Review | Published:

Regulation of apoptosis proteins in cancer cells by ubiquitin

Abstract

Ubiquitin inhibitors act at many levels to enhance apoptosis signaling. For TNF-related apoptosis-inducing ligand (TRAIL)-mediated apoptosis signaling, there are at least five mechanisms by which apoptosis are regulated by the ubiquitin–proteasome pathway. First, proteasome inhibitors can decrease Fas-like inhibitor protein (FLIP) protein levels in tumors, resulting in increased apoptosis signaling due to increased caspase-8 activation. This appears to involve the ubiquitin ligase TNF receptor activation factor-2 (TRAF2) and acts indirectly by causing cell-cycle arrest at a stage where there is high degradation of the FLIP–TRAF2 complex. Second, the regulation of the proapoptotic Bcl-2 family member BAX occurs indirectly. Apoptosis signaling and caspase activation results in a confirmation change in the normally monomeric BAX, which exposes the BH3 domain of BAX, leading to dimerization and resistance to ubiquitin degradation. BAX then translocates into the mitochondria, resulting in the release of proapoptotic mitochondrial factors such as cytochrome c and second mitochondria-derived activator of caspase (SMAC). This results in the activation of caspase-9 and formation of the apoptosome and efficient apoptosis signaling. A third mechanism of the regulation of TRAIL signaling in the ubiquitin–proteasome pathway is mediated by the inhibitor of apoptosis proteins (IAP) E3 ligases. These IAPs can directly bind to caspases but also can act as ubiquitin ligases for caspases, resulting in the degradation of these caspases. IAP binding to caspases can be inhibited by SMAC, which exhibits a caspase-9 homology domain. The fourth mechanism for apoptosis activation by proteasome inhibitors is through the stabilization of the inhibitor of the κB (IκB)/NF-κB complex and prevention of nuclear translocation of the antiapoptosis transcription factor NF-κB. During TRAIL-DR4, DR5 signaling, this pathway is activated by interactions of activated Fas-associated death domain with activated receptor-interacting protein (RIP), which in turn activates NF-κB-inducing kinase and phosphorylates IκB. Therefore, the inhibition of IκB degradation blocks this RIP-mediated antiapoptosis signaling event. Last, p53 protein levels, and susceptibility to apoptosis, can be deregulated by the human homolog Hdm2 (Mdm2) E3 ligase. This process is inhibited by p53 phosphorylation and by sequestration of Mdm2 by ARF. Better mechanisms to inhibit the ubiquitin–proteasome pathway targeted at the ubiquitin–proteasome degradation process itself, or more specifically at the E3 ligases known to modulate and downregulate proapoptosis pathways will lead to the enhancement of TRAIL apoptosis signaling and better cancer therapeutic outcomes act through this pathway.

Introduction

Protein stability is a key regulatory mechanism in the control of cell development, cell cycle, cell growth and apoptosis. The selective degradation or stabilization of intracellular proteins by ubiquitin-dependent pathways is essential for correcting the regulation of many cellular processes. Ubiquitin is covalently attached to substrate proteins by a protein complex usually including an activating enzyme (E1), a conjugating enzyme (E2) and a protein ligase (E3) (Orian et al., 1995; Sudakin et al., 1995; Haas and Siepmann, 1997). The addition of four or more ubiquitin moieties targets the substrates for destruction via the 26S proteasome, and free ubiquitin is recycled. The addition of fewer ubiquitin molecules can alter the substrate protein function or target the substrate protein to the endosome. This process can be opposed by the action of deubiquitinating enzymes, which remove ubiquitin from specific substrates, thus stabilizing them. This tightly regulated complex pathway is the key regulator of many important signaling pathways and plays an important role in many cellular processes including apoptosis. Many apoptosis regulatory proteins have been identified as target substrates for ubiquitination (Argentini et al., 2000; Buschmann et al., 2000; Huang et al., 2000; Suzuki et al., 2001; Wilson et al., 2002). In addition to being targets for ubiquitination, some apoptosis regulatory proteins exhibit ubiquitin ligase activity that can be ascribed to the RING finger domain that is part of their primary structure.

Components of the cell apoptosis machinery are frequently altered in cancer. Resistance to apoptosis is one of the major hurdles in the treatment of cancer. In this review, we describe how the ubiquitin–proteasome pathway can inhibit apoptosis by degrading proapoptotic proteins, and we highlight the potential of proteasome inhibitors as antitumor therapies that enhance apoptosis by blocking these pathways.

