1. Please check and comment entries here.
Table of Contents

    Topic review

    Tumor Suppressor Protein p53

    Subjects: Oncology
    View times: 35

    Definition

    Tumor suppressor 53 (p53) is a multifunctional protein that regulates cell cycle, DNA repair, apoptosis and metabolic pathways. In colorectal cancer (CRC), mutations of the gene occur in 60% of patients and are associated with a more aggressive tumor phenotype and resistance to anti-cancer therapy. In addition, inhibitor of apoptosis (IAP) proteins are distinguished biomarkers overexpressed in CRC that impact on a diverse set of signaling pathways associated with the regulation of apoptosis/autophagy, cell migration, cell cycle and DNA damage response. As these mechanisms are further firmly controlled by p53, a transcriptional and post-translational regulation of IAPs by p53 is expected to occur in cancer cells. Here, we aim to review the molecular regulatory mechanisms between IAPs and p53 and discuss the therapeutic potential of targeting their interrelationship by multimodal treatment options.

    1. Introduction

    Colorectal cancer (CRC) accounts for around 10% (more than 1.2 million cases) of annually diagnosed malignancies in the world. It is the fourth most mortal cancer with about 900,000 deaths per year and the incidence is predicted to increase approximately to 2.5 million new cases by 2035 [1]. CRC development is characterized by a multistep process involving a series of histological and morphological changes triggered by a sequential accumulation of specific genomic alterations [2]. Adenomatous polyposis coli (APC) gene mutations occurring in normal colon epithelial cells are among the early incidents of a complex tumorigenesis, resulting in abnormally growing benign precancerous polyps (adenomas and sessile serrated polyps) that, over time acquire the ability to invade the bowel wall and trigger low-grade dysplasia. Successively, promoted by Kirsten rat sarcoma virus (KRAS) oncogene activation and serine/threonine-protein kinase B-Raf (BRAF) mutations along with chromosomal (microsatellite) instabilities, high-grade dysplasia will develop that accelerate transformation to malignant progression and invasive carcinoma by further accumulating p53 mutations. Finally, these local malignancies may acquire the potential to metastasize to local lymph nodes and distant organs [1][3]. TP53 gene mutation frequency is 60% in colorectal cancers with the vast majority of mutations located in the DNA-binding domain of the protein. About 60% of TP53 mutations result in an abrogated function of one allele (loss of heterozygosity); however, this can have a dominant negative effect (DNE) and repress wild-type (wt)-p53 functions. By contrast, gain of function (GOF) mutations may induce tumor initiation and progression as well as cancer stemness, invasion, migration and therapy resistance [4][5][6].
    In dependence of tumor stage, location and lymph node status, CRC is treated by surgery, neoadjuvant (before surgery) or adjuvant chemotherapy (after surgery) with or without concurrent irradiation. By this, treatment of rectal adenocarcinoma represents a particularly good example for a successful implementation of multimodal concepts in cancer management. The establishment of neoadjuvant therapy based on chemoradiation (CRT) prior to surgical resection was a turning point in the treatment of this entity resulting in substantially reduced local recurrence rates and improved survival by the inclusion of oxaliplatin [7][8]. However, despite identical tumor histology and comparable tumor stages, patient’s response to neoadjuvant CRT ranges from a clinically and pathologically confirmed complete remission in 10–30% of cases to progression under treatment [9]. This variable tumor response further displays a strong prognostic impact and significantly correlates with disease-free (DFS) and overall survival (OS) [10][11], while a comprehensive understanding of the molecular basis that defines the individual therapy response is still at its early stage. Among the molecular tumor determinants associated with carcinogenesis, enhanced proliferation, invasion, migration and resistance to anticancer treatment, members of the inhibitor of apoptosis (IAP) family proteins, most pronounced cellular IAP (cIAP1, cIAP2), X chromosome linked IAP (XIAP) and Survivin, have gained increasing interest [12]. In this review, we aim to illustrate a mechanistic interrelationship between IAPs and p53 that may pave the way to develop new combinational therapies to overcome mutant p53 and IAPs based therapy resistance in CRC.

    2. Biology and Functions of p53, a Brief Introduction

    The p53 protein and poly(ADP-ribose) polymerases (PARPs) are considered to be the “guardians of the genome” due to their role in conserving genetic stability by preventing mutations and mediating tumor suppression via a tightly regulated network in response to stress signals, which results in either cell death or survival [4]. PARP-1, by a direct poly(ADP-ribosyl)ation of the p53 protein results in a nuclear accumulation and transcriptional activation of p21 [13] or is required for ATM-mediated p53 activation and gene expression [14]. In addition, epigenetic repression (p53) and activation (PARP-1) of DNA (cytosine-5)-methyltransferase 1 (DNMT1) activity, a key enzyme implicated in the silencing of DNA repair genes, may represent an indirect interrelationship between both proteins [15].
    The major structural part of the p53 protein covers a central DNA-binding domain (DBD), which is connected to the tetramerization domain by a linker region. The regulatory domain is located adjacent to the homo-oligomerization (OD) domain at the protein’s carboxy-terminal end. The vast majority of p53 mutations are located in the DNA-binding region [16]. Transcriptional functions of p53 are mediated by binding to variable consensus sequences in responsive elements in the promoter of target genes. Moreover, p53 also regulates genes partially or completely lacking these consensus sequences dependent on their secondary structure [17]. In addition, p53 directly binds and regulates proteins such as ataxia telangiectasia mutated (ATM) kinase and transcription factors such as Y-box-binding protein (YB-1) [18]. This diversity enables multiple regulatory functions in cellular pathways but needs to be tightly controlled. In that context, a negative feedback loop via murine double minute 2 homologue (MDM2) and MDM4 controls p53-mediated transcriptional and post-transcriptional activity and keeps p53 at low levels under physiological conditions [19][20]. Following DNA damage, p53 is activated by post-translational modifications, e.g., phosphorylation by phosphatidylinositol 3-kinase-related kinase (PIKK)-family members ATM, ataxia telangiectasia and Rad3-related (ATR) and DNA-dependent protein kinase catalytic subunit (DNA-PKcs) or indirect phosphorylation by ATM/ATR/DNA-PKcs substrates checkpoint kinases 1 (CHK1) and CHK2 [19][21]. These modifications result in p53 stabilization, activation and nuclear translocation, followed by p53-mediated transcription of a plethora of target genes, involved in cell cycle regulation, DNA damage repair and apoptosis [22].

