Open access peer-reviewed chapter

Regulation of Selective Proteolysis in Cancer

Written By

Pai-Sheng Chen

Submitted: June 11th, 2018 Reviewed: January 3rd, 2019 Published: February 19th, 2019

DOI: 10.5772/intechopen.83830

Chapter metrics overview

1,054 Chapter Downloads

View Full Metrics


Proteins are the fundamental building blocks of cells for diverse cellular and physiological functions. The dynamic equilibrium of protein turnover is balanced by protein synthesis and proteolysis. The newly synthesized proteins undergo proper folding into the three-dimensional conformations for executing biological functions and constructing cellular components like organelles. On the other hand, ubiquitin-proteasome system (UPS) and lysosome are two major proteolytic systems by which the unneeded, misfolded, or damaged proteins are selectively sent for clearance to maintain the quality and quantity of cellular proteins. Loss of the ability to maintain cellular proteolysis in control has been known to contribute as disease-causing factors. In this chapter, the function, regulation, and pathological roles of dysregulated proteolysis will be described in a concise view, focusing on the link between cancer and UPS.


  • ubiquitin-proteasome system
  • proteolysis
  • cancer

1. Introduction

Protein ubiquitination is a multistep process. It is initiated by an ATP-required activation and covalent binding of E1 ubiquitin-activating enzyme (E1) with ubiquitin [1]. The E1 then passes the ubiquitin to E2 ubiquitin-conjugating enzyme (E2) followed by forming complex with the E3 ubiquitin ligase (E3), which specifically recognizes substrate protein and catalyzes the ubiquitin transfer. Theoretically, E3s can function as oncogenes or tumor suppressors depending on the specificities on substrate proteins they targeted in cancer (Figure 1). For instance, MDM2 is oncogenic since it is the E3 for tumor-suppressive p53, while von Hippel–Lindau (VHL) disease tumor suppressor is tumor suppressive since it is the E3 for oncogenic HIF-1alpha (HIF-1α). However, alternative functions of E3 are also observed since multiple targets with diverse roles may be regulated by a common E3. Here, the selective ubiquitin-proteasome system (UPS) for p53, HIF-1α, and other cancer-related proteins are exemplified.

Figure 1.

Roles of E3s and DUBs in cancer.

Modification of substrate proteins by ubiquitination is the major way for selective proteolysis by proteasome. Ubiquitination is a reversible process controlled by the balance of ubiquitination and deubiquitination systems. This balance of ubiquitination is regulated by E3 ubiquitin ligases (E3s) [2] and deubiquitylating enzymes (DUBs) [3]. In addition to UPS-mediated protein degradation, ubiquitination is also involved in diverse non-proteolytic molecular and cellular functions, such as protein trafficking, activation, DNA repair, and apoptosis [4]. For example, K63-linked chains regulate DNA repair and NF-κB activation [5, 6, 7]. The TNF-α receptor–associated factor 6 (TRAF6) mediated K63-linked polyubiquitination of NF-κB essential modulator (NEMO) for IκB kinase (IKK) activation [8]. These studies indicate the versatile function of ubiquitination machinery. In this section, we focus on the role of ubiquitination in proteasomal degradation. There are seven lysine residues at positions 6, 11, 27, 29, 33, 48, and 63 of ubiquitin, which are utilized for polyubiquitination. These lysine residues serve as acceptors for other ubiquitins. Different types (mono, multi, poly) and links (K6, K11, K27, K29, K33, K48, K63) of ubiquitination determine the fate of tagged substrates [9, 10, 11]. For proteasome degradation, K48- and K11-linked polyubiquitination is the canonical signal that tags substrate proteins [12, 13, 14, 15, 16]. Recent studies showed that branched K48- and K11-linked chains enhanced proteasomal degradation, whereas homotypic K11 linkages prevent substrate protein recognition by proteasome [17]. K29-linked polyubiquitin is the most abundant atypical linkage in mammalian cells [18]. But little is known about its cellular function. The use of K29-linked chains as a degradation signal is also unclear as these chains may accumulate as a consequence of proteosomal stress induced by proteasome inhibition rather than via the accumulation of K29-linkage-modified proteasome substrates. The K63 linkage, while it can be recognized by the proteasome [19], is widely regarded as a non-degradative signaling modification that is known to regulate signal transduction and endocytosis [20, 21]. In addition, efficient proteasome activity has also been found to rely on the presence of K6-linked ubiquitination [22].


2. UPS-mediated proteolysis in cancer

2.1. Regulatory network for p53 degradation

Tumor protein p53 is a well-known tumor suppressor [23]. As a guardian of genome, p53 can sense DNA damages, activate repair systems, pause cell growth, or initiate apoptosis when necessary [23]. These functions establish a cellular protective machinery, thus loss of expression or tumor-suppressive activities of p53 are observed as a hallmark in cancer. Deregulation of p53 is orchestrated by multiple pathways, such as gene mutation and enhanced proteasomal degradation. As an E3 for p53, overexpression of MDM2 in human cancers has been linked to p53 degradation and tumorigenesis [24]. The transactivation domain of p53 is recognized by MDM2 and followed by MDM2-dependent ubiquitination and further proteasome degradation [25]. ATM serine/threonine kinase (ATM) is a DNA damage sensor participating in multiple mechanisms for p53 regulation. ATM-mediated phosphorylation of p53 at Ser15 is induced by genotoxic stress and therefore causes its escape from MDM recognition to further trigger cells to initiate DNA repair system through p53 [26, 27, 28]. Also, the MDM2-mediated p53 degradation is diminished when ATM-mediated MDM2 phosphorylation is induced by DNA damage [29]. Moreover, there is a negative feedback loop in which p53 activates the transcription of MDM2 [30]. Several inhibitors, such as Nutlin-3 and RG7112, were developed to disrupt the interaction between p53 and MDM2 and are currently undergoing clinical trials [31]. On the other hand, herpesvirus-associated ubiquitin-specific protease (HAUSP) is a deubiquitinase for p53. It removes ubiquitination and stabilizes p53 even in the presence of MDM2 [32]. Moreover, MDM2 is also stabilized by HAUSP through a p53-independent pathway [33, 34], suggesting a feedback regulatory loop between p53 and MDM2. The ATM-mediated phosphorylation, nuclear translocation, and stabilization of USP10 synergistically help nuclear HASUP stabilize p53 in the presence of DNA damage [35]. In addition, the constitutive photomorphogenesis protein 1 (COP1) forms an E3 ubiquitin ligase complex with cullin 4 (CUL4), DNA damage-binding protein 1 (DDB1), de-etiolated 1 (DET1), and ring-box 1 (RBX1) to target p53 [36]. Under genotoxic stress, ATM phosphorylates COP1 at Ser387 for degradation and subsequent p53 induction. Since p53 is targeted by COP1 for proteasomal degradation, downregulation or inactivation of COP1 subsequently activates p53 in cancer. Like MDM2, a transcriptional activation of COP1 by p53 forms a negative feedback loop [37]. Overexpression of COP1 is correlated with reduced p53 and has been observed in ovarian, breast, and liver cancers. P53-induced RING-H2 (Pirh2, also known as RCHY1) is another E3 ubiquitin ligase belonging to the RING finger family. Like MDM2, Pirh2 is considered as an oncogene to facilitate p53 protein degradation by UPS through a MDM2-independent manner [38, 39]. Notably, similar to the p53-MDM2 and p53-COP1 feedback loop, Pirh2 is also upregulated transcriptionally by p53. Interestingly, several researches suggested that Pirh2, but not MDM2, plays a major role in DNA damage-induced p53 degradation [38]. Moreover, in contrast to MDM2, Pirh2 can still recognize the p53 with Ser15 phosphorylation for UPS [40]. Elevated Pirh2 level has been found in human cancers and is correlated with unfavorable prognosis of cancer patients [41, 42]. The regulatory network for p53 degradation is illustrated in Figure 2.