Ubiquitination, apoptosis and cancer

The inhibition of apoptosis is a hallmark of cancer and autoimmune disease. Observations in the early 1990s began to reveal a link between ubiquitination and apoptosis or programmed cell death (Haas et al., 1995). Studies using the hawk moth (Manduca sexta) indicated that ubiquitin mRNA and ubiquitinated proteins were dramatically increased in the intersegmental muscle cells undergoing programmed cell death. In these dying cells, there were increased ubiquitinating enzyme activities as well as an increase in proteasome components and their proteolytic activities (Haas et al., 1995). Over the past few years, apoptosis-pathway molecules have been identified as substrates and targets for proteasome degradation leading to apoptosis resistance. Apoptosis molecules that are regulated by the ubiquitin–proteasome pathway include members of the Bcl-2 family (Chang et al., 1998; Breitschopf et al., 2000; Li and Dou, 2000), IAP family (Yang and Li, 2000; Yang et al., 2000; Suzuki et al., 2001; MacFarlane et al., 2002; Martin, 2002) and inhibitor of the κB (IκB) (Orian et al., 1995; Jesenberger and Jentsch, 2002; Kovalenko et al., 2003). Alteration of the stability of these proteins through the ubiquitin—proteasome-regulated pathway generally contributes to apoptosis resistance in cancer cells.

Regulation of TNF-related apoptosis-inducing ligand (TRAIL) apoptosis signaling by ubiquitination pathway

TRAIL/APO-2L has attracted attention for its ability to kill tumor cells preferentially, while most normal cells were resistant both in vitro and in vivo. TRAIL is a ligand for death domain (DR)-4 and 5, which are members of the tumor necrosis factor receptor superfamily (TNFRsF) that includes TNFR, CD95/APO-1 and TRAMP (Bodmer et al., 1997; Zhou et al., 2002). The functional analysis of the TRAIL receptor–ligand system has been complicated by the fact that a total of five different receptors for this cytokine have been identified. A very recent study by Johnson et al. (2003) shows that PS-341 treatment increases the expression of TRAIL receptors DR4 and DR5, and this increase in receptor protein levels is associated with the ubiquitination of the DR5 protein.

The ubiquitin–proteasome pathway has been proposed to regulate TRAIL apoptosis signaling by effecting levels of death domain adaptor molecules including Fas-associated death domain (FADD) and Fas-like inhibitor protein (FLIP). Crosslinking of the two apoptosis-inducing TRAIL receptors (TRAIL-R1 and TRAIL-R2) results in the recruitment of FADD and caspase-8 to the DISC (Srivastava, 2001; Baetu and Hiscott, 2002; Kiechle and Zhang, 2002; Rossi and Gaidano, 2003). The death effector domain of FADD interacts with the death effector domain of procaspase-8 and thereby recruits this proenzyme to the DISC. Procaspase-8 is activated at the DISC by proteolytical cleavage. Activated caspase-8 then initiates the apoptosis-executing caspase cascade. The activation of caspase-8 is regulated by FLIP that inhibits caspase-8 binding to FADD (Figure 1).

Figure 1
figure1

Ubiquitination of FLIP regulates TRAIL-induced apoptosis. The trimeric TRAIL results in the recruitment of death domain receptor 4 or death domain receptor 5 (DR4, DR5) molecules. The TRAIL-induced conformational changes of the DR4/DR5 death domain result in the recruitment of FADD and procaspase-8, which then signals apoptosis. FLIP contains a death domain that can compete with caspase-8 cytoplasmic binding domain of DR4, DR5. The intracellular levels of FLIP are regulated by the proteasome degradation pathway. One proposed pathway is binding of FLIP to TRAF2. TRAF2 contains a RING zinc-finger domain and possesses E3 ligase activity. FLIP degradation can be enhanced with the proteasome inhibitor PS-341 or by incubation with ligands for the PPAR-γ. It has been proposed that these can modulate cell cycle and result in increased TRAIL-mediated apoptosis

