Oxidative Medicine and Cellular Longevity
Volume 2017 (2017), Article ID 2969271, 18 pages
https://doi.org/10.1155/2017/2969271

Review Article

Unfolded Protein Response of the Endoplasmic Reticulum in Tumor Progression and Immunogenicity

Yoon Seon Yoo,1,2 Hye Gyeong Han,1,2 and Young Joo Jeon1,2

¹Department of Biochemistry, Chungnam National University School of Medicine, Daejeon 35015, Republic of Korea
²Department of Medical Science, Chungnam National University School of Medicine, Daejeon 35015, Republic of Korea

Correspondence should be addressed to Young Joo Jeon; yjjeon@cnu.ac.kr

Received 12 September 2017; Accepted 29 November 2017; Published 21 December 2017

Academic Editor: Dieter Wolf

Copyright © 2017 Yoon Seon Yoo et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Abstract.

The endoplasmic reticulum (ER) is a pivotal regulator of folding, quality control, trafficking, and targeting of secreted and transmembrane proteins, and accordingly, eukaryotic cells have evolved specialized machinery to ensure that the ER enables these proteins to acquire adequate folding and maturation in the presence of intrinsic and extrinsic insults. This adaptive capacity of the ER to intrinsic and extrinsic perturbations is important for maintaining protein homeostasis, which is termed proteostasis. Failure in adaptation to these perturbations leads to accumulation of misfolded or unassembled proteins in the ER, which is termed ER stress, resulting in the activation of unfolded protein response (UPR) of the ER and the execution of ER-associated degradation (ERAD) to restore homeostasis. Furthermore, both of the two axes play key roles in the control of tumor progression, inflammation, immunity, and aging. Therefore, understanding UPR of the ER and subsequent ERAD will provide new insights into the pathogenesis of many human diseases and contribute to therapeutic intervention in these diseases.

1. Introduction

The endoplasmic reticulum (ER) is a dynamic and specialized tubular-reticular network and extends throughout the cytoplasm in the form of connected sacs and branching tubules [1]. The ER network is heterogenous in its structure and adopts different morphologies in conjunction with different functions [2]. Interestingly, the ER is physically and functionally interconnected with every other cellular compartment and can sense intrinsic and extrinsic perturbations, combine these stress signals, and manage the cellular processes, indicating its role as a central coordinator for maintenance of cellular homeostasis [3]. The ER engages in various cellular functions involving the biosynthesis of lipid species such as cholesterol, triacylglycerol, and phospholipids, the degradation of glycogen, detoxification, and the maintenance of Ca²⁺ homeostasis [4–6]. Most importantly, the ER is involved in the synthesis, folding, maturation, and trafficking of secreted and transmembrane proteins, which constitute about one-third of all the proteins that are synthesized in the cell [5, 6]. These proteins participate in important cellular and organismal processes involving protein degradation, signal transduction, lipid metabolism, and cell-cell communications, suggesting that maintaining the integrity of these proteins is essential for life.

Protein quality control of the ER is composed of three axes, acceleration of adequate protein folding, activation of unfolded protein response (UPR), and protein clearance via ER-associated degradation (ERAD) [7]. Accumulation of misfolded and unassembled proteins can cause stress and damage, resulting in the activation of UPR to determine cell fate and function and in the subsequent restoration of protein homeostasis, which is termed proteostasis. Even with the assistance of dedicated protein folding machinery in the ER, a large portion of proteins entering the ER fails to obtain proper conformation due to mutations, unavailability of chaperones, or changes in the amounts of interacting partners and eventually must be eliminated [8, 9]. Eukaryotic cells have evolved ERAD for clearance of misfolded, unassembled, or tightly regulated proteins, resulting in the maintenance of proteostasis [10–13]. Intriguingly, a failure in the maintenance and/or restoration of the proteostasis leads to various protein misfolding diseases [14–17], implicating the importance of stringent protein quality control in the ER.

In this review, we not only discuss the molecular mechanisms of UPR of the ER and ERAD but also summarize advances in versatile aspects of these two axes. Furthermore, we provide current insights into how the adaptive capacity of the ER to intrinsic and extrinsic perturbations contributes to the modulation of malignancy, the regulation of cancer immunity, and the efficacy of therapies for cancer.

2. UPR of the ER

Numerous endogenous and exogenous stresses can disrupt ER protein folding environment, and unfolded or misfolded proteins accumulate in the ER, which activates UPR. While the UPR is also involved in mitochondria biology and apoptotic signal transduction, a main function of UPR is to maintain proteostasis under ER stress condition [18, 19]. In multicellular eukaryotes, UPR consists of three branches of ER transmembrane sensors, inositol-requiring protein 1 (IRE1), activating transcription factor 6 (ATF6), and protein kinase RNA- (PKR-) like ER kinase (PERK) (Figure 1). These sensors have two functional domains. The luminal domains of these sensors can sense the protein folding environment and their cytosolic domains can be connected to transcription and translation machinery. Under normal conditions, the luminal domains of these sensors are kept in an inactive state through the association with a chaperone, binding immunoglobulin protein (BiP; also known as GRP78), which belongs to the heat shock protein 70 family [20]. Upon ER stress, BiP dissociates from the ER sensors and is recruited to misfolded proteins, resulting in the activation of UPR [21–23]. It has been also known that unfolded proteins themselves can directly bind to IRE1 or PERK and this direct binding results in dimerization, oligomerization, and activation of UPR [24–27]. While the downstream response of UPR activation is a transient attenuation of global protein synthesis, an increase in a transcriptional program as well as the translation of many mRNAs including Atf4 is induced, all of which direct towards resolving the stress [28–31]. Furthermore, when protein misfolding is not resolved, prolonged UPR activation promotes apoptosis by inducing the expression of proapoptotic genes via PERK-eIF2α-ATF4-CHOP axis [32, 33].

Figure 1: Unfolded protein response (UPR) of the endoplasmic reticulum (ER). UPR is composed of three branches of ER transmembrane sensors, IRE1, PERK, and ATF6. Upon ER stress, BiP is released from the ER sensors and is recruited to misfolded proteins, leading to the activation of UPR. Activated ER sensors transmit the stress signal into the cytosol and nucleus and subsequently operate the coordinated stress response, the UPR.

2.1. PERK

Upon ER stress, BiP dissociates from PERK, permitting PERK homodimerization and autophosphorylation to activate its cytoplasmic kinase domain. The activated cytosolic kinase domain of PERK in turn phosphorylates the α subunit of eukaryotic translation initiation factor 2 (eIF2α) at serine 51, which inhibits guanine nucleotide exchange factor (eIF2B) and lowers global mRNA translation, thereby attenuating the entrance of newly synthesized proteins into the ER and facilitating the cell to resolve the stress [34]. Although global mRNA translation is reduced under ER stress conditions, certain species of mRNA are favorably translated, involving activation transcription factor 4 (ATF4; also known as CREB2), which transactivates various genes, including C/EBP homologous protein (Chop), ER oxidoreductin 1 (Ero1), and growth arrest and DNA damage-inducible protein (Gadd34) [35–37]. Among them, CHOP is involved in ER stress-induced apoptosis under excessive and chronic activation of PERK [37, 38]. At early time points after ER stress, transcription of Chop is suppressed by several ways, including histone methylation and Toll-like receptor (TLR) signaling [39, 40]. However, if ER stress cannot be resolved, ATF4 and CHOP function together as a heterodimer, which increases protein synthesis, protein misfolding, oxidative stress, and finally apoptosis [17, 30].

2.2. IRE1

IRE1 possesses a serine/threonine kinase activity as well as endoribonuclease activity within the cytoplasmic domain [41]. Upon sensing the ER stress, IRE1 is released from BiP and activated, leading to the nonconventional splicing of a single mRNA that encodes X-box binding protein 1 (XBP1). As a result, a translational frameshift is generated and spliced Xbp1 (XBP1s) isoform is produced [25, 42]. As a transcription factor, XBP1s induces the transcription of a wide range of targets, involving molecular chaperones and enzymes that together assist folding of polypeptides [43, 44]. In addition, XBP1s induces the expression of genes involved in membrane expansion and lipid synthesis [45]. Interestingly, once translated, an unspliced form, XBP1u negatively regulates XBP1s by promoting its proteasome-mediated degradation [46]. When ER stress persists, IRE1 is in a hyperactive state, resulting in the cleavage of many other RNAs besides Xbp1, involving precursors of apoptosis-inhibitory microRNAs, which in turn promotes programmed cell death [47–49].

2.3. ATF6

ATF6 contains a bZIP transcription factor within its cytosolic domain. Under stress-free conditions, the luminal domain of ATF6 is kept inactive via association with BiP. Upon ER stress, BiP is released from ATF6 and ATF6 is transported to the Golgi apparatus, where it is processed by the Golgi enzyme site 1 protease (S1P) and S2P, leading to the transport of its cleaved cytosolic p50 fragment into the nucleus. The cytosolic p50 fragment then induces the expression of genes such as Xbp1 to increase the capacity of the ER to resolve ER stress as well as genes required for ERAD [50–52]. Intriguingly, XBP1s and ATF6 can heterodimerize and also induce the expression of genes involved in ERAD [52, 53].

3. ERAD

The ER participates in the synthesis of the secretory proteins, of the luminal proteins of the ER, Golgi apparatus, endosomes, and lysosomes, and of membrane proteins. Protein synthesis in the ER is a complicated process involving targeting of ribosomes loaded with nascent polypeptide to the ER membrane, cotranslational translocation of nascent polypeptide, and co- and posttranslational folding and maturation of the polypeptide chain [2, 54]. The co- and posttranslational folding and maturation of polypeptides commence during translocation and are assisted by molecular chaperones residing in the ER [55–58]. Chaperones associate with folding intermediates, accelerate their proper folding and assembly, and prevent their improper aggregation. In addition, modifications involving N-linked glycosylation, proline cis-trans isomerization, and disulfide bond formation support proper folding of translocated polypeptides in the ER [59–62]. Nevertheless, protein maturation is not a perfect process and produces improper polypeptides, which can cause cellular stress and cytotoxicity [16, 17] and therefore must be eliminated. Eukaryotic cells have evolved ERAD to eliminate misfolded, unassembled, or metabolically regulated proteins by the cytosolic ubiquitin proteasome system (UPS) [10–13] (Figure 2). Since the late 1980s, it has been elucidated that ERAD is an elaborate and multistep process that recognizes, extracts, and ubiquitinates proteins for degradation by the cytosolic 26S proteasome [7, 13, 63–65]. In ERAD, proteins to be integrated into ER membrane or translocated into the lumen can be ultimately subject to UPS. E3 ubiquitin ligases in ERAD are spatially separated from their substrates, in part, by the ER membrane, suggesting that proofreading step is required to sort out potential ERAD targets. Polypeptides that have failed to acquire a native structure are subject to ERAD. These polypeptides are delivered to the ERAD E3 ubiquitin ligases and ubiquitinated on the cytosolic side of ER membrane. Then, the ubiquitinated substrates are subsequently extracted from the ER membrane and released into the cytoplasm for the proteasome-mediated degradation.

(a)

(b)

Figure 2: ER-associated degradation (ERAD). (a) ERAD functions to eliminate terminally misfolded, unassembled, or tightly regulated proteins by the cytosolic ubiquitin proteasome system (UPS). (1) Protein translocation into the ER through translocon. (2) Protein folding and maturation. Proteins translocated into the ER are subject to cotranslational and posttranslational folding. (3) Substrate recognition. Proteins failing to acquire their native conformation are recognized for ERAD. (4) Retrotranslocation and ubiquitination. Recognition of ERAD substrates facilitates the assembly of retrotranslocon and initiates ERAD E3 ubiquitin ligase-mediated polyubiquitination of substrates. (5) Proteasomal degradation. Carbohydrate and ubiquitin chains are removed from the retrotranslocated substrates. The retrotranslocated substrates are then inserted into the narrow channel of the proteasome, resulting in the degradation of substrates. (b) Retrotranslocation. ERAD substrate is recruited to retrotranslocon complex, which involves SEL1L, OS-9, Derlin, E3 ubiquitin ligase, and p97/Npl4/Ufd1 complex. Blue pentagon indicates N-glycan and green circle indicates ubiquitin.

Recently, it is also demonstrated that ERAD plays a role in the control of degradation of some properly folded ER proteins [9]. In addition, certain viruses exploit ERAD to degrade host proteins such as major histocompatibility class I (MHC I) heavy chain and CD4 molecules, thereby escaping immune surveillance [66–68]. The human cytomegalovirus (HCMV) encodes ER membrane adaptor proteins such as US2 and US11, which bind to MHC I molecules and deliver them to ERAD [66]. Similarly, the human immunodeficiency virus- (HIV-1-) encoded adaptor protein, Vpu leads to the proteasome-mediated degradation of CD4 [67, 68]. Collectively, as a sophisticated ER protein quality control mechanism, ERAD not only functions as the gateway for the flux of proteins into the secretory pathway or membrane incorporation but also impacts intracellular organelle function and cellular communication with the extracellular environment [28]. Genetic ablation of the components involved in ERAD results in embryonic lethality in mice, indicating the importance of ERAD in the maintenance of organismal homeostasis [69–71]. A failure of the ERAD process to remove misfolded or unfolded proteins results in the accumulation of these proteins, a condition referenced as ER stress and is closely associated with a variety of human diseases, involving cancer, neurodegeneration, infectious diseases, and metabolic diseases [72].