Figure 2.

Regulatory network for p53 degradation.

2.2. Ubiquitination system of HIF-1α

During tumorigenesis, the increased tumor mass leads to the reduction of available intratumoral oxygen, which is theoretically a survival stress to normal cells. However, cancer cells develop several mechanisms to face this stressful condition, such as the activation of hypoxia-inducible factor 1α (HIF-1α). Through transcriptional regulation of downstream genes, accumulation of HIF-1α is not only observed on facilitating angiogenesis at the initiation of rapid tumor growth (also called angiogenic switch) but also enhances metastasis and malignant progression of cancer [43]. Expression of HIF-1α is tightly controlled by ubiquitination in coordination with hypoxia (Figure 3). Inactivation of Von Hippel-Lindau (VHL) in familial kidney cancer syndrome contributes to oncogenic effects [44]. At the molecular level, VHL interacts with cullin 2, elongin B, elongin C, and Rbx [45, 46, 47, 48]. This complex then targets HIF-1α for ubiquitination and proteasomal degradation [48, 49]. Under normoxia, prolyl hydroxylase (PHD) hydroxylates HIF-1α and facilitates its binding through N-TAD domain with VHL complex, leading to sustained ubiquitination and subsequent degradation of HIF-1α. The PHD-mediated post-translational modification (PTM) is abolished when cells encounter hypoxia during tumor growth. The stabilized HIF-1α is then accumulated in cancer cells and translocated to nucleus in complex with HIF-1β and other cofactors, resulting in transcriptional activation of downstream genes [50]. The transactivation activity of HIF-1α is also regulated by factor inhibiting HIF-1 (FIH-1). Through interaction with the ID and C-TAD domains, FIH-1 represses HIF-1α-mediated transactivation in association with histone deacetylase [51]. The HIF-1α-independent functions of VHL have also been pointed out. Besides HIF-1α, growing evidence has identified several targets of VHL with oncogenic properties in cancer [52]. It is well known that both downregulation of VHL and accumulation of HIF-1α are associated with cancer progression [53]. In addition to VHL, the chaperone-dependent E3 carboxy terminus of Hsp70-interacting protein (CHIP) is also identified to ubiquitinate HIF-1α for protein degradation [54]. Cellular response to hypoxia is also modulated by the E3s seven in absentia homolog (Siah) family proteins [55]. As another layer for HIF-1α regulation, Siah proteins are accumulated by transcriptional regulation and post-translational modification (PTM) under hypoxia [55]. The increased Siah proteins subsequently activate the degradation of PHDs and factors inhibiting HIF-1 (FIH) reduce prolyl hydroxylation of HIF-1α and consequently prevent VHL-mediated degradation [51, 55]. In addition to this regulation, there are several mechanisms known to cooperatively activate HIF-1α. For example, HIF-1α is stabilized when its ubiquitination is removed by ubiquitin-specific protease-19 (USP19) [56]. Siah proteins ubiquitinate the HIF-1α inhibitor, homeodomain-interacting protein kinase 2 (HIPK2), for degradation and thus enhance HIF-1α activity [57]. Siah2 also enhances the ubiquitination and degradation of large tumor suppressor kinase 2 (LATS2) resulting in suppressed HIPPO pathway and activated Yes-associated protein 1 (YAP1) that subsequently stabilizes HIF-1α [58]. Parkin is a recently-identified E3 for HIF-1α [59]. It facilitates HIF-1α polyubiquitination at K477 for proteasomal degradation through the interaction with HIF-1α. Alternatively, under the stimulations by hypoxia or growth factors, the induced HIF-1α brings Parkin and Dicer together, following by ubiquitination and autophagic degradation of Dicer, and eventually enhances cancer metastasis [60]. The findings exemplify the dual role of E3, which in this case, the target substrate (HIF-1α or Dicer) determines the fate of cellular function (Figure 4).

Figure 3.

Regulation of HIF-1α ubiquitination.

Figure 4.

Dual role of Parkin in cancer metastasis.

2.3. Cellular signaling regulated by UPS

Networks of signaling pathways coordinately orchestrate the cellular functions. Dysregulation of signal transduction pathways, especially those controlling oncogenic behaviors, is tightly regulated and also controlled by UPS. E3s play as modulators through regulating the proteolysis of key proteins in signaling networks. Several E3s can mediate substrate degradation to modulate PI3K/Akt/mTOR and RAS/MAPK, which are two central pathways, coordinately to control a broad range of tumor-promoting functions.