Ubiquitination of FLIP is associated with death signaling pathway

PS-341, a proteasome inhibitor, can induce a substantial reduction in c-FLIP, and has been successfully combined with the death ligand TRAIL to promote tumor cells' apoptosis synergistically (Sayers et al., 2003). Apoptosis sensitization is mediated by the effects of ubiquitin on an unidentified cellular target, resulting in post-transcriptional reduction in FLIP protein levels (Fukazawa et al., 2001). This synergistic effect is correlated with reductions in the antiapoptotic protein c-FLIP. Thus, the decrease in c-FLIP, in the presence of a proteasome inhibitor, PS-341, is surprising since c-FLIP is reported to be degraded by the proteasome in some cells (Sayers et al., 2003). One explanation is that the concentrations of PS-341 that sensitize cells to TRAIL degradation also inhibits cell-cycle progression, and this cell-cycle inhibition results in an accumulation of cells in S-G2/M phase. Perturbation of cell cycle has been reported to modulate the level of c-FLIP with peak levels occurring during G1 followed by a reduction in S-G2/M. Thus, ubiquitin may indirectly affect c-FLIP levels by modulating cell cycle. FLIP has also been shown to interact with TNF receptor activation factor-2 (TRAF2), which contains a RING finger domain known to possess E3 ligase activity (Park et al., 2001). Whether TRAF2 is responsible for FLIP degradation is unknown. Another clue to the regulation of TRAIL apoptosis signaling has been reported to be mediated by a variety of natural and synthetic ligands of peroxisome proliferator-activated receptor-gamma (PPARγ), which can sensitize human prostate and ovarian cancer cells to TRAIL-mediated apoptosis (Kim et al., 2002). The interaction of the ubiquitin–proteasome pathway with FLIP, TRAF and PPARγ in the regulation of TRAIL apoptosis signaling will be an important subject for future investigation.

Bcl-2 modulation of TRAIL apoptosis signaling

TRAIL apoptosis signaling has been reported to be modulated by Bcl-2 family members. TRAIL proapoptotic signaling can be blocked by the overexpression of Bcl-2 or Bcl-XL in many cancer cells. Other Bcl-2 family members, including Bax and Bid, have been reported to play a critical role in TRAIL-mediated apoptosis of cancer cells, and they are also regulated by the ubiquitin–proteasome pathway (He et al., 1998; Martinou et al., 1998; Basu and Haldar, 2002; Brichese et al., 2002). In a nonapoptotic cell, Bax is a monomeric protein exhibiting a diffuse cytosolic distribution. The three-dimensional structure of monomeric Bax predicts that the hydrophobic C terminus will interact with and mask the BH3 domain, which will inhibit Bax–Bax dimer formation and mitochondrial membrane targeting (Suzuki et al., 2000). In response to apoptotic stimuli, Bax undergoes a conformational change to expose not only domains vital for its apoptosis function but also reveals previously masked epitopes, thus allowing the recognition of the active form by conformation-specific antibodies (Figure 2). Proteasome inhibitor-induced apoptosis of B-CLL cells involves caspase-independent conformational change of Bax and its translocation to mitochondria, which correlates with mitochondria membrane depolarization and the release of cytochrome c (Dewson et al., 2003). A similar mechanism may exist for other proapoptotic family members. In Bax-negative TRAIL-resistant HC-4 colon cancer cells, the combination of PS-341 and TRAIL signaling overcomes the block in the activation of the mitochondrial pathway and causes second mitochondria-derived activator of caspase (SMAC, also known as DIABLO) and cytochrome c release followed by apoptosis (Johnson et al., 2003). Furthermore, fibroblasts lacking Bak are significantly more resistant to undergo apoptosis when exposed to the combination of PS-341 and TRAIL (Johnson et al., 2003). Taken together, these findings indicate that PS-341 enhances TRAIL-induced apoptosis by increasing the cleavage of caspase-8, and by causing Bak-dependent release of mitochondrial proapoptotic proteins.

Figure 2
figure2

Apoptosis induces a conformational change in BAX that inhibits proteasome degradation. In nonapoptotic cells, BAX dimerization is inhibited because the hydrophobic C terminus of BAX masks the BH3 domain. This configuration is also more susceptible to proteasome degradation. After apoptosis signaling, a caspase-dependent conformational change results in resistance to proteasome degradation and translocation of BAX to the mitochondria, which correlates the release of cytochrome c that amplifies the apoptosis signaling events

Ubiquitination of IAP proteins and TRAIL signaling pathway

Inhibitor of apoptosis proteins (IAPs) are a family of proteins defined by baculovirus repeat (BIR) domains and, in some cases, a RING zinc-finger domain (Takahashi et al., 1998). Among the eight mammalian (IAPs), four of these proteins including cIAP1, cIAP2, X-linked IAP (XIAP) and neuronal apoptosis inhibitory protein (NAIP) have three BIR domains (Liston et al., 1996). Other IAPs including ubiquitin conjugation domain (BRUCE), survivin, Drosophila BIR and IAP-like protein-1 (ILP-1) only have one BIR domain (Wang et al., 2003). XIAP, survivin, cIAP1 and cIAP2 block apoptosis by directly inhibiting caspases. XIAP is one of the most potent inhibitors of apoptosis through its ability to inhibit caspase-3, -7 and -9. The BIR3 domain of XIAP directly binds to the small subunit of caspase-9, whereas the linker region between the BIR1 and BIR2 domains binds with caspase-3 or -7, and prevents their active sites from binding with substrates (Srinivasula et al., 2001).