    3. Structure and Function of the Inhibitor of Apoptosis Protein Family (IAP)

    The IAP family was first described in 1993 as a class of baculoviral proteins characterized by a functional baculovirus IAP repeat (BIR) domain [23] that prevented apoptosis of insect cells during viral infection [24]. Since their discovery, BIR containing (BIRC) proteins were reported in yeast, insects and mammalians. The human IAP family currently covers eight members, including neuronal apoptosis inhibitory protein (NAIP/BIRC1), cellular IAP1 (cIAP1/BIRC2), cellular IAP2 (cIAP2/BIRC3), X-chromosome-linked IAP (XIAP/BIRC4), Survivin (BIRC5), BIR repeat-containing ubiquitin-conjugating enzyme (BRUCE/Apollon/BIRC6), LIVIN (BIRC7) and human IAP-like 2 (hILP2/BIRC8) [25][26]. The family members differ substantially in protein size and functional domains but share at least one of the family-defining BIR domain facilitating protein-protein interactions with other factors. Additional functional domains include a centrally located ubiquitin associated (UBA) domain present in cIAP1 cIAP2, XIAP and hILP2 to allow these proteins to bind to poly-ubiquitin chains, or a carboxy-terminal localized really interesting new gene (RING) domain conferring ubiquitin ligase activity and mediating signal transduction, protein-protein interactions, and transcription [25]. Further, a caspase activation and recruitment domain (CARD), unique to cIAP1/2, helps to control their ubiquitin ligase activity and stability. In addition, NAIP includes a NAIP-C2TA-HETE-TEP1 nucleotide-binding and oligomerization domain (NACHT) which functions in apoptosis inhibition and major histocompatibility complex (MHC) class II transcriptional activation and a leucine-rich repeat (LRR) domain with a role in signaling pathways of innate immunity and host-pathogen recognition [27]. Finally, Survivin carries an amphipathic α-helical coiled-coil domain at the C-terminus, common in microtubule-associated proteins [28].
    Although IAPs are primarily considered as sole inhibitors of apoptosis, growing evidence evolves regarding their vital impact as transduction intermediates in a diverse set of signaling pathways associated with the regulation of cell migration, cell cycle and DNA damage response. Some of these functions will briefly be described below and are summarized in Figure 1.
    Figure 1. Inhibitor of apoptosis proteins (IAPs) are multifunctional proteins that regulate a variety of key cellular mechanisms such as apoptosis, cell division, invasion and metastasis, autophagy, DNA double-strand break (DSB) repair, cancer progression, immune response and inflammasome formation. Moreover, IAPs are associated with radiation and chemotherapy resistance and are considered to be valuable prognostic and predictive biomarkers in colorectal cancer (CRC). Please see the text for a more detailed discussion. Abbreviations: DNA-PKcs, DNA-dependent protein kinase, catalytic subunit; NF-κB, nuclear factor kappa B; SMAC, second mitochondrial activator of caspases.

    3.1. cIAP1 and cIAP2

    Primarily, cIAP1 was discovered by its involvement in inflammation/apoptosis signaling interacting with tumor necrosis factor receptor-2 associated factors (TNFR2-TRAFs) via its N-terminal BIR domain [29]. The major biological activities of cIAP1 cover a positive regulation of the canonical transcription factor nuclear factor kappaB (NF-κB) activation pathways. By this, cIAP1 complexes with TRAF2, Src homology 2 domain-containing protein tyrosine phosphatase 1 (SHP1), Src and myeloid differentiation primary response 88 (MyD88) to promote canonical activation of NF-κB [30]. Briefly, in the TNFR1 complex, cIAP1/2 serve as ubiquitin ligases for receptor-interacting serine/threonine-protein kinase 1 (RIPK1), which is needed for TNF-α mediated NF-κB and mitogen-activated protein kinase (MAPK) signaling, gene expression, differentiation, mitosis and inhibition of both, caspase-dependent and -independent cell death. In addition, cIAP1/2 limit the non-canonical NF-κB activation pathway. Here, cIAP1/2 act as ubiquitin ligases that target NF-κB-inducing kinase (NIK) for degradation. Further, upon viral infection, cIAP1/2 ubiquitinate TRAF3/6 which is an essential factor for NF-κB deregulation, while attenuation of cIAP1/2 impedes an antiviral response via inhibition of the virus-triggered activation of NF-κB, interferon regulatory factor 3 (IRF3) and interferon-beta (IFN-β) induction [31].
    The expression level of cIAP1 is regulated by a variety of transcriptional and post-translational mechanisms including microRNAs (miRNAs) and proteasomal degradation. For instance, microRNA-29c (miR-29c) binds to the 3′-UTR of cIAP1 effectively downregulating its mRNA and protein levels [32]. Further, downregulation of ubiquitin thioesterase OTU domain ubiquitin aldehyde binding 1 (OTUB1) enhances the degradation of cIAP1 and inhibits the TNF-related weak inducer of apoptosis (TWEAK)-induced MAPK and NF-κB pathway [33]. Upon genotoxic stress, a bilateral cell death regulation involving the ripoptosome (RIP1/3-FADD-Caspase-8-c-FLIP) complex is reported: cIAP1 and cIAP2 directly ubiquitinate RIP1 which associates with the pro-survival transforming growth factor beta-activated kinase 1 (TAK1) and triggers proteolytic degradation of the ripoptosome complex while attenuation of IAPs allows the proper formation of the complex [34].
    Beside its function in concert with cIAP1, cIAP2 is involved in response to metal stress, DNA repair and together with XIAP and Survivin in exosomal secretion [35][36][37]. The latter may not only serve as warning signals, but may also play a role in providing protection to the cancer cells against potential dangers in the tumor microenvironment [38].
    As a response to histone deacetylase (HDAC) inhibitor Panobinostat treatment, cIAP2 exhibits the highest upregulation in line with a decreased level of DNA double-strand break (DSB) repair protein meiotic recombination 11 homolog (MRE11). Moreover, cIAP2 directly interacts with MRE11, promotes its ubiquitination and directs it to degradation that in turn delays DNA DSB repair resulting in increased radiation sensitivity [35].