3.1. Recognition

Substrate recognition must be tightly controlled, because this is the commitment step for substrate degradation in ERAD [9]. A number of proteins synthesized in the ER are cotranslationally modified by attachment of high-mannose “core” glycans, with the structure Glc₃Man₉GlcNAc₂ ((Glc) glucose, (Man) mannose, (GlcNAc) N-acetylglucosamine), to consensus asparagine residues within canonical N-glycosylation sites (NxS/T) [73]. The ER quality control system uses these glycans in monitoring conformational maturation, directing correctly folded proteins to ER exit, or directing misfolded proteins to ERAD. The lectin-type chaperone, calnexin or calreticulin binds to Glc₁Man₉GlcNAc₂ produced by deglucosylation of core glycans and facilitates folding of immature glycoproteins [73]. Further deglucosylation of final glucose from N-glycan inhibits additional binding of the glycoproteins to calnexin or calreticulin, allowing ER exit of the proteins. Interestingly, incompletely folded proteins are subject to reglucosylation by UDP-glucose : glycoprotein glucosyltransferase (UGGT). These glycoproteins reassociate with calnexin or calreticulin and undergo further rounds of folding [74, 75].

Terminally misfolded proteins must escape from calnexin/calreticulin cycle for ERAD. This escape is regulated by mannosidases that progressively remove terminal mannose residues from core glycans, permitting them to associate with mannose-specific lectins for ERAD [74, 76]. Further trimming of terminal mannoses by ER mannosidase I (ERManI) [77, 78], the ER degradation-enhancing α-mannosidase-like proteins 1 (EDEM1) [79, 80], EDEM3 [81, 82], or Golgi-resident mannosidase α class 1C member 1 (Man1C1) [83] leads to the discrimination of terminally misfolded proteins from their maturation-competent counterparts. ER-resident lectins, osteosarcoma 9 (OS-9), and XTP3-B/Erlectin then recognize these mannose-trimmed proteins through mannose-6-phosphate receptor homology (MRH) domains and recruit them to protein penetration channel, retrotranslocon [84–86]. Silencing both lectins attenuates the degradation of model substrates, while knockdown of either lectin has marginal effects in stabilizing ERAD substrates, suggesting that there may be some redundancy between them [82, 85, 87].

Whereas oligosaccharides are common for substrate recognition step, features besides glycan trimming can contribute to the targeting of folding-defective proteins to ERAD. The nonlectin chaperone BiP associates with glycoproteins as well as nonglycosylated proteins for targeting to ERAD [88, 89]. In addition, EDEM1 is involved in targeting of unglycosylated proteins to ERAD [90]. Redox-driven protein disulfide isomerase (PDI) that is characterized by thioredoxin-like motifs [91] is also involved in ERAD [92]. Interaction of chaperones with ERAD substrates permits the association of substrates with PDI.

3.2. Retrotranslocation

Energy-dependent protein extraction across the ER membrane back into the cytoplasm is a step known as dislocation or retrotranslocation [93] (Figure 2(b)). Importantly, no evidence indicates that the ER lumen contains any components involved in UPS such as E1 ubiquitin-activating enzyme, E2 ubiquitin-conjugating enzyme, or the proteasome, implicating that retrotranslocation is an essential step for degradation of ERAD substrates. Intriguingly, the processes of retrotranslocation and proteasomal degradation should be also tightly coupled, because many ERAD substrates are highly hydrophobic and easily aggregate in an aqueous environment. Therefore, a number of adaptors that recognize a diverse set of features through which substrates are committed to ERAD are essential for recruitment of ERAD substrates to retrotranslocons. As one of the most abundant proteins, p97/valosin-containing protein (VCP), a homohexameric enzyme, is a member of the type II AAA+ protein family of ATPases and consists of two AAA domains, D1 and D2, that are assembled in a head-to-tail manner, as well as an N-terminal domain that plays a role in substrate recognition [94–97]. In addition, the C-terminal domain of p97/VCP associates with a large number of adaptors, explaining the diversity of p97/VCP interacting partners [96]. p97/VCP has been demonstrated to be implicated in chromatin remodeling, autophagosome maturation, proteasome-mediated degradation, and ER membrane fusion [98, 99]. Importantly, p97/VCP is crucial for the clearance of misfolded proteins by affecting a large number of protein homeostatic mechanisms. p97/VCP couples ATP hydrolysis to unfolding of ERAD substrates and functions in the retrotranslocation of nearly all ERAD substrates, along with cofactors recruited through p97/VCP-binding domains, involving VIM, VBR, and SHP [94].

Several studies suggest that Derlins are part of the retrotranslocon channel [100, 101]. Mammalian cells have three Derlins, Derlin-1, Derlin-2, and Derlin-3. As a rhomboid-like protein, Derlin-1 has six membrane-spanning domains and homo- or heterooligomerizes with Derlin-2 and Derlin-3 [102–107]. Derlins are related to rhomboid proteases such as ER-resident intramembrane protein RHBDL4, which cleaves unstable single-membrane-spanning or polytopic membrane proteins [108]. However, Derlins are deficient in proteolytic activity, implicating that these proteins associate with ERAD substrates and target them to p97/VCP for retrotranslocation and to E3 ubiquitin ligases for ubiquitination [109].

Suppressor/enhancer of Lin12-like (SEL1L) recruits luminal substrate recognition factors, involving OS-9, XTP3-B, EDEMs, ERdj5, and PDI to components of the retrotranslocon [64, 110]. In addition, SEL1L serves as a scaffold for the formation of a complex with integral membrane ERAD components that include Derlin-1, Derlin-2, ancient ubiquitous protein 1 (AUP1), ubiquitin regulatory X (UBX) domain-containing protein 8 (UBXD8), and VCP-interacting membrane protein (VIMP) [85, 86, 111–115], which in turn recruits the p97/VCP, thereby leading to substrate retrotranslocation. Furthermore, SEL1L is not only required for the transfer of substrates from ER lectins to E3 ubiquitin ligase hydroxymethylglutaryl reductase degradation protein 1 (HRD1) but also crucial for the stabilization of HRD1, suggesting that SEL1L is important for ERAD substrate recruitment, retrotranslocation, and ubiquitination [85, 116–120].

Erlin1/2, heterotetrameric complex located in the ER membrane rapidly associates with inositol 1,4,5-triphosphate receptors (IP₃R) for activation of IP₃R and links IP₃R to the ER-resident E3 ubiquitin ligase RNF170, indicating its role in the degradation of membrane-integrated substrates [121].

3.3. Ubiquitination

Ubiquitination is a reversible process that conjugates ubiquitin to target proteins, which in most, but not all, cases leads to proteasome-mediated degradation of ubiquitinated proteins and is conserved in all eukaryotes. Ubiquitin is covalently attached to target proteins by a sequential enzymatic system consisting of E1 ubiquitin-activating, E2 ubiquitin-conjugating, and E3 ubiquitin-ligating enzymes [13, 122]. Additionally, removal of ubiquitin catalyzed by deubiquitinating enzymes also plays key roles in the control of numerous biological pathways [123]. In the initial step of ubiquitination, an E1 ubiquitin-activating enzyme activates ubiquitin and forms a thioester bond with ubiquitin. In the next step, an E2 ubiquitin-conjugating enzyme transfers the ubiquitin from the E1 to the target protein, which is assisted by an E3 ubiquitin ligase. Ubiquitin is normally conjugated via its C-terminus to lysine or, in some cases, to serine, threonine, or cysteine residues on the target proteins [63, 124–126]. Once ubiquitinated, ubiquitin can be further extended by the additional ubiquitin moieties on one of the lysine residues within ubiquitin, involving K6, K11, K27, K29, K33, K48, and K63 or its N-terminus [127–130]. The linkages of ubiquitin chains confer diverse structural properties to ubiquitin chains, providing a different binding platform for various processes.

In mammalian cells, more than a dozen E3 ubiquitin ligases have been demonstrated to be involved in ERAD. Several ERAD E3 ubiquitin ligases are transmembrane proteins, involving HRD1, glycoprotein 78 (gp78), membrane-associated RING (really interesting new gene) finger protein 6 (MARCH6), and RNF5 [63, 131–138]. In addition, cytoplasmic E3 ubiquitin ligases involving parkin, CHIP, SCF complexes with the F-box proteins Fbx2, Fbx6, and β-TrCP1/2, Smurf1, and Nrdp1/FLRF have been demonstrated to be involved in ERAD [139–145]. ERAD E3 ubiquitin ligases accomplish ERAD substrate processing in parallel with multiple E3 ubiquitin ligases, by conjugating ubiquitin to different sites of a substrate at the same time, by an initial monoubiquitination and extension by E4 ubiquitin ligase, or via sequential rounds of ubiquitination and deubiquitination, suggesting that various strategies have been evolved for optimal efficiency of ERAD [136, 146, 147]. For instance, gp78 and Trc8 cooperate as E3 ubiquitin ligase pairs to degrade HMG-CoAR [146]. RNF5 functions sequentially with CHIP to degrade misfolded CFTRΔF508 and also serves as a primer for gp78-mediated chain elongation [135, 136]. In addition to the ubiquitination of ERAD substrates, ERAD E3 ubiquitin ligases may ubiquitinate other ERAD components to recruit p97/VCP or other ERAD components that possess ubiquitin-binding domains (UBDs), involving gp78, AUP1, ubiquitin-associated- (UBA-) domain-containing protein 2 (UBAC2), and UBX domain-containing protein 8 (UBXD8) [63, 148, 149]. Interestingly, ERAD E3 ubiquitin ligases ubiquitinate each other and form a negative feedback loop, thereby leading to fine-tuning of ERAD [64, 136, 146, 150, 151].

The ER membrane-embedded E3 ubiquitin ligases comprise a part of the retrotranslocon, and inhibiting ubiquitination attenuates retrotranslocation of ERAD substrates, suggesting that retrotranslocation is tightly coupled with ubiquitination [109]. Derlin-1 and Derlin-2 are closely linked to E3 ubiquitin ligases such as HRD1, gp78, and RNF5 to form huge complexes spanning the ER membrane [103, 104, 115, 135, 152–154]. Additionally, it is speculated that Hrd1p in yeast functions as an essential part of retrotranslocon with its cofactors, thereby recruiting ERAD substrates and in turn promoting their retrotranslocation from the ER [155].

Most of the p97/VCP cofactors possess UBDs and associate directly with ubiquitinated substrates. p97/VCP and its cofactors, Npl4 and Ufd1, cooperatively produce a driving force for the retrotranslocation of ERAD substrates [156, 157]. The ERAD substrate is slightly exposed to the ER surface through the retrotranslocon, which in turn is subject to E3 ubiquitin ligase-mediated polyubiquitination, and further retrotranslocated by the p97/Npl4/Ufd1 complex, which can recognize the polyubiquitinated substrate, suggesting that polyubiquitination serves as a binding site that promotes p97/VCP-mediated substrate extraction. To summarize, membrane-embedded ERAD components such as UBXD2, UBXD8, and VIMP, ERAD E3 ubiquitin ligases, such as gp78 and HRD1, and Derlins have p97/VCP-binding motifs, implicating that p97/VCP provides a platform for these factors to regulate ubiquitination at the sites of retrotranslocation [63, 102, 104, 158–160].

3.4. Proteasome-Mediated Degradation

p97/VCP is also closely linked to proteasome-mediated degradation of ERAD substrates [94]. p97/VCP plays a key role in linking retrotranslocated substrates to cytoplasmic cofactors involved in further processing of substrates. The deglycosylating enzyme NGly1 localized in the cytoplasm is recruited to retrotranslocon complexes through direct binding to p97/VCP and cleaves N-linked glycans from retrotranslocated ERAD substrates [161, 162]. In addition, deubiquitinating enzymes (DUBs), involving YOD1 (OTUD2), VCIP135, USP13, and Ataxin-3, associate with p97/VCP either directly or indirectly and are implicated in ERAD [163–165]. Recently, it is demonstrated that impairment of p97/VCP-associated deubiquitination or expression of dominant-negative YOD1 attenuates retrotranslocation and degradation of ERAD substrates, whereas expression of p97/VCP-associating DUB restores them [163], indicating that sequential rounds of ubiquitination and deubiquitination are essential for efficient ERAD process.

Retrotranslocated substrates need to be rapidly degraded to prevent misfolded proteins from aggregating in the cytoplasm. A chaperone complex consisting of Bag6-Ubl4A-Trc35 and a cochaperone SGTA is involved in this process. Bag6 is a cytosolic chaperone and forms a large homooligomer through proline-rich domain. The proline-rich domain is sufficient for binding to the hydrophobic segments of misfolded proteins and maintaining them in a soluble state [166]. The holdase activity of Bag6 is required to maintain some retrotranslocated substrates in a competent state for proteasome-mediated degradation [167]. Ubl4A, an adaptor of Bag6, associates with SGTA via its noncanonical ubiquitin-like (UBL) domain [168]. Bag6 also associates with proteasome and adaptor proteins of proteasome, suggesting that Bag6 transfers retrotranslocated substrates to proteasome for degradation.

4. UPR of the ER and Cancer

UPR of the ER has been demonstrated in diverse human cancers. In fact, it has been documented that UPR of the ER plays a crucial role in the control of tumor progression and affects tumor microenvironment involving immune cells and endothelial cells [6]. UPR of the ER modulates the expression and/or the function of oncogenes or tumor-suppressive genes in cancer, which leads to an increase in protein synthesis, resulting in an increased necessity of protein-folding capacity of the ER and subsequent activation of UPR to improve the adaptive capacity of the ER. However, persistent activation of UPR consequently affects cancer cell survival, metastasis, angiogenesis, immunogenicity, and drug resistance [6].