In addition to direct inhibition of caspase activation, the majority of the caspase-inhibiting IAPs possess a carboxyl-terminal RING zinc-finger motif and exhibit E3 ligase activity (Liston et al., 1996). It was found that these IAPs have RING zinc-finger-dependent E3 activities that catalyse autoubiquitination in vitro and in cells. The overexpression of cIAP1 caused its autoubiquitination and degradation (Yang et al., 2000). cIAP2 can promote monoubiquitination of caspase-3 and -7, and that XIAP catalyses the ubiquitination and degradation of caspase-3 (Suzuki et al., 2001; MacFarlane et al., 2002). This is supported by the observation that IAPs catalysed their own ubiquitination in vitro, and this activity requires the RING domain. Overexpressed wild-type c-IAP1, but not a RING domain mutant c-IAP1, was spontaneously ubiquitinated and degraded, and stably expressed XIAP lacking the RING domain was relatively resistant to apoptosis-induced degradation and, correspondingly, more effective at preventing apoptosis than wild-type XIAP (Yang et al., 2000).

It has been shown that XIAP is able to promote the ubiquitination and degradation of SMAC. SMAC is an apoptosis regulator that is synthesized as a 239 amino-acid precursor protein. The first 55 amino acids of the molecules are required for mitochondria transport, after which this portion of the molecule is cleaved (Du et al., 2000; Srinivasula et al., 2000). The amino-terminal of SMAC has a motif similar to that of the small subunit of caspase-9, which is responsible for binding of caspase-9 to the groove on the surface of the BIR3 domain of XIAP (Liu et al., 2000; Srinivasula et al., 2001). This four amino-acid motif is similar to the RHG motif of Drosophila proteins Reaper, Hid and Grim, raising the possibility that SMAC may be able to promote the ubiquitination and degradation of XIAP (Silke et al., 2000; Wu et al., 2001; Holley et al., 2002; Olson et al., 2003).

XIAP levels are reduced after exposure to TRAIL because TRAIL induces the release of SMAC/DIABLO from mitochondria, which binds to and inactivates XIAP. Leverkus et al. (2003) showed that proteasome inhibitors enhance TRAIL-mediated apoptosis of primary keratinocytes by enhancing the release of proapoptotic molecules such as SMAC/DIABLO and cytochrome c from mitochondria (Figure 3). It was found that TRAIL could not induce apoptosis effectively in Bax-deficient cells, apparently due to the formation of p20/p12 caspase-3 but not the fully active p17/p12 dimer. This incompletely processed p20/p12 caspase-3 was associated with XIAP due to the lack of the release of SMAC from mitochondria in the absence of Bax (Leverkus et al., 2003). When SMAC was expressed in the cytoplasm of these Bax−/− cells, TRAIL could induce the generation of p17/p12 caspase-3 as well as apoptosis, indicating that the interaction of SMAC with XIAP is required for full activation of caspase and a cell death program.

Figure 3
figure3

Proteasome-regulated stability of IAPs. The IAP consists of a family including cIAPs, XIAP, as well as NAIP, ILP, and, in Drosophilae, BIR- and ubiquitin-conjugated domain (DRUCE). These IAPs can directly inhibit caspases by blocking their active sites. In addition, many of the IAPs have RING zinc-finger domains that can act as E3 ligases, resulting in the proteasome degradation of caspases. During apoptosis, mitochondrial release of SMAC, which has an amino-terminal motif that is similar to the small subunit of caspase-9, binds to the XIAP BIR3 domain. It has been proposed that binding of SMAC to XIAP prevents binding to caspase-9 but promotes autophosphorylation of XIAP and degradation of XIAP, thereby promoting caspase activation and apoptosis

Alternatively, XIAP might exert its caspase-inhibitory role by its ubiquitin–protein ligase activity (Yang and Li, 2000; Yang et al., 2000; Suzuki et al., 2001; MacFarlane et al., 2002). Blocking the degradation of XIAP-targeted proapoptosis molecules such as caspase-3 might also be responsible for the enhancement of TRAIL-mediated apoptosis. Inhibition may thereby lead to increased levels of active caspase-3 and subsequent apoptotic cell death in cells expressing high levels of XIAP. Another possibility of the action of the proteasomal inhibitor may be decreased XIAP-mediated degradation of SMAC/DIABLO as they are key molecules that regulate mitochondria-mediated apoptosis. Future studies are required to delineate these points.