    3.2. XIAP

    Human XIAP was initially discovered as an IAP-like apoptosis inhibitor protein by its homology to baculovirus IAP genes [39]. XIAP is an archetypical IAP protein that, in contrast to other family members, inhibits the active catalytic sites of caspases-3 and caspases-7 in a direct manner and interferes with the dimerization and activation of caspase-9. This prevents their downstream effector functions, including the release of mitochondrial IAP antagonists such as second mitochondrial activator of caspases (SMAC/Diablo) and the serine peptidase HtrA2/Omi, that bind XIAP’s BIR domains, releasing active caspases into the cytosol [40]. In addition, recent studies presented XIAP as a multifunctional protein involved in cellular and metabolic regulatory circuits such as invasion, migration, necroptosis, oxidative stress, inflammasome formation and autophagy [41][42][43][44][45][46][47][48][49][50]. XIAP´s BIR domain 1 mediates activation of stress-responsive signaling pathways, such as Jun Kinase (JNK) [51] and MAPK phosphorylation cascade that in turn activate NF-κB. XIAP also activates NF-κB by promoting the translocation of the p65 subunit to the nucleus and by degradation of the NF-κB inhibitor IκB [52]. XIAP empowers interleukin-17 mediated NF-κB activation and caspase-3 inhibition that drives colon tumor formation [44]. In line with Survivin, XIAP represents a radiation resistance factor and attenuation of the protein triggers radiation response in CRC cancer cell lines [53][54].

    3.3. Survivin

    Survivin is among the most studied members of the IAP family. The protein was discovered in the late nineties as its smallest member involved in fetal development and cancer progression. Survivin is downregulated in most terminally differentiated cells and re-expressed in the majority of solid and liquid human tumors investigated [55][56]. Survivin is a prime example of a multifunctional protein involved in a variety of regulatory circuits in tumor cells [37][57]. By this, a present conception is that most IAPs, except for XIAP, block apoptosis by mechanisms other than direct caspase inhibition [58], but via cooperative interactions with other partners. Thus, an association of Survivin with hepatitis B X-interacting protein (HBXIP) and/or XIAP inhibits caspases, while binding to SMAC/Diablo counteracts this activity. Moreover, Survivin is expressed in a cell cycle regulated manner participating in cell division as an interactor of chromosomal passenger complex (CPC) proteins INCENP, Borealin and Aurora-B [59][60]. In malignant cells, however, Survivin is regulated independently of mitosis by a variety of oncogenic pathways. Further, Survivin is a predominantly nucleocytoplasmic protein; however, shuttling to or from other compartments like the nucleus is mediated by Exportin-1, irradiation and post-translational regulations such as homodimerization and acetylation of residue K129 [61][62]. Survivin is subjected to multiple post-translational modifications that mostly are decision-makers on its functions and fate of the host. For instance, phosphorylation of residue T34 by p34(cdc2)-cyclin B1 facilitates proper Survivin-caspase-9 interaction that results in inhibition of apoptosis [63]. In addition, Survivin is a radiation-inducible factor mediating the cellular radiation response in a multitude of tumors including colorectal cancer [64][65][66]. By this, Survivin accumulates in the nucleus and interacts with a prime non-homologous end joining repair factor DNA-dependent protein kinase (DNA-PKcs) [67][68]. Survivin forms a heterotetramer complex with DNA-PKcs that results in a conformational change on the DNA-PKcs phosphoinositide 3-kinase domain with enhanced enzymatic activity and detection of differentially abundant phosphopeptides and proteins implicated in the DNA damage response [69].

    3.4. BRUCE/Apollon

    With a molecular weight of 528 kDa, BRUCE/Apollon is a huge E3 ubiquitin transferase whose mutation causes embryonic lethality [70][71]. In concert with other IAPs, one of the main functions of BRUCE is inhibition of apoptosis [70]. BRUCE binds and ubiquitinates SMAC/Diablo, caspase-9 and mitochondrial serine peptidase HtrA2/Omi to prevent apoptosis by facilitating their proteasomal degradation [72][73]. Early in mitosis, BRUCE binds the anaphase-promoting complex/cyclosome (APC/C) and enables the degradation of Cyclin A by ubiquitination independent of cyclin-dependent kinases (CDKs) [74]. However, in the late phase of cytokinesis, BRUCE is an essential component of the midbody ring and the tubular recycling system under the regulation of mitotic kinesin-like protein-1 to regulate cytokinetic abscission [75]. Upon DNA damage, BRUCE acts as a scaffold to form a complex with ubiquitin-specific peptidase 8 (USP8) and breast cancer susceptibility gene C terminus-repeat inhibitor of human telomerase repeat transcriptase expression-1 (BRIT1) at the DSBs, which is vital for the formation of BRIT1 DNA damage foci [76]. BRUCE further regulates ATR-directed signaling pathways in DNA replication stress via interaction with pre-mRNA-processing factor 19, while depletion of BRUCE causes a stalled DNA replication, prevents the activation of ATR and inhibits the phosphorylation of CHK1 and replication protein A [77].

