apoptosis (Aggarwal et al., 2007). Sustained checkpoint activation provides a selective pressure for p53 mutation or inactivation, and alterations in the p53 pathway are commonly observed in cyclin D1T286A-driven lymphomas (Figure 5.3B), (Aggarwal et al., 2007; Gladden et al., 2006). Furthermore, p53 heterozygosity substantially accelerates cyclin D1T286A-driven tumor formation, suggesting that intact checkpoint function is essential to prevent neoplastic growth (Aggarwal et al., 2007).
These findings led to the question of whether loss of the DSB checkpoint responder ATM in the presence of nuclear cyclin D1T286A would sensitize cells to chromosomal instability, resulting in translocation events that facilitate cell transformation and tumor outgrowth (Figure 5.3C). My preliminary work suggests that loss of ATM may, in fact, accelerate cyclin D1T286A-driven lymphomagenesis in a mouse model. Furthermore, initial analysis of several E-D1T286A/ATM-/- murine tumors revealed clonal chromosomal translocations, in contrast to gross aneuploidy previously observed in E-D1T286A-expressing lymphomas (Aggarwal et al., 2007). This phenotype also contrasts that of ATM-/- mice, where tumors arise in CD4+CD8+ immature T-cells coincident with V(D)J recombination; as a result, these tumors typically harbor chromosome 14 translocations involving the T-cell receptor / locus, with t(12;14) as a frequently observed alteration (Liyanage et al., 2000). The fact that preliminary E-D1T286A/ATM-/- cytogenetic phenotype differs from E-D1T286A or ATM-/- tumors suggests potential synergy between nuclear cyclin D1 function and impaired checkpoint response in tumor initiation.
The few E-D1T286A/ATM-/- tumors analyzed in preliminary cytogenetic experiments exhibit clonal translocations commonly involving chromosome 15. The well-characterized t(12;15) translocation, for example, juxtaposes the c-Myc and immunoglobulin heavy-chain (IgH) loci, promoting aberrant c-Myc overexpression (Ramiro et al., 2004). While mature B-lymphocytes are prone to DSBs in Ig loci during class switch recombination (CSR) and somatic hypermutation (SHM), two processes necessary for antibody diversification (Robbiani et al., 2008), we observed t(12;15) translocations in both B- and T-cells suggesting that an alternative mechanism underlies DSB induction and translocations within these regions. Interestingly, the murine c-Myc locus resides on chromosome 15, and an origin of replication was mapped approximately 1kb upstream of c-Myc, conserved in human, mouse, and xenopus (Ghosh et al., 2006; Girard-Reydet et al., 2004). Thus, it is possible that nuclear cyclin D1-dependent CUL4 repression/CDT1 stabilization promotes DNA re-replication in developing B- and T-cells, and frequent replication fork collision/collapse leads to DSB induction. In the absence of ATM, such breaks would not be efficiently sensed and repaired, resulting in chromosomal translocations. 
While my proposed mechanism could occur theoretically at any replication origin, productive chromosome alterations that facilitate loss of tumor suppressor proteins or activation of oncogenes should be clonally selected for tumor growth. Preliminary evidence in support of this model is that clonal translocations typically involve chromosome 15, potentially leading to c-Myc deregulation. Since clonal t(12;15) is present in E-D1T286A/ATM-/- tumors, we also considered whether DNA re-replication could promote breaks downstream of the IgH locus in an AID (Activation-induced cytidine deaminase)-independent manner, since AID is primarily expressed in germinal center B-cells and is involved in CSR and SHM processes during B-cell development. Interestingly, an origin was mapped between the murine 3’IgH regulatory region and the downstream gene, Crip; further analysis revealed that sequences downstream of this region are replicated bi-directionally, while upstream sequences including IgH C, D, and J segments replicate in a single direction (Norio et al., 2005; Zhou et al., 2002). Bi-directional fork movement in the context of DNA re-replication could generate significant DNA damage, permitting chromosome translocations involving this region as well.
To formally examine these hypothesis, future work is focused on demonstrating enrichment of origin binding proteins (CDC45, MCM3, MCM7) within putative replication origins when cyclin D1T286A is expressed in mouse and human cell culture systems. This simplified experimental system should permit analysis of a single S-phase in the presence of acutely overexpressed cyclin D1T286A, and origin binding protein enrichment at c-Myc has been previously established (Ghosh et al., 2006), validating this approach. I will also examine origin binding protein enrichment at the c-Myc locus in vivo by assessing CDC45, MCM, and ORC binding to this region by ChIP in non-transgenic control and E-D1T286A-expressing splenocytes. Together, these descriptive and mechanistic studies are of significant importance with regard to human cancer, as cyclin D1 deregulation is a hallmark of MCL, and t(11;14) translocations responsible for ectopic cyclin D1 expression frequently disrupt the ATM locus on chromosome 11 as well. Future work will include allograft transplant of E-D1T286A or E-D1T286A/ATM-/- tumor cells into recipient mice, followed by treatment with various chemotherapeutic regimens to determine how these genetic alterations influence sensitivity to conventional DNA damaging chemotherapy, thereby elucidating effective regimens for tumor regression.

Concluding remarks.
Previous and current studies characterizing cyclin D1T286A transgenic mice or Fbx4-deficient mice invoke a model where impaired cyclin D1 phosphorylation or Fbx4 ligase inactivation facilitates accumulation of active, nuclear cyclin D1/CDK4 kinase following the G1/S transition. In S-phase, cyclin D1/CDK4 phosphorylates MEP50, thereby triggering increased PRMT5-dependent methyltransferase activity, subsequent CUL4 repression, and CDT1 stabilization, ultimately driving DNA re-replication. Initial DNA damage checkpoint responses counter cell transformation; however, alterations in the p53 tumor suppressor pathway ultimately permit aneuploidy and neoplasia. Similarly, in the absence of checkpoint responders such as ATM, nuclear cyclin D1/CDK4 drives DNA re-replication; however, an impaired checkpoint response to DNA damage likely facilitates chromosomal translocations which could be clonally selected based on growth or survival advantages for transformed cells (Figure 5.4).
Collectively, the analyses presented in this thesis highlight the functional significance of cyclin D1 deregulation and provide novel insight into oncogenic functions of nuclear cyclin D1/CDK4 kinase. Ultimately, elucidating mechanisms of cyclin D1-driven genomic instability will facilitate development of novel detection and therapeutic approaches for diagnosing and treating cyclin D1 overexpressing cancers. Considering these findings, could cyclin D1 overexpression or inactivating mutations in Fbx4 serve as biomarkers for cancer therapy? Preliminary results suggest yes. Inactivating Fbx4 mutations in esophageal carcinoma correlate with an intense histone arginine di-methylation signature by IHC; this mark could, in fact, serve as a biomarker for deregulated cyclin D1/CDK4 function in human tumors. Analysis of a broad range of tumors known to harbor alterations in Fbx4 or constitutively nuclear cyclin D1 will further validate this IHC approach.
Furthermore, mouse models revealed that cyclin D1/CDK4 contributes to neoplasia at least in part through perturbations in DNA replication and loss of genomic integrity; exploiting this phenotype is an attractive therapeutic strategy. For example, one scenario might be to take advantage of the fact that tumors harboring mutations in cyclin D1 or Fbx4 have a compromised DNA damage checkpoint. In theory, treatment of normal cells with chemotherapeutic agents that generate DNA crosslinks should trigger an intra S-phase checkpoint, thereby providing an opportunity for repair. Expression of stabilized cyclin D1 should promote maintenance of CDT1 and MCM complexes on chromatin, promoting continued origin firing without repair, ultimately resulting in mitotic catastrophe. Further work is required to investigate how alterations in cyclin D1 proteolysis might influence cellular responses to DNA damaging therapeutics in vivo. 
Alternatively, drugs that directly target CDKs have been developed, and various CDK inhibitors are currently in clinical trials. The small molecule CDK4/6 inhibitor PD 0332991 is highly specific for these CDKs and elicits cell cycle arrest in vitro and in xenograft models in vivo, as evidenced by reduced Rb phosphorylation, downregulation of E2F target genes, and reduced Ki-67 staining, an IHC marker of proliferation (Fry et al., 2004). Importantly, the mode of action of PD 0332991 is cytostatic, leading to cell cycle arrest, but not apoptosis, suggesting low toxicity for normal cells; however, alone, it is insufficient to elicit tumor killing, highlighting the requirement for combination therapy. In addition to direct CDK inhibition, modulation of signaling responses downstream of CDK-mediated phosphorylation also has therapeutic promise. For instance, recent identification of MEP50 as a novel CDK4 substrate and subsequent data demonstrating that MEP50 phosphorylation triggers increased PRMT5 enzymatic activity suggest that targeting the PRMT5 methyltransferase may be an effective therapeutic approach. Preliminary evidence revealed that treatment with commercially available PRMT inhibitors sensitizes transformed cyclin D1T286A lymphoma cells to apoptosis in vitro. Analysis of these “proof-of-concept” inhibitors in mice, as well as development of potent, specific small molecule inhibitors of PRMT5 is of great clinical relevance.
Phosphorylation-dependent nuclear export and subsequent degradation of cyclin D1 is essential to maintain cellular homeostasis. Disruption of this regulatory pathway has been extensively shown to promote neoplastic transformation; however, the precise mechanism of this event has been elusive. Significantly, recent work revealed that nuclear accumulation of active cyclin D1/CDK4 complexes generates genomic instability through PRMT5-dependent transcriptional repression, CDT1 stabilization, and DNA re-replication.  Furthermore, Fbx4 functions as a haploinsufficient tumor suppressor in vivo, and mutations targeting SCFFbx4 in human cancer implicate ligase function in cyclin D1 overexpression and subsequent nuclear accumulation of active cyclin D1/CDK4 kinase. Importantly, GSK3 functions as the master switch, turning on cyclin D1 destruction at the G1/S transition by regulating both ligase activation and cyclin D1 phosphorylation. Given the phenotypic outcome of accumulated nuclear cyclin D1/CDK4, further mechanistic investigation is required for development of novel therapeutic strategies to promote tumor cell death in cancers overexpressing cyclin D1.

Figure 5.1.

Figure 5.2.
Figure 5.3

Figure 5.4

CHAPTER VI:
Materials and Methods

Chapter 2
Cell Culture Conditions. NIH 3T3, MDA-MB 231, MCF7, U20S, ATRflox/- CET immortalized ear fibroblasts, and 293T cells were maintained in complete DMEM media containing glutamine, 1% penicillin/streptomycin, and 10% FBS. (MDA-MB 231 and MCF7 cells were also supplemented with 10g/mL insulin).  KYSE 520 and TE3/TE7 cells were maintained in RPMI media containing glutamine, antibiotics, and 10% FBS. NIH 3T3 cells stably expressing FLAG-D1, FLAG-D1T286A, FLAG-D1290-95A, αB crystallin shRNA, and Fbx4 shRNA have been previously described (Benzeno and Diehl, 2004; Diehl et al., 1997; Lin et al., 2006). For synchronization, NIH 3T3 cells and their derivatives were cultured in media containing 0.1% fetal bovine serum for 24 hours followed by addition of FBS for 15 hours for a synchronous S-phase population as determined by FACS. For transient plasmid expression, cells were transfected using Lipofectamine/Plus (Invitrogen).