Transcription factor-mediated and ubiquitin-regulated apoptosis of TRAIL signaling pathway

The NF-κB family (RelA[p65], cRel, RelB, p50 and p52) form heterodimers that act as transcription factors and upregulate antiapoptosis-associated genes. In quiescent cells, NF-κB dimers are bound to members of the IκB family and are sequestered in the cytoplasm. Malignant transformation, viral infection or cytokine signaling have been associated with the phosphorylation of IκB at specific serine sites by the IκB kinase (IKK) complex and degradation via the ubiquitin–proteasome pathway. NF-κB can then translocate to the nucleus and activate transcription of various genes including antiapoptotic genes (Brummerlkamp et al., 2003; Trompouki et al., 2003). TRAIL-mediated apoptosis signaling may be abrogated by the concomitant activation of NF-κB and upregulation of antiapoptosis genes. This activation involves the recruitment of the receptor-interacting protein (RIP) through FADD or TRADD. Upon localization to the death receptor complex, RIP activates NF-κB-inducing kinase (NIK), which in turn phosphorylates and activates IKK. This leads to phosphorylation and subsequent degradation of IκB and translocation of NF-κB to the nucleus. The antiapoptosis or proapoptosis signaling of TRAIL through NF-κB is currently of high interest, since TRAIL-mediated activation of NF-κB has also been shown to have either no effect for survival or apoptotic responses following TRAIL stimulation (Leverkus et al., 2003), or exhibit proapoptotic effects caused by the upregulation of TRAIL receptor-DR5 (Rivera-Walsh et al., 2001).

p53 is a transcription factor that is regulated by ubiquitin pathway

Wild-type p53 is expressed at low levels in most cells because of its short half-life under normal conditions. DNA damage, hypoxia and inappropriate oncogene signaling lead to stabilization and an increase in the level of p53. In its active form, p53 can bind DNA in a sequence-specific manner and activate transcription of target genes. p53 levels are regulated in large part by human homolog Hdm2 (Mdm2), the product of a p53-inducible gene. Mdm2 can interact with the N terminus of p53, which also contains the major acidic transcriptional activation of p53 that is downregulated by its target gene product Mdm2 (Gottifredi and Prives, 2001). Mdm2 forms an autoregulatory loop with p53 by binding to its N-terminal domain, inhibiting its transcriptional activity and increasing its degradation by the ubiquitin–proteasome pathway (Figure 4). Mdm2 is a RING finger-dependent ubiquitin protein ligase for p53 and itself. Mdm2 also inhibits p53 by nuclear export through a mechanism involving either the nuclear export signal (NES) of Mdm2 or the RING finger of Mdm2 and the NES of p53. The NES of p53 is masked in the transcriptionally active heterodimer, but is exposed in the monomeric form of p53. MdmX, an Mdm2 homolog that lacks an NES, stabilizes p53 by promoting its retention in the nucleus.

Figure 4
figure4

TRAIL antiapoptosis signaling through FADD, RIP and NF-κB. Signaling of DR4 and DR5 results in the recruitment of RIP to an FADD. RIP can then activate NIK, which phosphorylates IKK leading to proteasome degradation of phosphorylated IκB as well as nuclear translocation and antiapoptosis signaling of NF-κB

There is a high frequency of p14 (ARF) gene deletion in human cancers, accounting for some 40% of cancers overall, which suggests that ARF would be a strong candidate for therapeutic application. ARF can block nuclear cytoplasmic shuttling of Mdm2 leading to increased nuclear p53 levels. Mdm2 can also bind to and attenuate Rb function, and ARF can therefore lead to increased Rb activity and growth suppression. However, the possible dependence of ARF activity on p53 and Rb function presents a potential limitation to its application, as these functions are often impaired in cancer. Under certain circumstances, such as oncogene activation, the expression of an Mdm2 binding protein, ARF, is increased, which directly inhibits Mdm2's E3 activity and therefore increases intracellular p53 level (Bothner et al., 2001).