    3.5. LIVIN

    LIVIN (37/39 kDa) plays a key role in a multitude of cellular mechanisms and stress responses including radiation response, invasion, hypoxia-resistance and autophagy [78][79][80][81][82]. LIVIN inhibits apoptosis by binding both, caspase-3/7 as well as TNFα-induced DEVD-like caspase. In addition, LIVIN indirectly inhibits caspase-9 via apoptotic protease activating factor-1 (Apaf-1) [83]. However, reports on a bidirectional regulation between caspases and LIVIN revealed that the truncation of LIVIN (28/30 kDa) by caspase-3/7 transforms it to a pro-apoptotic protein [84][85]. Chemosensitivity studies in colon cancer cells further revealed LIVIN as a drug resistance gene against etoposide (VP-16) and 5-fluorouracil (5-FU) [86], while attenuation of the protein significantly decreases the size of colon cancer xenograft tumors [86][87]. Upon irradiation, LIVIN overexpression is associated with cellular radioresistance, whereas attenuation of LIVIN decreases radiation-induced cell invasion ability and enhances radiation response [78][79].

    3.6. NAIP and hILP-2

    NAIP was first discovered as spinal muscular atrophy (SMA) related gene whose deletion or mutation was restricted to SMA patients [88]. However, comprehensive studies over the last years indicate the relevance of NAIP in a variety of different molecular mechanisms and diseases such as cytokinesis, inflammasome formation, amyloid-β toxicity, amyotrophic lateral sclerosis (ALS) and Parkinson’s disease [89][90][91][92][93]. The least-studied member of IAP family, hILP-2 is discovered as a protein owing a high sequence homology to XIAP but no inhibitory effect on TNF-mediated apoptosis. Nevertheless, it can inhibit Bcl-2-associated X protein (Bax) or caspase-9 and Apaf-1 triggered apoptosis. Moreover, hILP-2 directly interacts with cleaved caspase-9 [94] and attenuation of hILP-2 triggers apoptosis and inhibits migration [95].