Cell treatments and inhibitors. Acute ATR deletion was achieved by treating cells with 200nM 4-OH tamoxifen for 24h. Genotoxic stress was induced by γ-irradiation using a 137Cs Source (MS Nordion Gammacell 40 Exactor), 2mM HU, 10 or 20μg/mL bleomycin, or 5μM aphidicolin. For proteasome inhibition experiments, cells were treated with 10M MG132 (Calbiochem) and 2mM HU for 4 hours.  The following small-molecule inhibitors were used according to manufacturer’s recommendation: GSK3β inhibitors LiCl (Sigma) and SB216763 (Calbiochem), and p38 inhibitor SB203580 (Calbiochem); cells were pre-treated with inhibitors for 30 minutes prior to DNA damage induction.  For ATM inhibition, cells were pre-treated with KU-55933 (Sigma) for 30 minutes. For Chk1 or Chk2 inhibition, cells were pre-treated with SB218078 (Calbiochem) or Chk2 inhibitor II (Sigma) for 30 minutes, followed by DNA damage. The pJ3M-kdGSK3β construct was described previously (Diehl et al., 1998). Chromatin fractionation was described previously (Gladden and Diehl, 2003).

Cell Cycle Analysis. NIH3T3 cells stably expressing cyclin D1 or D1T286A were treated with CPT for 24 or 48 hours.  Cells were washed with PBS, fixed with ethanol, and stained with propidium iodide for 30 minutes, prior to FACS analysis.  Cell cycle profiles based on DNA content were established using FlowJo software, and the sub-G1 population of cells served as a readout for apoptotic cells following CPT treatment in three independent experiments.

Immunoblotting. Cells were lysed in Tween 20 buffer containing protease and phosphatase inhibitors (50mM HEPES (pH 8.0), 150mM NaCl, 2.5mM EGTA, 1mM EDTA, 0.1% Tween 20, 1mM DTT, 0.1mM PMSF, 20U/mL aprotinin, 10mM β-glycerophosphate, 5g/mL leupeptin, and 1mM NaF). M2 Agarose beads were utilized to precipitate Flag-tagged protein. Antibodies used for direct western include cyclin D1-72-13G, phospho-T286 rabbit polyclonal antibody, cyclin D2, cyclin E, CDK4, myc 9E10 (Santa Cruz), γ-H2AX, phospho-p38, Chk1, phospho-p53 (Ser 15), phospho-ATM (Cell Signaling), ATM, ATR (Serotec), Chk2, GSK3 (BD Transduction Labs), β-actin (AC15, Sigma), αB Crystallin (SPA 223, Stressgen), and Fbx4 (Rockland).

In vitro Kinase Assay. Equal concentration of endogenous GSK3 was precipitated from NIH3T3 cells that were untreated or subjected to IR using protein A-sepharose beads in GSK3 immunoprecipitation buffer: 50mM Tris-HCl pH 7.5, 1mM EDTA, 1mM EGTA, 10% TritonX-100, 0.27M sucrose, 1mM DTT and protease/phosphatase inhibitors. GSK3-bound beads were washed with kinase buffer: 20mM Tris-HCl, 40mM MgCl2, 2.5mM EGTA, 1mM DTT, protease/phosphatase inhibitors, and 20M ATP. Reactions were carried out in the presence of 32P-ATP for 20 minutes at 30C, resolved by SDS-PAGE and transferred to a nitrocellulose membrane. Immunoblot for GSK3 indicated equal precipitation of kinase.

Metaphase Spreads. Splenocytes were isolated from transgenic Eμ-D1T286A or non-transgenic control mouse spleens and cultured for 48 hours in RPMI medium supplemented with L-Glutamine, LPS, β-Mercaptoethanol, Concavalin A, and FBS.  Cultures were treated with 100M mitomycin-C (MMC) for 24h to induce damage. For HU experiments, cultures were treated for 2 hours with 2mM HU, washed, and re-suspended in fresh media. Cells were arrested in metaphase by treatment with colcemid (Gibco) for 2 hours. Cells were treated with hypotonic KCl solution and fixed with methanol-acetic acid.  Metaphase spreads were dropped, permitted to dry overnight, then stained with Giemsa (Sigma). The average number of chromatid breaks per cell was scored in 16 metaphase spreads following HU treatment or 25 total spreads for MMC treatment. NIH3T3 cells expressing vector control, Fbx4shRNA, or cyclin D1T286A were synchronized by serum starvation, treated with 2mM HU for 3 hours during S-phase, followed by 4 hours of 0.5M nocodazole treatment to arrest cells in metaphase. Metaphase spreads were prepared as described for splenocytes, and chromatid breaks in 20 metaphases for each treatment were scored in a blind fashion.

RDS Assays. Cells were synchronized by serum starvation for 24 hours in the presence of 20nCi [14C]thymidine (Amersham).  Cells were subsequently washed with PBS, and DMEM containing 10% serum was added to trigger S-phase entry.  S-phase populations were exposed to 10 Gy γIR as indicated and a recovery period at 37ºC for 30 minutes.  Cells were then pulsed with 20μCi [3H]thymidine (PerkinElmer) for 2 hours, and total DNA was isolated by high salt extraction.  Equal concentrations of purified DNA were subjected to scintillation counting, and DNA synthesis was expressed as the ratio of [3H]/[14C] incorporation, normalized to non-irradiated cells. 

shRNA/siRNA. Luciferase control and Chk2-specific siRNAs (Dharmacon smartpool) were kindly provided by Dr. Roger Greenberg, University of Pennsylvania. ATR-specific siRNA smartpool was purchased from Dharmacon. siRNAs were transfected into U20S cells using HiPerfect (Qiagen). Cells were irradiated 60h post siRNA transfection and harvested for western blotting. For Chk1 shRNA experiments, scramble control or Chk1 shRNA lentivirus were used to infect NIH3T3 cells.  72 hours post infection, cells were plated and synchronized by serum starvation.  Cells were released into complete media, and after 15 hours, cells were treated with HU to induce replication stress in S-phase.  
Statistical Analysis.  Error bars indicate represent ± standard deviation.  The Student’s t-test was used to compare groups in RDS assays, metaphase spread analysis, and sensitivity to CPT assays. Western blotting quantification was performed by densitometric analysis. Error bars indicate represent ± standard deviation of the densitometry of 3 independent experiments.

Chapter 3
Targeting Vector Construction and Generation of Fbx4-/- Mice. The engineered construct targeting murine Fbx4 exon 2 was developed by Vega Biolab (Philadelphia, PA). The targeting vector was electroporated into ES cells, and clones harboring a correctly targeted allele with a normal karyotype were injected in B6 blastocysts by the University of Pennsylvania core facility for generation of chimeric mice with germline transmission; these mice were intercrossed with EIIa-Cre transgenic mice to generate Fbx4 heterozygous progeny. Genotypes were verified by PCR using tail DNA (Extraction and PCR performed with Extract-N-Amp Tissue PCR Kit, Sigma-Aldrich). The following PCR primer sequences were used for genotyping: 
1loxP-F, 5’-GGCAGAGCTTGAGTTTGCAACATTTCAGGTG-3’; 
3loxP-R, 5’-TCCTGATCTTTGGAAATTCTTCCTCTGAGT-3’. 
Aged mice were examined daily for signs of distress or palpable tumor mass according to IACUC guidelines, and tumor-free survival was assessed by Kaplan-Meier analysis.

Southern Hybridization. 5’ and 3’ probes in the genomic region flanking Fbx4 exon 1-3  (site of homologous recombination, see Figure 3.1) were generated.  The 5’ probe generates a band of 15kb for normal Fbx4 locus, and a 10kb band for an allele that has undergone homologous recombination.

Histology/Immunohistochemistry. Tissues were fixed in 10% buffered formalin overnight, followed by dehydration in a series of ethanol and paraffin embedding/sectioning. Antibodies utilized for IHC staining were purified cyclin D1 (72-13G, Santa Cruz) and Ki-67 (Novocastra). 

MEFs and Cell Culture. Mouse embryos extracted at day 14 of gestation were maintained on a 3T9 passage protocol (9x105 cells passaged every three days on 60mm plates), cultured in DMEM containing 10% fetal bovine serum, 2mM glutamine, 0.1mM non-essential amino acids, 55M -mercaptoethanol, and 10g/mL gentamicin. For growth curves, 1x105 cells were plated on 35mm dishes in duplicate and counted every 24 hours by using a hemocytometer. Cell counts were also verified using the Countess Automated Cell Counter (Invitrogen). Primary MEFs were immortalized by lentiviral-mediated infection with p19Arf shRNA vector, generously provided by Dr. Eric Brown’s laboratory, University of Pennsylvania. Immortalized cells and NIH 3T3 fibroblasts were cultured in complete DMEM containing 10% fetal bovine serum, glutamine, and 1% penicillin/streptomycin. Cell synchronization was achieved by serum starvation for 36-48 hours, followed by serum stimulation to allow cell cycle entry. 

Western analysis. Whole tissues and cultured cells were lysed in Tween 20 buffer containing 50mM HEPES (pH 8.0), 150mM NaCl, 2.5mM EGTA, 1mM EDTA, 0.1% Tween 20, and protease/phosphatase inhibitors (1mM PMSF, 20U/mL aprotinin, 5mg/mL leupeptin, 1mM DTT, 0.4mM NaF, and 10mM -glycerophosphate). Whole tissues were homogenized in Tween 20 buffer with inhibitors, and tissue and cell culture lysates were sonicated prior to lysate clearing by centrifugation at 4C for 30 minutes. Lysate protein concentration was determined by BCA assay, and proteins were resolved by SDS-PAGE, transferred to nitrocellulose membranes, and subjected to immunoblotting. Antibodies utilized include Fbx4 rabbit polyclonal antibodies (YenZym, Rockland), cyclin D1 mouse monoclonal D1-72-13G, H4R3 (Abcam), H2AX (Cell Signaling), p53 mouse monoclonal pab421, p21 and cyclin A (Santa Cruz), and -Actin (Sigma).

Immunofluorescence. Primary MEFs were plated at optimal densities on glass coverslips. For cyclin D1 localization, passage 3 primary MEFs were grown to 70-80% density, then fixed in 1:1 methanol:acetone for 10 minutes at -20C. Coverslips were dried then rehydrated with PBS, blocked with 10% fetal calf serum in PBS for 30 minutes, then incubated with cyclin D1-72-13G primary antibody for 1 hour. Coverslips were washed 3x with PBS, followed by incubation with anti-mouse Alexa Fluor 546 secondary antibody for 30 minutes. Coverslips were washed 3x with PBS, dried by 70% and 100% ethanol washes and mounted with Vectashield mounting media containing DAPI (Vector Laboratories). For H2AX foci visualization, passage 3 cells maintained under atmospheric (21%) oxygen conditions or physiological (3%) oxygen tension were plated on glass coverslips at a density of 1x105 cells/well of a 6-well plate and cultured for 24 hours. Cells were fixed and blocked as described above, followed by incubation with H2AX antibodies for 2 hours. Coverslips were washed 3x with PBS, then incubated with anti-rabbit Alexa Fluor 488 secondary antibody for 30 minutes. Coverslips were washed and mounted as described above, and visualized with a Nikon Eclipse 80i fluorescent microscope.

Metaphase Spreads. Immortalized MEFs were synchronized by serum starvation, followed by serum stimulation to allow progression into S-phase. Cells in mid-S-phase were pulsed with HU for 2 hours, followed by HU washout and incubation in complete DMEM media. Cells were arrested in metaphase by treatment with colcemid for 2 hours, harvested, treated with hypotonic KCl solution and fixed with methanol-acetic acid. Metaphase spreads were dropped onto glass slides and permitted to dry, followed by Giemsa staining (Sigma). The average number of chromatid breaks per cell was scored in two independent biological replicate cell lines for each genotype.