2.3.1. RAS/MAPK pathway

RAS oncogenes encode the highly-conserved RAS proteins as GTPases functioning in oncogenic transformation through the activation of MAPK pathway [61, 62]. Similar to p53, RAS mutations have been identified in human cancers, while stabilized RAS protein at post-translational level is also observed. The E3 ubiquitin-protein ligase, neural precursor cell-expressed developmentally downregulated protein 4 (NEDD4), is known to ubiquitinate RAS proteins for proteasomal degradation. NEDD4 targets KRAS, HRAS, NRAS for UPS, while its transcription is also activated by RAS signaling, which in turn, serves as a negative feedback to prevent the hyperactivation of RAS pathway [63]. More interestingly, this feedback mechanism is disrupted in cells expressing oncogenic RAS with activating mutation, exemplifying how an oncogenic protein (RAS) can escape from E3 (NEDD)-mediated degradation in cancer. Moreover, PTEN, a PI3K/Akt inhibitor, is another NEDD4 target, meaning that overexpression of NEDD4 facilitates PTEN degradation and activation of PI3K/Akt pathway. Thus, NEDD4 is supposedly to act as a tumor suppressor, but in cells expressing activating mutated RAS, NEDD4 no longer suppresses for RAS suppression and the concomitant PI3K/Akt activation corporately to amplify oncogenic signaling. In this case, NEDD4 functions as an oncogenic E3 [63]. However, a sustained RAS activation might be observed in lung cancer due to an elevated expression of DUB OTU domain-containing ubiquitin aldehyde-binding protein 1 (OTUB1), a deubiquitinase removing the ubiquitination of RAS and promoting the activation of RAS-mediated oncogenic downstream [64]. In addition to targeting RAS, several E3s are also identified to regulate downstream molecules of RAS. For example, ring finger protein 149 (RNF149) is an E3 targeting BRAF, a downstream kinase of RAS [65]. These machineries expend the complexity from reciprocal regulation in RAS/MAPK pathway.

2.3.2. PI3K/Akt pathway

PI3K/Akt pathway is induced by extracellular signaling such as activation of receptor tyrosine kinase (RTK) or G protein-coupled receptors (GPCRs). The regulatory subunit p85 and catalytic subunit p110 form heterodimer of PI3K [66]. In addition to p85, the p110 subunit also binds to Grb2 or insulin receptor substrate (IRS), and the competition from free p85 binding serves as an inhibitor for PI3K signaling [66, 67]. P85β is one of the variants of p85 subunits and is a target of the SCF-F-box and leucine-rich repeat protein 2 (FBXL2) complex [68]. SCF-FBXL2 enhances free p85β degradation through UPS and, consequently, disrupts the inhibitory effect of p85β pool on PI3K activation. Another layer of the regulation on this mechanism is controlled by dephosphorylation of p85β by PTPL1, which facilitates p85β degradation through enhanced interaction with FBXL2 [68]. The mechanistic target of rapamycin (mTOR) is a core component of PI3K/Akt pathway. The expression of mTOR is regulated by SCF-FBXW7 complex that triggers the ubiquitination and degradation of mTOR [69]. Loss of a single copy of FBXW7 in several breast cancer cell lines is observed to be incompatible with the loss of a single copy of PTEN, a negative regulator of mTOR, which further confirmed the significance of the stabilization of FBXW7-mediated mTOR in tumorigenesis. Therefore, loss of SCF-FBXW7 may motivate anabolic processes for tumor progression [69]. In addition to mTOR regulation, the F-box protein FBXW7 is a well-known tumor suppressor which recognizes and facilitates UPS of other oncogenic substrates including c-Myc, Notch1, and cyclin E [70, 71, 72, 73, 74, 75, 76, 77, 78, 79, 80, 81]. The mechanism of cyclin E regulation will be discussed later in this chapter.

2.4. E3s as cell cycle regulators

Uncontrolled cell growth is one of the hallmarks of cancer [82]. Cell cycle progression is a fundamental process for cell survival and proliferation. Properly regulated cell cycle progression is required for the maintenance of genome stability, organismal development, tissue homeostasis that when deregulation coordinately leads to defect of cell growth control. Signals that control cell cycle entry, progression, and arrest are commonly malfunctioned in cancer, and the subsequent disruption of DNA replication, DNA repair, and chromosomal segregation often lead to genomic instability [83]. There is aberrant degradation caused by improper E3 activity in cancer. For example, abnormal expression of E3s regulates the degradation of cyclins and cyclin-dependent kinase (CDK) inhibitor proteins by UPS. The Anaphase Promoting Complex/Cyclosome (APC/C) and Skp1, Cullin1 F-box (SCF) E3 complexes that regulate cell cycle progression are the best-studied E3s, which further form different complexes with specific co-activators and provide the functional diversity of E3s to recognize different substrates at different phases for orchestrating cell cycle progression. In mitosis and G1 phase of cell cycle, APC/C is active to suppress mitotic CDKs [84]. In contrast, various SCF complexes formed by different protein partners contribute to multifaceted functions during the cell cycle progression. Here, we will discuss these two ubiquitin-protein ligases, and how they cooperatively regulate cell cycle progression.

APC/C is a well-studied E3 that recognizes the D-box sequence of substrate proteins and ubiquitinates them for proteasome degradation [85]. The recognition of substrates by APC/C is known to rely on a short linear motif called degron (derived from degradation motif) including KEN-box, D-box, and ABBA motif [86]. KEN-box is the prominent signal among APC/C degron and is contained in substrate proteins, such as CDC20 and securin. The substrates of APC/C cover numerous cell cycle regulators. Thus, APC/C contributes to the cell cycle regulation, especially during M phase to G1 phase. Cell division cycle 20 (CDC20) or CDC20-like protein 1 (CDH1) are two known activators for APC/C activation [87]. Interestingly, the two activators show opposite functions in cancer as oncogenic CDC20 or tumor-suppressive CDH1 interact with APC/C to exert their spatial and temporal functions during cell cycle [87]. It is widely observed that CDC20 is highly expressed in human malignancies and associates with poor prognosis of cancer patients [88, 89, 90, 91, 92, 93]. Mechanistically, CDC20 recognizes securin [94], Cyclin A [95, 96], Cyclin B1 [97, 98], Nek2A [99], Mcl-1 [100], and p21 [101] as it substrates for ubiquitination. Cdc20 is primarily active in mid to late mitosis to promote ubiquitination and degradation of securin and cyclins to coordinately facilitate mitotic progression [87]. Thereafter, CDC20 degradation is triggered through ubquitination by APC/C-CDH1 or by itself in late M phase. As a result, the APC/C complex shifts from APC/C-CDC20 to APC/C-CDH1. APC/C-CDH1 is activated at late mitosis phase to degrade mitotic regulators, such as cyclins and kinases, and thereby promotes cells to exit from M phase and enter G1 phase to further prevent premature S phase entry [87]. Mutation or abnormal expression of CDH1 leads to genomic instability and premature S-phase entry [87, 102].