    The entry is from 10.3390/cancers13040624

    References

    1. Dekker, E.; Tanis, P.J.; Vleugels, J.L.A.; Kasi, P.M.; Wallace, M.B. Colorectal cancer. Lancet 2019, 394, 1467–1480.
    2. Vogelstein, B.; Fearon, E.R.; Hamilton, S.R.; Kern, S.E.; Preisinger, A.C.; Leppert, M.; Nakamura, Y.; White, R.; Smits, A.M.; Bos, J.L. Genetic alterations during colorectal-tumor development. N. Engl. J. Med. 1988, 319, 525–532.
    3. Simon, K. Colorectal cancer development and advances in screening. Clin. Interv. Aging 2016, 11, 967–976.
    4. Nakayama, M.; Oshima, M. Mutant p53 in colon cancer. J. Mol. Cell Biol. 2019, 11, 267–276.
    5. Solomon, H.; Dinowitz, N.; Pateras, I.S.; Cooks, T.; Shetzer, Y.; Molchadsky, A.; Charni, M.; Rabani, S.; Koifman, G.; Tarcic, O.; et al. Mutant p53 gain of function underlies high expression levels of colorectal cancer stem cells markers. Oncogene 2018, 37, 1669–1684.
    6. Schulz-Heddergott, R.; Moll, U.M. Gain-of-Function (GOF) Mutant p53 as Actionable Therapeutic Target. Cancers 2018, 10, 188.
    7. Van Gijn, W.; Marijnen, C.A.; Nagtegaal, I.D.; Kranenbarg, E.M.; Putter, H.; Wiggers, T.; Rutten, H.J.; Pahlman, L.; Glimelius, B.; van de Velde, C.J. Preoperative radiotherapy combined with total mesorectal excision for resectable rectal cancer: 12-year follow-up of the multicentre, randomised controlled TME trial. Lancet Oncol. 2011, 12, 575–582.
    8. Rodel, C.; Liersch, T.; Becker, H.; Fietkau, R.; Hohenberger, W.; Hothorn, T.; Graeven, U.; Arnold, D.; Lang-Welzenbach, M.; Raab, H.R.; et al. Preoperative chemoradiotherapy and postoperative chemotherapy with fluorouracil and oxaliplatin versus fluorouracil alone in locally advanced rectal cancer: Initial results of the German CAO/ARO/AIO-04 randomised phase 3 trial. Lancet Oncol. 2012, 13, 679–687.
    9. Fokas, E.; Glynne-Jones, R.; Appelt, A.; Beets-Tan, R.; Beets, G.; Haustermans, K.; Marijnen, C.; Minsky, B.D.; Ludmir, E.; Quirke, P.; et al. Outcome measures in multimodal rectal cancer trials. Lancet Oncol. 2020, 21, e252–e264.
    10. Fokas, E.; Fietkau, R.; Hartmann, A.; Hohenberger, W.; Grutzmann, R.; Ghadimi, M.; Liersch, T.; Strobel, P.; Grabenbauer, G.G.; Graeven, U.; et al. Neoadjuvant rectal score as individual-level surrogate for disease-free survival in rectal cancer in the CAO/ARO/AIO-04 randomized phase III trial. Ann. Oncol. 2018, 29, 1521–1527.
    11. Fokas, E.; Strobel, P.; Fietkau, R.; Ghadimi, M.; Liersch, T.; Grabenbauer, G.G.; Hartmann, A.; Kaufmann, M.; Sauer, R.; Graeven, U.; et al. Tumor Regression Grading After Preoperative Chemoradiotherapy as a Prognostic Factor and Individual-Level Surrogate for Disease-Free Survival in Rectal Cancer. J. Natl. Cancer Inst. 2017, 109, djx095.
    12. Mohamed, M.S.; Bishr, M.K.; Almutairi, F.M.; Ali, A.G. Inhibitors of apoptosis: Clinical implications in cancer. Apoptosis 2017, 22, 1487–1509.
    13. Alvarez-Gonzalez, R. Genomic maintenance: The p53 poly(ADP-ribosyl)ation connection. Sci. Signal. 2007, 2007, pe68.
    14. Gajewski, S.; Hartwig, A. PARP1 Is Required for ATM-Mediated p53 Activation and p53-Mediated Gene Expression after Ionizing Radiation. Chem. Res. Toxicol. 2020, 33, 1933–1940.
    15. Christmann, M.; Kaina, B. Epigenetic regulation of DNA repair genes and implications for tumor therapy. Mutat. Res. 2019, 780, 15–28.
    16. Joerger, A.C.; Fersht, A.R. The tumor suppressor p53: From structures to drug discovery. Cold Spring Harb. Perspect. Biol. 2010, 2, a000919.
    17. Brazda, V.; Fojta, M. The Rich World of p53 DNA Binding Targets: The Role of DNA Structure. Int. J. Mol. Sci. 2019, 20, 5605.
    18. Inoue, K.; Fry, E.A.; Frazier, D.P. Transcription factors that interact with p53 and Mdm2. Int. J. Cancer 2016, 138, 1577–1585.
    19. Murray, D.; Mirzayans, R. Cellular Responses to Platinum-Based Anticancer Drugs and UVC: Role of p53 and Implications for Cancer Therapy. Int. J. Mol. Sci. 2020, 21, 5766.
    20. Dobbelstein, M.; Levine, A.J. Mdm2: Open questions. Cancer Sci. 2020, 111, 2203–2211.
    21. Achanta, G.; Pelicano, H.; Feng, L.; Plunkett, W.; Huang, P. Interaction of p53 and DNA-PK in response to nucleoside analogues: Potential role as a sensor complex for DNA damage. Cancer Res. 2001, 61, 8723–8729.
    22. Riley, T.; Sontag, E.; Chen, P.; Levine, A. Transcriptional control of human p53-regulated genes. Nat. Rev. Mol. Cell Biol. 2008, 9, 402–412.
    23. Birnbaum, M.J.; Clem, R.J.; Miller, L.K. An apoptosis-inhibiting gene from a nuclear polyhedrosis virus encoding a polypeptide with Cys/His sequence motifs. J. Virol. 1994, 68, 2521–2528.
    24. Crook, N.E.; Clem, R.J.; Miller, L.K. An apoptosis-inhibiting baculovirus gene with a zinc finger-like motif. J. Virol. 1993, 67, 2168–2174.
    25. Miura, K.; Fujibuchi, W.; Ishida, K.; Naitoh, T.; Ogawa, H.; Ando, T.; Yazaki, N.; Watanabe, K.; Haneda, S.; Shibata, C.; et al. Inhibitor of apoptosis protein family as diagnostic markers and therapeutic targets of colorectal cancer. Surg. Today 2011, 41, 175–182.
    26. Hrdinka, M.; Yabal, M. Inhibitor of apoptosis proteins in human health and disease. Genes Immun. 2019, 20, 641–650.