In vivo Ubiquitylation. Immortalized MEFs were synchronized by serum starvation and stimulated for 15 hours to enter early S-phase. Cells were treated for 3 hours with 10M MG-132 (Calbiochem) for proteasome inhibition and harvested in Tween 20 buffer containing proteasome and phosphatase inhibitors, 1mM DTT, 10mM N-ethylmaleimide (NEM), and 20M MG-132, followed by immunoprecipitation with cyclin D1 72-13G antibody as described above. IP samples were resolved by SDS-PAGE on 10% poly-acrylamide gels and immunoblotted with phospho-cyclin D1 T286 antibodies and total cyclin D1 to visualize ubiquitylated cyclin D1 laddering.

In vitro Ubiquitylation. NIH 3T3 cells were synchronized by serum starvation, followed by serum stimulation to enter S-phase. Cells were either untreated or subjected to 10Gy IR, followed by 30 minutes of recovery. Cells were then harvested in Tween 20 buffer, and 1mg of total protein was immunoprecipitated with Fbx4 antibodies and protein A sepharose beads for 4 hours at 4C. Fbx4-containing endogenous SCF complexes were washed 4x with Tween-20 buffer, then 3x with kinase buffer (20mM Tris, 40mM MgCl2, 2.5mM EGTA). Beads containing SCFFbx4 complexes were then mixed with Sf9-produced purified cyclin D1 substrate, ATP, ubiquitin, E1, E2, MG-132, ubiquitin aldehyde, okadaic acid, energy regeneration buffer (20x: 10mM ATP, 20mM Hepes pH 7.4, 10mM MgOAc, 300mM creatine phosphate, 0.5mg/mL creatine phosphokinase) and kinase buffer to a final volume of 50L. Reactions were carried out at 37C for 30 minutes, followed by SDS-PAGE for visualization of ubiquitylated cyclin D1 by western blot.

In vitro Transformation Assays. Primary passage 3 MEFs were transduced with pBabe-Puro (empty vector) or pBabe-RasV12 retrovirus. 24 hours post infection, cells were trypsinized and re-plated at a density of 1.5x105 cells/35mm plate in duplicate. Cells were cultured in DMEM containing 5% serum, with media changed every 2 days for 21 days, followed by giemsa stain to visualize foci. For spontaneous transformation of p19shRNA-immortalized MEFs, early passage cells were plated at a density of 1.5x105/35mm plate and cultured as described above.

Statistical Analysis. Prism GraphPad Software was utilized for generating Kaplan-Meier mouse tumor free survival plots and statistical analysis of survival and tumor onset (log rank test). All other statistical analyses utilized a two-tailed student’s t-test, with p-values less than 0.05 indicating statistical significance. Error bars represent the mean  the standard deviation. 

Chapter 4
Cell Culture, Transfections and Plasmids. Cell culture conditions and transfections were performed as previously noted (Aggarwal et al., 2007). The human PRMT5 Myc-tagged vector was generated by PCR using primers designed for directional cloning into pCS2-MT plasmid (in frame with six Myc epitope tags at the N terminus of PRMT5), using PRMT5 cDNA (Open Biosystems) as the template. Human GST-tagged MEP50 vector was generated by PCR of human MEP50 with primers designed for directional cloning into pGEX-4T-1 plasmid (in frame with GST tag at the N terminus of MEP50), using pOTB7-MEP50 (Open Biosystems) as the template. Human MEP50 Myc-tagged vector was generated by PCR of human MEP50 with primers designed for directional cloning into pcDNA3 plasmid (in frame with 1X Myc tag at the N terminus of MEP50).  Site directed mutagenesis was performed using QuikChange Site-Directed Mutagenesis Kit (Stratagene). All clones were sequenced in their entirety.

Chromatin Immunoprecipitation Assay. Cells were cross-linked in 0.75% formaldehyde for 10 min and quenched with 0.18mM glycine for 5 min. Cells were harvested in cell lysis buffer (50mM HEPES, pH 7.5, 140mM NaCl, 1mM EDTA pH8.0, 1.0% Triton-X-100, 0.1% SDS, 0.1% deoxycholate, fresh protease inhibitors) and chromatin sheared to an average size of 350-600 bp by sonication (3x10min cycles of 30x30 sec each) at constant output (high setting, Bioruptor). Sonicated DNA was diluted 3-fold in dilution buffer (20mM Tris-HCl pH 8.0, 2mM EDTA pH 8.0, 1% Triton X-100, 150mM NaCl, protease inhibitors) and pre-cleared with protein A plus-agarose (Millipore). Lysates were rotated at 4°C overnight with 5μg of polyclonal antibodies specific for H4R3 (Abcam) or H3R8 anti-sera as described below, PRMT5 (Abcam), M2 monoclonal (Sigma), Brg1 (Santa Cruz) or normal rabbit IgG (Cell Signaling). After washing, complexes were eluted in 130μl of Elution buffer (1% SDS + 100mM NaHCO3), followed by addition of 100μg of proteinase K at 65°C overnight for cross-link reversal. The DNA was purified by phenol-chloroform extraction and precipitated DNA was analyzed by qPCR. CUL4A and CUL4B primers are described in Figure 4.3. 

Immunoprecipitation and Immunoblot Analysis. Cells were harvested in Tween 20 buffer (50mM HEPES (pH 8.0), 150mM NaCl, 2.5mM EGTA, 1mM EDTA, 0.1% Tween 20, protease, and phosphatase inhibitors (1mM PMSF, 20 U/ml aprotinin, 5g/ml leupeptin, 1mM DTT, 0.4M NaF, and 10M β-glycerophosphate), and protein concentration of samples was determined by BCA assay. Cyclin D1, PRMT5 and MEP50 were precipitated using M2 anti-flag agarose (to recognize flag-tagged proteins, Sigma-Aldrich), PRMT5 rabbit polyclonal antibody (Abcam) or c-myc (9E10) and MEP50 rabbit polyclonal antibody (Bethyl Lab. Inc), respectively. Proteins were resolved by SDS-PAGE, transferred to nitrocellulose membranes, and analyzed by immunoblotting. Antibodies used in these studies were as follows: Fbx4 rabbit polyclonal antibody (Rockland), cyclin D1 mouse monoclonal D1-72-13G, cyclin D1 mouse anti-human (AB3, Calbiochem); β-actin (Sigma-Aldrich); CUL4A rabbit polyclonal (Bethyl Lab. Inc); CUL4B rabbit polyclonal (ProteinTech Group Inc); pSP/TP mouse monoclonal (16B4, Biomol); CDK4 (C-22 or H-22), Cdt1, Brg1, RB, PRMT5 mouse monoclonal, PRMT2, mSin3a (Santa Cruz); H4R3 rabbit polyclonal-ChIP grade, H3K4 rabbit polyclonal, Histone H3 rabbit polyclonal, PRMT1 and PRMT7, Histone H4 rabbit polyclonal (Abcam). Anti-H3R8 (corresponding to symmetric di-methylated R8 of human histone H3) antibody was generated by immunizing rabbits with peptide containing modified histone H3 (YenZym Antibodies, LLC) followed by affinity purification. Antibody binding was visualized by chemiluminescence (Perkin Elmer). 

Immunohistochemistry. Human esophageal tumors harboring Fbx4 mutations and normal esophageal tissue were obtained with informed consent and with Institutional Review Board approval from the University of Pennsylvania (IRB #803902), Fox Chase Cancer Center (IRB #98-53), and University of Fukui, Japan (IRB #H190228-155). Tissue was fixed in 10% buffered formalin and dehydrated, paraffin embedded and sectioned. Tissue sections were immunostained using rabbit polyclonal H4R3 (Abcam) at 1:500 dilution as the primary antibody. 

In vitro kinase assay and in vitro methyltransferase assay. Purified GST-MEP50 wild type, GST-T5A, S176A, S264A, S306A mutants and GST-RB were used as substrates for in vitro kinase reactions (Matsushime et al., 1994). Cyclin D1/CDK4 kinase complexes were purified from Sf9 cells. For assessment of PRMT5 methyltransferase activity, PRMT5 complexes were collected from the indicated sources by immunoprecipitation. Beads were washed in Tween 20 buffer followed by methyltransferase buffer (15mM HEPES (pH7.9), 100mM KCl, 5mM MgCl2, 20% Glycerol, 1mM EDTA, 0.25mM DTT and 0.5mM PMSF). The methylation reaction included PRMT5 immune complexes on beads, 1g recombinant Histone H4 (NEB), and 2.75Ci S-adenosyl-L-(methyl-3H)methionine (Amersham Pharmacia) in a total volume of 25l for 1.5 hours at 30oC.  The reaction mixture was resolved on SDS-polyacrylamide gel, and modified histone H4 was detected by fluorography.  For coupled kinase assay/methyltransferase assays, cyclin/CDK complexes were purified from Sf9 cells and mixed with purified recombinant PRMT5/MEP50 (produced in Sf9 cells, BPS Bioscience) or PRMT5 precipitated from HeLa cells immobilized on beads. In vitro kinase assays were performed for 30 minutes at 30°C in kinase buffer. PRMT5-containing beads were washed into methyltransferase buffer, followed by in vitro methylation reaction for 2 hours at 30°C with 2μg H4 substrate and 2.75μCi 3H-SAM. Methyl-H4 was visualized by fluorography/scintillation counting for 3H incorporation. 

Foci Formation. Low passage NIH3T3 cells were transfected with control or murine PRMT5 siRNA (Dharmacon) using HiPerfect (Qiagen). 24 hours post siRNA delivery, cells were transfected with 2ug of the indicated plasmids (pBabe Ras V12 or pBabe cyclin D1T286A) or empty vector (pBabe puro) using Lipofectamine/Plus Reagent (Invitrogen). Transfected cells were counted after 48 hours, and 2x105 cells were plated in duplicate on 35mm plates in DMEM containing 5% FBS. Cells were re-fed with DMEM plus 5% FBS every 2 days for 14 days, followed by staining with Giemsa to visualize foci formation. Foci were counted for duplicate plates and error bars represent  standard deviation and * indicates p-value < 0.05, as determined by student’s t-test. For MEP50 overexpression assays, NIH3T3 cells were transfected with 2ug of wild type MEP50 or phosphorylation-deficient MEP50 mutant T5A or S264A, along with empty vector or Ras V12 and cyclin D1T286A constructs as indicated. Cells were seeded and foci formation was analyzed as described above. 

Real time quantitative PCR analysis of gene expression. RNA isolation was performed using standard protocols. DNA fragments were generated by reverse transcriptase PCR (RT-PCR; SuperscriptTM, Invitrogen). Mixed primer/probe sets for human CUL4A/B and 18S rRNA were used to measure the levels of these transcripts using the Applied Biosystems 7900HT sequence detection system according to the manufacturer's instructions.  Primers used for detection of human CUL4A and CUL4B were described previously (Aggarwal et al., 2007). 

siRNA and Methyltransferase Inhibitor experiments. siGENOME SMARTpool targeting PRMT5 and MEP50 or control siRNA were purchased from Dharmacon. HeLa cells were treated with indicated siRNA or methyltransferase inhibitors, MTA (Sigma) or AMI-1 (Calbiochem), for 24 hours followed by transfection and synchronization of cells in G1 and S-phases. For targeting 3’ UTR region of human MEP50, siRNA (5’ CUCCUUACCAUUAAACUGA 3’) was designed and purchased from Sigma.