4.1. UPR of the ER and Tumorigenesis

During malignant transformation, tumor cells are exposed to not only extrinsic stresses such as nutrient deprivation, accumulation of acidic waste, and hypoxia but also intrinsic stresses such as alteration in chromosome number, activation of oncogenes, inactivation of tumor-suppressive genes, and accelerated secretion, thereby triggering exacerbated protein synthesis, which results in a cellular state of ER stress and subsequently activates UPR of the ER [169–172]. It is also suggested that chronic UPR at later stages leads to adaptation of tumor to extrinsic and intrinsic perturbations and confers resistance to ER stress-induced apoptosis on tumor, while transient UPR at early stages of tumorigenesis often impedes tumor progression [173].

Oncogenic RAS-mediated transformation of melanocytes activates UPR, which induces cell cycle arrest coupled with vacuolization and ER expansion, resulting in premature senescence [174]. In models of RET-induced fibroblast transformation, UPR activation plays a protective role against oncogene-induced malignant progression through the proapoptotic CHOP pathway [175]. In a model of KRAS-transformed lung tumor, high caloric diet-induced ER stress hinders tumor growth [176]. In addition, depletion of XBP1s is known to promote tumorigenesis, suggesting that UPR of the ER may play a tumor-suppressive role [177].

PERK has been described in the initiation and progression of various tumors. Depletion of PERK leads to tumor progression [178, 179]. ATF4-CHOP axis in PERK pathway promotes protein synthesis and in turn accelerates ROS production from oxidative protein folding in the ER. The treatment of antioxidant and depletion of RPL24 reduced apoptosis by decreasing ROS production and protein synthesis, indicating that PERK is involved in tumor regression [30]. In contrast, PERK facilitates tumor growth through the stabilization of NRF2, the modulation of redox homeostasis as well as of metabolism, and the regulation of lipid biosynthesis [178, 180–184]. Intriguingly, eIF2α phosphorylation by PERK facilitates LC3 lipidation, autophagy initiation, and subsequent survival [185]. Additionally, it is also demonstrated that ATF4-CHOP axis induces the expression of numerous genes involved in autophagophore formation and maturation, including Atg5, Atg12, Atg16l1, and Becn1 [186].

IRE1 is also involved in tumor progression. JNK activation by IRE1 suppresses antiapoptotic BCL2 activity and accelerates the action of proapoptotic BIM, leading to cell death [6]. In addition, IRE1-dependent decay of mRNA (RIDD) activates proapoptotic caspase-2 in MEFs and facilitates the expression of gene encoding thioredoxin-interacting protein (Txnip) in pancreatic β cells [48, 49]. On the contrary, IRE1-mediated activation of STAT3 and NF-κB upregulates the expression of antiapoptotic proteins, involving BCL2 family members, caspase-8 inhibitor c-FLIP, MCL1, and inhibitor of apoptosis protein (IAP) [187]. IRE1-XBP1 axis is also demonstrated to correlate with poor prognosis in glioblastoma and pre-B acute lymphoblastic leukemia [188–192]. Additionally, mutated forms of IRE1 facilitate tumor progression, although some of these mutants have intact kinase and endoribonuclease activity [47, 193, 194].

ATF6-dependent p58 (IPK) restricts apoptosis during oncogenic transformation via the inhibition of PERK [175]. ATF6 also facilitates the survival of glucose restriction-resistant squamous carcinoma cells [195].

In order to provide sufficient oxygen and nutrients, growing cancer cells produce proangiogenic factors to initiate vascularization. Several studies indicate that UPR facilitates angiogenesis. PERK upregulates the expression of the vessel growth and stabilization factors VCIP and PDGFRB [179]. In addition, PERK-mediated upregulation of fibroblast growth factor 2 (FGF2), vascular endothelial growth factor (VEGF), and interleukin-6 (IL-6) and downregulation of antiangiogenic cytokines remarkably promote tumor growth and vascularization [196]. IRE1-XBP1 axis also facilitates angiogenesis via the association of XBP1s with hypoxia-inducing factor 1α (HIF1α), a key regulator of VEGF in triple negative breast cancer (TNBC) cells [197]. Intriguingly, VEGF signaling also activates UPR in endothelial cells through a phospholipase Cγ-mTORC1 pathway, indicating that VEGF signaling and UPR may operate a positive feedback loop for angiogenesis [198].

UPR of the ER has begun to be elucidated in metastasis. Metastasis is a complicated process in which cancer cells migrate from the original tumor site, infiltrate extracellular matrix (ECM) and stromal cell layers, penetrate the lymphatic circulatory systems, colonize foreign tissues, and grow into new tumor mass [172, 199–201]. PERK-ATF4 axis activates lysosome-associated membrane protein 3 (LAMP3), thereby facilitating metastasis of hypoxic breast cancer cells [202, 203]. The upregulation of ATF4 in esophageal squamous carcinoma leads to an increase in metastasis through the regulation of matrix metalloproteinases [204]. Intriguingly, ATF4-mediated gene expression is potentially correlated with the expression of genes involved in epithelial-to-mesenchymal transition (EMT) [199].

IRE1-XBP1 axis is also implicated in metastasis. TNBC cell lines constitutively express XBP1s, and silencing of Xbp1 potentially inhibits metastasis [197]. XBP1s drives TNBC tumorigenicity and invasiveness by assembling a transcriptional complex with HIF1α that upregulates the expression of HIF1α targets such as PDK1 and GLUT1. On the contrary, while inhibition of IRE1 in malignant glioma correlates with the downregulation of proangiogenic factors such as VEGF-A, IL-1β, IL-6, and IL-8, it induces a significant upregulation of proteins linked to mesenchymal differentiation and glioma invasiveness such as SPARC, decorin, and thrombospondin-1, demonstrating that IRE1 in malignant glioma promotes the formation of functional tumor blood vessels and attenuates tumor cell invasion as well as vessel cooption [205, 206]. Therefore, a comprehensive analysis of IRE1-XBP1 axis is required to determine the relationship between invasiveness and angiogenesis. Additionally, the different consequences of UPR activation likely result from an interplay between particular axes of signaling pathways within specific tumor contexts.

4.2. UPR of the ER and Cancer Immunogenicity

It is of great importance to explore the crosstalk between UPR of the ER in tumor cells, the release of damage-associated molecules, and the activation of immune responses for the understanding of anti-tumor immunity. Through a process “transmissible ER stress,” ER stress enables cancer cells to secrete some factors that promote macrophage activation and induce a proinflammatory response in the microenvironment of tumors [207]. This process represses the antigen-presenting capacity of bone-marrow-derived dendritic cells (DCs) and inhibits T cell proliferation, which promotes the upregulation of immunosuppressive molecules [208], suggesting that ER stress signaling may facilitate immune escape. On the contrary, ER stress also triggers immunogenic cell death (ICD) and antitumor immunity [209]. The ICD provokes release of damage-associated molecular patterns (DAMPs), involving surface exposure of calreticulin, ATP secretion, and passive release of high-mobility group box 1 (HMGB1), suggesting that DAMPs serve as signals of danger and facilitate antitumor immunity [173, 210, 211]. PERK-eIF2α axis is associated with the exposure of calreticulin in non-small-cell lung carcinoma (NSCLC) and is correlated with ICD and antitumor immunity [212]. Photodynamic therapy increases the surface exposure of calreticulin as well as ATP secretion via PERK signaling in human bladder carcinoma, leading to engulfment of cancer cells by dendritic cells (DCs) [213]. In addition, radiation and anthracycline treatment induce lethal ER stress characterized by ROS production, an increase in the level of cytosolic Ca²⁺, and the excessive activation of UPR, thereby leading to the activation of inflammasome and subsequent ICD [48, 214]. However, IRE1-XBP1 axis is demonstrated to prevent the induction of ICD in metastatic colorectal cancer cells exposed to chemotherapy [215].

The tumor microenvironment is a complex environment consisting of stromal cells such as fibroblasts and endothelial cells and infiltrating immune cells such as CD8 T cells, Tregs, myeloid-derived suppressor cells (MDSCs), and DCs. Recently, it has begun to emerge as a new research area to elucidate the relationship between ER stress response in tumor-associated immune cells and tumor progression [199]. IRE1-XBP1 axis is essential for the differentiation of plasma cells and some dendritic cells [216–218]. ER stress response driven by XBP1 hyperactivation promotes neutrophil-infiltrating acute lung injury [219]. Additionally, XBP1 is required for the production of IL-6 in macrophages [220]. Persistent activation of IRE1-XBP1 axis is found in ovarian tumor-infiltrating DCs [221]. Intriguingly, the ovarian tumor-infiltrating DCs facilitate ROS production and consequential disruption of ER homeostasis, thereby leading to the control of antitumor immunity. In addition, the status of ROS-promoted lipid peroxidation has been suggested as a biomarker of disease recurrence in breast cancer patients [222]. Consistently, tumor-infiltrating DCs lacking XBP1 acquire immunostimulatory and antitumoral characteristics in vivo [223–225]. Pharmacological inhibition of IRE1 in bone-marrow-derived macrophages stimulated by IL-6 and IL-4 attenuates macrophage-mediated cell invasion in vitro [226]. Interestingly, IL-4 and IL-6 synergistically activate IRE1-XBP1 axis in macrophages. In addition, pharmacological induction of ER stress triggers the upregulation of the lectin-type oxidized LDL receptor-1 (LOX-1) in neutrophils and can induce transformation of neutrophils into immunosuppressive cells [227, 228]. These studies suggest that IRE1-XBP1 axis plays a key role in the control of tumor-associated myeloid cells.

CHOP is known to be upregulated in tumor-infiltrating MDSCs [229]. Tumor-infiltrating MDSCs devoid of CHOP show reduced immunosuppressive activity toward T cells due to defective expression of arginase. In addition, UPR activation in tumor-infiltrating MDSCs promotes apoptosis through death receptor 5 (DR5) and caspase-8 activation [230]. To summarize, these findings suggest that UPR activation plays a pivotal role in fine-tuning of tumor-associated immune responses.

4.3. UPR of the ER and Therapies for Cancer

UPR-activating or inhibiting strategies have begun to emerge as new pharmacological tools for cancer treatment. A large number of anticancer drugs induce UPR activation and facilitate the development of chemosensitivity or chemoresistance in a context-dependent manner [199]. Anticancer drugs involving paclitaxel, the epidermal growth factor receptor (EGFR) inhibitor cetuximab, and the BRAF (V600E) inhibitor vemurafenib induce PERK-mediated eIF2α phosphorylation [137, 215, 231]. In addition, nonsteroidal anti-inflammatory drugs promote UPR-mediated apoptosis and are used in combined anticancer therapies [232, 233].

Recently, targeting of UPR of the ER in cancer cells has been demonstrated to inhibit survival or promote cell death [173, 234, 235]. Therapies targeting UPR are applied for multiple myeloma or B cell-associated hematologic malignancies [173]. In addition, inhibition of PERK-eIF2α axis promotes cell death of therapy-resistant hypoxic glioblastoma and colon carcinoma cells [236], suggesting that combination of cancer therapy and UPR targeting may be desirable for cancer treatment.

ER stress-induced autophagy may facilitate therapeutic resistance. Autophagy induced by IRE1-JNK pathway facilitates sorafenib resistance in hepatocellular carcinoma cell lines [237, 238]. It is also demonstrated that a marked increase in cytoprotective autophagy induced by PERK develops vemurafenib resistance in melanoma [231]. Intriguingly, simultaneous inhibition of BRAF (V600E) and PERK sensitizes chemoresistant melanoma to ER stress-induced apoptosis, suggesting that the balance between autophagy mediated by UPR and chemotherapy is required for the overcome of chemoresistance.

Because ER stress modulates the functions of tumor-associated immune cells and subsequently protumoral or antitumor immune responses, it is noteworthy to consider UPR-targeting therapies in immune cells. Targeting IRE1-XBP axis in DCs has been demonstrated to be effective for cancer treatment [221, 239, 240]. Depletion of Xbp1 or silencing of Ire1 in preclinical models remarkably transforms DCs into immunostimulatory cells, thereby promoting survival through T-cell-mediated antitumor immunity [221]. In addition, transplanted MDSCs devoid of DNA damage-inducible transcript 3 (Ddit3) show enhanced antigen-presenting capacity and T cell stimulatory effects [229]. Intriguingly, ERO1α is demonstrated to upregulate programmed death ligand 1 (PDL1) in TNBCs [241]. These studies suggest that the combination of UPR-modulating therapies with immunotherapies might be effective for cancer treatment.

5. ERAD and Cancer

ERAD in tumor progression and immunogenicity is not well known. Several studies suggest that the high degree of cell division and high mutation rates in cancer cells lead to an accumulation of misfolded proteins, which activates ERAD [242, 243]. Higher expression of SEL1L in pancreatic cancer cells leads to not only G1 phase cell cycle arrest via the induction of a phosphatase and tensin homolog (PTEN) but also reduction in invasiveness by modulating genes related to cell-matrix interactions [244, 245]. Additionally, low expression of SEL1L in breast cancer patients has been reported to correlate with poor prognosis [246]. However, in the context of colorectal cancer, while basal expression of SEL1L in normal mucosa of the epithelial lining is low, SEL1L is upregulated in adenoma and adenocarcinoma cells [247]. Therefore, it is pivotal to elucidate the mechanism by which SEL1L is involved in tumor progression within a specific tumor context and the effects of changes in SEL1L expression in tumor cells on ERAD substrates or ER homeostasis to clarify the role of SEL1L in cancer pathogenesis.