S-phase kinase-associated protein (Skp), cullin, and F-box domain containing proteins (F-box proteins) form SCF E3 ubiquitin ligase complex. Aurora kinase A and Cyclin E are substrates for SCF-FBXW7, thus inactivation of this complex causes defect in DNA repair system and sustained cell growth [103] suggesting the tumor-suppressive roles of SCF-FBXW7. As a key factor in SCF complex, dysregulation of F-box protein is frequently observed to affect SCF activity. SCF-Skp2 regulates cell cycle from G1/S to G2/M phase by targeting multiple CDK inhibitors for UPS degradation and consequently leads to enhanced cell cycle progression and tumorigenesis [104, 105, 106, 107, 108, 109, 110, 111]. Interestingly, SCF-βTRCP complex also mediates the crosstalk between APC/C and SCF complex during cell cycle. Degradation of the APC/C inhibitor Emi1 during prometaphase and degradation of the Cdk1 inhibitor Wee1 during prophase are facilitated by SCF-βTRCP in cell cycle progression [112, 113]. Activation of SCF-Skp2 complex during G1 to S phase degrades cyclin-dependent kinase inhibitors (CKIs), thereby induces CDK activity. The CDK2-mediated phosphorylation and inactivation of CDH1 further stabilize Skp2 by phosphorylation. SKP2 is also a D-box-containing target of APC/C-CDH1 as an autoregulatory loop [114]. It is also noticed that the casein kinase I (CKI)-phosphorylated MDM2 is targeted and degraded by SCF-βTRCP complex and results in p53 stabilization and cell cycle arrest [115].

Parkin is another E3 participating in cell cycle regulation. Mutations and loss of copy number of PARK2, a gene encoding Parkin, are observed in cancer, which implies its tumor-suppressive role [116, 117, 118, 119, 120, 121]. Loss of Parkin expression respectively results in the elevation of Cyclin D and Cyclin E owing to the suppression of FBXW7-containing Parkin-cullin-RING or F-box only protein 4 (FBXO4)-containing complexes [117]. In animal models, Park2+/−ApcMin/+ mice have higher rate of tumor formation than ApcMin/+ mice, which may result from the accumulation of Cyclin E and uncontrolled cell growth when Parkin expression is lost [118, 120]. Similar associations between PARK2 mutations and Cyclin D, Cyclin E, and CDK4 are also observed in human cancers [117]. Therefore, Parkin may also regulate several cell cycle or mitotic regulators including CDC20, CDH1, Aurora kinase A, Aurora kinase B, NEK2, PLK1, Cyclin B1, and securin, suggesting its function in maintaining genomic stability and growth control to suppress tumor formation [119].


3. Conclusions

Malfunction of UPS machinery, especially the target selection factor E3, has been observed in cancer for a period of time. Abnormal expression, mutation, distribution of E3s, or even the degradation of themselves may affect the affinity or activity on substrate recognition and ubiquitination, and thus consequently regulate proteasomal degradation and cellular behaviors depending on the normal functions of dysregulated targets. Although we have focused on the selective proteolysis through UPS, E3-mediated ubiquitination is not the only way for proteasomal degradation and also, proteasomal degradation is not the only fate for ubiquitinated proteins. Oftentimes, these proteins undergo autophagic degradation, intracellular localization, functional inhibition, or activation. Moreover, the lysosomal and autolysosomal (autophagy-lysosomal) degradation, which are not described in detail in this chapter, are responsible for another side of selective proteolysis. In concert with the landscape of post-translational modification, the crosstalk and cooperation among these proteolysis systems enable our cells to maintain biological functions in control. Simply speaking, proteolysis serves as a dead end for protein, thus the selection of target substrates should be tightly controlled. This chapter introduces several pathways as examples of selective UPS. In addition, there are several clinical trials for drugs designed to target proteolysis. As we know more about the mechanisms, we are moving a step forward in developing strategies to fix the proteolytic chaos of cells.



We would like to thank Jie-Ning Li, Yu-Jhen Lyu, and Yuchieh Chien for their help on content editing.


Conflict of interest

The author declares that there is no conflict of interest.