CHAPTER VII:
REFERENCES

Agami, R., and Bernards, R. (2000). Distinct initiation and maintenance mechanisms cooperate to induce G1 cell cycle arrest in response to DNA damage. Cell 102, 55-66.
Aggarwal, P., Lessie, M.D., Lin, D.I., Pontano, L., Gladden, A.B., Nuskey, B., Goradia, A., Wasik, M.A., Klein-Szanto, A.J., Rustgi, A.K., et al. (2007). Nuclear accumulation of cyclin D1 during S phase inhibits Cul4-dependent Cdt1 proteolysis and triggers p53-dependent DNA rereplication. Genes Dev 21, 2908-2922.
Aggarwal, P., Vaites, L.P., Kim, J.K., Mellert, H., Gurung, B., Nakagawa, H., Herlyn, M., Hua, X., Rustgi, A.K., McMahon, S.B., et al. (2010). Nuclear cyclin D1/CDK4 kinase regulates CUL4 expression and triggers neoplastic growth via activation of the PRMT5 methyltransferase. Cancer Cell 18, 329-340.
Aleem, E., Kiyokawa, H., and Kaldis, P. (2005). Cdc2-cyclin E complexes regulate the G1/S phase transition. Nat Cell Biol 7, 831-836.
Alt, J.R., Cleveland, J.L., Hannink, M., and Diehl, J.A. (2000). Phosphorylation-dependent regulation of cyclin D1 nuclear export and cyclin D1-dependent cellular transformation. Genes Dev 14, 3102-3114.
Amador, V., Ge, S., Santamaria, P.G., Guardavaccaro, D., and Pagano, M. (2007). APC/C(Cdc20) controls the ubiquitin-mediated degradation of p21 in prometaphase. Mol Cell 27, 462-473.
Baldin, V., Cans, C., Knibiehler, M., and Ducommun, B. (1997). Phosphorylation of human CDC25B phosphatase by CDK1-cyclin A triggers its proteasome-dependent degradation. J Biol Chem 272, 32731-32734.
Bani-Hani, K., Martin, I.G., Hardie, L.J., Mapstone, N., Briggs, J.A., Forman, D., and Wild, C.P. (2000). Prospective study of cyclin D1 overexpression in Barrett's esophagus: association with increased risk of adenocarcinoma. J Natl Cancer Inst 92, 1316-1321.
Barbash, O., Lee, E.K., and Diehl, J.A. (2011). Phosphorylation-dependent regulation of SCF(Fbx4) dimerization and activity involves a novel component, 14-3-3varepsilon. Oncogene 30, 1995-2002.
Barbash, O., Zamfirova, P., Lin, D.I., Chen, X., Yang, K., Nakagawa, H., Lu, F., Rustgi, A.K., and Diehl, J.A. (2008). Mutations in Fbx4 Inhibit Dimerization of the SCF(Fbx4) Ligase and Contribute to Cyclin D1 Overexpression in Human Cancer. Cancer Cell 14, 68-78.
Barnes, D.M., and Gillett, C.E. (1998). Cyclin D1 in breast cancer. Breast Cancer Res Treat 52, 1-15.
Bartek, J., Lukas, C., and Lukas, J. (2004). Checking on DNA damage in S phase. Nat Rev Mol Cell Biol 5, 792-804.
Bartkova J, H.Z., Koed K, Kramer A, Tort F, Zieger K, Guldberg P, Sehested M, Nesland JM, Lukas C, Orntoft T, Lukas J, Bartek J (2005). DNA damage response as a candidate anti-cancer barrier in early human tumorigenesis Nature 434, 864-870.
Bartkova, J., Lukas, J., Muller, H., Lutzhoft, D., Strauss, M., and Bartek, J. (1994a). Cyclin D1 protein expression and function in human breast cancer. Int J Cancer 57, 353-361.
Bartkova, J., Lukas, J., Muller, H., Strauss, M., Gusterson, B., and Bartek, J. (1995). Abnormal patterns of D-type cyclin expression and G1 regulation in human head and neck cancer. Cancer Res 55, 949-956.
Bartkova, J., Lukas, J., Strauss, M., and Bartek, J. (1994b). The PRAD-1/cyclin D1 oncogene product accumulates aberrantly in a subset of colorectal carcinomas. Int J Cancer 58, 568-573.
Bell, S.P., and Dutta, A. (2002). DNA replication in eukaryotic cells. Annu Rev Biochem 71, 333-374.
Benzeno, S., and Diehl, J.A. (2004). C-terminal sequences direct cyclin D1-CRM1 binding. J Biol Chem 279, 56061-56066.
Benzeno, S., Lu, F., Guo, M., Barbash, O., Zhang, F., Herman, J.G., Klein, P.S., Rustgi, A., and Diehl, J.A. (2006). Identification of mutations that disrupt phosphorylation-dependent nuclear export of cyclin D1. Oncogene 25, 6291-6303.
Berthet, C., Klarmann, K.D., Hilton, M.B., Suh, H.C., Keller, J.R., Kiyokawa, H., and Kaldis, P. (2006). Combined loss of Cdk2 and Cdk4 results in embryonic lethality and Rb hypophosphorylation. Dev Cell 10, 563-573.
Bianchi, V., Pontis, E., and Reichard, P. (1986). Changes of deoxyribonucleoside triphosphate pools induced by hydroxyurea and their relation to DNA synthesis. J Biol Chem 261, 16037-16042.
Bienvenu, F., Jirawatnotai, S., Elias, J.E., Meyer, C.A., Mizeracka, K., Marson, A., Frampton, G.M., Cole, M.F., Odom, D.T., Odajima, J., et al. (2010). Transcriptional role of cyclin D1 in development revealed by a genetic-proteomic screen. Nature 463, 374-378.
Bornstein, G., Bloom, J., Sitry-Shevah, D., Nakayama, K., Pagano, M., and Hershko, A. (2003). Role of the SCFSkp2 ubiquitin ligase in the degradation of p21Cip1 in S phase. J Biol Chem 278, 25752-25757.
Brandeis, M., Rosewell, I., Carrington, M., Crompton, T., Jacobs, M.A., Kirk, J., Gannon, J., and Hunt, T. (1998). Cyclin B2-null mice develop normally and are fertile whereas cyclin B1-null mice die in utero. Proc Natl Acad Sci U S A 95, 4344-4349.
Brown, E.J., and Baltimore, D. (2003). Essential and dispensable roles of ATR in cell cycle arrest and genome maintenance. Genes Dev 17, 615-628.
Busino, L., Donzelli, M., Chiesa, M., Guardavaccaro, D., Ganoth, D., Dorrello, N.V., Hershko, A., Pagano, M., and Draetta, G.F. (2003). Degradation of Cdc25A by beta-TrCP during S phase and in response to DNA damage. Nature 426, 87-91.
Calbo, J., Parreno, M., Sotillo, E., Yong, T., Mazo, A., Garriga, J., and Grana, X. (2002). G1 cyclin/cyclin-dependent kinase-coordinated phosphorylation of endogenous pocket proteins differentially regulates their interactions with E2F4 and E2F1 and gene expression. J Biol Chem 277, 50263-50274.
Carrano, A.C., Eytan, E., Hershko, A., and Pagano, M. (1999). SKP2 is required for ubiquitin-mediated degradation of the CDK inhibitor p27. Nat Cell Biol 1, 193-199.
Casanovas, O., Miro, F., Estanyol, J.M., Itarte, E., Agell, N., and Bachs, O. (2000). Osmotic stress regulates the stability of cyclin D1 in a p38SAPK2-dependent manner. J Biol Chem 275, 35091-35097.
Cenciarelli, C., Chiaur, D.S., Guardavaccaro, D., Parks, W., Vidal, M., and Pagano, M. (1999). Identification of a family of human F-box proteins. Curr Biol 9, 1177-1179.
Cesaro, E., De Cegli, R., Medugno, L., Florio, F., Grosso, M., Lupo, A., Izzo, P., and Costanzo, P. (2009). The Kruppel-like zinc finger protein ZNF224 recruits the arginine methyltransferase PRMT5 on the transcriptional repressor complex of the aldolase A gene. J Biol Chem 284, 32321-32330.
Chen, Y., Knosel, T., Kristiansen, G., Pietas, A., Garber, M.E., Matsuhashi, S., Ozaki, I., and Petersen, I. (2003). Loss of PDCD4 expression in human lung cancer correlates with tumour progression and prognosis. J Pathol 200, 640-646.
Cheng, D., Yadav, N., King, R.W., Swanson, M.S., Weinstein, E.J., and Bedford, M.T. (2004). Small molecule regulators of protein arginine methyltransferases. J Biol Chem 279, 23892-23899.
Ciechanover, A. (1994). The ubiquitin-proteasome proteolytic pathway. Cell 79, 13-21.
Ciemerych, M.A., Kenney, A.M., Sicinska, E., Kalaszczynska, I., Bronson, R.T., Rowitch, D.H., Gardner, H., and Sicinski, P. (2002). Development of mice expressing a single D-type cyclin. Genes Dev 16, 3277-3289.
Clurman, B.E., Sheaff, R.J., Thress, K., Groudine, M., and Roberts, J.M. (1996). Turnover of cyclin E by the ubiquitin-proteasome pathway is regulated by cdk2 binding and cyclin phosphorylation. Genes Dev 10, 1979-1990.
Connell-Crowley, L., Harper, J.W., and Goodrich, D.W. (1997). Cyclin D1/Cdk4 regulates retinoblastoma protein-mediated cell cycle arrest by site-specific phosphorylation. Mol Biol Cell 8, 287-301.
Costanzo, V., Robertson, K., Ying, C.Y., Kim, E., Avvedimento, E., Gottesman, M., Grieco, D., and Gautier, J. (2000). Reconstitution of an ATM-dependent checkpoint that inhibits chromosomal DNA replication following DNA damage. Mol Cell 6, 649-659.
Cross, D.A., Alessi, D.R., Cohen, P., Andjelkovich, M., and Hemmings, B.A. (1995). Inhibition of glycogen synthase kinase-3 by insulin mediated by protein kinase B. Nature 378, 785-789.
De Boer, L., Oakes, V., Beamish, H., Giles, N., Stevens, F., Somodevilla-Torres, M., Desouza, C., and Gabrielli, B. (2008). Cyclin A/cdk2 coordinates centrosomal and nuclear mitotic events. Oncogene 27, 4261-4268.
Deshaies, R.J. (1999). SCF and Cullin/Ring H2-based ubiquitin ligases. Annu Rev Cell Dev Biol 15, 435-467.
Diehl, J.A. (2002). Cycling to Cancer with Cyclin Dl. Cancer Biol Ther 1, 226-231.
Diehl, J.A., Cheng, M., Roussel, M.F., and Sherr, C.J. (1998). Glycogen synthase kinase-3beta regulates cyclin D1 proteolysis and subcellular localization. Genes Dev 12, 3499-3511.
Diehl, J.A., Zindy, F., and Sherr, C.J. (1997). Inhibition of cyclin D1 phosphorylation on threonine-286 prevents its rapid degradation via the ubiquitin-proteasome pathway. Genes Dev 11, 957-972.
Dohner, H., Stilgenbauer, S., Dohner, K., Bentz, M., and Lichter, P. (1999). Chromosome aberrations in B-cell chronic lymphocytic leukemia: reassessment based on molecular cytogenetic analysis. J Mol Med 77, 266-281.
Dorrello, N.V., Peschiaroli, A., Guardavaccaro, D., Colburn, N.H., Sherman, N.E., and Pagano, M. (2006). S6K1- and betaTRCP-mediated degradation of PDCD4 promotes protein translation and cell growth. Science 314, 467-471.