Under hypoxic conditions, OS-9-mediated degradation of HIF1α is crucial in the downregulation of genes that promote cell survival, proliferation, angiogenesis, and metastasis [248–250], suggesting that OS-9 is important in the regulation of tumor progression.

gp78 induces a signaling cascade to mediate tumorigenesis and is linked with various types of cancers [251–253]. gp78 is highly expressed in bladder carcinoma tissues, and colorectal cancer patients with higher expression of gp78 have less survival and high risk of cancer recurrence, suggesting that gp78 is closely related to increased risk of cancer with lower survival rate [254–256].

The role of gp78 in metastasis is largely unknown. However, several studies suggest the involvement of gp78 in metastasis. Expression of gp78 is modulated by cell-cell contact, and loss of this balance is linked to metastasis [257]. Intriguingly, an inverse correlation between gp78 and E-cadherin has been reported in patients with bladder carcinomas as well as gastric cancers [258–260]. gp78-mediated degradation of metastasis suppressor protein Kangai1 (KAI1) promotes metastasis [261, 262]. Additionally, gp78 activates ROCK2, an important metastasis-associated protein, indicating the involvement of gp78 in metastasis [263].

CHIP, cytosolic E3 ubiquitin ligase involved in ERAD is inversely correlated with malignancy in breast cancers and depletion of CHIP results in an increase in the growth of subcutaneous tumors, indicating its role as a tumor suppressor [264].

6. Conclusion

Organisms are continuously exposed to extrinsic and intrinsic stresses that destroy proteostasis and subsequently result in protein misfolding and aggregation, thereby leading to the state of ER stress. To restore proteostasis, eukaryotic cells have evolved UPR of the ER and ERAD as key adaptive responses. Intriguingly, a failure in these adaptive responses leads to various protein misfolding diseases, involving cancer. UPR of the ER not only acts as a guardian of tumor progression at an early stage but also serves as a key player for maintenance of tumors under chronic ER stress. Additionally, UPR has been described to manipulate immune cells in tumor microenvironment, resulting in antitumor immunity or immune escape. Importantly, UPR activation also develops chemosensitivity or chemoresistance in a context-dependent manner. Overall, UPR of the ER is involved in the modulation of tumor growth, metastasis, and angiogenesis; the interaction of tumor and stromal cells; and the regulation of inflammatory/immune responses. Therefore, to elucidate the precise molecular mechanisms by which UPR of the ER and ERAD coordinate tumor progression at different stages and modulate the communication between tumor and tumor microenvironment remarkably contributes to novel therapeutic interventions.

Conflicts of Interest

The authors have declared that no conflict of interest exists.

Acknowledgments

This work was supported by a grant from the National Research Foundation of Korea (NRF-2016R1A2B4006566). Yoon Seon Yoo and Hye Gyeong Han were the recipients of the BK21 Plus fellowship.

References

G. Griffiths, G. Warren, P. Quinn, O. Mathieu-Costello, and H. Hoppeler, “Density of newly synthesized plasma membrane proteins in intracellular membranes. I. Stereological studies,” Journal of Cell Biology, vol. 98, no. 6, pp. 2133–2141, 1984.

O. Baumann and B. Walz, “Endoplasmic reticulum of animal cells and its organization into structural and functional domains,” International Review of Cytology, vol. 205, pp. 149–214, 2001.

D. T. Rutkowski and R. S. Hegde, “Regulation of basal cellular physiology by the homeostatic unfolded protein response,” The Journal of Cell Biology, vol. 189, no. 5, pp. 783–794, 2010.

M. J. Berridge, P. Lipp, and M. D. Bootman, “The versatility and universality of calcium signalling,” Nature Reviews Molecular Cell Biology, vol. 1, no. 1, pp. 11–21, 2000.

S. A. Oakes and F. R. Papa, “The role of endoplasmic reticulum stress in human pathology,” Annual Review of Pathology: Mechanisms of Disease, vol. 10, no. 1, pp. 173–194, 2015.

M. Wang and R. J. Kaufman, “The impact of the endoplasmic reticulum protein-folding environment on cancer development,” Nature Reviews Cancer, vol. 14, no. 9, pp. 581–597, 2014.

D. Morito and K. Nagata, “Pathogenic hijacking of ER-associated degradation: is ERAD flexible?” Molecular Cell, vol. 59, no. 3, pp. 335–344, 2015.

F. U. Hartl and M. Hayer-Hartl, “Converging concepts of protein folding in vitro and in vivo,” Nature Structural & Molecular Biology, vol. 16, no. 6, pp. 574–581, 2009.

A. Ruggiano, O. Foresti, and P. Carvalho, “Quality control: ER-associated degradation: protein quality control and beyond,” Journal of Cell Biology, vol. 204, no. 6, pp. 869–879, 2014.

J. L. Brodsky and R. J. Wojcikiewicz, “Substrate-specific mediators of ER associated degradation (ERAD),” Current Opinion in Cell Biology, vol. 21, no. 4, pp. 516–521, 2009.

R. Y. Hampton, “ER-associated degradation in protein quality control and cellular regulation,” Current Opinion in Cell Biology, vol. 14, no. 4, pp. 476–482, 2002.

S. S. Vembar and J. L. Brodsky, “One step at a time: endoplasmic reticulum-associated degradation,” Nature Reviews Molecular Cell Biology, vol. 9, no. 12, pp. 944–957, 2008.

A. Hershko and A. Ciechanover, “The ubiquitin system,” Annual Review of Biochemistry, vol. 67, no. 1, pp. 425–479, 1998.

C. Lopez-Otin, M. A. Blasco, L. Partridge, M. Serrano, and G. Kroemer, “The hallmarks of aging,” Cell, vol. 153, no. 6, pp. 1194–1217, 2013.

R. C. Taylor and A. Dillin, “Aging as an event of proteostasis collapse,” Cold Spring Harbor Perspectives in Biology, vol. 3, no. 5, pp. 1–17, 2011.

F. Chiti and C. M. Dobson, “Protein misfolding, functional amyloid, and human disease,” Annual Review of Biochemistry, vol. 75, no. 1, pp. 333–366, 2006.

M. Wang and R. J. Kaufman, “Protein misfolding in the endoplasmic reticulum as a conduit to human disease,” Nature, vol. 529, no. 7586, pp. 326–335, 2016.

T. K. Rainbolt, J. M. Saunders, and R. L. Wiseman, “Stress-responsive regulation of mitochondria through the ER unfolded protein response,” Trends in Endocrinology & Metabolism, vol. 25, no. 10, pp. 528–537, 2014.

N. Hiramatsu, W. C. Chiang, T. D. Kurt, C. J. Sigurdson, and J. H. Lin, “Multiple mechanisms of unfolded protein response-induced cell death,” The American Journal of Pathology, vol. 185, no. 7, pp. 1800–1808, 2015.

J. H. Otero, B. Lizak, and L. M. Hendershot, “Life and death of a BiP substrate,” Seminars in Cell & Developmental Biology, vol. 21, no. 5, pp. 472–478, 2010.

A. Bertolotti, Y. Zhang, L. M. Hendershot, H. P. Harding, and D. Ron, “Dynamic interaction of BiP and ER stress transducers in the unfolded-protein response,” Nature Cell Biology, vol. 2, no. 6, pp. 326–332, 2000.

J. Shen, E. L. Snapp, J. Lippincott-Schwartz, and R. Prywes, “Stable binding of ATF6 to BiP in the endoplasmic reticulum stress response,” Molecular and Cellular Biology, vol. 25, no. 3, pp. 921–932, 2005.

D. Pincus, M. W. Chevalier, T. Aragon et al., “BiP binding to the ER-stress sensor Ire1 tunes the homeostatic behavior of the unfolded protein response,” PLoS Biology, vol. 8, no. 7, article e1000415, 2010.

B. M. Gardner and P. Walter, “Unfolded proteins are Ire1-activating ligands that directly induce the unfolded protein response,” Science, vol. 333, no. 6051, pp. 1891–1894, 2011.

A. V. Korennykh, P. F. Egea, A. A. Korostelev et al., “The unfolded protein response signals through high-order assembly of Ire1,” Nature, vol. 457, no. 7230, pp. 687–693, 2008.

H. Li, A. V. Korennykh, S. L. Behrman, and P. Walter, “Mammalian endoplasmic reticulum stress sensor IRE1 signals by dynamic clustering,” Proceedings of the National Academy of Sciences of the United States of America, vol. 107, no. 37, pp. 16113–16118, 2010.

P. Walter and D. Ron, “The unfolded protein response: from stress pathway to homeostatic regulation,” Science, vol. 334, no. 6059, pp. 1081–1086, 2011.

D. Ron and P. Walter, “Signal integration in the endoplasmic reticulum unfolded protein response,” Nature Reviews Molecular Cell Biology, vol. 8, no. 7, pp. 519–529, 2007.

G. V. Di Prisco, W. Huang, S. A. Buffington et al., “Translational control of mGluR-dependent long-term depression and object-place learning by eIF2α,” Nature Neuroscience, vol. 17, no. 8, pp. 1073–1082, 2014.

J. Han, S. H. Back, J. Hur et al., “ER-stress-induced transcriptional regulation increases protein synthesis leading to cell death,” Nature Cell Biology, vol. 15, no. 5, pp. 481–490, 2013.

H. P. Harding, Y. Zhang, D. Scheuner, J. J. Chen, R. J. Kaufman, and D. Ron, “Ppp1r15 gene knockout reveals an essential role for translation initiation factor 2 alpha (eIF2α) dephosphorylation in mammalian development,” Proceedings of the National Academy of Sciences of the United States of America, vol. 106, no. 6, pp. 1832–1837, 2009.

D. T. Rutkowski, S. M. Arnold, C. N. Miller et al., “Adaptation to ER stress is mediated by differential stabilities of pro-survival and pro-apoptotic mRNAs and proteins,” PLoS Biology, vol. 4, no. 11, article e374, 2006.

J. H. Lin, H. Li, D. Yasumura et al., “IRE1 signaling affects cell fate during the unfolded protein response,” Science, vol. 318, no. 5852, pp. 944–949, 2007.

H. P. Harding, Y. Zhang, and D. Ron, “Protein translation and folding are coupled by an endoplasmic-reticulum-resident kinase,” Nature, vol. 397, no. 6716, pp. 271–274, 1999.

K. M. Vattem and R. C. Wek, “Reinitiation involving upstream ORFs regulates ATF4 mRNA translation in mammalian cells,” Proceedings of the National Academy of Sciences of the United States of America, vol. 101, no. 31, pp. 11269–11274, 2004.

H. P. Harding, Y. Zhang, H. Zeng et al., “An integrated stress response regulates amino acid metabolism and resistance to oxidative stress,” Molecular Cell, vol. 11, no. 3, pp. 619–633, 2003.

S. J. Marciniak, C. Y. Yun, S. Oyadomari et al., “CHOP induces death by promoting protein synthesis and oxidation in the stressed endoplasmic reticulum,” Genes & Development, vol. 18, no. 24, pp. 3066–3077, 2004.

B. Song, D. Scheuner, D. Ron, S. Pennathur, and R. J. Kaufman, “Chop deletion reduces oxidative stress, improves β cell function, and promotes cell survival in multiple mouse models of diabetes,” The Journal of Clinical Investigation, vol. 118, no. 10, pp. 3378–3389, 2008.

N. S. Chitnis, D. Pytel, E. Bobrovnikova-Marjon et al., “miR-211 is a prosurvival microRNA that regulates chop expression in a PERK-dependent manner,” Molecular Cell, vol. 48, no. 3, pp. 353–364, 2012.

C. W. Woo, L. Kutzler, S. R. Kimball, and I. Tabas, “Toll-like receptor activation suppresses ER stress factor CHOP and translation inhibition through activation of eIF2B,” Nature Cell Biology, vol. 14, no. 2, pp. 192–200, 2012.

W. Tirasophon, A. A. Welihinda, and R. J. Kaufman, “A stress response pathway from the endoplasmic reticulum to the nucleus requires a novel bifunctional protein kinase/endoribonuclease (Ire1p) in mammalian cells,” Genes & Development, vol. 12, no. 12, pp. 1812–1824, 1998.

H. Yoshida, T. Matsui, A. Yamamoto, T. Okada, and K. Mori, “XBP1 mRNA is induced by ATF6 and spliced by IRE1 in response to ER stress to produce a highly active transcription factor,” Cell, vol. 107, no. 7, pp. 881–891, 2001.

A. H. Lee, N. N. Iwakoshi, and L. H. Glimcher, “XBP-1 regulates a subset of endoplasmic reticulum resident chaperone genes in the unfolded protein response,” Molecular and Cellular Biology, vol. 23, no. 21, pp. 7448–7459, 2003.

J. R. Hassler, D. L. Scheuner, S. Wang et al., “The IRE1α/XBP1s pathway is essential for the glucose response and protection of β cells,” PLoS Biology, vol. 13, no. 10, article e1002277, 2015.

M. Calfon, H. Zeng, F. Urano et al., “IRE1 couples endoplasmic reticulum load to secretory capacity by processing the XBP-1 mRNA,” Nature, vol. 415, no. 6867, pp. 92–96, 2002.

H. Yoshida, M. Oku, M. Suzuki, and K. Mori, “pXBP1(U) encoded in XBP1 pre-mRNA negatively regulates unfolded protein response activator pXBP1(S) in mammalian ER stress response,” Journal of Cell Biology, vol. 172, no. 4, pp. 565–575, 2006.

R. Ghosh, L. Wang, E. S. Wang et al., “Allosteric inhibition of the IRE1α RNase preserves cell viability and function during endoplasmic reticulum stress,” Cell, vol. 158, no. 3, pp. 534–548, 2014.