  1. 1. Ciechanover A. Intracellular protein degradation: From a vague idea through the lysosome and the ubiquitin-proteasome system and onto human diseases and drug targeting. Cell Death and Differentiation. 2005;12(9):1178-1190
  2. 2. Chhangani D, Joshi AP, Mishra A. E3 ubiquitin ligases in protein quality control mechanism. Molecular Neurobiology. 2012;45(3):571-585
  3. 3. Eletr ZM, Wilkinson KD. Regulation of proteolysis by human deubiquitinating enzymes. Biochimica et Biophysica Acta-Molecular Cell Research. 2014;1843(1):114-128
  4. 4. Mukhopadhyay D, Riezman H. Proteasome-independent functions of ubiquitin in endocytosis and signaling. Science. 2007;315(5809):201-205
  5. 5. Liu PD, Gan WJ, Su SY, Hauenstein AV, Fu TM, Brasher B, et al. K63-linked polyubiquitin chains bind to DNA to facilitate DNA damage repair. Science Signaling. 2018;11(533)
  6. 6. Jackson SP, Durocher D. Regulation of DNA damage responses by ubiquitin and SUMO. Molecular Cell. 2013;49(5):795-807
  7. 7. Chen ZJJ. Ubiquitin signalling in the NF-kappa B pathway. Nature Cell Biology. 2005;7(8):758-765
  8. 8. Sun LJ, Deng L, Ea CK, Xia ZP, Chen ZJJ. The TRAF6 ubiquitin ligase and TAK1 kinase mediate IKK activation by BCL10 and MALT1 in T lymphocytes. Molecular Cell. 2004;14(3):289-301
  9. 9. Raasi S, Varadan R, Fushman D, Pickart CM. Diverse polyubiquitin interaction properties of ubiquitin-associated domains. Nature Structural & Molecular Biology. 2005;12(8):708-714
  10. 10. Varadan R, Assfalg M, Raasi S, Pickart C, Fushman D. Structural determinants for selective recognition of a lys48-linked polyubiquitin chain by a UBA domain. Molecular Cell. 2005;18(6):687-698
  11. 11. Hicke L, Schubert HL, Hill CP. Ubiquitin-binding domains. Nature Reviews Molecular Cell Biology. 2005;6(8):610-621
  12. 12. Koegl M, Hoppe T, Schlenker S, Ulrich HD, Mayer TU, Jentsch S. A novel ubiquitination factor, E4, is involved in multiubiquitin chain assembly. Cell. 1999;96(5):635-644
  13. 13. Crosas B, Hanna J, Kirkpatrick DS, Zhang DP, Tone Y, Hathaway NA, et al. Ubiquitin chains are remodeled at the proteasome by opposing ubiquitin ligase and deubiquitinating activities. Cell. 2006;127(7):1401-1413
  14. 14. Kim HT, Kim KP, Lledias F, Kisselev AF, Scaglione KM, Skowyra D, et al. Certain pairs of ubiquitin-conjugating enzymes (E2s) and ubiquitin-protein ligases (E3s) synthesize nondegradable forked ubiquitin chains containing all possible isopeptide linkages. Journal of Biological Chemistry. 2007;282(24):17375-17386
  15. 15. Johnson ES, Ma PCM, Ota IM, Varshavsky A. A Proteolytic pathway that recognizes ubiquitin as a degradation signal. Journal of Biological Chemistry. 1995;270(29):17442-17456
  16. 16. Kirkpatrick DS, Hathaway NA, Hanna J, Elsasser S, Rush J, Finley D, et al. Quantitative analysis of in vitro ubiquitinated cyclin B1 reveals complex chain topology. Nature Cell Biology. 2006;8(7):700-U121
  17. 17. Grice GL, Lobb IT, Weekes MP, Gygi SP, Antrobus R, Nathan JA. The proteasome distinguishes between heterotypic and Homotypic Lysine-11-linked polyubiquitin chains. Cell Reports. 2015;12(4):545-553
  18. 18. Dammer EB, Na CH, Xu P, Seyfried NT, Duong DM, Cheng DM, et al. Polyubiquitin linkage profiles in three models of proteolytic stress suggest the etiology of Alzheimer disease. Journal of Biological Chemistry. 2011;286(12):10457-10465
  19. 19. Ohtake F, Tsuchiya H, Saeki Y, Tanaka K. K63 ubiquitylation triggers proteasomal degradation by seeding branched ubiquitin chains. Proceedings of the National Academy of Sciences of the United States of America. 2018;115(7):E1401-E1408
  20. 20. Husnjak K, Dikic I. Ubiquitin-binding proteins: Decoders of ubiquitin-mediated cellular functions. Annual Review of Biochemistry. 2012;81:291-322
  21. 21. Komander D, Rape M. The ubiquitin code. Annual Review of Biochemistry. 2012;81:203-229
  22. 22. Shang F, Deng GJ, Liu Q, Guo WM, Haas AL, Crosas B, et al. Lys(6)-modified ubiquitin inhibits ubiquitin-dependent protein degradation. Journal of Biological Chemistry. 2005;280(21):20365-20374
  23. 23. Bieging KT, Mello SS, Attardi LD. Unravelling mechanisms of p53-mediated tumour suppression. Nature Reviews Cancer. 2014;14(5):359-370
  24. 24. Wade M, Li YC, Wahl GM. MDM2, MDMX and p53 in oncogenesis and cancer therapy. Nature Reviews Cancer. 2013;13(2):83-96
  25. 25. Vousden KH, Lu X. Live or let die: The cell’s response to p53. Nature Reviews Cancer. 2002;2(8):594-604
  26. 26. Banin S, Moyal L, Shieh S, Taya Y, Anderson CW, Chessa L, et al. Enhanced phosphorylation of p53 by ATM in response to DNA damage. Science. 1998;281(5383):1674-1677
  27. 27. Canman CE, Lim DS, Cimprich KA, Taya Y, Tamai K, Sakaguchi K, et al. Activation of the ATM kinase by ionizing radiation and phosphorylation of p53. Science. 1998;281(5383):1677-1679
  28. 28. Shieh SY, Ikeda M, Taya Y, Prives C. DNA damage-induced phosphorylation of p53 alleviates inhibition by MDM2. Cell. 1997;91(3):325-334
  29. 29. Maya R, Balass M, Kim ST, Shkedy D, Leal JF, Shifman O, et al. ATM-dependent phosphorylation of Mdm2 on serine 395: Role in p53 activation by DNA damage. Genes & Development. 2001;15(9):1067-1077
  30. 30. Lahav G, Rosenfeld N, Sigal A, Geva-Zatorsky N, Levine AJ, Elowitz MB, et al. Dynamics of the p53-Mdm2 feedback loop in individual cells. Nature Genetics. 2004;36(2):147-150
  31. 31. Zhao YJ, Aguilar A, Bernard D, Wang SM. Small-molecule inhibitors of the MDM2-p53 protein-protein interaction (MDM2 inhibitors) in clinical trials for cancer treatment. Journal of Medicinal Chemistry. 2015;58(3):1038-1052
  32. 32. Li MY, Chen DL, Shiloh A, Luo JY, Nikolaev AY, Qin J, et al. Deubiquitination of p53 by HAUSP is an important pathway for p53 stabilization. Nature. 2002;416(6881):648-653
  33. 33. Li MY, Brooks CL, Kon N, Gu W. A dynamic role of HAUSP in the p53-Mdm2 pathway. Molecular Cell. 2004;13(6):879-886