Draetta, G., Luca, F., Westendorf, J., Brizuela, L., Ruderman, J., and Beach, D. (1989). Cdc2 protein kinase is complexed with both cyclin A and B: evidence for proteolytic inactivation of MPF. Cell 56, 829-838.
Evans, T., Rosenthal, E.T., Youngblom, J., Distel, D., and Hunt, T. (1983). Cyclin: a protein specified by maternal mRNA in sea urchin eggs that is destroyed at each cleavage division. Cell 33, 389-396.
Falck, J., Petrini, J.H., Williams, B.R., Lukas, J., and Bartek, J. (2002). The DNA damage-dependent intra-S phase checkpoint is regulated by parallel pathways. Nat Genet 30, 290-294.
Fantl, V., Stamp, G., Andrews, A., Rosewell, I., and Dickson, C. (1995). Mice lacking cyclin D1 are small and show defects in eye and mammary gland development. Genes Dev 9, 2364-2372.
Fero, M.L., Randel, E., Gurley, K.E., Roberts, J.M., and Kemp, C.J. (1998). The murine gene p27Kip1 is haplo-insufficient for tumour suppression. Nature 396, 177-180.
Fero, M.L., Rivkin, M., Tasch, M., Porter, P., Carow, C.E., Firpo, E., Polyak, K., Tsai, L.H., Broudy, V., Perlmutter, R.M., et al. (1996). A syndrome of multiorgan hyperplasia with features of gigantism, tumorigenesis, and female sterility in p27(Kip1)-deficient mice. Cell 85, 733-744.
Friesen, W.J., Paushkin, S., Wyce, A., Massenet, S., Pesiridis, G.S., Van Duyne, G., Rappsilber, J., Mann, M., and Dreyfuss, G. (2001). The methylosome, a 20S complex containing JBP1 and pICln, produces dimethylarginine-modified Sm proteins. Mol Cell Biol 21, 8289-8300.
Fry, D.W., Harvey, P.J., Keller, P.R., Elliott, W.L., Meade, M., Trachet, E., Albassam, M., Zheng, X., Leopold, W.R., Pryer, N.K., et al. (2004). Specific inhibition of cyclin-dependent kinase 4/6 by PD 0332991 and associated antitumor activity in human tumor xenografts. Mol Cancer Ther 3, 1427-1438.
Gavet, O., and Pines, J. Progressive activation of CyclinB1-Cdk1 coordinates entry to mitosis. Dev Cell 18, 533-543.
Geng, Y., Yu, Q., Sicinska, E., Das, M., Bronson, R.T., and Sicinski, P. (2001). Deletion of the p27Kip1 gene restores normal development in cyclin D1-deficient mice. Proc Natl Acad Sci U S A 98, 194-199.
Geng, Y., Yu, Q., Sicinska, E., Das, M., Schneider, J., Bhattacharya, S., Rideout, W., Bronson, R.T., Gardner, H., and Sicinski, P. (2003). Cyclin E Ablation in the Mouse. Cell 114, 431-443.
Ghosh, M., Kemp, M., Liu, G., Ritzi, M., Schepers, A., and Leffak, M. (2006). Differential binding of replication proteins across the human c-myc replicator. Mol Cell Biol 26, 5270-5283.
Ghosh, S.K., Paik, W.K., and Kim, S. (1988). Purification and molecular identification of two protein methylases I from calf brain. Myelin basic protein- and histone-specific enzyme. J Biol Chem 263, 19024-19033.
Gillett, C., Fantl, V., Smith, R., Fisher, C., Bartek, J., Dickson, C., Barnes, D., and Peters, G. (1994). Amplification and overexpression of cyclin D1 in breast cancer detected by immunohistochemical staining. Cancer Res 54, 1812-1817.
Girard-Reydet, C., Gregoire, D., Vassetzky, Y., and Mechali, M. (2004). DNA replication initiates at domains overlapping with nuclear matrix attachment regions in the xenopus and mouse c-myc promoter. Gene 332, 129-138.
Gladden, A.B., and Diehl, J.A. (2003). The cyclin D1-dependent kinase associates with the pre-replication complex and modulates RB.MCM7 binding. J Biol Chem 278, 9754-9760.
Gladden, A.B., Woolery, R., Aggarwal, P., Wasik, M.A., and Diehl, J.A. (2006). Expression of constitutively nuclear cyclin D1 in murine lymphocytes induces B-cell lymphoma. Oncogene 25, 998-1007.
Glotzer, M., Murray, A.W., and Kirschner, M.W. (1991). Cyclin is degraded by the ubiquitin pathway. Nature 349, 132-138.
Gorgoulis VG, V.L., Karakaidos P, Zacharatos P, Kotsinas A, Liloglou T, Venere M, Ditullio RA Jr, Kastrinakas NG, Levy B, Kletsas D, Yoneta A, Herlyn M, Kittas C, Halazonetis TD (2005). Activation of the DNA damage checkpoint and genomic instability in human precancerous lesions Nature 434, 907-913.
Gregory, M.A., and Hann, S.R. (2000). c-Myc proteolysis by the ubiquitin-proteasome pathway: stabilization of c-Myc in Burkitt's lymphoma cells. Mol Cell Biol 20, 2423-2435.
Guardavaccaro, D., Kudo, Y., Boulaire, J., Barchi, M., Busino, L., Donzelli, M., Margottin-Goguet, F., Jackson, P.K., Yamasaki, L., and Pagano, M. (2003). Control of meiotic and mitotic progression by the F box protein beta-Trcp1 in vivo. Dev Cell 4, 799-812.
Hao, B., Oehlmann, S., Sowa, M.E., Harper, J.W., and Pavletich, N.P. (2007). Structure of a Fbw7-Skp1-cyclin E complex: multisite-phosphorylated substrate recognition by SCF ubiquitin ligases. Mol Cell 26, 131-143.
Harbour, J.W., Luo, R.X., Dei Santi, A., Postigo, A.A., and Dean, D.C. (1999). Cdk phosphorylation triggers sequential intramolecular interactions that progressively block Rb functions as cells move through G1. Cell 98, 859-869.
Harper, J.W., Burton, J.L., and Solomon, M.J. (2002). The anaphase-promoting complex: it's not just for mitosis any more. Genes Dev 16, 2179-2206.
Hatakeyama, M., Brill, J.A., Fink, G.R., and Weinberg, R.A. (1994). Collaboration of G1 cyclins in the functional inactivation of the retinoblastoma protein. Genes Dev 8, 1759-1771.
Havens, C.G., and Walter, J.C. (2009). Docking of a specialized PIP Box onto chromatin-bound PCNA creates a degron for the ubiquitin ligase CRL4Cdt2. Mol Cell 35, 93-104.
Herman, J.G., Merlo, A., Mao, L., Lapidus, R.G., Issa, J.P., Davidson, N.E., Sidransky, D., and Baylin, S.B. (1995). Inactivation of the CDKN2/p16/MTS1 gene is frequently associated with aberrant DNA methylation in all common human cancers. Cancer Res 55, 4525-4530.
Hershko, A., and Ciechanover, A. (1998). The ubiquitin system. Annu Rev Biochem 67, 425-479.
Hibberts, N.A., Simpson, D.J., Bicknell, J.E., Broome, J.C., Hoban, P.R., Clayton, R.N., and Farrell, W.E. (1999). Analysis of cyclin D1 (CCND1) allelic imbalance and overexpression in sporadic human pituitary tumors. Clin Cancer Res 5, 2133-2139.
Higa, L.A., Banks, D., Wu, M., Kobayashi, R., Sun, H., and Zhang, H. (2006). L2DTL/CDT2 interacts with the CUL4/DDB1 complex and PCNA and regulates CDT1 proteolysis in response to DNA damage. Cell Cycle 5, 1675-1680.
Higa, L.A., Mihaylov, I.S., Banks, D.P., Zheng, J., and Zhang, H. (2003). Radiation-mediated proteolysis of CDT1 by CUL4-ROC1 and CSN complexes constitutes a new checkpoint. Nat Cell Biol 5, 1008-1015.
Hirai, H., and Sherr, C.J. (1996). Interaction of D-type cyclins with a novel myb-like transcription factor, DMP1. Mol Cell Biol 16, 6457-6467.
Hitomi, M., Yang, K., Stacey, A.W., and Stacey, D.W. (2008). Phosphorylation of cyclin D1 regulated by ATM or ATR controls cell cycle progression. Mol Cell Biol 28, 5478-5493.
Hosokawa, Y., and Arnold, A. (1998). Mechanism of cyclin D1 (CCND1, PRAD1) overexpression in human cancer cells: analysis of allele-specific expression. Genes Chromosomes Cancer 22, 66-71.
Hosokawa, Y., Joh, T., Maeda, Y., Arnold, A., and Seto, M. (1999). Cyclin D1/PRAD1/BCL-1 alternative transcript [B] protein product in B-lymphoid malignancies with t(11;14)(q13;q32) translocation. Int J Cancer 81, 616-619.
Hou, Z., Peng, H., Ayyanathan, K., Yan, K.P., Langer, E.M., Longmore, G.D., and Rauscher, F.J., 3rd (2008). The LIM protein AJUBA recruits protein arginine methyltransferase 5 to mediate SNAIL-dependent transcriptional repression. Mol Cell Biol 28, 3198-3207.
Hu, J., McCall, C.M., Ohta, T., and Xiong, Y. (2004). Targeted ubiquitination of CDT1 by the DDB1-CUL4A-ROC1 ligase in response to DNA damage. Nat Cell Biol 6, 1003-1009.
Huang, H., Regan, K.M., Wang, F., Wang, D., Smith, D.I., van Deursen, J.M., and Tindall, D.J. (2005). Skp2 inhibits FOXO1 in tumor suppression through ubiquitin-mediated degradation. Proc Natl Acad Sci U S A 102, 1649-1654.
Huard, J.M., Forster, C.C., Carter, M.L., Sicinski, P., and Ross, M.E. (1999). Cerebellar histogenesis is disturbed in mice lacking cyclin D2. Development 126, 1927-1935.
Hwang, H.C., and Clurman, B.E. (2005). Cyclin E in normal and neoplastic cell cycles. Oncogene 24, 2776-2786.
Ikeguchi, M., Sakatani, T., Ueta, T., and Kaibara, N. (2001). Cyclin D1 expression and retinoblastoma gene protein (pRB) expression in esophageal squamous cell carcinoma. J Cancer Res Clin Oncol 127, 531-536.
Inuzuka, H., Shaik, S., Onoyama, I., Gao, D., Tseng, A., Maser, R.S., Zhai, B., Wan, L., Gutierrez, A., Lau, A.W., et al. (2011). SCF(FBW7) regulates cellular apoptosis by targeting MCL1 for ubiquitylation and destruction. Nature 471, 104-109.
Iwasaki, H., and Yada, T. (2007). Protein arginine methylation regulates insulin signaling in L6 skeletal muscle cells. Biochem Biophys Res Commun 364, 1015-1021.
Jansson, M., Durant, S.T., Cho, E.C., Sheahan, S., Edelmann, M., Kessler, B., and La Thangue, N.B. (2008). Arginine methylation regulates the p53 response. Nat Cell Biol 10, 1431-1439.
Jazayeri, A., Falck, J., Lukas, C., Bartek, J., Smith, G.C., Lukas, J., and Jackson, S.P. (2006). ATM- and cell cycle-dependent regulation of ATR in response to DNA double-strand breaks. Nat Cell Biol 8, 37-45.
Jeffrey, P.D., Russo, A.A., Polyak, K., Gibbs, E., Hurwitz, J., Massague, J., and Pavletich, N.P. (1995). Mechanism of CDK activation revealed by the structure of a cyclinA-CDK2 complex. Nature 376, 313-320.