A. G. Lerner, J. P. Upton, P. V. Praveen et al., “IRE1α induces thioredoxin-interacting protein to activate the NLRP3 inflammasome and promote programmed cell death under irremediable ER stress,” Cell Metabolism, vol. 16, no. 2, pp. 250–264, 2012.

J. P. Upton, L. Wang, D. Han et al., “IRE1α cleaves select microRNAs during ER stress to derepress translation of proapoptotic caspase-2,” Science, vol. 338, no. 6108, pp. 818–822, 2012.

K. Haze, H. Yoshida, H. Yanagi, T. Yura, and K. Mori, “Mammalian transcription factor ATF6 is synthesized as a transmembrane protein and activated by proteolysis in response to endoplasmic reticulum stress,” Molecular Biology of the Cell, vol. 10, no. 11, pp. 3787–3799, 1999.

K. Lee, W. Tirasophon, X. Shen et al., “IRE1-mediated unconventional mRNA splicing and S2P-mediated ATF6 cleavage merge to regulate XBP1 in signaling the unfolded protein response,” Genes & Development, vol. 16, no. 4, pp. 452–466, 2002.

K. Yamamoto, T. Sato, T. Matsui et al., “Transcriptional induction of mammalian ER quality control proteins is mediated by single or combined action of ATF6α and XBP1,” Developmental Cell, vol. 13, no. 3, pp. 365–376, 2007.

M. D. Shoulders, L. M. Ryno, J. C. Genereux et al., “Stress-independent activation of XBP1s and/or ATF6 reveals three functionally diverse ER proteostasis environments,” Cell Reports, vol. 3, no. 4, pp. 1279–1292, 2013.

K. E. Matlack, W. Mothes, and T. A. Rapoport, “Protein translocation: tunnel vision,” Cell, vol. 92, no. 3, pp. 381–390, 1998.

A. Helenius, T. Marquardt, and I. Braakman, “The endoplasmic reticulum as a protein-folding compartment,” Trends in Cell Biology, vol. 2, no. 8, pp. 227–231, 1992.

A. Helenius, E. S. Trombetta, D. N. Hebert, and J. F. Simons, “Calnexin, calreticulin and the folding of glycoproteins,” Trends in Cell Biology, vol. 7, no. 5, pp. 193–200, 1997.

L. Ellgaard, M. Molinari, and A. Helenius, “Setting the standards: quality control in the secretory pathway,” Science, vol. 286, no. 5446, pp. 1882–1888, 1999.

A. Zapun, C. A. Jakob, D. Y. Thomas, and J. J. Bergeron, “Protein folding in a specialized compartment: the endoplasmic reticulum,” Structure, vol. 7, no. 8, pp. R173–R182, 1999.

C. S. Sevier and C. A. Kaiser, “Formation and transfer of disulphide bonds in living cells,” Nature Reviews Molecular Cell Biology, vol. 3, no. 11, pp. 836–847, 2002.

B. P. Tu and J. S. Weissman, “Oxidative protein folding in eukaryotes: mechanisms and consequences,” Journal of Cell Biology, vol. 164, no. 3, pp. 341–346, 2004.

H. Saibil, “Chaperone machines for protein folding, unfolding and disaggregation,” Nature Reviews Molecular Cell Biology, vol. 14, no. 10, pp. 630–642, 2013.

M. Schroder and R. J. Kaufman, “The mammalian unfolded protein response,” Annual Review of Biochemistry, vol. 74, no. 1, pp. 739–789, 2005.

J. C. Christianson and Y. Ye, “Cleaning up in the endoplasmic reticulum: ubiquitin in charge,” Nature Structural & Molecular Biology, vol. 21, no. 4, pp. 325–335, 2014.

J. A. Olzmann, R. R. Kopito, and J. C. Christianson, “The mammalian endoplasmic reticulum-associated degradation system,” Cold Spring Harbor Perspectives in Biology, vol. 5, no. 9, pp. 1–16, 2013.

C. Xu and D. T. Ng, “Glycosylation-directed quality control of protein folding,” Nature Reviews Molecular Cell Biology, vol. 16, no. 12, pp. 742–752, 2015.

E. J. Wiertz, T. R. Jones, L. Sun, M. Bogyo, H. J. Geuze, and H. L. Ploegh, “The human cytomegalovirus US11 gene product dislocates MHC class I heavy chains from the endoplasmic reticulum to the cytosol,” Cell, vol. 84, no. 5, pp. 769–779, 1996.

K. Fujita, S. Omura, and J. Silver, “Rapid degradation of CD4 in cells expressing human immunodeficiency virus type 1 Env and Vpu is blocked by proteasome inhibitors,” Journal of General Virology, vol. 78, no. 3, pp. 619–625, 1997.

U. Schubert, L. C. Anton, I. Bacik et al., “CD4 glycoprotein degradation induced by human immunodeficiency virus type 1 Vpu protein requires the function of proteasomes and the ubiquitin-conjugating pathway,” Journal of Virology, vol. 72, no. 3, pp. 2280–2288, 1998.

N. Yagishita, K. Ohneda, T. Amano et al., “Essential role of synoviolin in embryogenesis,” Journal of Biological Chemistry, vol. 280, no. 9, pp. 7909–7916, 2005.

A. B. Francisco, R. Singh, S. Li et al., “Deficiency of suppressor enhancer Lin12 1 like (SEL1L) in mice leads to systemic endoplasmic reticulum stress and embryonic lethality,” Journal of Biological Chemistry, vol. 285, no. 18, pp. 13694–13703, 2010.

Y. Eura, H. Yanamoto, Y. Arai, T. Okuda, T. Miyata, and K. Kokame, “Derlin-1 deficiency is embryonic lethal, Derlin-3 deficiency appears normal, and Herp deficiency is intolerant to glucose load and ischemia in mice,” PLoS One, vol. 7, no. 3, article e34298, 2012.

C. J. Guerriero and J. L. Brodsky, “The delicate balance between secreted protein folding and endoplasmic reticulum-associated degradation in human physiology,” Physiological Reviews, vol. 92, no. 2, pp. 537–576, 2012.

A. Helenius and M. Aebi, “Roles of N-linked glycans in the endoplasmic reticulum,” Annual Review of Biochemistry, vol. 73, no. 1, pp. 1019–1049, 2004.

M. Aebi, R. Bernasconi, S. Clerc, and M. Molinari, “N-glycan structures: recognition and processing in the ER,” Trends in Biochemical Sciences, vol. 35, no. 2, pp. 74–82, 2010.

D. N. Hebert, R. Bernasconi, and M. Molinari, “ERAD substrates: which way out?” Seminars in Cell & Developmental Biology, vol. 21, no. 5, pp. 526–532, 2010.

G. Z. Lederkremer, “Glycoprotein folding, quality control and ER-associated degradation,” Current Opinion in Structural Biology, vol. 19, no. 5, pp. 515–523, 2009.

D. S. Gonzalez, K. Karaveg, A. S. Vandersall-Nairn, A. Lal, and K. W. Moremen, “Identification, expression, and characterization of a cDNA encoding human endoplasmic reticulum mannosidase I, the enzyme that catalyzes the first mannose trimming step in mammalian Asn-linked oligosaccharide biosynthesis,” Journal of Biological Chemistry, vol. 274, no. 30, pp. 21375–21386, 1999.

L. O. Tremblay and A. Herscovics, “Cloning and expression of a specific human alpha 1,2-mannosidase that trims Man9GlcNAc2 to Man8GlcNAc2 isomer B during N-glycan biosynthesis,” Glycobiology, vol. 9, no. 10, pp. 1073–1078, 1999.

S. Olivari, T. Cali, K. E. Salo, P. Paganetti, L. W. Ruddock, and M. Molinari, “EDEM1 regulates ER-associated degradation by accelerating de-mannosylation of folding-defective polypeptides and by inhibiting their covalent aggregation,” Biochemical and Biophysical Research Communications, vol. 349, no. 4, pp. 1278–1284, 2006.

N. Hosokawa, L. O. Tremblay, B. Sleno et al., “EDEM1 accelerates the trimming of alpha1,2-linked mannose on the C branch of N-glycans,” Glycobiology, vol. 20, no. 5, pp. 567–575, 2010.

K. Hirao, Y. Natsuka, T. Tamura et al., “EDEM3, a soluble EDEM homolog, enhances glycoprotein endoplasmic reticulum-associated degradation and mannose trimming,” Journal of Biological Chemistry, vol. 281, no. 14, pp. 9650–9658, 2006.

N. Hosokawa, Y. Kamiya, D. Kamiya, K. Kato, and K. Nagata, “Human OS-9, a lectin required for glycoprotein endoplasmic reticulum-associated degradation, recognizes mannose-trimmed N-glycans,” Journal of Biological Chemistry, vol. 284, no. 25, pp. 17061–17068, 2009.

N. Hosokawa, Z. You, L. O. Tremblay, K. Nagata, and A. Herscovics, “Stimulation of ERAD of misfolded null Hong Kong α1-antitrypsin by Golgi α1,2-mannosidases,” Biochemical and Biophysical Research Communications, vol. 362, no. 3, pp. 626–632, 2007.

R. Bernasconi, T. Pertel, J. Luban, and M. Molinari, “A dual task for the Xbp1-responsive OS-9 variants in the mammalian endoplasmic reticulum: inhibiting secretion of misfolded protein conformers and enhancing their disposal,” Journal of Biological Chemistry, vol. 283, no. 24, pp. 16446–16454, 2008.

J. C. Christianson, T. A. Shaler, R. E. Tyler, and R. R. Kopito, “OS-9 and GRP94 deliver mutant α1-antitrypsin to the Hrd1-SEL1L ubiquitin ligase complex for ERAD,” Nature Cell Biology, vol. 10, no. 3, pp. 272–282, 2008.

N. Hosokawa, I. Wada, K. Nagasawa, T. Moriyama, K. Okawa, and K. Nagata, “Human XTP3-B forms an endoplasmic reticulum quality control scaffold with the HRD1-SEL1L ubiquitin ligase complex and BiP,” Journal of Biological Chemistry, vol. 283, no. 30, pp. 20914–20924, 2008.

R. Bernasconi, C. Galli, V. Calanca, T. Nakajima, and M. Molinari, “Stringent requirement for HRD1, SEL1L, and OS-9/XTP3-B for disposal of ERAD-LS substrates,” Journal of Cell Biology, vol. 188, no. 2, pp. 223–235, 2010.

R. K. Plemper, S. Bohmler, J. Bordallo, T. Sommer, and D. H. Wolf, “Mutant analysis links the translocon and BiP to retrograde protein transport for ER degradation,” Nature, vol. 388, no. 6645, pp. 891–895, 1997.

R. Ushioda, J. Hoseki, and K. Nagata, “Glycosylation-independent ERAD pathway serves as a backup system under ER stress,” Molecular Biology of the Cell, vol. 24, no. 20, pp. 3155–3163, 2013.

M. Shenkman, B. Groisman, E. Ron, E. Avezov, L. M. Hendershot, and G. Z. Lederkremer, “A shared endoplasmic reticulum-associated degradation pathway involving the EDEM1 protein for glycosylated and nonglycosylated proteins,” Journal of Biological Chemistry, vol. 288, no. 4, pp. 2167–2178, 2013.

J. L. Brodsky and W. R. Skach, “Protein folding and quality control in the endoplasmic reticulum: recent lessons from yeast and mammalian cell systems,” Current Opinion in Cell Biology, vol. 23, no. 4, pp. 464–475, 2011.

S. Grubb, L. Guo, E. A. Fisher, and J. L. Brodsky, “Protein disulfide isomerases contribute differentially to the endoplasmic reticulum-associated degradation of apolipoprotein B and other substrates,” Molecular Biology of the Cell, vol. 23, no. 4, pp. 520–532, 2012.

R. Y. Hampton and T. Sommer, “Finding the will and the way of ERAD substrate retrotranslocation,” Current Opinion in Cell Biology, vol. 24, no. 4, pp. 460–466, 2012.

H. Meyer, M. Bug, and S. Bremer, “Emerging functions of the VCP/p97 AAA-ATPase in the ubiquitin system,” Nature Cell Biology, vol. 14, no. 2, pp. 117–123, 2012.

T. Huyton, V. E. Pye, L. C. Briggs et al., “The crystal structure of murine p97/VCP at 3.6A,” Journal of Structural Biology, vol. 144, no. 3, pp. 337–348, 2003.

B. DeLaBarre and A. T. Brunger, “Complete structure of p97/valosin-containing protein reveals communication between nucleotide domains,” Nature Structural Biology, vol. 10, no. 10, pp. 856–863, 2003.

I. Dreveny, H. Kondo, K. Uchiyama, A. Shaw, X. Zhang, and P. S. Freemont, “Structural basis of the interaction between the AAA ATPase p97/VCP and its adaptor protein p47,” The EMBO Journal, vol. 23, no. 5, pp. 1030–1039, 2004.

H. Meyer and C. C. Weihl, “The VCP/p97 system at a glance: connecting cellular function to disease pathogenesis,” Journal of Cell Science, vol. 127, no. 18, pp. 3877–3883, 2014.

P. H. Vekaria, T. Home, S. Weir, F. J. Schoenen, and R. Rao, “Targeting p97 to disrupt protein homeostasis in cancer,” Frontiers in Oncology, vol. 6, p. 181, 2016.

M. Mehnert, T. Sommer, and E. Jarosch, “Der1 promotes movement of misfolded proteins through the endoplasmic reticulum membrane,” Nature Cell Biology, vol. 16, no. 1, pp. 77–86, 2014.