  34. 34. Cummins JM, Rago C, Kohli M, Kinzler KW, Lengauer C, Vogelstein B. Disruption of HAUSP gene stabilizes p53. Nature. 2004;428(6982)
  35. 35. Yuan J, Luo KT, Zhang LZ, Cheville JC, Lou ZK. USP10 regulates p53 localization and stability by deubiquitinating p53. Cell. 2010;140(3):384-396
  36. 36. Dornan D, Wertz I, Shimizu H, Arnott D, Frantz GD, Dowd P, et al. The ubiquitin ligase COP1 is a critical negative regulator of p53. Nature. 2004;429(6987):86-92
  37. 37. Brooks CL, Gu W. p53 ubiquitination: Mdm2 and beyond. Molecular Cell. 2006;21(3):307-315
  38. 38. Leng RP, Lin YP, Ma WL, Wu H, Lemmers B, Chung S, et al. Pirh2, a p53-induced ubiquitin-protein ligase, promotes p53 degradation. Cell. 2003;112(6):779-791
  39. 39. Sheng Y, Laister RC, Lemak A, Wu B, Tai E, Duan S, et al. Molecular basis of Pirh2-mediated p53 ubiquitylation. Nature Structural & Molecular Biology. 2008;15(12):1334-1342
  40. 40. Wang ZH, Yang B, Dong LL, Peng BW, He XH, Liu WH. A novel oncoprotein Pirh2: Rising from the shadow of MDM2. Cancer Science. 2011;102(5):909-917
  41. 41. Duan WR, Gao L, Druhan LJ, Zhu WG, Morrison C, Otterson GA, et al. Expression of Pirh2, a newly identified ubiquitin protein ligase, in lung cancer. Journal of the National Cancer Institute. 2004;96(22):1718-1721
  42. 42. Logan IR, Gaughan L, McCracken SRC, Sapountzi V, Leung HY, Robson CN. Human PIRH2 enhances androgen receptor signaling through inhibition of histone deacetylase 1 and is overexpressed in prostate cancer. Molecular and Cellular Biology. 2006;26(17):6502-6510
  43. 43. Semenza GL. Targeting HIF-1 for cancer therapy. Nature Reviews. Cancer. 2003;3(10):721-732
  44. 44. Latif F, Tory K, Gnarra J, Yao M, Duh FM, Orcutt ML, et al. Identification of the von Hippel-Lindau disease tumor suppressor gene. Science. 1993;260(5112):1317-1320
  45. 45. Kibel A, Iliopoulos O, DeCaprio JA, and Kaelin WG, Jr., Binding of the von Hippel-Lindau tumor suppressor protein to Elongin B and C. Science 1995;269(5229):1444-1446
  46. 46. Pause A, Lee S, Worrell RA, Chen DY, Burgess WH, Linehan WM, et al. The von Hippel-Lindau tumor-suppressor gene product forms a stable complex with human CUL-2, a member of the Cdc53 family of proteins. Proceedings of the National Academy of Sciences of the United States of America. 1997;94(6):2156-2161
  47. 47. Kamura T, Koepp DM, Conrad MN, Skowyra D, Moreland RJ, Iliopoulos O, et al. Rbx1, a component of the VHL tumor suppressor complex and SCF ubiquitin ligase. Science. 1999;284(5414):657-661
  48. 48. Ohh M, Park CW, Ivan N, Hoffman MA, Kim TY, Huang LE, et al. Ubiquitination of hypoxia-inducible factor requires direct binding to the beta-domain of the von Hippel-Lindau protein. Nature Cell Biology. 2000;2(7):423-427
  49. 49. Tanimoto K, Makino Y, Pereira T, Poellinger L. Mechanism of regulation of the hypoxia-inducible factor-1 alpha by the von Hippel-Lindau tumor suppressor protein. The EMBO Journal. 2000;19(16):4298-4309
  50. 50. Triner D, Shah YM. Hypoxia-inducible factors: A central link between inflammation and cancer. Journal of Clinical Investigation. 2016;126(10):3689-3698
  51. 51. Mahon PC, Hirota K, Semenza GL. FIH-1: A novel protein that interacts with HIF-1 alpha and VHL to mediate repression of HIF-1 transcriptional activity. Genes & Development. 2001;15(20):2675-2686
  52. 52. Zhang J, Zhang Q. VHL and hypoxia signaling: Beyond HIF in cancer. Biomedicine. 2018;6(1): pii: E35. DOI: 10.3390/biomedicines6010035.
  53. 53. Kaelin WG Jr, The VHL. Tumor suppressor gene: Insights into oxygen sensing and cancer. Transactions of the American Clinical and Climatological Association. 2017;128:298-307
  54. 54. Bento CF, Fernandes R, Ramalho J, Marques C, Shang F, Taylor A, et al. The chaperone-dependent ubiquitin ligase CHIP targets HIF-1 alpha for degradation in the presence of methylglyoxal. PLoS One. 2010;5(11):e15062. DOI: 10.1371/journal.pone.0015062
  55. 55. Nakayama K, Frew IJ, Hagensen M, Skals M, Habelhah H, Bhoumik A, et al. Siah2 regulates stability of prolyl-hydroxylases, controls HIF1 alpha abundance, and modulates physiological responses to hypoxia. Cell. 2004;117(7):941-952
  56. 56. Altun M, Zhao B, Velasco K, Liu HY, Hassink G, Paschke J, et al. Ubiquitin-specific protease 19 (USP19) regulates hypoxia-inducible factor 1 alpha (HIF-1 alpha) during hypoxia. Journal of Biological Chemistry. 2012;287(3):1962-1969
  57. 57. Calzado MA, de la Vega L, Moller A, Bowtell DDL, Schmitz ML. An inducible autoregulatory loop between HIPK2 and Siah2 at the apex of the hypoxic response. Nature Cell Biology. 2009;11(1):85-U180
  58. 58. Ma B, Chen Y, Chen L, Cheng HC, Mu CL, Li J, et al. Hypoxia regulates Hippo signalling through the SIAH2 ubiquitin E3 ligase. Nature Cell Biology. 2015;17(1):95-103
  59. 59. Liu J, Zhang C, Zhao YH, Yue XT, Wu H, Huang S, et al. Parkin targets HIF-1 alpha for ubiquitination and degradation to inhibit breast tumor progression. Nature Communications. 2017;8
  60. 60. Lai HH, Li JN, Wang MY, Huang HY, Croce CM, Sun HL, et al. HIF-1alpha promotes autophagic proteolysis of dicer and enhances tumor metastasis. The Journal of Clinical Investigation. 2018;128(2):625-643
  61. 61. Simanshu DK, Nissley DV, McCormick F. RAS proteins and their regulators in human disease. Cell. 2017;170(1):17-33
  62. 62. Pylayeva-Gupta Y, Grabocka E, Bar-Sagi D. RAS oncogenes: Weaving a tumorigenic web. Nature Reviews Cancer. 2011;11(11):761-774
  63. 63. Zeng TL, Wang Q, Fu JY, Lin Q, Bi J, Ding WC, et al. Impeded Nedd4-1-mediated RAS degradation underlies Ras-driven tumorigenesis. Cell Reports. 2014;7(3):871-882
  64. 64. Baietti MF, Simicek M, Abbasi Asbagh L, Radaelli E, Lievens S, Crowther J, et al. OTUB1 triggers lung cancer development by inhibiting RAS monoubiquitination. EMBO Molecular Medicine. 2016;8(3):288-303