Much less is known of how the physiological stress, hypoxia, effects Mdm2 and p53. Under hypoxic conditions, p53 is stabilized by mitochondria through a redox-dependent mechanism and by hypoxia-inducible factor (HIF)-1α, whereas p53 induces the degradation of HIF-1α. The response to hypoxia in vivo also involves the glucocorticoid receptor (GR). Recently, evidence has been growing for crosstalk between the p53- and GR-mediated responses to stress. p53 and GR mutually inhibit each others activity by cytoplasmic sequestration in a ligand-dependent manner, which leads to increased degradation through the proteasome pathway by the recruitment of the E3 ubiquitin ligase Mdm2 (Sengupta et al., 2000).

Deubiquitination enzymes including the herpesvirus-associated ubiquitin-specific protease (HAUSP) can specifically remove ubiquitin from p53 and stabilize it, even in the presence of Mdm2. The overexpression of HAUSP suppressed the growth of H460 human lung carcinoma cells in a p53-dependent manner, presumably through p53-induced growth arrest and/or apoptosis (Li et al., 2002). It is not yet known whether or how HAUSP is regulated by p53-inducing stimuli such as DNA damage and oncogene activation. On the other hand, increased p53 upregulates Mdm2 gene transcription, forming a negative feedback to prevent the unwanted high level of p53 in the cells. The importance of regulating p53 level by Mdm2 was illustrated by the observation that the early embryonic lethality of Mdm2-deficient mice was completely rescued by the simultaneous deletion of p53 (de Rozieres et al., 2000). Surprisingly, the deletion of the Mdm2 homologue MdmX also led to embryonic death, which could be rescued by loss of p53 (Gu et al., 2002). Although MdmX has a RING zinc-finger and can bind to p53, it does not possess E3 activity toward p53. Recent studies indicated that MdmX formed a heterodimer with Mdm2 that led to its stabilization. This is likely because Mdm2 promotes its own ubiquitination and degradation in the absence of MdmX. However, overexpressed MdmX might compete with Mdm2 in binding with p53 and therefore inhibit Mdm2-mediated p53 ubiquitination and degradation. Like mutations of p53 itself, abnormalities in these regulating mechanisms such as enhanced p53 degradation due to the expression of E6 after infection of oncogenic HPV, loss of ARF and amplification of Mdm2 or MdmX gene have been demonstrated to play critical roles in the development of certain tumors (Wesierska-Gadek et al., 2002). It is conceivable that dysfunction of p53 pathway may contribute to the development of most, if not all, human cancer.

TRAIL death receptor DR5 is induced in cells undergoing p53-dependent apoptosis, and therefore the upregulation of DR5 receptor by p53 might have synergistic effects on the induction of apoptosis of cancer cells. The ubiquitin-regulated p53 pathway might indirectly affect the TRAIL-mediated apoptosis of cancer cells. We recently found that prevention of p53 degradation leads to the enhancement of TRAIL-mediated apoptosis, and that the release of p53 sequestered in the cytosol by vimentin during TRAIL signaling and subsequent nuclear translocation is one of the mechanisms underlying the TRAIL-mediated apoptosis (Figure 5) (Yang et al., unpublished observation). P53 can also affect TRAIL signaling by promoting the downmodulation of FLIP. The decrease of FLIP by p53 could be prevented by a proteasome inhibitor, suggesting that p53 enhances the degradation of FLIP via a ubiquitin–proteasome pathway (Fukazawa et al., 2001).

Figure 5
figure5

Regulation of p53 by Mdm2 ubiquitin–proteasome degradation. Mdm2 binds with the transactivating (TA) domain of p53, which inhibits its transcriptional activity. Mdm2 is a RING zinc-finger-dependent ubiquitin protein ligase for p53, which results in the degradation of p53. Other Mdm2-like proteins, such as MdmX, which lacks the NES of Mdm2, stabilizes p53 retention in the nucleus. Mdm2 activity on p53 can be inhibited by two ways: (1) the phosphorylation of the transactivating domain of p53 inhibits binding to Mdm2 and promotes its stability, and (2) The p14 ARF can bind to Mdm2 and inhibits its ability to act as an E3 ligase on p53, thereby stabilizing p53

References

  1. Argentini M, Barboule N and Wasylyk B . (2000). Oncogene, 19, 3849–3857.

  2. Baetu TM and Hiscott J . (2002). Cytokine Growth Factor Rev., 13, 199–207.

  3. Basu A and Haldar S . (2002). Int. J. Oncol., 21, 597–601.

  4. Bodmer JL, Burns K, Schneider P, Hofmann K, Steiner V, Thome M, Bornand T, Hahne M, Schroter M, Becker K, Wilson A, French LE, Browning JL, MacDonald HR and Tschopp J . (1997). Immunity, 6, 79–88.