J. Wahlman, G. N. DeMartino, W. R. Skach, N. J. Bulleid, J. L. Brodsky, and A. E. Johnson, “Real-time fluorescence detection of ERAD substrate retrotranslocation in a mammalian in vitro system,” Cell, vol. 129, no. 5, pp. 943–955, 2007.

E. J. Greenblatt, J. A. Olzmann, and R. R. Kopito, “Derlin-1 is a rhomboid pseudoprotease required for the dislocation of mutant α-1 antitrypsin from the endoplasmic reticulum,” Nature Structural & Molecular Biology, vol. 18, no. 10, pp. 1147–1152, 2011.

B. N. Lilley and H. L. Ploegh, “A membrane protein required for dislocation of misfolded proteins from the ER,” Nature, vol. 429, no. 6994, pp. 834–840, 2004.

Y. Ye, Y. Shibata, C. Yun, D. Ron, and T. A. Rapoport, “A membrane protein complex mediates retro-translocation from the ER lumen into the cytosol,” Nature, vol. 429, no. 6994, pp. 841–847, 2004.

Y. Ye, Y. Shibata, M. Kikkert, S. van Voorden, E. Wiertz, and T. A. Rapoport, “Recruitment of the p97 ATPase and ubiquitin ligases to the site of retrotranslocation at the endoplasmic reticulum membrane,” Proceedings of the National Academy of Sciences of the United States of America, vol. 102, no. 40, pp. 14132–14138, 2005.

Y. Oda, T. Okada, H. Yoshida, R. J. Kaufman, K. Nagata, and K. Mori, “Derlin-2 and Derlin-3 are regulated by the mammalian unfolded protein response and are required for ER-associated degradation,” Journal of Cell Biology, vol. 172, no. 3, pp. 383–393, 2006.

B. N. Lilley and H. L. Ploegh, “Multiprotein complexes that link dislocation, ubiquitination, and extraction of misfolded proteins from the endoplasmic reticulum membrane,” Proceedings of the National Academy of Sciences of the United States of America, vol. 102, no. 40, pp. 14296–14301, 2005.

L. Fleig, N. Bergbold, P. Sahasrabudhe, B. Geiger, L. Kaltak, and M. K. Lemberg, “Ubiquitin-dependent intramembrane rhomboid protease promotes ERAD of membrane proteins,” Molecular Cell, vol. 47, no. 4, pp. 558–569, 2012.

J. L. Brodsky, “Cleaning up: ER-associated degradation to the rescue,” Cell, vol. 151, no. 6, pp. 1163–1167, 2012.

J. M. Williams, T. Inoue, L. Banks, and B. Tsai, “The ERdj5-Sel1L complex facilitates cholera toxin retrotranslocation,” Molecular Biology of the Cell, vol. 24, no. 6, pp. 785–795, 2013.

B. Mueller, B. N. Lilley, and H. L. Ploegh, “SEL1L, the homologue of yeast Hrd3p, is involved in protein dislocation from the mammalian ER,” Journal of Cell Biology, vol. 175, no. 2, pp. 261–270, 2006.

B. Mueller, E. J. Klemm, E. Spooner, J. H. Claessen, and H. L. Ploegh, “SEL1L nucleates a protein complex required for dislocation of misfolded glycoproteins,” Proceedings of the National Academy of Sciences of the United States of America, vol. 105, no. 34, pp. 12325–12330, 2008.

Y. Iida, T. Fujimori, K. Okawa, K. Nagata, I. Wada, and N. Hosokawa, “SEL1L protein critically determines the stability of the HRD1-SEL1L endoplasmic reticulum-associated degradation (ERAD) complex to optimize the degradation kinetics of ERAD substrates,” Journal of Biological Chemistry, vol. 286, no. 19, pp. 16929–16939, 2011.

E. J. Klemm, E. Spooner, and H. L. Ploegh, “Dual role of ancient ubiquitous protein 1 (AUP1) in lipid droplet accumulation and endoplasmic reticulum (ER) protein quality control,” Journal of Biological Chemistry, vol. 286, no. 43, pp. 37602–37614, 2011.

J. C. Christianson, J. A. Olzmann, T. A. Shaler et al., “Defining human ERAD networks through an integrative mapping strategy,” Nature Cell Biology, vol. 14, no. 1, pp. 93–105, 2011.

R. G. Gardner, G. M. Swarbrick, N. W. Bays et al., “Endoplasmic reticulum degradation requires lumen to cytosol signaling. Transmembrane control of Hrd1p by Hrd3p,” The Journal of Cell Biology, vol. 151, no. 1, pp. 69–82, 2000.

P. Carvalho, V. Goder, and T. A. Rapoport, “Distinct ubiquitin-ligase complexes define convergent pathways for the degradation of ER proteins,” Cell, vol. 126, no. 2, pp. 361–373, 2006.

V. Denic, E. M. Quan, and J. S. Weissman, “A luminal surveillance complex that selects misfolded glycoproteins for ER-associated degradation,” Cell, vol. 126, no. 2, pp. 349–359, 2006.

R. Gauss, E. Jarosch, T. Sommer, and C. Hirsch, “A complex of Yos9p and the HRD ligase integrates endoplasmic reticulum quality control into the degradation machinery,” Nature Cell Biology, vol. 8, no. 8, pp. 849–854, 2006.

S. Sun, G. Shi, X. Han et al., “Sel1L is indispensable for mammalian endoplasmic reticulum-associated degradation, endoplasmic reticulum homeostasis, and survival,” Proceedings of the National Academy of Sciences of the United States of America, vol. 111, no. 5, pp. E582–E591, 2014.

J. P. Lu, Y. Wang, D. A. Sliter, M. M. Pearce, and R. J. Wojcikiewicz, “RNF170 protein, an endoplasmic reticulum membrane ubiquitin ligase, mediates inositol 1,4,5-trisphosphate receptor ubiquitination and degradation,” The Journal of Biological Chemistry, vol. 286, no. 27, pp. 24426–24433, 2011.

Y. J. Jeon, J. H. Park, and C. H. Chung, “Interferon-stimulated gene 15 in the control of cellular responses to genotoxic stress,” Molecules and Cells, vol. 40, no. 2, pp. 83–89, 2017.

K. D. Wilkinson, “Regulation of ubiquitin-dependent processes by deubiquitinating enzymes,” The FASEB Journal, vol. 11, no. 14, pp. 1245–1256, 1997.

Y. Shimizu, Y. Okuda-Shimizu, and L. M. Hendershot, “Ubiquitylation of an ERAD substrate occurs on multiple types of amino acids,” Molecular Cell, vol. 40, no. 6, pp. 917–926, 2010.

X. Wang, R. A. Herr, W. J. Chua, L. Lybarger, E. J. Wiertz, and T. H. Hansen, “Ubiquitination of serine, threonine, or lysine residues on the cytoplasmic tail can induce ERAD of MHC-I by viral E3 ligase mK3,” The Journal of Cell Biology, vol. 177, no. 4, pp. 613–624, 2007.

S. Ishikura, A. M. Weissman, and J. S. Bonifacino, “Serine residues in the cytosolic tail of the T-cell antigen receptor alpha-chain mediate ubiquitination and endoplasmic reticulum-associated degradation of the unassembled protein,” The Journal of Biological Chemistry, vol. 285, no. 31, pp. 23916–23924, 2010.

F. Ikeda and I. Dikic, “Atypical ubiquitin chains: new molecular signals. 'Protein modifications: beyond the usual Suspects' review series,” EMBO Reports, vol. 9, no. 6, pp. 536–542, 2008.

Y. Kulathu and D. Komander, “Atypical ubiquitylation - the unexplored world of polyubiquitin beyond Lys48 and Lys63 linkages,” Nature Reviews. Molecular Cell Biology, vol. 13, no. 8, pp. 508–523, 2012.

C. M. Pickart and D. Fushman, “Polyubiquitin chains: polymeric protein signals,” Current Opinion in Chemical Biology, vol. 8, no. 6, pp. 610–616, 2004.

W. Li and Y. Ye, “Polyubiquitin chains: functions, structures, and mechanisms,” Cellular and Molecular Life Sciences, vol. 65, no. 15, pp. 2397–2406, 2008.

E. Nadav, A. Shmueli, H. Barr, H. Gonen, A. Ciechanover, and Y. Reiss, “A novel mammalian endoplasmic reticulum ubiquitin ligase homologous to the yeast Hrd1,” Biochemical and Biophysical Research Communications, vol. 303, no. 1, pp. 91–97, 2003.

M. Kikkert, R. Doolman, M. Dai et al., “Human HRD1 is an E3 ubiquitin ligase involved in degradation of proteins from the endoplasmic reticulum,” The Journal of Biological Chemistry, vol. 279, no. 5, pp. 3525–3534, 2004.

S. Fang, M. Ferrone, C. Yang, J. P. Jensen, S. Tiwari, and A. M. Weissman, “The tumor autocrine motility factor receptor, gp78, is a ubiquitin protein ligase implicated in degradation from the endoplasmic reticulum,” Proceedings of the National Academy of Sciences of the United States of America, vol. 98, no. 25, pp. 14422–14427, 2001.

G. Hassink, M. Kikkert, S. van Voorden et al., “TEB4 is a C4HC3 RING finger-containing ubiquitin ligase of the endoplasmic reticulum,” Biochemical Journal, vol. 388, no. 2, pp. 647–655, 2005.

J. M. Younger, L. Chen, H. Y. Ren et al., “Sequential quality-control checkpoints triage misfolded cystic fibrosis transmembrane conductance regulator,” Cell, vol. 126, no. 3, pp. 571–582, 2006.

D. Morito, K. Hirao, Y. Oda et al., “Gp78 cooperates with RMA1 in endoplasmic reticulum-associated degradation of CFTRDeltaF508,” Molecular Biology of the Cell, vol. 19, no. 4, pp. 1328–1336, 2008.

Y. J. Jeon, S. Khelifa, B. Ratnikov et al., “Regulation of glutamine carrier proteins by RNF5 determines breast cancer response to ER stress-inducing chemotherapies,” Cancer Cell, vol. 27, no. 3, pp. 354–369, 2015.

V. Tomati, E. Sondo, A. Armirotti et al., “Genetic inhibition of the ubiquitin ligase Rnf5 attenuates phenotypes associated to F508del cystic fibrosis mutation,” Scientific Reports, vol. 5, no. 1, p. 12138, 2015.

Y. Imai, M. Soda, S. Hatakeyama et al., “CHIP is associated with Parkin, a gene responsible for familial Parkinson’s disease, and enhances its ubiquitin ligase activity,” Molecular Cell, vol. 10, no. 1, pp. 55–67, 2002.

G. C. Meacham, C. Patterson, W. Zhang, J. M. Younger, and D. M. Cyr, “The Hsc70 co-chaperone CHIP targets immature CFTR for proteasomal degradation,” Nature Cell Biology, vol. 3, no. 1, pp. 100–105, 2001.

Y. Yoshida, T. Chiba, F. Tokunaga et al., “E3 ubiquitin ligase that recognizes sugar chains,” Nature, vol. 418, no. 6896, pp. 438–442, 2002.

Y. Yoshida, F. Tokunaga, T. Chiba, K. Iwai, K. Tanaka, and T. Tai, “Fbs2 is a new member of the E3 ubiquitin ligase family that recognizes sugar chains,” The Journal of Biological Chemistry, vol. 278, no. 44, pp. 43877–43884, 2003.

J. G. Magadan, F. J. Perez-Victoria, R. Sougrat, Y. Ye, K. Strebel, and J. S. Bonifacino, “Multilayered mechanism of CD4 downregulation by HIV-1 Vpu involving distinct ER retention and ERAD targeting steps,” PLoS Pathogens, vol. 6, no. 4, article e1000869, 2010.

X. Guo, S. Shen, S. Song et al., “The E3 ligase Smurf1 regulates Wolfram syndrome protein stability at the endoplasmic reticulum,” The Journal of Biological Chemistry, vol. 286, no. 20, pp. 18037–18047, 2011.

W. H. Fry, C. Simion, C. Sweeney, and K. L. Carraway III, “Quantity control of the ErbB3 receptor tyrosine kinase at the endoplasmic reticulum,” Molecular and Cellular Biology, vol. 31, no. 14, pp. 3009–3018, 2011.

Y. Jo, P. C. Lee, P. V. Sguigna, and R. A. DeBose-Boyd, “Sterol-induced degradation of HMG CoA reductase depends on interplay of two Insigs and two ubiquitin ligases, gp78 and Trc8,” Proceedings of the National Academy of Sciences of the United States of America, vol. 108, no. 51, pp. 20503–20508, 2011.

T. Zhang, Y. Xu, Y. Liu, and Y. Ye, “gp78 functions downstream of Hrd1 to promote degradation of misfolded proteins of the endoplasmic reticulum,” Molecular Biology of the Cell, vol. 26, no. 24, pp. 4438–4450, 2015.

A. Stein, A. Ruggiano, P. Carvalho, and T. A. Rapoport, “Key steps in ERAD of luminal ER proteins reconstituted with purified components,” Cell, vol. 158, no. 6, pp. 1375–1388, 2014.

J. Stevenson, E. Y. Huang, and J. A. Olzmann, “Endoplasmic reticulum-associated degradation and lipid homeostasis,” Annual Review of Nutrition, vol. 36, no. 1, pp. 511–542, 2016.

A. Shmueli, Y. C. Tsai, M. Yang, M. A. Braun, and A. M. Weissman, “Targeting of gp78 for ubiquitin-mediated proteasomal degradation by Hrd1: cross-talk between E3s in the endoplasmic reticulum,” Biochemical and Biophysical Research Communications, vol. 390, no. 3, pp. 758–762, 2009.