  65. 65. Hong SW, Jin DH, Shin JS, Moon JH, Na YS, Jung KA, et al. Ring finger protein 149 is an E3 ubiquitin ligase active on wild-type v-Raf murine sarcoma viral oncogene homolog B1 (BRAF). Journal of Biological Chemistry. 2012;287(28):24017-24025
  66. 66. Hemmings BA, Restuccia DF. PI3K-PKB/Akt Pathway. Cold Spring Harbor Perspectives in Biology. 2012;4(9):a011189. DOI: 10.1101/cshperspect.a011189
  67. 67. Luo J, Field SJ, Lee JY, Engelman JA, Cantley LC. The p85 regulatory subunit of phosphoinositide 3-kinase down-regulates IRS-1 signaling via the formation of a sequestration complex. Journal of Cell Biology. 2005;170(3):455-464
  68. 68. Kuchay S, Duan SS, Schenkein E, Peschiaroli A, Saraf A, Florens L, et al. FBXL2-and PTPL1-mediated degradation of p110-free p85 beta regulatory subunit controls the PI(3)K signalling cascade. Nature Cell Biology. 2013;15(5):472-480
  69. 69. Mao JH, Kim IJ, Wu D, Climent J, Kang HC, DelRosario R, et al. FBXW7 targets mTOR for degradation and cooperates with PTEN in tumor suppression. Science. 2008;321(5895):1499-1502
  70. 70. Yada M, Hatakeyama S, Kamura T, Nishiyama M, Tsunematsu R, Imaki H, et al. Phosphorylation-dependent degradation of c-Myc is mediated by the F-box protein Fbw7. EMBO Journal. 2004;23(10):2116-2125
  71. 71. King B, Trimarchi T, Reavie L, Xu LY, Mullenders J, Ntziachristos P, et al. The ubiquitin ligase FBXW7 modulates leukemia-initiating cell activity by regulating MYC stability. Cell. 2013;153(7):1552-1566
  72. 72. Reavie L, Buckley SM, Loizou E, Takeishi S, Aranda-Orgilles B, Ndiaye-Lobry D, et al. Regulation of c-Myc ubiquitination controls chronic myelogenous leukemia initiation and progression. Cancer Cell. 2013;23(3):362-375
  73. 73. Ntziachristos P, Lim JS, Sage J, Aifantis I. From fly wings to targeted cancer therapies: A centennial for notch signaling. Cancer Cell. 2014;25(3):318-334
  74. 74. Weng AP, Ferrando AA, Lee W, Morris JP, Silverman LB, Sanchez-Irizarry C, et al. Activating mutations of NOTCH1 in human T cell acute lymphoblastic leukemia. Science. 2004;306(5694):269-271
  75. 75. Malyukova A, Dohda T, von der Lehr N, Akhondi S, Corcoran M, Heyman M, et al. The tumor suppressor gene hCDC4 is frequently mutated in human T-cell acute lymphoblastic leukemia with functional consequences for notch signaling. Cancer Research. 2007;67(12):5611-5616
  76. 76. Strohmaier H, Spruck CH, Kaiser P, Won KA, Sangfelt O, Reed SI. Human F-box protein hCdc4 targets cyclin E for proteolysis and is mutated in a breast cancer cell line. Nature. 2001;413(6853):316-322
  77. 77. Minella AC, Grim JE, Welcker M, Clurman BE. P53 and SCFFbw7 cooperatively restrain cyclin E-associated genome instability. Oncogene. 2007;26(48):6948-6953
  78. 78. Rajagopalan H, Jallepalli PV, Rago C, Velculescu VE, Kinzler KW, Vogelstein B, et al. Inactivation of hCDC4 can cause chromosomal instability. Nature. 2004;428(6978):77-81
  79. 79. Hubalek MM, Widschwendter A, Erdel M, Gschwendtner A, Fiegl HM, Muller HM, et al. Cyclin E dysregulation and chromosomal instability in endometrial cancer. Oncogene. 2004;23(23):4187-4192
  80. 80. Koepp DM, Schaefer LK, Ye X, Keyomarsi K, Chu C, Harper JW, et al. Phosphorylation-dependent ubiquitination of cyclin E by the SCFFbw7 ubiquitin ligase. Science. 2001;294(5540):173-177
  81. 81. Welcker M, Singer J, Loeb KR, Grim J, Bloecher A, Gurien-West M, et al. Multisite phosphorylation by Cdk2 and GSK3 controls cyclin E degradation. Molecular Cell. 2003;12(2):381-392
  82. 82. Hanahan D, Weinberg RA. Hallmarks of cancer: The next generation. Cell. 2011;144(5):646-674
  83. 83. Jeggo PA, Pearl LH, Carr AM. DNA repair, genome stability and cancer: A historical perspective. Nature Reviews Cancer. 2016;16(1):35-42
  84. 84. Zachariae W, Nasmyth K. Whose end is destruction: Cell division and the anaphase-promoting complex. Genes & Development. 1999;13(16):2039-2058
  85. 85. Morgan DO. The D box meets its match. Molecular Cell. 2013;50(5):609-610
  86. 86. He J, Chao WCH, Zhang ZG, Yang J, Cronin N, Barford D. Insights into degron recognition by APC/C coactivators from the structure of an Acm1-Cdh1 complex. Molecular Cell. 2013;50(5):649-660
  87. 87. Pines J. Cubism and the cell cycle: The many faces of the APC/C. Nature Reviews Molecular Cell Biology. 2011;12(7):427-438
  88. 88. Jiang JH, Jedinak A, Sliva D. Ganodermanontriol (GDNT) exerts its effect on growth and invasiveness of breast cancer cells through the down-regulation of CDC20 and uPA. Biochemical and Biophysical Research Communications. 2011;415(2):325-329
  89. 89. Rajkumar T, Sabitha K, Vijayalakshmi N, Shirley S, Bose MV, Gopal G, et al. Identification and validation of genes involved in cervical tumourigenesis. BMC Cancer. 2011;11:80. DOI: 10.1186/1471-2407-11-80
  90. 90. Marucci G, Morandi L, Magrini E, Farnedi A, Franceschi E, Miglio R, et al. Gene expression profiling in glioblastoma and immunohistochemical evaluation of IGFBP-2 and CDC20. Virchows Archiv. 2008;453(6):599-609
  91. 91. Ouellet V, Guyot MC, Le Page C, Filali-Mouhim A, Lussier C, Tonin PN, et al. Tissue array analysis of expression microarray candidates identifies markers associated with tumor grade and outcome in serous epithelial ovarian cancer. International Journal of Cancer. 2006;119(3):599-607
  92. 92. Zaravinos A, Lambrou GI, Boulalas I, Delakas D, Spandidos DA. Identification of common differentially expressed genes in urinary bladder Cancer. PLoS One. 2011;6(4):e18135. DOI: 10.1371/journal.pone.0018135
  93. 93. Kim JM, Sohn HY, Yoon SY, Oh JH, Yang JO, Kim JH, et al. Identification of gastric cancer-related genes using a cDNA microarray containing novel expressed sequence tags expressed in gastric cancer cells. Clinical Cancer Research. 2005;11(2):473-482