Jin, J., Shirogane, T., Xu, L., Nalepa, G., Qin, J., Elledge, S.J., and Harper, J.W. (2003). SCFbeta-TRCP links Chk1 signaling to degradation of the Cdc25A protein phosphatase. Genes Dev 17, 3062-3074.
Jin, M., Inoue, S., Umemura, T., Moriya, J., Arakawa, M., Nagashima, K., and Kato, H. (2001). Cyclin D1, p16 and retinoblastoma gene product expression as a predictor for prognosis in non-small cell lung cancer at stages I and II. Lung Cancer 34, 207-218.
Kalaszczynska, I., Geng, Y., Iino, T., Mizuno, S., Choi, Y., Kondratiuk, I., Silver, D.P., Wolgemuth, D.J., Akashi, K., and Sicinski, P. (2009). Cyclin A is redundant in fibroblasts but essential in hematopoietic and embryonic stem cells. Cell 138, 352-365.
Karin, M., and Greten, F.R. (2005). NF-kappaB: linking inflammation and immunity to cancer development and progression. Nat Rev Immunol 5, 749-759.
Kasper, J.S., Kuwabara, H., Arai, T., Ali, S.H., and DeCaprio, J.A. (2005). Simian virus 40 large T antigen's association with the CUL7 SCF complex contributes to cellular transformation. J Virol 79, 11685-11692.
Kastan, M.B., and Bartek, J. (2004). Cell-cycle checkpoints and cancer. Nature 432, 316-323.
Kato, J., Matsushime, H., Hiebert, S.W., Ewen, M.E., and Sherr, C.J. (1993). Direct binding of cyclin D to the retinoblastoma gene product (pRb) and pRb phosphorylation by the cyclin D-dependent kinase CDK4. Genes Dev 7, 331-342.
Khanna, K.K., and Jackson, S.P. (2001). DNA double-strand breaks: signaling, repair and the cancer connection. Nat Genet 27, 247-254.
Kida, A., Kakihana, K., Kotani, S., Kurosu, T., and Miura, O. (2007). Glycogen synthase kinase-3beta and p38 phosphorylate cyclin D2 on Thr280 to trigger its ubiquitin/proteasome-dependent degradation in hematopoietic cells. Oncogene 26, 6630-6640.
Knudsen, K.E., Cavenee, W.K., and Arden, K.C. (1999). D-type cyclins complex with the androgen receptor and inhibit its transcriptional transactivation ability. Cancer Res 59, 2297-2301.
Koepp, D.M., Schaefer, L.K., Ye, X., Keyomarsi, K., Chu, C., Harper, J.W., and Elledge, S.J. (2001). Phosphorylation-dependent ubiquitination of cyclin E by the SCFFbw7 ubiquitin ligase. Science 294, 173-177.
Kossatz, U., Dietrich, N., Zender, L., Buer, J., Manns, M.P., and Malek, N.P. (2004). Skp2-dependent degradation of p27kip1 is essential for cell cycle progression. Genes Dev 18, 2602-2607.
Kozar, K., Ciemerych, M.A., Rebel, V.I., Shigematsu, H., Zagozdzon, A., Sicinska, E., Geng, Y., Yu, Q., Bhattacharya, S., Bronson, R.T., et al. (2004). Mouse development and cell proliferation in the absence of D-cyclins. Cell 118, 477-491.
Kraft, C., Herzog, F., Gieffers, C., Mechtler, K., Hagting, A., Pines, J., and Peters, J.M. (2003). Mitotic regulation of the human anaphase-promoting complex by phosphorylation. EMBO J 22, 6598-6609.
Krause, C.D., Yang, Z.H., Kim, Y.S., Lee, J.H., Cook, J.R., and Pestka, S. (2007). Protein arginine methyltransferases: evolution and assessment of their pharmacological and therapeutic potential. Pharmacol Ther 113, 50-87.
Kudo, Y., Guardavaccaro, D., Santamaria, P.G., Koyama-Nasu, R., Latres, E., Bronson, R., Yamasaki, L., and Pagano, M. (2004). Role of F-box protein betaTrcp1 in mammary gland development and tumorigenesis. Mol Cell Biol 24, 8184-8194.
Kushner, J.A., Ciemerych, M.A., Sicinska, E., Wartschow, L.M., Teta, M., Long, S.Y., Sicinski, P., and White, M.F. (2005). Cyclins D2 and D1 are essential for postnatal pancreatic beta-cell growth. Mol Cell Biol 25, 3752-3762.
Kwak, Y.T., Li, R., Becerra, C.R., Tripathy, D., Frenkel, E.P., and Verma, U.N. (2005). IkappaB kinase alpha regulates subcellular distribution and turnover of cyclin D1 by phosphorylation. J Biol Chem 280, 33945-33952.
Lacroix, M., Messaoudi, S.E., Rodier, G., Le Cam, A., Sardet, C., and Fabbrizio, E. (2008). The histone-binding protein COPR5 is required for nuclear functions of the protein arginine methyltransferase PRMT5. EMBO Rep 9, 452-458.
Lamb, J., Ladha, M.H., McMahon, C., Sutherland, R.L., and Ewen, M.E. (2000). Regulation of the functional interaction between cyclin D1 and the estrogen receptor. Mol Cell Biol 20, 8667-8675.
Lamb, J., Ramaswamy, S., Ford, H.L., Contreras, B., Martinez, R.V., Kittrell, F.S., Zahnow, C.A., Patterson, N., Golub, T.R., and Ewen, M.E. (2003). A mechanism of cyclin D1 action encoded in the patterns of gene expression in human cancer. Cell 114, 323-334.
Lan, Z., Sever-Chroneos, Z., Strobeck, M.W., Park, C.H., Baskaran, R., Edelmann, W., Leone, G., and Knudsen, E.S. (2002). DNA damage invokes mismatch repair-dependent cyclin D1 attenuation and retinoblastoma signaling pathways to inhibit CDK2. J Biol Chem 277, 8372-8381.
Landis, M.W., Pawlyk, B.S., Li, T., Sicinski, P., and Hinds, P.W. (2006). Cyclin D1-dependent kinase activity in murine development and mammary tumorigenesis. Cancer Cell 9, 13-22.
Latres, E., Malumbres, M., Sotillo, R., Martin, J., Ortega, S., Martin-Caballero, J., Flores, J.M., Cordon-Cardo, C., and Barbacid, M. (2000). Limited overlapping roles of P15(INK4b) and P18(INK4c) cell cycle inhibitors in proliferation and tumorigenesis. EMBO J 19, 3496-3506.
Lee, J.Y., Yu, S.J., Park, Y.G., Kim, J., and Sohn, J. (2007). Glycogen synthase kinase 3beta phosphorylates p21WAF1/CIP1 for proteasomal degradation after UV irradiation. Mol Cell Biol 27, 3187-3198.
Lee, T.H., Perrem, K., Harper, J.W., Lu, K.P., and Zhou, X.Z. (2006). The F-box protein FBX4 targets PIN2/TRF1 for ubiquitin-mediated degradation and regulates telomere maintenance. J Biol Chem 281, 759-768.
Leng, X., Noble, M., Adams, P.D., Qin, J., and Harper, J.W. (2002). Reversal of growth suppression by p107 via direct phosphorylation by cyclin D1/cyclin-dependent kinase 4. Mol Cell Biol 22, 2242-2254.
Li, Y., and Hao, B. Structural basis of dimerization-dependent ubiquitination by the SCF(Fbx4) ubiquitin ligase. J Biol Chem 285, 13896-13906.
Lin, D.I., Barbash, O., Kumar, K.G., Weber, J.D., Harper, J.W., Klein-Szanto, A.J., Rustgi, A., Fuchs, S.Y., and Diehl, J.A. (2006). Phosphorylation-dependent ubiquitination of cyclin D1 by the SCF(FBX4-alphaB crystallin) complex. Mol Cell 24, 355-366.
Lin, D.I., Lessie, M.D., Gladden, A.B., Bassing, C.H., Wagner, K.U., and Diehl, J.A. (2008). Disruption of cyclin D1 nuclear export and proteolysis accelerates mammary carcinogenesis. Oncogene 27, 1231-1242.
Liu, F., and Matsuura, I. (2005). Inhibition of Smad antiproliferative function by CDK phosphorylation. Cell Cycle 4, 63-66.
Liu, J., Suresh Kumar, K.G., Yu, D., Molton, S.A., McMahon, M., Herlyn, M., Thomas-Tikhonenko, A., and Fuchs, S.Y. (2007). Oncogenic BRAF regulates beta-Trcp expression and NF-kappaB activity in human melanoma cells. Oncogene 26, 1954-1958.
Liu, J.S., Kuo, S.R., and Melendy, T. (2003). Comparison of checkpoint responses triggered by DNA polymerase inhibition versus DNA damaging agents. Mutat Res 532, 215-226.
Liyanage, M., Weaver, Z., Barlow, C., Coleman, A., Pankratz, D.G., Anderson, S., Wynshaw-Boris, A., and Ried, T. (2000). Abnormal rearrangement within the alpha/delta T-cell receptor locus in lymphomas from Atm-deficient mice. Blood 96, 1940-1946.
Lu, F., Gladden, A.B., and Diehl, J.A. (2003). An alternatively spliced cyclin D1 isoform, cyclin D1b, is a nuclear oncogene. Cancer Res 63, 7056-7061.
Lundin, C., Erixon, K., Arnaudeau, C., Schultz, N., Jenssen, D., Meuth, M., and Helleday, T. (2002). Different roles for nonhomologous end joining and homologous recombination following replication arrest in mammalian cells. Mol Cell Biol 22, 5869-5878.
Mailand, N., Bekker-Jensen, S., Bartek, J., and Lukas, J. (2006). Destruction of Claspin by SCFbetaTrCP restrains Chk1 activation and facilitates recovery from genotoxic stress. Mol Cell 23, 307-318.
Malumbres, M., Sotillo, R., Santamaria, D., Galan, J., Cerezo, A., Ortega, S., Dubus, P., and Barbacid, M. (2004). Mammalian cells cycle without the D-type cyclin-dependent kinases Cdk4 and Cdk6. Cell 118, 493-504.
Manchado, E., Eguren, M., and Malumbres, M. The anaphase-promoting complex/cyclosome (APC/C): cell-cycle-dependent and -independent functions. Biochem Soc Trans 38, 65-71.
Margottin-Goguet, F., Hsu, J.Y., Loktev, A., Hsieh, H.M., Reimann, J.D., and Jackson, P.K. (2003). Prophase destruction of Emi1 by the SCF(betaTrCP/Slimb) ubiquitin ligase activates the anaphase promoting complex to allow progression beyond prometaphase. Dev Cell 4, 813-826.
Marzec, M., Kasprzycka, M., Lai, R., Gladden, A.B., Wlodarski, P., Tomczak, E., Nowell, P., Deprimo, S.E., Sadis, S., Eck, S., et al. (2006). Mantle cell lymphoma cells express predominantly cyclin D1a isoform and are highly sensitive to selective inhibition of CDK4 kinase activity. Blood 108, 1744-1750.
Matsushime, H., Quelle, D.E., Shurtleff, S.A., Shibuya, M., Sherr, C.J., and Kato, J.Y. (1994). D-type cyclin-dependent kinase activity in mammalian cells. Mol Cell Biol 14, 2066-2076.
Matsushime, H., Roussel, M.F., Ashmun, R.A., and Sherr, C.J. (1991). Colony-stimulating factor 1 regulates novel cyclins during the G1 phase of the cell cycle. Cell 65, 701-713.
Matsuura, I., Denissova, N.G., Wang, G., He, D., Long, J., and Liu, F. (2004). Cyclin-dependent kinases regulate the antiproliferative function of Smads. Nature 430, 226-231.