P. Ballar, A. U. Ors, H. Yang, and S. Fang, “Differential regulation of CFTRDeltaF508 degradation by ubiquitin ligases gp78 and Hrd1,” The International Journal of Biochemistry & Cell Biology, vol. 42, no. 1, pp. 167–173, 2010.

M. Knop, A. Finger, T. Braun, K. Hellmuth, and D. H. Wolf, “Der1, a novel protein specifically required for endoplasmic reticulum degradation in yeast,” The EMBO Journal, vol. 15, no. 4, pp. 753–763, 1996.

M. Kothe, Y. Ye, J. S. Wagner et al., “Role of p97 AAA-ATPase in the retrotranslocation of the cholera toxin A1 chain, a non-ubiquitinated substrate,” The Journal of Biological Chemistry, vol. 280, no. 30, pp. 28127–28132, 2005.

R. Gauss, T. Sommer, and E. Jarosch, “The Hrd1p ligase complex forms a linchpin between ER-lumenal substrate selection and Cdc48p recruitment,” The EMBO Journal, vol. 25, no. 9, pp. 1827–1835, 2006.

P. Carvalho, A. M. Stanley, and T. A. Rapoport, “Retrotranslocation of a misfolded luminal ER protein by the ubiquitin-ligase Hrd1p,” Cell, vol. 143, no. 4, pp. 579–591, 2010.

E. Rabinovich, A. Kerem, K. U. Frohlich, N. Diamant, and S. Bar-Nun, “AAA-ATPase p97/Cdc48p, a cytosolic chaperone required for endoplasmic reticulum-associated protein degradation,” Molecular and Cellular Biology, vol. 22, no. 2, pp. 626–634, 2002.

Y. Ye, H. H. Meyer, and T. A. Rapoport, “The AAA ATPase Cdc48/p97 and its partners transport proteins from the ER into the cytosol,” Nature, vol. 414, no. 6864, pp. 652–656, 2001.

J. Liang, C. Yin, H. Doong et al., “Characterization of erasin (UBXD2): a new ER protein that promotes ER-associated protein degradation,” Journal of Cell Science, vol. 119, no. 19, pp. 4011–4024, 2006.

M. Suzuki, T. Otsuka, Y. Ohsaki et al., “Derlin-1 and UBXD8 are engaged in dislocation and degradation of lipidated ApoB-100 at lipid droplets,” Molecular Biology of the Cell, vol. 23, no. 5, pp. 800–810, 2012.

P. Ballar, Y. Shen, H. Yang, and S. Fang, “The role of a novel p97/valosin-containing protein-interacting motif of gp78 in endoplasmic reticulum-associated degradation,” The Journal of Biological Chemistry, vol. 281, no. 46, pp. 35359–35368, 2006.

I. Kim, J. Ahn, C. Liu et al., “The Png1-Rad23 complex regulates glycoprotein turnover,” The Journal of Cell Biology, vol. 172, no. 2, pp. 211–219, 2006.

G. Li, G. Zhao, X. Zhou, H. Schindelin, and W. J. Lennarz, “The AAA ATPase p97 links peptide N-glycanase to the endoplasmic reticulum-associated E3 ligase autocrine motility factor receptor,” Proceedings of the National Academy of Sciences of the United States of America, vol. 103, no. 22, pp. 8348–8353, 2006.

R. Ernst, B. Mueller, H. L. Ploegh, and C. Schlieker, “The otubain YOD1 is a deubiquitinating enzyme that associates with p97 to facilitate protein dislocation from the ER,” Molecular Cell, vol. 36, no. 1, pp. 28–38, 2009.

M. E. Sowa, E. J. Bennett, S. P. Gygi, and J. W. Harper, “Defining the human deubiquitinating enzyme interaction landscape,” Cell, vol. 138, no. 2, pp. 389–403, 2009.

Q. Wang, L. Li, and Y. Ye, “Regulation of retrotranslocation by p97-associated deubiquitinating enzyme ataxin-3,” The Journal of Cell Biology, vol. 174, no. 7, pp. 963–971, 2006.

Y. Xu, Y. Liu, J. G. Lee, and Y. Ye, “A ubiquitin-like domain recruits an oligomeric chaperone to a retrotranslocation complex in endoplasmic reticulum-associated degradation,” The Journal of Biological Chemistry, vol. 288, no. 25, pp. 18068–18076, 2013.

Q. Wang, Y. Liu, N. Soetandyo, K. Baek, R. Hegde, and Y. Ye, “A ubiquitin ligase-associated chaperone holdase maintains polypeptides in soluble states for proteasome degradation,” Molecular Cell, vol. 42, no. 6, pp. 758–770, 2011.

Y. Xu, M. Cai, Y. Yang, L. Huang, and Y. Ye, “SGTA recognizes a noncanonical ubiquitin-like domain in the Bag6-Ubl4A-Trc35 complex to promote endoplasmic reticulum-associated degradation,” Cell Reports, vol. 2, no. 6, pp. 1633–1644, 2012.

D. Hanahan and R. A. Weinberg, “Hallmarks of cancer: the next generation,” Cell, vol. 144, no. 5, pp. 646–674, 2011.

Y. Ma and L. M. Hendershot, “The role of the unfolded protein response in tumour development: friend or foe?” Nature Reviews. Cancer, vol. 4, no. 12, pp. 966–977, 2004.

D. Ruggero, “Translational control in cancer etiology,” Cold Spring Harbor Perspectives in Biology, vol. 5, no. 2, pp. 1–27, 2013.

H. Urra, E. Dufey, T. Avril, E. Chevet, and C. Hetz, “Endoplasmic reticulum stress and the hallmarks of cancer,” Trends in Cancer, vol. 2, no. 5, pp. 252–262, 2016.

H. Vanacker, J. Vetters, L. Moudombi, C. Caux, S. Janssens, and M. C. Michallet, “Emerging role of the unfolded protein response in tumor immunosurveillance,” Trends in Cancer, vol. 3, no. 7, pp. 491–505, 2017.

C. Denoyelle, G. Abou-Rjaily, V. Bezrookove et al., “Anti-oncogenic role of the endoplasmic reticulum differentially activated by mutations in the MAPK pathway,” Nature Cell Biology, vol. 8, no. 10, pp. 1053–1063, 2006.

A. L. Huber, J. Lebeau, P. Guillaumot et al., “p58IPK-mediated attenuation of the proapoptotic PERK-CHOP pathway allows malignant progression upon low glucose,” Molecular Cell, vol. 49, no. 6, pp. 1049–1059, 2013.

G. Ramadori, G. Konstantinidou, N. Venkateswaran et al., “Diet-induced unresolved ER stress hinders KRAS-driven lung tumorigenesis,” Cell Metabolism, vol. 21, no. 1, pp. 117–125, 2015.

L. Niederreiter, T. M. Fritz, T. E. Adolph et al., “ER stress transcription factor Xbp1 suppresses intestinal tumorigenesis and directs intestinal stem cells,” The Journal of Experimental Medicine, vol. 210, no. 10, pp. 2041–2056, 2013.

M. Bi, C. Naczki, M. Koritzinsky et al., “ER stress-regulated translation increases tolerance to extreme hypoxia and promotes tumor growth,” The EMBO Journal, vol. 24, no. 19, pp. 3470–3481, 2005.

J. D. Blais, C. L. Addison, R. Edge et al., “Perk-dependent translational regulation promotes tumor cell adaptation and angiogenesis in response to hypoxic stress,” Molecular and Cellular Biology, vol. 26, no. 24, pp. 9517–9532, 2006.

S. Dey, C. M. Sayers, I. I. Verginadis et al., “ATF4-dependent induction of heme oxygenase 1 prevents anoikis and promotes metastasis,” The Journal of Clinical Investigation, vol. 125, no. 7, pp. 2592–2608, 2015.

S. B. Cullinan, D. Zhang, M. Hannink, E. Arvisais, R. J. Kaufman, and J. A. Diehl, “Nrf2 is a direct PERK substrate and effector of PERK-dependent cell survival,” Molecular and Cellular Biology, vol. 23, no. 20, pp. 7198–7209, 2003.

C. A. Del Vecchio, Y. Feng, E. S. Sokol et al., “De-differentiation confers multidrug resistance via noncanonical PERK-Nrf2 signaling,” PLoS Biology, vol. 12, no. 9, article e1001945, 2014.

W. Zhang, V. Hietakangas, S. Wee, S. C. Lim, J. Gunaratne, and S. M. Cohen, “ER stress potentiates insulin resistance through PERK-mediated FOXO phosphorylation,” Genes & Development, vol. 27, no. 4, pp. 441–449, 2013.

D. Pytel, I. Majsterek, and J. A. Diehl, “Tumor progression and the different faces of the PERK kinase,” Oncogene, vol. 35, no. 10, pp. 1207–1215, 2016.

Y. Kouroku, E. Fujita, I. Tanida et al., “ER stress (PERK/eIF2α phosphorylation) mediates the polyglutamine-induced LC3 conversion, an essential step for autophagy formation,” Cell Death and Differentiation, vol. 14, no. 2, pp. 230–239, 2007.

W. B'Chir, A. C. Maurin, V. Carraro et al., “The eIF2α/ATF4 pathway is essential for stress-induced autophagy gene expression,” Nucleic Acids Research, vol. 41, no. 16, pp. 7683–7699, 2013.

S. I. Grivennikov and M. Karin, “Dangerous liaisons: STAT3 and NF-κB collaboration and crosstalk in cancer,” Cytokine & Growth Factor Reviews, vol. 21, no. 1, pp. 11–19, 2010.

O. Pluquet, N. Dejeans, M. Bouchecareilh et al., “Posttranscriptional regulation of PER1 underlies the oncogenic function of IREα,” Cancer Research, vol. 73, no. 15, pp. 4732–4743, 2013.

B. Kharabi Masouleh, H. Geng, C. Hurtz et al., “Mechanistic rationale for targeting the unfolded protein response in pre-B acute lymphoblastic leukemia,” Proceedings of the National Academy of Sciences of the United States of America, vol. 111, no. 21, pp. E2219–E2228, 2014.

N. Mimura, M. Fulciniti, G. Gorgun et al., “Blockade of XBP1 splicing by inhibition of IRE1α is a promising therapeutic option in multiple myeloma,” Blood, vol. 119, no. 24, pp. 5772–5781, 2012.

I. Papandreou, N. C. Denko, M. Olson et al., “Identification of an Ire1alpha endonuclease specific inhibitor with cytotoxic activity against human multiple myeloma,” Blood, vol. 117, no. 4, pp. 1311–1314, 2011.

M. Ri, E. Tashiro, D. Oikawa et al., “Identification of Toyocamycin, an agent cytotoxic for multiple myeloma cells, as a potent inhibitor of ER stress-induced XBP1 mRNA splicing,” Blood Cancer Journal, vol. 2, no. 7, article e79, 2012.

C. Greenman, P. Stephens, R. Smith et al., “Patterns of somatic mutation in human cancer genomes,” Nature, vol. 446, no. 7132, pp. 153–158, 2007.

Z. Xue, Y. He, K. Ye, Z. Gu, Y. Mao, and L. Qi, “A conserved structural determinant located at the interdomain region of mammalian inositol-requiring enzyme 1α,” The Journal of Biological Chemistry, vol. 286, no. 35, pp. 30859–30866, 2011.

D. M. Schewe and J. A. Aguirre-Ghiso, “ATF6α-Rheb-mTOR signaling promotes survival of dormant tumor cells in vivo,” Proceedings of the National Academy of Sciences of the United States of America, vol. 105, no. 30, pp. 10519–10524, 2008.

Y. Wang, G. N. Alam, Y. Ning et al., “The unfolded protein response induces the angiogenic switch in human tumor cells through the PERK/ATF4 pathway,” Cancer Research, vol. 72, no. 20, pp. 5396–5406, 2012.

X. Chen, D. Iliopoulos, Q. Zhang et al., “XBP1 promotes triple-negative breast cancer by controlling the HIF1α pathway,” Nature, vol. 508, no. 7494, pp. 103–107, 2014.

E. Karali, S. Bellou, D. Stellas et al., “VEGF Signals through ATF6 and PERK to promote endothelial cell survival and angiogenesis in the absence of ER stress,” Molecular Cell, vol. 54, no. 4, pp. 559–572, 2014.

J. R. Cubillos-Ruiz, S. E. Bettigole, and L. H. Glimcher, “Tumorigenic and immunosuppressive effects of endoplasmic reticulum stress in cancer,” Cell, vol. 168, no. 4, pp. 692–706, 2017.

D. Senft and Z. A. Ronai, “Adaptive stress responses during tumor metastasis and dormancy,” Trends in Cancer, vol. 2, no. 8, pp. 429–442, 2016.

M. A. Nieto, R. Y. Huang, R. A. Jackson, and J. P. Thiery, “EMT: 2016,” Cell, vol. 166, no. 1, pp. 21–45, 2016.

H. Mujcic, A. Nagelkerke, K. M. Rouschop et al., “Hypoxic activation of the PERK/eIF2α arm of the unfolded protein response promotes metastasis through induction of LAMP3,” Clinical Cancer Research, vol. 19, no. 22, pp. 6126–6137, 2013.

A. Nagelkerke, J. Bussink, H. Mujcic et al., “Hypoxia stimulates migration of breast cancer cells via the PERK/ATF4/LAMP3-arm of the unfolded protein response,” Breast Cancer Res, vol. 15, no. 1, article R2, pp. 467–480, 2013.