  94. 94. Zur A, Brandeis M. Securin degradation is mediated by fzy and fzr, and is required for complete chromatid separation but not for cytokinesis. EMBO Journal. 2001;20(4):792-801
  95. 95. Geley S, Kramer E, Gieffers C, Gannon J, Peters JM, Hunt T. Anaphase-promoting complex/cyclosome-dependent proteolysis of human cyclin a starts at the beginning of mitosis and is not subject to the spindle assembly checkpoint. Journal of Cell Biology. 2001;153(1):137-147
  96. 96. Ohtoshi A, Maeda T, Higashi H, Ashizawa S, Hatakeyama M. Human p55(CDC)/Cdc20 associates with cyclin a and is phosphorylated by the cyclin A-Cdk2 complex. Biochemical and Biophysical Research Communications. 2000;268(2):530-534
  97. 97. Lim HH, Goh PY, Surana U. Cdc20 is essential for the cyclosome-mediated proteolysis of both Pds1 and Clb2 during M phase in budding yeast. Current Biology. 1998;8(4):231-234
  98. 98. Shirayama M, Toth A, Galova M, Nasmyth K. APC(Cdc20) promotes exit from mitosis by destroying the anaphase inhibitor Pds1 and cyclin Clb5. Nature. 1999;402(6758):203-207
  99. 99. Hames RS, Wattam SL, Yamano H, Bacchieri R, Fry AM. APC/C-mediated destruction of the centrosomal kinase Nek2A occurs in early mitosis and depends upon a cyclin A-type D-box. EMBO Journal. 2001;20(24):7117-7127
  100. 100. Harley ME, Allan LA, Sanderson HS, Clarke PR. Phosphorylation of Mcl-1 by CDK1-cyclin B1 initiates its Cdc20-dependent destruction during mitotic arrest. EMBO Journal. 2010;29(14):2407-2420
  101. 101. Amador V, Ge S, Santamaria PG, Guardavaccaro D, Pagano M. APC/C-Cdc20 controls the ubiquitin-mediated degradation of p21 in prometaphase. Molecular Cell. 2007;27(3):462-473
  102. 102. Sudo T, Ota Y, Kotani S, Nakao M, Takami Y, Takeda S, et al. Activation of Cdh1-dependent APC is required for G(1) cell cycle arrest and DNA damage-induced G(2) checkpoint in vertebrate cells. EMBO Journal. 2001;20(22):6499-6508
  103. 103. Grim JE, Knoblaugh SE, Guthrie KA, Hagar A, Swanger J, Hespelt J, et al. Fbw7 and p53 cooperatively suppress advanced and chromosomally unstable intestinal cancer. Molecular and Cellular Biology. 2012;32(11):2160-2167
  104. 104. Carrano AC, Eytan E, Hershko A, Pagano M. SKP2 is required for ubiquitin-mediated degradation of the CDK inhibitor p27. Nature Cell Biology. 1999;1(4):193-199
  105. 105. Carrano AC, Pagano M. Role of the F-box protein Skp2 in adhesion-dependent cell cycle progression. Journal of Cell Biology. 2001;153(7):1381-1389
  106. 106. Latres E, Chiarle R, Schulman BA, Pavletich NP, Pellicer A, Inghirami C, et al. Role of the F-box protein Skp2 in lymphomagenesis. Proceedings of the National Academy of Sciences of the United States of America. 2001;98(5):2515-2520
  107. 107. Loda M, Cukor B, Tam SW, Lavin P, Fiorentino M, Draetta GF, et al. Increased proteasome-dependent degradation of the cyclin-dependent kinase inhibitor p27 in aggressive colorectal carcinomas. Nature Medicine. 1997;3(2):231-234
  108. 108. Shim EH, Johnson L, Noh HL, Kim YJ, Sun H, Zeiss C, et al. Expression of the F-box protein SKP2 induces hyperplasia, dysplasia, and low-grade carcinoma in the mouse prostate. Cancer Research. 2003;63(7):1583-1588
  109. 109. Delogu S, Wang CM, Cigliano A, Utpatel K, Sini M, Longerich T, et al. SKP2 cooperates with N-Ras or Akt to induce liver tumor development in mice. Oncotarget. 2015;6(4):2222-2234
  110. 110. Lin HK, Chen ZB, Wang GC, Nardella C, Lee SW, Chan CH, et al. Skp2 targeting suppresses tumorigenesis by Arf-p53-independent cellular senescence. Nature. 2010;464(7287):374-U66
  111. 111. Zhao HL, Bauzon F, Fu H, Lu ZL, Cui JH, Nakayama K, et al. Skp2 deletion unmasks a p27 safeguard that blocks tumorigenesis in the absence of pRb and p53 tumor suppressors. Cancer Cell. 2013;24(5):645-659
  112. 112. Vodermaier HC. APC/C and SCF: Controlling each other and the cell cycle. Current Biology. 2004;14(18):R787-R796
  113. 113. Cappell SD, Mark KG, Garbett D, Pack LR, Rape M, Meyer T. EMI1 switches from being a substrate to an inhibitor of APC/C-CDH1 to start the cell cycle. Nature. 2018;558(7709):313-317
  114. 114. Nakayama KI, Nakayama K. Ubiquitin ligases: Cell-cycle control and cancer. Nature Reviews Cancer. 2006;6(5):369-381
  115. 115. Inuzuka H, Tseng A, Gao DM, Zhai B, Zhang Q, Shaik S, et al. Phosphorylation by casein kinase I promotes the turnover of the Mdm2 oncoprotein via the SCF beta-TRCP ubiquitin ligase. Cancer Cell. 2010;18(2):147-159
  116. 116. Beroukhim R, Mermel CH, Porter D, Wei G, Raychaudhuri S, Donovan J, et al. The landscape of somatic copy-number alteration across human cancers. Nature. 2010;463(7283):899-905
  117. 117. Gong YX, Zack TI, Morris LGT, Lin K, Hukkelhoven E, Raheja R, et al. Pan-cancer genetic analysis identifies PARK2 as a master regulator of G1/S cyclins. Nature Genetics. 2014;46(6):588-594
  118. 118. Veeriah S, Taylor BS, Meng S, Fang F, Yilmaz E, Vivanco I, et al. Somatic mutations of the Parkinson’s disease-associated gene PARK2 in glioblastoma and other human malignancies. Nature Genetics. 2010;42(1):77-U98
  119. 119. Lee SB, Kim JJ, Nam HJ, Gao BW, Yin P, Qin B, et al. Parkin regulates mitosis and genomic stability through Cdc20/Cdh1. Molecular Cell. 2015;60(1):21-34
  120. 120. Poulogiannis G, McIntyre RE, Dimitriadi M, Apps JR, Wilson CH, Ichimura K, et al. PARK2 deletions occur frequently in sporadic colorectal cancer and accelerate adenoma development in Apc mutant mice. Proceedings of the National Academy of Sciences of the United States of America. 2010;107(34):15145-15150
  121. 121. Yeo CWS, Ng FSL, Chai C, Tan JMM, Koh GRH, Chong YK, et al. Parkin pathway activation mitigates glioma cell proliferation and predicts patient survival. Cancer Research. 2012;72(10):2543-2553

Written By

Pai-Sheng Chen

Submitted: June 11th, 2018 Reviewed: January 3rd, 2019 Published: February 19th, 2019