McMahon, C., Suthiphongchai, T., DiRenzo, J., and Ewen, M.E. (1999). P/CAF associates with cyclin D1 and potentiates its activation of the estrogen receptor. Proc Natl Acad Sci U S A 96, 5382-5387.
Meraldi, P., Lukas, J., Fry, A.M., Bartek, J., and Nigg, E.A. (1999). Centrosome duplication in mammalian somatic cells requires E2F and Cdk2-cyclin A. Nat Cell Biol 1, 88-93.
Miller, J.J., Summers, M.K., Hansen, D.V., Nachury, M.V., Lehman, N.L., Loktev, A., and Jackson, P.K. (2006). Emi1 stably binds and inhibits the anaphase-promoting complex/cyclosome as a pseudosubstrate inhibitor. Genes Dev 20, 2410-2420.
Moore, J.D., Kirk, J.A., and Hunt, T. (2003). Unmasking the S-phase-promoting potential of cyclin B1. Science 300, 987-990.
Moreno-Bueno, G., Rodriguez-Perales, S., Sanchez-Estevez, C., Hardisson, D., Sarrio, D., Prat, J., Cigudosa, J.C., Matias-Guiu, X., and Palacios, J. (2003). Cyclin D1 gene (CCND1) mutations in endometrial cancer. Oncogene 22, 6115-6118.
Murphy, M., Stinnakre, M.G., Senamaud-Beaufort, C., Winston, N.J., Sweeney, C., Kubelka, M., Carrington, M., Brechot, C., and Sobczak-Thepot, J. (1997). Delayed early embryonic lethality following disruption of the murine cyclin A2 gene. Nat Genet 15, 83-86.
Nakayama, K., Ishida, N., Shirane, M., Inomata, A., Inoue, T., Shishido, N., Horii, I., and Loh, D.Y. (1996). Mice lacking p27(Kip1) display increased body size, multiple organ hyperplasia, retinal dysplasia, and pituitary tumors. Cell 85, 707-720.
Nakayama, K., Nagahama, H., Minamishima, Y.A., Matsumoto, M., Nakamichi, I., Kitagawa, K., Shirane, M., Tsunematsu, R., Tsukiyama, T., Ishida, N., et al. (2000). Targeted disruption of Skp2 results in accumulation of cyclin E and p27(Kip1), polyploidy and centrosome overduplication. Embo J 19, 2069-2081.
Nakayama, K.I., and Nakayama, K. (2005). Regulation of the cell cycle by SCF-type ubiquitin ligases. Semin Cell Dev Biol 16, 323-333.
Nateri, A.S., Riera-Sans, L., Da Costa, C., and Behrens, A. (2004). The ubiquitin ligase SCFFbw7 antagonizes apoptotic JNK signaling. Science 303, 1374-1378.
Nigg, E.A. (2001). Mitotic kinases as regulators of cell division and its checkpoints. Nat Rev Mol Cell Biol 2, 21-32.
Nishitani, H., Sugimoto, N., Roukos, V., Nakanishi, Y., Saijo, M., Obuse, C., Tsurimoto, T., Nakayama, K.I., Nakayama, K., Fujita, M., et al. (2006). Two E3 ubiquitin ligases, SCF-Skp2 and DDB1-Cul4, target human Cdt1 for proteolysis. Embo J 25, 1126-1136.
Norio, P., Kosiyatrakul, S., Yang, Q., Guan, Z., Brown, N.M., Thomas, S., Riblet, R., and Schildkraut, C.L. (2005). Progressive activation of DNA replication initiation in large domains of the immunoglobulin heavy chain locus during B cell development. Mol Cell 20, 575-587.
O'Connell, B.C., and Harper, J.W. (2007). Ubiquitin proteasome system (UPS): what can chromatin do for you? Curr Opin Cell Biol 19, 206-214.
Oberg, C., Li, J., Pauley, A., Wolf, E., Gurney, M., and Lendahl, U. (2001). The Notch intracellular domain is ubiquitinated and negatively regulated by the mammalian Sel-10 homolog. J Biol Chem 276, 35847-35853.
Okabe, H., Lee, S.H., Phuchareon, J., Albertson, D.G., McCormick, F., and Tetsu, O. (2006). A critical role for FBXW8 and MAPK in cyclin D1 degradation and cancer cell proliferation. PLoS ONE 1, e128.
Onoyama, I., Tsunematsu, R., Matsumoto, A., Kimura, T., de Alboran, I.M., Nakayama, K., and Nakayama, K.I. (2007). Conditional inactivation of Fbxw7 impairs cell-cycle exit during T cell differentiation and results in lymphomatogenesis. J Exp Med 204, 2875-2888.
Ortega, S., Prieto, I., Odajima, J., Martin, A., Dubus, P., Sotillo, R., Barbero, J.L., Malumbres, M., and Barbacid, M. (2003). Cyclin-dependent kinase 2 is essential for meiosis but not for mitotic cell division in mice. Nat Genet.
Ougolkov, A., Zhang, B., Yamashita, K., Bilim, V., Mai, M., Fuchs, S.Y., and Minamoto, T. (2004). Associations among beta-TrCP, an E3 ubiquitin ligase receptor, beta-catenin, and NF-kappaB in colorectal cancer. J Natl Cancer Inst 96, 1161-1170.
Pal, S., and Sif, S. (2007). Interplay between chromatin remodelers and protein arginine methyltransferases. J Cell Physiol 213, 306-315.
Pal, S., Vishwanath, S.N., Erdjument-Bromage, H., Tempst, P., and Sif, S. (2004). Human SWI/SNF-associated PRMT5 methylates histone H3 arginine 8 and negatively regulates expression of ST7 and NM23 tumor suppressor genes. Mol Cell Biol 24, 9630-9645.
Parisi, T., Beck, A.R., Rougier, N., McNeil, T., Lucian, L., Werb, Z., and Amati, B. (2003). Cyclins E1 and E2 are required for endoreplication in placental trophoblast giant cells. Embo J 22, 4794-4803.
Parrinello, S., Samper, E., Krtolica, A., Goldstein, J., Melov, S., and Campisi, J. (2003). Oxygen sensitivity severely limits the replicative lifespan of murine fibroblasts. Nat Cell Biol 5, 741-747.
Pavletich, N.P. (1999). Mechanisms of cyclin-dependent kinase regulation: structures of Cdks, their cyclin activators, and Cip and INK4 inhibitors. J Mol Biol 287, 821-828.
Pawar, S.A., Sarkar, T.R., Balamurugan, K., Sharan, S., Wang, J., Zhang, Y., Dowdy, S.F., Huang, A.M., and Sterneck, E. (2010). C/EBP{delta} targets cyclin D1 for proteasome-mediated degradation via induction of CDC27/APC3 expression. Proc Natl Acad Sci U S A 107, 9210-9215.
Peschiaroli, A., Dorrello, N.V., Guardavaccaro, D., Venere, M., Halazonetis, T., Sherman, N.E., and Pagano, M. (2006). SCFbetaTrCP-mediated degradation of Claspin regulates recovery from the DNA replication checkpoint response. Mol Cell 23, 319-329.
Peters, J.M. (2006). The anaphase promoting complex/cyclosome: a machine designed to destroy. Nat Rev Mol Cell Biol 7, 644-656.
Petersen, B.O., Lukas, J., Sorensen, C.S., Bartek, J., and Helin, K. (1999). Phosphorylation of mammalian CDC6 by cyclin A/CDK2 regulates its subcellular localization. Embo J 18, 396-410.
Petersen, B.O., Wagener, C., Marinoni, F., Kramer, E.R., Melixetian, M., Lazzerini Denchi, E., Gieffers, C., Matteucci, C., Peters, J.M., and Helin, K. (2000). Cell cycle- and cell growth-regulated proteolysis of mammalian CDC6 is dependent on APC-CDH1. Genes Dev 14, 2330-2343.
Petroski, M.D., and Deshaies, R.J. (2005). Function and regulation of cullin-RING ubiquitin ligases. Nat Rev Mol Cell Biol 6, 9-20.
Pfleger, C.M., and Kirschner, M.W. (2000). The KEN box: an APC recognition signal distinct from the D box targeted by Cdh1. Genes Dev 14, 655-665.
Pines, J., and Hunter, T. (1991). Human cyclins A and B1 are differentially located in the cell and undergo cell cycle-dependent nuclear transport. J Cell Biol 115, 1-17.
Pontano, L.L., Aggarwal, P., Barbash, O., Brown, E.J., Bassing, C.H., and Diehl, J.A. (2008). Genotoxic stress-induced cyclin D1 phosphorylation and proteolysis are required for genomic stability. Mol Cell Biol 28, 7245-7258.
Pontano, L.L., and Diehl, J.A. (2008). Speeding through cell cycle roadblocks: Nuclear cyclin D1-dependent kinase and neoplastic transformation. Cell Div 3, 12.
Pontano, L.L., and Diehl, J.A. (2009). DNA damage-dependent cyclin D1 proteolysis: GSK3beta holds the smoking gun. Cell Cycle 8, 824-827.
Popov, N., Schulein, C., Jaenicke, L.A., and Eilers, M. Ubiquitylation of the amino terminus of Myc by SCF(beta-TrCP) antagonizes SCF(Fbw7)-mediated turnover. Nat Cell Biol 12, 973-981.
Quelle, D.E., Ashmun, R.A., Shurtleff, S.A., Kato, J.Y., Bar-Sagi, D., Roussel, M.F., and Sherr, C.J. (1993). Overexpression of mouse D-type cyclins accelerates G1 phase in rodent fibroblasts. Genes Dev 7, 1559-1571.
Raman, M., Earnest, S., Zhang, K., Zhao, Y., and Cobb, M.H. (2007). TAO kinases mediate activation of p38 in response to DNA damage. Embo J 26, 2005-2014.
Ramiro, A.R., Jankovic, M., Eisenreich, T., Difilippantonio, S., Chen-Kiang, S., Muramatsu, M., Honjo, T., Nussenzweig, A., and Nussenzweig, M.C. (2004). AID is required for c-myc/IgH chromosome translocations in vivo. Cell 118, 431-438.
Rane, S.G., Dubus, P., Mettus, R.V., Galbreath, E.J., Boden, G., Reddy, E.P., and Barbacid, M. (1999). Loss of Cdk4 expression causes insulin-deficient diabetes and Cdk4 activation results in beta-islet cell hyperplasia. Nat Genet 22, 44-52.
Reinhardt, H.C., and Yaffe, M.B. (2009). Kinases that control the cell cycle in response to DNA damage: Chk1, Chk2, and MK2. Curr Opin Cell Biol 21, 245-255.
Robbiani, D.F., Bothmer, A., Callen, E., Reina-San-Martin, B., Dorsett, Y., Difilippantonio, S., Bolland, D.J., Chen, H.T., Corcoran, A.E., Nussenzweig, A., et al. (2008). AID is required for the chromosomal breaks in c-myc that lead to c-myc/IgH translocations. Cell 135, 1028-1038.
Rosenberg, A.R., Zindy, F., Le Deist, F., Mouly, H., Metezeau, P., Brechot, C., and Lamas, E. (1995). Overexpression of human cyclin A advances entry into S phase. Oncogene 10, 1501-1509.
Russo, A.A., Jeffrey, P.D., and Pavletich, N.P. (1996). Structural basis of cyclin-dependent kinase activation by phosphorylation. Nat Struct Biol 3, 696-700.
Saha, A., and Deshaies, R.J. (2008). Multimodal activation of the ubiquitin ligase SCF by Nedd8 conjugation. Mol Cell 32, 21-31.
Sansam, C.L., Shepard, J.L., Lai, K., Ianari, A., Danielian, P.S., Amsterdam, A., Hopkins, N., and Lees, J.A. (2006). DTL/CDT2 is essential for both CDT1 regulation and the early G2/M checkpoint. Genes Dev 20, 3117-3129.