    27. Ng, A.; Xavier, R.J. Leucine-rich repeat (LRR) proteins: Integrators of pattern recognition and signaling in immunity. Autophagy 2011, 7, 1082–1084.
    28. LaCasse, E.C.; Baird, S.; Korneluk, R.G.; MacKenzie, A.E. The inhibitors of apoptosis (IAPs) and their emerging role in cancer. Oncogene 1998, 17, 3247–3259.
    29. Rothe, M.; Pan, M.G.; Henzel, W.J.; Ayres, T.M.; Goeddel, D.V. The TNFR2-TRAF signaling complex contains two novel proteins related to baculoviral inhibitor of apoptosis proteins. Cell 1995, 83, 1243–1252.
    30. Busca, A.; Konarski, Y.; Gajanayaka, N.; O’Hara, S.; Angel, J.; Kozlowski, M.; Kumar, A. cIAP1/2-TRAF2-SHP-1-Src-MyD88 Complex Regulates Lipopolysaccharide-Induced IL-27 Production through NF-kappaB Activation in Human Macrophages. J. Immunol. 2018, 200, 1593–1606.
    31. Mao, A.P.; Li, S.; Zhong, B.; Li, Y.; Yan, J.; Li, Q.; Teng, C.; Shu, H.B. Virus-triggered ubiquitination of TRAF3/6 by cIAP1/2 is essential for induction of interferon-beta (IFN-beta) and cellular antiviral response. J. Biol. Chem. 2010, 285, 9470–9476.
    32. Huang, L.-G.; Li, J.-P.; Pang, X.-M.; Chen, C.-Y.; Xiang, H.-Y.; Feng, L.-B.; Su, S.-Y.; Li, S.-H.; Zhang, L.; Liu, J.-L. MicroRNA-29c Correlates with Neuroprotection Induced by FNS by Targeting Both Birc2 and Bak1 in Rat Brain after Stroke. CNS Neurosci. Ther. 2015, 21, 496–503.
    33. Goncharov, T.; Niessen, K.; de Almagro, M.C.; Izrael-Tomasevic, A.; Fedorova, A.V.; Varfolomeev, E.; Arnott, D.; Deshayes, K.; Kirkpatrick, D.S.; Vucic, D. OTUB1 modulates c-IAP1 stability to regulate signalling pathways. EMBO J. 2013, 32, 1103–1114.
    34. Tenev, T.; Bianchi, K.; Darding, M.; Broemer, M.; Langlais, C.; Wallberg, F.; Zachariou, A.; Lopez, J.; MacFarlane, M.; Cain, K.; et al. The Ripoptosome, a Signaling Platform that Assembles in Response to Genotoxic Stress and Loss of IAPs. Mol. Cell 2011, 43, 432–448.
    35. Nicholson, J.; Jevons, S.J.; Groselj, B.; Ellermann, S.; Konietzny, R.; Kerr, M.; Kessler, B.M.; Kiltie, A.E. E3 Ligase cIAP2 Mediates Downregulation of MRE11 and Radiosensitization in Response to HDAC Inhibition in Bladder Cancer. Cancer Res. 2017, 77, 3027–3039.
    36. Nudel, K.; Massari, P.; Genco, C.A. Neisseria gonorrhoeae Modulates Cell Death in Human Endocervical Epithelial Cells through Export of Exosome-Associated cIAP2. Infect. Immun. 2015, 83, 3410–3417.
    37. Valenzuela, M.M.; Ferguson Bennit, H.R.; Gonda, A.; Diaz Osterman, C.J.; Hibma, A.; Khan, S.; Wall, N.R. Exosomes Secreted from Human Cancer Cell Lines Contain Inhibitors of Apoptosis (IAP). Cancer Microenviron. 2015, 8, 65–73.
    38. Khan, S.; Bennit, H.F.; Wall, N.R. The emerging role of exosomes in survivin secretion. Histol. Histopathol. 2015, 30, 43–50.
    39. Duckett, C.S.; Nava, V.E.; Gedrich, R.W.; Clem, R.J.; Van Dongen, J.L.; Gilfillan, M.C.; Shiels, H.; Hardwick, J.M.; Thompson, C.B. A conserved family of cellular genes related to the baculovirus iap gene and encoding apoptosis inhibitors. EMBO J. 1996, 15, 2685–2694.
    40. Silke, J.; Vucic, D. IAP family of cell death and signaling regulators. Methods Enzymol. 2014, 545, 35–65.
    41. Xu, J.; Hua, X.; Yang, R.; Jin, H.; Li, J.; Zhu, J.; Tian, Z.; Huang, M.; Jiang, G.; Huang, H.; et al. XIAP Interaction with E2F1 and Sp1 via its BIR2 and BIR3 domains specific activated MMP2 to promote bladder cancer invasion. Oncogenesis 2019, 8, 71.
    42. Delbue, D.; Mendonca, B.S.; Robaina, M.C.; Lemos, L.G.T.; Lucena, P.I.; Viola, J.P.B.; Magalhaes, L.M.; Crocamo, S.; Oliveira, C.A.B.; Teixeira, F.R.; et al. Expression of nuclear XIAP associates with cell growth and drug resistance and confers poor prognosis in breast cancer. Biochim. Biophys. Acta Mol. Cell Res. 2020, 1867, 118761.
    43. Wicki, S.; Gurzeler, U.; Wei-Lynn Wong, W.; Jost, P.J.; Bachmann, D.; Kaufmann, T. Loss of XIAP facilitates switch to TNFalpha-induced necroptosis in mouse neutrophils. Cell Death Dis. 2016, 7, e2422.
    44. Liao, Y.; Zhao, J.; Bulek, K.; Tang, F.; Chen, X.; Cai, G.; Jia, S.; Fox, P.L.; Huang, E.; Pizarro, T.T.; et al. Inflammation mobilizes copper metabolism to promote colon tumorigenesis via an IL-17-STEAP4-XIAP axis. Nat. Commun. 2020, 11, 900.
    45. Zilu, S.; Qian, H.; Haibin, W.; Chenxu, G.; Deshuai, L.; Qiang, L.; Linfeng, H.; Jun, T.; Minxuan, X. Effects of XIAP on high fat diet-induced hepatic steatosis: A mechanism involving NLRP3 inflammasome and oxidative stress. Aging 2019, 11, 12177–12201.
    46. Lu, M.; Qin, X.; Yao, J.; Yang, Y.; Zhao, M.; Sun, L. MiR-134-5p targeting XIAP modulates oxidative stress and apoptosis in cardiomyocytes under hypoxia/reperfusion-induced injury. IUBMB Life 2020, 72, 2154–2166.
    47. Damgaard, R.B.; Nachbur, U.; Yabal, M.; Wong, W.W.; Fiil, B.K.; Kastirr, M.; Rieser, E.; Rickard, J.A.; Bankovacki, A.; Peschel, C.; et al. The ubiquitin ligase XIAP recruits LUBAC for NOD2 signaling in inflammation and innate immunity. Mol. Cell 2012, 46, 746–758.
    48. Andree, M.; Seeger, J.M.; Schull, S.; Coutelle, O.; Wagner-Stippich, D.; Wiegmann, K.; Wunderlich, C.M.; Brinkmann, K.; Broxtermann, P.; Witt, A.; et al. BID-dependent release of mitochondrial SMAC dampens XIAP-mediated immunity against Shigella. EMBO J. 2014, 33, 2171–2187.