  5. Bothner B, Lewis WS, DiGiammarino EL, Weber JD, Bothner SJ and Kriwacki RW . (2001). J. Mol. Biol., 314, 263–277.

  6. Breitschopf K, Zeiher AM and Dimmeler S . (2000). J. Biol. Chem., 275, 21648–21652.

  7. Brichese L, Barboule N, Heliez C and Valette A . (2002). Exp. Cell Res., 278, 101–111.

  8. Brummerlkamp T, Nijman SB, Dirac AG and Bernards R . (2003). Nature, 424, 797–801.

  9. Buschmann T, Fuchs SY, Lee CG, Pan ZQ and Ronai Z . (2000). Cell, 101, 753–762.

  10. Chang YC, Lee YS, Tejima T, Tanaka K, Omura S, Heintz NH, Mitsui Y and Magae J . (1998). Cell Growth Differ., 9, 79–84.

  11. de Rozieres S, Maya R, Oren M and Lozano G . (2000). Oncogene, 19, 1691–1697.

  12. Dewson G, Snowden RT, Almond JB, Dyer MJ and Cohen GM . (2003). Oncogene, 22, 2643–2654.

  13. Du C, Fang M, Li Y, Li L and Wang X . (2000). Cell, 102, 33–42.

  14. Fukazawa T, Fujiwara T, Uno F, Teraishi F, Kadowaki Y, Itoshima T, Takata Y, Kagawa S, Roth JA, Tschopp J and Tanaka N . (2001). Oncogene, 20, 5225–5231.

  15. Gottifredi V and Prives C . (2001). Science, 292, 1851–1852.

  16. Gu J, Kawai H, Nie L, Kitao H, Wiederschain D, Jochemsen AG, Parant J, Lozano G and Yuan ZM . (2002). J. Biol. Chem., 277, 19251–19254.

  17. Haas AL, Baboshina O, Williams B and Schwartz LM . (1995). J. Biol. Chem., 270, 9407–9412.

  18. Haas AL and Siepmann TJ . (1997). FASEB J., 11, 1257–1268.

  19. He H, Qi XM, Grossmann J and Distelhorst CW . (1998). J. Biol. Chem., 273, 25015–25019.

  20. Holley CL, Olson MR, Colon-Ramos DA and Kornbluth S . (2002). Nat. Cell Biol., 4, 439–444.

  21. Huang H, Joazeiro CA, Bonfoco E, Kamada S, Leverson JD and Hunter T . (2000). J. Biol. Chem., 275, 26661–26664.

  22. Jesenberger V and Jentsch S . (2002). Nat. Rev. Mol. Cell. Biol., 3, 112–121.

  23. Johnson TR, Stone K, Nikrad M, Yeh T, Zong WX, Thompson CB, Nesterov A and Kraft AS . (2003). Oncogene, 22, 4953–4963.

  24. Kiechle FL and Zhang X . (2002). Clin. Chim. Acta, 326, 27–45.

  25. Kim Y, Suh N, Sporn M and Reed JC . (2002). J. Biol. Chem., 277, 22320–22329.

  26. Kovalenko A, Chable-Bessia C, Cantarella G, Israel A, Wallach D and Courtois G . (2003). Nature, 424, 801–805.

  27. Leverkus M, Sprick MR, Wachter T, Mengling T, Baumann B, Serfling E, Brocker EB, Goebeler M, Neumann M and Walczak H . (2003). Mol. Cell. Biol., 23, 777–790.

  28. Li B and Dou QP . (2000). Proc. Natl. Acad. Sci. USA, 97, 3850–3855.

  29. Li M, Chen D, Shiloh A, Luo J, Nikolaev AY, Qin J and Gu W . (2002). Nature, 416, 648–653.

  30. Liston P, Roy N, Tamai K, Lefebvre C, Baird S, Cherton-Horvat G, Farahani R, McLean M, Ikeda JE, MacKenzie A and Korneluk RG . (1996). Nature, 379, 349–353.

  31. Liu Z, Sun C, Olejniczak ET, Meadows RP, Betz SF, Oost T, Herrmann J, Wu JC and Fesik SW . (2000). Nature, 408, 1004–1008.