Santamaria, D., Barriere, C., Cerqueira, A., Hunt, S., Tardy, C., Newton, K., Caceres, J.F., Dubus, P., Malumbres, M., and Barbacid, M. (2007). Cdk1 is sufficient to drive the mammalian cell cycle. Nature 448, 811-815.
Santra, M.K., Wajapeyee, N., and Green, M.R. (2009). F-box protein FBXO31 mediates cyclin D1 degradation to induce G1 arrest after DNA damage. Nature 459, 722-725.
Satyanarayana, A., Berthet, C., Lopez-Molina, J., Coppola, V., Tessarollo, L., and Kaldis, P. (2008). Genetic substitution of Cdk1 by Cdk2 leads to embryonic lethality and loss of meiotic function of Cdk2. Development 135, 3389-3400.
Sherr, C.J. (1994). G1 phase progression: cycling on cue. Cell 79, 551-555.
Sherr, C.J. (1996). Cancer cell cycles. Science 274, 1672-1677.
Sicinska, E., Aifantis, I., Le Cam, L., Swat, W., Borowski, C., Yu, Q., Ferrando, A.A., Levin, S.D., Geng, Y., von Boehmer, H., et al. (2003). Requirement for cyclin D3 in lymphocyte development and T cell leukemias. Cancer Cell 4, 451-461.
Sicinski, P., Donaher, J.L., Parker, S.B., Li, T., Fazeli, A., Gardner, H., Haslam, S.Z., Bronson, R.T., Elledge, S.J., and Weinberg, R.A. (1995). Cyclin D1 provides a link between development and oncogenesis in the retina and breast. Cell 82, 621-630.
Skowyra, D., Craig, K.L., Tyers, M., Elledge, S.J., and Harper, J.W. (1997). F-box proteins are receptors that recruit phosphorylated substrates to the SCF ubiquitin-ligase complex. Cell 91, 209-219.
Solvason, N., Wu, W.W., Parry, D., Mahony, D., Lam, E.W., Glassford, J., Klaus, G.G., Sicinski, P., Weinberg, R., Liu, Y.J., et al. (2000). Cyclin D2 is essential for BCR-mediated proliferation and CD5 B cell development. Int Immunol 12, 631-638.
Sorensen, C.S., Lukas, C., Kramer, E.R., Peters, J.M., Bartek, J., and Lukas, J. (2001). A conserved cyclin-binding domain determines functional interplay between anaphase-promoting complex-Cdh1 and cyclin A-Cdk2 during cell cycle progression. Mol Cell Biol 21, 3692-3703.
Stambolic, V., Ruel, L., and Woodgett, J.R. (1996). Lithium inhibits glycogen synthase kinase-3 activity and mimics wingless signalling in intact cells. Curr Biol 6, 1664-1668.
Stiff, T., Walker, S.A., Cerosaletti, K., Goodarzi, A.A., Petermann, E., Concannon, P., O'Driscoll, M., and Jeggo, P.A. (2006). ATR-dependent phosphorylation and activation of ATM in response to UV treatment or replication fork stalling. Embo J 25, 5775-5782.
Strausfeld, U.P., Howell, M., Descombes, P., Chevalier, S., Rempel, R.E., Adamczewski, J., Maller, J.L., Hunt, T., and Blow, J.J. (1996). Both cyclin A and cyclin E have S-phase promoting (SPF) activity in Xenopus egg extracts. J Cell Sci 109 ( Pt 6), 1555-1563.
Sundqvist, A., Bengoechea-Alonso, M.T., Ye, X., Lukiyanchuk, V., Jin, J., Harper, J.W., and Ericsson, J. (2005). Control of lipid metabolism by phosphorylation-dependent degradation of the SREBP family of transcription factors by SCF(Fbw7). Cell Metab 1, 379-391.
Suzuki, H., Chiba, T., Suzuki, T., Fujita, T., Ikenoue, T., Omata, M., Furuichi, K., Shikama, H., and Tanaka, K. (2000). Homodimer of two F-box proteins betaTrCP1 or betaTrCP2 binds to IkappaBalpha for signal-dependent ubiquitination. J Biol Chem 275, 2877-2884.
Tang, X., Orlicky, S., Lin, Z., Willems, A., Neculai, D., Ceccarelli, D., Mercurio, F., Shilton, B.H., Sicheri, F., and Tyers, M. (2007). Suprafacial orientation of the SCFCdc4 dimer accommodates multiple geometries for substrate ubiquitination. Cell 129, 1165-1176.
Tetzlaff, M.T., Yu, W., Li, M., Zhang, P., Finegold, M., Mahon, K., Harper, J.W., Schwartz, R.J., and Elledge, S.J. (2004). Defective cardiovascular development and elevated cyclin E and Notch proteins in mice lacking the Fbw7 F-box protein. Proc Natl Acad Sci U S A 101, 3338-3345.
Tsutsui, T., Hesabi, B., Moons, D.S., Pandolfi, P.P., Hansel, K.S., Koff, A., and Kiyokawa, H. (1999). Targeted disruption of CDK4 delays cell cycle entry with enhanced p27(Kip1) activity. Mol Cell Biol 19, 7011-7019.
Tsutsumi, T., Kuwabara, H., Arai, T., Xiao, Y., and Decaprio, J.A. (2008). Disruption of the Fbxw8 gene results in pre- and postnatal growth retardation in mice. Mol Cell Biol 28, 743-751.
Tsvetkov, L.M., Yeh, K.H., Lee, S.J., Sun, H., and Zhang, H. (1999). p27(Kip1) ubiquitination and degradation is regulated by the SCF(Skp2) complex through phosphorylated Thr187 in p27. Curr Biol 9, 661-664.
Vaziri, C., Saxena, S., Jeon, Y., Lee, C., Murata, K., Machida, Y., Wagle, N., Hwang, D.S., and Dutta, A. (2003). A p53-dependent checkpoint pathway prevents rereplication. Mol Cell 11, 997-1008.
Watanabe, N., Arai, H., Iwasaki, J., Shiina, M., Ogata, K., Hunter, T., and Osada, H. (2005). Cyclin-dependent kinase (CDK) phosphorylation destabilizes somatic Wee1 via multiple pathways. Proc Natl Acad Sci U S A 102, 11663-11668.
Watanabe, N., Arai, H., Nishihara, Y., Taniguchi, M., Hunter, T., and Osada, H. (2004). M-phase kinases induce phospho-dependent ubiquitination of somatic Wee1 by SCFbeta-TrCP. Proc Natl Acad Sci U S A 101, 4419-4424.
Weinberg, R.A. (1995). The retinoblastoma protein and cell cycle control. Cell 81, 323-330.
Welcker, M., and Clurman, B.E. (2008). FBW7 ubiquitin ligase: a tumour suppressor at the crossroads of cell division, growth and differentiation. Nat Rev Cancer 8, 83-93.
Welcker, M., Larimore, E.A., Frappier, L., and Clurman, B.E. Nucleolar targeting of the fbw7 ubiquitin ligase by a pseudosubstrate and glycogen synthase kinase 3. Mol Cell Biol 31, 1214-1224.
Welcker, M., Orian, A., Jin, J., Grim, J.A., Harper, J.W., Eisenman, R.N., and Clurman, B.E. (2004). The Fbw7 tumor suppressor regulates glycogen synthase kinase 3 phosphorylation-dependent c-Myc protein degradation. Proc Natl Acad Sci U S A 101, 9085-9090.
Welcker, M., Singer, J., Loeb, K.R., Grim, J., Bloecher, A., Gurien-West, M., Clurman, B.E., and Roberts, J.M. (2003). Multisite phosphorylation by Cdk2 and GSK3 controls cyclin E degradation. Mol Cell 12, 381-392.
Winget, J.M., and Mayor, T. The diversity of ubiquitin recognition: hot spots and varied specificity. Mol Cell 38, 627-635.
Winston, J.T., Koepp, D.M., Zhu, C., Elledge, S.J., and Harper, J.W. (1999a). A family of mammalian F-box proteins. Curr Biol 9, 1180-1182.
Winston, J.T., Strack, P., Beer-Romero, P., Chu, C.Y., Elledge, S.J., and Harper, J.W. (1999b). The SCFbeta-TRCP-ubiquitin ligase complex associates specifically with phosphorylated destruction motifs in IkappaBalpha and beta-catenin and stimulates IkappaBalpha ubiquitination in vitro. Genes Dev 13, 270-283.
Xiao, W., Zhang, Q., Habermacher, G., Yang, X., Zhang, A.Y., Cai, X., Hahn, J., Liu, J., Pins, M., Doglio, L., et al. (2008). U19/Eaf2 knockout causes lung adenocarcinoma, B-cell lymphoma, hepatocellular carcinoma and prostatic intraepithelial neoplasia. Oncogene 27, 1536-1544.
Xu, X., Sarikas, A., Dias-Santagata, D.C., Dolios, G., Lafontant, P.J., Tsai, S.C., Zhu, W., Nakajima, H., Nakajima, H.O., Field, L.J., et al. (2008). The CUL7 E3 ubiquitin ligase targets insulin receptor substrate 1 for ubiquitin-dependent degradation. Mol Cell 30, 403-414.
Yang, J., Bardes, E.S., Moore, J.D., Brennan, J., Powers, M.A., and Kornbluth, S. (1998). Control of cyclin B1 localization through regulated binding of the nuclear export factor CRM1. Genes Dev 12, 2131-2143.
Yu, H. (2007). Cdc20: a WD40 activator for a cell cycle degradation machine. Mol Cell 27, 3-16.
Yu, Z.K., Gervais, J.L., and Zhang, H. (1998). Human CUL-1 associates with the SKP1/SKP2 complex and regulates p21(CIP1/WAF1) and cyclin D proteins. Proc Natl Acad Sci U S A 95, 11324-11329.
Zeng, Z., Wang, W., Yang, Y., Chen, Y., Yang, X., Diehl, J.A., Liu, X., and Lei, M. (2010). Structural basis of selective ubiquitination of TRF1 by SCFFbx4. Dev Cell 18, 214-225.
Zhang, H., Ozaki, I., Mizuta, T., Hamajima, H., Yasutake, T., Eguchi, Y., Ideguchi, H., Yamamoto, K., and Matsuhashi, S. (2006). Involvement of programmed cell death 4 in transforming growth factor-beta1-induced apoptosis in human hepatocellular carcinoma. Oncogene 25, 6101-6112.
Zhang, W., and Koepp, D.M. (2006). Fbw7 isoform interaction contributes to cyclin E proteolysis. Mol Cancer Res 4, 935-943.
Zhao, Q., Rank, G., Tan, Y.T., Li, H., Moritz, R.L., Simpson, R.J., Cerruti, L., Curtis, D.J., Patel, D.J., Allis, C.D., et al. (2009). PRMT5-mediated methylation of histone H4R3 recruits DNMT3A, coupling histone and DNA methylation in gene silencing. Nat Struct Mol Biol 16, 304-311.
Zhou, J., Ashouian, N., Delepine, M., Matsuda, F., Chevillard, C., Riblet, R., Schildkraut, C.L., and Birshtein, B.K. (2002). The origin of a developmentally regulated Igh replicon is located near the border of regulatory domains for Igh replication and expression. Proc Natl Acad Sci U S A 99, 13693-13698.
Zwijsen, R.M., Buckle, R.S., Hijmans, E.M., Loomans, C.J., and Bernards, R. (1998). Ligand-independent recruitment of steroid receptor coactivators to estrogen receptor by cyclin D1. Genes Dev 12, 3488-3498.
Zwijsen, R.M., Wientjens, E., Klompmaker, R., van der Sman, J., Bernards, R., and Michalides, R.J. (1997). CDK-independent activation of estrogen receptor by cyclin D1. Cell 88, 405-415.