    49. Huang, X.; Wu, Z.; Mei, Y.; Wu, M. XIAP inhibits autophagy via XIAP-Mdm2-p53 signalling. EMBO J. 2013, 32, 2204–2216.
    50. Lin, F.; Ghislat, G.; Luo, S.; Renna, M.; Siddiqi, F.; Rubinsztein, D.C. XIAP and cIAP1 amplifications induce Beclin 1-dependent autophagy through NFkappaB activation. Hum. Mol. Genet. 2015, 24, 2899–2913.
    51. Gyrd-Hansen, M.; Meier, P. IAPs: From caspase inhibitors to modulators of NF-kappaB, inflammation and cancer. Nat. Rev. Cancer 2010, 10, 561–574.
    52. Jin, H.S.; Lee, D.H.; Kim, D.H.; Chung, J.H.; Lee, S.J.; Lee, T.H. cIAP1, cIAP2, and XIAP act cooperatively via nonredundant pathways to regulate genotoxic stress-induced nuclear factor-kappaB activation. Cancer Res. 2009, 69, 1782–1791.
    53. Moussata, D.; Amara, S.; Siddeek, B.; Decaussin, M.; Hehlgans, S.; Paul-Bellon, R.; Mornex, F.; Gerard, J.P.; Romestaing, P.; Rodel, F.; et al. XIAP as a radioresistance factor and prognostic marker for radiotherapy in human rectal adenocarcinoma. Am. J. Pathol. 2012, 181, 1271–1278.
    54. Hehlgans, S.; Petraki, C.; Reichert, S.; Cordes, N.; Rodel, C.; Rodel, F. Double targeting of Survivin and XIAP radiosensitizes 3D grown human colorectal tumor cells and decreases migration. Radiother. Oncol. 2013, 108, 32–39.
    55. Ambrosini, G.; Adida, C.; Altieri, D.C. A novel anti-apoptosis gene, survivin, expressed in cancer and lymphoma. Nat. Med. 1997, 3, 917–921.
    56. Altieri, D.C. Validating survivin as a cancer therapeutic target. Nat. Rev. Cancer 2003, 3, 46–54.
    57. Barrera-Vazquez, O.S.; Cancio-Lonches, C.; Hernandez-Gonzalez, O.; Chavez-Munguia, B.; Villegas-Sepulveda, N.; Gutierrez-Escolano, A.L. The feline calicivirus leader of the capsid protein causes survivin and XIAP downregulation and apoptosis. Virology 2019, 527, 146–158.
    58. Eckelman, B.P.; Salvesen, G.S.; Scott, F.L. Human inhibitor of apoptosis proteins: Why XIAP is the black sheep of the family. EMBO Rep. 2006, 7, 988–994.
    59. Li, F.; Ambrosini, G.; Chu, E.Y.; Plescia, J.; Tognin, S.; Marchisio, P.C.; Altieri, D.C. Control of apoptosis and mitotic spindle checkpoint by survivin. Nature 1998, 396, 580–584.
    60. Vader, G.; Kauw, J.J.W.; Medema, R.H.; Lens, S.M.A. Survivin mediates targeting of the chromosomal passenger complex to the centromere and midbody. EMBO Rep. 2006, 7, 85–92.
    61. Wang, H.; Holloway, M.P.; Ma, L.; Cooper, Z.A.; Riolo, M.; Samkari, A.; Elenitoba-Johnson, K.S.; Chin, Y.E.; Altura, R.A. Acetylation directs survivin nuclear localization to repress STAT3 oncogenic activity. J. Biol. Chem. 2010, 285, 36129–36137.
    62. Engelsma, D.; Rodriguez, J.A.; Fish, A.; Giaccone, G.; Fornerod, M. Homodimerization antagonizes nuclear export of survivin. Traffic 2007, 8, 1495–1502.
    63. Wheatley, S.P.; Altieri, D.C. Survivin at a glance. J. Cell Sci. 2019, 132.
    64. Lu, B.; Mu, Y.; Cao, C.; Zeng, F.; Schneider, S.; Tan, J.; Price, J.; Chen, J.; Freeman, M.; Hallahan, D.E. Survivin as a therapeutic target for radiation sensitization in lung cancer. Cancer Res. 2004, 64, 2840–2845.
    65. Chakravarti, A.; Zhai, G.G.; Zhang, M.; Malhotra, R.; Latham, D.E.; Delaney, M.A.; Robe, P.; Nestler, U.; Song, Q.; Loeffler, J. Survivin enhances radiation resistance in primary human glioblastoma cells via caspase-independent mechanisms. Oncogene 2004, 23, 7494–7506.
    66. Rodel, F.; Hoffmann, J.; Distel, L.; Herrmann, M.; Noisternig, T.; Papadopoulos, T.; Sauer, R.; Rodel, C. Survivin as a radioresistance factor, and prognostic and therapeutic target for radiotherapy in rectal cancer. Cancer Res. 2005, 65, 4881–4887.
    67. Capalbo, G.; Dittmann, K.; Weiss, C.; Reichert, S.; Hausmann, E.; Rodel, C.; Rodel, F. Radiation-induced survivin nuclear accumulation is linked to DNA damage repair. Int. J. Radiat. Oncol. Biol. Phys. 2010, 77, 226–234.
    68. Wang, X.; Beitler, J.J.; Huang, W.; Chen, G.; Qian, G.; Magliocca, K.; Patel, M.R.; Chen, A.Y.; Zhang, J.; Nannapaneni, S.; et al. Honokiol Radiosensitizes Squamous Cell Carcinoma of the Head and Neck by Downregulation of Survivin. Clin. Cancer Res. 2018, 24, 858–869.
    69. Gullulu, O.; Hehlgans, S.; Mayer, B.E.; Gossner, I.; Petraki, C.; Hoffmann, M.; Dombrowsky, M.J.; Kunzmann, P.; Hamacher, K.; Strebhardt, K.; et al. A spatial and functional interaction of a heterotetramer Survivin-DNA-PKcs complex in DNA damage response. Cancer Res. 2021.
    70. Hauser, H.P.; Bardroff, M.; Pyrowolakis, G.; Jentsch, S. A giant ubiquitin-conjugating enzyme related to IAP apoptosis inhibitors. J. Cell Biol. 1998, 141, 1415–1422.
    71. Hitz, C.; Vogt-Weisenhorn, D.; Ruiz, P.; Wurst, W.; Floss, T. Progressive loss of the spongiotrophoblast layer of Birc6/Bruce mutants results in embryonic lethality. Genesis 2005, 42, 91–103.
    72. Hao, Y.; Sekine, K.; Kawabata, A.; Nakamura, H.; Ishioka, T.; Ohata, H.; Katayama, R.; Hashimoto, C.; Zhang, X.; Noda, T.; et al. Apollon ubiquitinates SMAC and caspase-9, and has an essential cytoprotection function. Nat. Cell Biol. 2004, 6, 849–860.