  32. MacFarlane M, Merrison W, Bratton SB and Cohen GM . (2002). J. Biol. Chem., 277, 36611–36616.

  33. Martin SJ . (2002). Cell, 109, 793–796.

  34. Martinou I, Missotten M, Fernandez PA, Sadoul R and Martinou JC . (1998). Neuroreport, 9, 15–19.

  35. Olson MR, Holley CL, Yoo SJ, Huh JR, Hay BA and Kornbluth S . (2003). J. Biol. Chem., 278, 4028–4034.

  36. Orian A, Whiteside S, Israel A, Stancovski I, Schwartz AL and Ciechanover A . (1995). J. Biol. Chem., 270, 21707–21714.

  37. Park SJ, Kim YY, Ju JW, Han BG, Park SI and Park BJ . (2001). Biochem. Biophys. Res. Commun., 289, 1205–1210.

  38. Rivera-Walsh I, Waterfield M, Xiao G, Fong A and Sun SC . (2001). J. Biol. Chem., 276, 40385–40388.

  39. Rossi D and Gaidano G . (2003). Haematologica, 88, 212–218.

  40. Sayers TJ, Brooks AD, Koh CY, Ma W, Seki N, Raziuddin A, Blazar BR, Zhang X, Elliott PJ and Murphy WJ . (2003). Blood, 102, 303–310.

  41. Sengupta S, Vonesch JL, Waltzinger C, Zheng H and Wasylyk B . (2000). EMBO J., 19, 6051–6064.

  42. Silke J, Verhagen AM, Ekert PG and Vaux DL . (2000). Cell. Death Differ., 7, 1275.

  43. Srinivasula SM, Datta P, Fan XJ, Fernandes-Alnemri T, Huang Z and Alnemri ES . (2000). J. Biol. Chem., 275, 36152–36157.

  44. Srinivasula SM, Hegde R, Saleh A, Datta P, Shiozaki E, Chai J, Lee RA, Robbins PD, Fernandes-Alnemri T, Shi Y and Alnemri ES . (2001). Nature, 410, 112–116.

  45. Srivastava RK . (2001). Neoplasia, 3, 535–546.

  46. Sudakin V, Ganoth D, Dahan A, Heller H, Hershko J, Luca FC, Ruderman JV and Hershko A . (1995). Mol. Cell. Biol., 6, 185–197.

  47. Suzuki Y, Nakabayashi Y and Takahashi R . (2001). Proc. Natl. Acad. Sci. USA, 98, 8662–8667.

  48. Suzuki M, Youle RJ and Tjandra N . (2000). Cell, 103, 645–654.

  49. Takahashi R, Deveraux Q, Tamm I, Welsh K, Assa-Munt N, Salvesen GS and Reed JC . (1998). J. Biol. Chem., 273, 7787–7790.

  50. Trompouki E, Hatzivassiliou E, Tsichritzis T, Farmer H, Ashworth A and Mosialos G . (2003). Nature, 424, 793–796.

  51. Wang Q, Wang X and Evers BM . (2003). J. Biol. Chem..

  52. Wesierska-Gadek J, Schloffer D, Kotala V and Horky M . (2002). Int. J. Cancer, 101, 128–136.

  53. Wilson R, Goyal L, Ditzel M, Zachariou A, Baker DA, Agapite J, Steller H and Meier P . (2002). Nat. Cell Biol., 4, 445–450.

  54. Wu JW, Cocina AE, Chai J, Hay BA and Shi Y . (2001). Mol. Cell, 8, 95–104.

  55. Yang Y, Fang S, Jensen JP, Weissman AM and Ashwell JD . (2000). Science, 288, 874–877.

  56. Yang YL and Li XM . (2000). Cell Res., 10, 169–177.

  57. Zhou T, Mountz JD and Kimberly RP . (2002). Immunol. Res., 26, 323–336.

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Acknowledgements

We thank Ms Carol Humber for excellent secretarial work. This work is supported by NIH Grants R01 AG 11653, N01, RO1 AI 42900 and CA 20408, a Birmingham VAMC Merit Review Grant, and a grant from Sankyo Inc. Huang-Ge Zhang is a recipient of Arthritis Foundation Investigator Award.

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Correspondence to John D Mountz.

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Keywords

  • cancer
  • apoptosis
  • ubiquitin proteasome inhibitor
  • TRAIL
  • BAX
  • NF-κB; p53
  • Mdm2

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