    73. Sekine, K.; Hao, Y.; Suzuki, Y.; Takahashi, R.; Tsuruo, T.; Naito, M. HtrA2 cleaves Apollon and induces cell death by IAP-binding motif in Apollon-deficient cells. Biochem. Biophys. Res. Commun. 2005, 330, 279–285.
    74. Kikuchi, R.; Ohata, H.; Ohoka, N.; Kawabata, A.; Naito, M. APOLLON protein promotes early mitotic CYCLIN A degradation independent of the spindle assembly checkpoint. J. Biol. Chem. 2014, 289, 3457–3467.
    75. Pohl, C.; Jentsch, S. Final stages of cytokinesis and midbody ring formation are controlled by BRUCE. Cell 2008, 132, 832–845.
    76. Ge, C.; Che, L.; Ren, J.; Pandita, R.K.; Lu, J.; Li, K.; Pandita, T.K.; Du, C. BRUCE regulates DNA double-strand break response by promoting USP8 deubiquitination of BRIT1. Proc. Natl. Acad. Sci. USA 2015, 112, E1210–E1219.
    77. Ge, C.; Vilfranc, C.L.; Che, L.; Pandita, R.K.; Hambarde, S.; Andreassen, P.R.; Niu, L.; Olowokure, O.; Shah, S.; Waltz, S.E.; et al. The BRUCE-ATR Signaling Axis Is Required for Accurate DNA Replication and Suppression of Liver Cancer Development. Hepatology 2019, 69, 2608–2622.
    78. Sun, J.G.; Liao, R.X.; Zhang, S.X.; Duan, Y.Z.; Zhuo, W.L.; Wang, X.X.; Wang, Z.X.; Li, D.Z.; Chen, Z.T. Role of inhibitor of apoptosis protein Livin in radiation resistance in nonsmall cell lung cancer. Cancer Biother. Radiopharm. 2011, 26, 585–592.
    79. Wu, S.Q.; Xu, Q.B.; Sheng, W.Y.; Su, L.Y.; Zhu, L.W. A novel role for Livin in the response to ultraviolet B radiation and pterygium development. Int. J. Mol. Med. 2020, 45, 1103–1111.
    80. Chen, F.; Yang, D.; Wang, S.; Che, X.; Wang, J.; Li, X.; Zhang, Z.; Chen, X.; Song, X. Livin regulates prostate cancer cell invasion by impacting the NF-kappaB signaling pathway and the expression of FN and CXCR4. IUBMB Life 2012, 64, 274–283.
    81. Li, F.; Yin, X.; Luo, X.; Li, H.Y.; Su, X.; Wang, X.Y.; Chen, L.; Zheng, K.; Ren, G.S. Livin promotes progression of breast cancer through induction of epithelial-mesenchymal transition and activation of AKT signaling. Cell Signal. 2013, 25, 1413–1422.
    82. Hsieh, C.H.; Lin, Y.J.; Wu, C.P.; Lee, H.T.; Shyu, W.C.; Wang, C.C. Livin contributes to tumor hypoxia-induced resistance to cytotoxic therapies in glioblastoma multiforme. Clin. Cancer Res. 2015, 21, 460–470.
    83. Kasof, G.M.; Gomes, B.C. Livin, a novel inhibitor of apoptosis protein family member. J. Biol. Chem. 2001, 276, 3238–3246.
    84. Nachmias, B.; Ashhab, Y.; Bucholtz, V.; Drize, O.; Kadouri, L.; Lotem, M.; Peretz, T.; Mandelboim, O.; Ben-Yehuda, D. Caspase-mediated cleavage converts Livin from an antiapoptotic to a proapoptotic factor: Implications for drug-resistant melanoma. Cancer Res. 2003, 63, 6340–6349.
    85. Nachmias, B.; Lazar, I.; Elmalech, M.; Abed-El-Rahaman, I.; Asshab, Y.; Mandelboim, O.; Perlman, R.; Ben-Yehuda, D. Subcellular localization determines the delicate balance between the anti- and pro-apoptotic activity of Livin. Apoptosis 2007, 12, 1129–1142.
    86. Wang, X.; Xu, J.; Ju, S.; Ni, H.; Zhu, J.; Wang, H. Livin gene plays a role in drug resistance of colon cancer cells. Clin. Biochem. 2010, 43, 655–660.
    87. Oh, B.Y.; Lee, R.A.; Kim, K.H. siRNA targeting Livin decreases tumor in a xenograft model for colon cancer. World J. Gastroenterol. 2011, 17, 2563–2571.
    88. Roy, N.; Mahadevan, M.S.; McLean, M.; Shutler, G.; Yaraghi, Z.; Farahani, R.; Baird, S.; Besner-Johnston, A.; Lefebvre, C.; Kang, X.; et al. The gene for neuronal apoptosis inhibitory protein is partially deleted in individuals with spinal muscular atrophy. Cell 1995, 80, 167–178.
    89. Abadia-Molina, F.; Moron-Calvente, V.; Baird, S.D.; Shamim, F.; Martin, F.; MacKenzie, A. Neuronal apoptosis inhibitory protein (NAIP) localizes to the cytokinetic machinery during cell division. Sci. Rep. 2017, 7, 39981.
    90. Karki, R.; Lee, E.; Place, D.; Samir, P.; Mavuluri, J.; Sharma, B.R.; Balakrishnan, A.; Malireddi, R.K.S.; Geiger, R.; Zhu, Q.; et al. IRF8 Regulates Transcription of Naips for NLRC4 Inflammasome Activation. Cell 2018, 173, 920–933.
    91. Lesne, S.; Gabriel, C.; Nelson, D.A.; White, E.; Mackenzie, E.T.; Vivien, D.; Buisson, A. Akt-dependent expression of NAIP-1 protects neurons against amyloid- toxicity. J. Biol. Chem. 2005, 280, 24941–24947.
    92. Kano, O.; Tanaka, K.; Kanno, T.; Iwasaki, Y.; Ikeda, J.E. Neuronal apoptosis inhibitory protein is implicated in amyotrophic lateral sclerosis symptoms. Sci. Rep. 2018, 8, 6.
    93. Crocker, S.J.; Wigle, N.; Liston, P.; Thompson, C.S.; Lee, C.J.; Xu, D.; Roy, S.; Nicholson, D.W.; Park, D.S.; MacKenzie, A.; et al. NAIP protects the nigrostriatal dopamine pathway in an intrastriatal 6-OHDA rat model of Parkinson’s disease. Eur. J. Neurosci. 2001, 14, 391–400.
    94. Richter, B.W.; Mir, S.S.; Eiben, L.J.; Lewis, J.; Reffey, S.B.; Frattini, A.; Tian, L.; Frank, S.; Youle, R.J.; Nelson, D.L.; et al. Molecular cloning of ILP-2, a novel member of the inhibitor of apoptosis protein family. Mol. Cell Biol. 2001, 21, 4292–4301.
    95. Zhu, L.; Zhou, W.; Zhu, X.; Xiang, S.; Wang, S.; Peng, Y.; Lu, B.; Tang, P.; Chen, Q.; Wu, M.; et al. Inhibitor of apoptosis proteinlike protein2: A novel growth accelerator for breast cancer cells. Oncol. Rep. 2018, 40, 2047–2055.
    More