H. Zhu, X. Chen, B. Chen et al., “Activating transcription factor 4 promotes esophageal squamous cell carcinoma invasion and metastasis in mice and is associated with poor prognosis in human patients,” PLoS One, vol. 9, no. 7, article e103882, 2014.

G. Auf, A. Jabouille, S. Guerit et al., “Inositol-requiring enzyme 1alpha is a key regulator of angiogenesis and invasion in malignant glioma,” Proceedings of the National Academy of Sciences of the United States of America, vol. 107, no. 35, pp. 15553–15558, 2010.

N. Dejeans, O. Pluquet, S. Lhomond et al., “Autocrine control of glioma cells adhesion and migration through IRE1α-mediated cleavage of SPARC mRNA,” Journal of Cell Science, vol. 125, Part 18, pp. 4278–4287, 2012.

N. R. Mahadevan, J. Rodvold, H. Sepulveda, S. Rossi, A. F. Drew, and M. Zanetti, “Transmission of endoplasmic reticulum stress and pro-inflammation from tumor cells to myeloid cells,” Proceedings of the National Academy of Sciences of the United States of America, vol. 108, no. 16, pp. 6561–6566, 2011.

N. R. Mahadevan, V. Anufreichik, J. J. Rodvold, K. T. Chiu, H. Sepulveda, and M. Zanetti, “Cell-extrinsic effects of tumor ER stress imprint myeloid dendritic cells and impair CD8⁺ T cell priming,” PLoS One, vol. 7, no. 12, article e51845, 2012.

J. Pol, E. Vacchelli, F. Aranda et al., “Trial Watch: Immunogenic cell death inducers for anticancer chemotherapy,” Oncoimmunology, vol. 4, no. 4, article e1008866, 2015.

D. V. Krysko, A. D. Garg, A. Kaczmarek, O. Krysko, P. Agostinis, and P. Vandenabeele, “Immunogenic cell death and DAMPs in cancer therapy,” Nature Reviews. Cancer, vol. 12, no. 12, pp. 860–875, 2012.

A. R. van Vliet, S. Martin, A. D. Garg, and P. Agostinis, “The PERKs of damage-associated molecular patterns mediating cancer immunogenicity: from sensor to the plasma membrane and beyond,” Seminars in Cancer Biology, vol. 33, pp. 74–85, 2015.

J. Fucikova, E. Becht, K. Iribarren et al., “Calreticulin expression in human non-small cell lung cancers correlates with increased accumulation of antitumor immune cells and favorable prognosis,” Cancer Research, vol. 76, no. 7, pp. 1746–1756, 2016.

A. D. Garg, D. V. Krysko, T. Verfaillie et al., “A novel pathway combining calreticulin exposure and ATP secretion in immunogenic cancer cell death,” The EMBO Journal, vol. 31, no. 5, pp. 1062–1079, 2012.

O. Kepp, L. Menger, E. Vacchelli et al., “Crosstalk between ER stress and immunogenic cell death,” Cytokine & Growth Factor Reviews, vol. 24, no. 4, pp. 311–318, 2013.

C. Pozzi, A. Cuomo, I. Spadoni et al., “The EGFR-specific antibody cetuximab combined with chemotherapy triggers immunogenic cell death,” Nature Medicine, vol. 22, no. 6, pp. 624–631, 2016.

S. E. Bettigole and L. H. Glimcher, “Endoplasmic reticulum stress in immunity,” Annual Review of Immunology, vol. 33, no. 1, pp. 107–138, 2015.

N. N. Iwakoshi, A. H. Lee, P. Vallabhajosyula, K. L. Otipoby, K. Rajewsky, and L. H. Glimcher, “Plasma cell differentiation and the unfolded protein response intersect at the transcription factor XBP-1,” Nature Immunology, vol. 4, no. 4, pp. 321–329, 2003.

A. M. Reimold, N. N. Iwakoshi, J. Manis et al., “Plasma cell differentiation requires the transcription factor XBP-1,” Nature, vol. 412, no. 6844, pp. 300–307, 2001.

R. Hu, Z. F. Chen, J. Yan et al., “Endoplasmic reticulum stress of neutrophils is required for ischemia/reperfusion-induced acute lung injury,” Journal of Immunology, vol. 195, no. 10, pp. 4802–4809, 2015.

F. Martinon, X. Chen, A. H. Lee, and L. H. Glimcher, “TLR activation of the transcription factor XBP1 regulates innate immune responses in macrophages,” Nature Immunology, vol. 11, no. 5, pp. 411–418, 2010.

J. R. Cubillos-Ruiz, P. C. Silberman, M. R. Rutkowski et al., “ER stress sensor XBP1 controls anti-tumor immunity by disrupting dendritic cell homeostasis,” Cell, vol. 161, no. 7, pp. 1527–1538, 2015.

A. C. Herrera, V. J. Victorino, F. C. Campos et al., “Impact of tumor removal on the systemic oxidative profile of patients with breast cancer discloses lipid peroxidation at diagnosis as a putative marker of disease recurrence,” Clinical Breast Cancer, vol. 14, no. 6, pp. 451–459, 2014.

D. L. Herber, W. Cao, Y. Nefedova et al., “Lipid accumulation and dendritic cell dysfunction in cancer,” Nature Medicine, vol. 16, no. 8, pp. 880–886, 2010.

F. Hossain, A. A. Al-Khami, D. Wyczechowska et al., “Inhibition of fatty acid oxidation modulates immunosuppressive functions of myeloid-derived suppressor cells and enhances cancer therapies,” Cancer Immunology Research, vol. 3, no. 11, pp. 1236–1247, 2015.

R. Ramakrishnan, V. A. Tyurin, F. Veglia et al., “Oxidized lipids block antigen cross-presentation by dendritic cells in cancer,” Journal of Immunology, vol. 192, no. 6, pp. 2920–2931, 2014.

D. Yan, H. W. Wang, R. L. Bowman, and J. A. Joyce, “STAT3 and STAT6 signaling pathways synergize to promote Cathepsin secretion from macrophages via IRE1α activation,” Cell Reports, vol. 16, no. 11, pp. 2914–2927, 2016.

T. Condamine, G. A. Dominguez, J. I. Youn et al., “Lectin-type oxidized LDL receptor-1 distinguishes population of human polymorphonuclear myeloid-derived suppressor cells in cancer patients,” Science Immunology, vol. 1, no. 2, pp. 1–15, 2016.

C. H. Tang, S. Ranatunga, C. L. Kriss et al., “Inhibition of ER stress-associated IRE-1/XBP-1 pathway reduces leukemic cell survival,” The Journal of Clinical Investigation, vol. 124, no. 6, pp. 2585–2598, 2014.

P. T. Thevenot, R. A. Sierra, P. L. Raber et al., “The stress-response sensor chop regulates the function and accumulation of myeloid-derived suppressor cells in tumors,” Immunity, vol. 41, no. 3, pp. 389–401, 2014.

T. Condamine, V. Kumar, I. R. Ramachandran et al., “ER stress regulates myeloid-derived suppressor cell fate through TRAIL-R-mediated apoptosis,” The Journal of Clinical Investigation, vol. 124, no. 6, pp. 2626–2639, 2014.

X. H. Ma, S. F. Piao, S. Dey et al., “Targeting ER stress-induced autophagy overcomes BRAF inhibitor resistance in melanoma,” The Journal of Clinical Investigation, vol. 124, no. 3, pp. 1406–1417, 2014.

G. B. Park, D. Y. Hur, and D. Kim, “Combining CAL-101 with celecoxib enhances apoptosis of EBV-transformed B-cells through MAPK-induced ER stress,” Anticancer Research, vol. 35, no. 5, pp. 2699–2708, 2015.

A. Strasser and H. Puthalakath, “Fold up or perish: unfolded protein response and chemotherapy,” Cell Death and Differentiation, vol. 15, no. 2, pp. 223–225, 2007.

B. Kharabi Masouleh, E. Chevet, J. Panse et al., “Drugging the unfolded protein response in acute leukemias,” Journal of Hematology & Oncology, vol. 8, no. 1, p. 87, 2015.

L. Vincenz, R. Jager, M. O'Dwyer, and A. Samali, “Endoplasmic reticulum stress and the unfolded protein response: targeting the Achilles heel of multiple myeloma,” Molecular Cancer Therapeutics, vol. 12, no. 6, pp. 831–843, 2013.

K. M. Rouschop, L. J. Dubois, T. G. Keulers et al., “PERK/eIF2α signaling protects therapy resistant hypoxic cells through induction of glutathione synthesis and protection against ROS,” Proceedings of the National Academy of Sciences of the United States of America, vol. 110, no. 12, pp. 4622–4627, 2013.

M. Ogata, S. Hino, A. Saito et al., “Autophagy is activated for cell survival after endoplasmic reticulum stress,” Molecular and Cellular Biology, vol. 26, no. 24, pp. 9220–9231, 2006.

Y. H. Shi, Z. B. Ding, J. Zhou et al., “Targeting autophagy enhances sorafenib lethality for hepatocellular carcinoma via ER stress-related apoptosis,” Autophagy, vol. 7, no. 10, pp. 1159–1172, 2011.

J. R. Cubillos-Ruiz, S. Fiering, and J. R. Conejo-Garcia, “Nanomolecular targeting of dendritic cells for ovarian cancer therapy,” Future Oncology, vol. 5, no. 8, pp. 1189–1192, 2009.

J. R. Cubillos-Ruiz, J. R. Baird, A. J. Tesone et al., “Reprogramming tumor-associated dendritic cells in vivo using miRNA mimetics triggers protective immunity against ovarian cancer,” Cancer Research, vol. 72, no. 7, pp. 1683–1693, 2012.

T. Tanaka, T. Kajiwara, T. Torigoe, Y. Okamoto, N. Sato, and Y. Tamura, “Cancer-associated oxidoreductase ERO1-α drives the production of tumor-promoting myeloid-derived suppressor cells via oxidative protein folding,” Journal of Immunology, vol. 194, no. 4, pp. 2004–2010, 2015.

I. Saez and D. Vilchez, “The mechanistic links between proteasome activity, aging and age-related diseases,” Current Genomics, vol. 15, no. 1, pp. 38–51, 2014.

D. Vilchez, I. Saez, and A. Dillin, “The role of protein clearance mechanisms in organismal ageing and age-related diseases,” Nature Communications, vol. 5, p. 5659, 2014.

M. Cattaneo, E. Fontanella, C. Canton, D. Delia, and I. Biunno, “SEL1L affects human pancreatic cancer cell cycle and invasiveness through modulation of PTEN and genes related to cell-matrix interactions,” Neoplasia, vol. 7, no. 11, pp. 1030–1038, 2005.

H. Kim, A. Bhattacharya, and L. Qi, “Endoplasmic reticulum quality control in cancer: friend or foe,” Seminars in Cancer Biology, vol. 33, pp. 25–33, 2015.

R. Orlandi, M. Cattaneo, F. Troglio et al., “SEL1L expression decreases breast tumor cell aggressiveness in vivo and in vitro,” Cancer Research, vol. 62, no. 2, pp. 567–574, 2002.

H. Ashktorab, W. Green, G. Finzi et al., “SEL1L, an UPR response protein, a potential marker of colonic cell transformation,” Digestive Diseases and Sciences, vol. 57, no. 4, pp. 905–912, 2012.

J. H. Baek, P. C. Mahon, J. Oh et al., “OS-9 interacts with hypoxia-inducible factor 1alpha and prolyl hydroxylases to promote oxygen-dependent degradation of HIF-1α,” Molecular Cell, vol. 17, no. 4, pp. 503–512, 2005.

M. Ivan, K. Kondo, H. Yang et al., “HIFα targeted for VHL-mediated destruction by proline hydroxylation: implications for O₂ sensing,” Science, vol. 292, no. 5516, pp. 464–468, 2001.

K. Yanagisawa, H. Konishi, C. Arima et al., “Novel metastasis-related gene CIM functions in the regulation of multiple cellular stress-response pathways,” Cancer Research, vol. 70, no. 23, pp. 9949–9958, 2010.

L. A. Liotta, R. Mandler, G. Murano et al., “Tumor cell autocrine motility factor,” Proceedings of the National Academy of Sciences of the United States of America, vol. 83, no. 10, pp. 3302–3306, 1986.

I. R. Nabi and A. Raz, “Cell shape modulation alters glycosylation of a metastatic melanoma cell-surface antigen,” International Journal of Cancer, vol. 40, no. 3, pp. 396–402, 1987.

V. Joshi, A. Upadhyay, A. Kumar, and A. Mishra, “Gp78 E3 ubiquitin ligase: essential functions and contributions in proteostasis,” Frontiers in Cellular Neuroscience, vol. 11, p. 259, 2017.

S. Silletti, J. Yao, J. Sanford et al., “Autocrine motility factor-receptor in human bladder-carcinoma - gene-expression, loss of cell-contact regulation and chromosomal mapping,” International Journal of Oncology, vol. 3, no. 5, pp. 801–807, 1993.

S. Nakamori, H. Watanabe, M. Kameyama et al., “Expression of autocrine motility factor receptor in colorectal cancer as a predictor for disease recurrence,” Cancer, vol. 74, no. 7, pp. 1855–1862, 1994.

K. Maruyama, H. Watanabe, H. Shiozaki et al., “Expression of autocrine motility factor receptor in human esophageal squamous cell carcinoma,” International Journal of Cancer, vol. 64, no. 5, pp. 316–321, 1995.

S. Silletti, J. P. Yao, K. J. Pienta, and A. Raz, “Loss of cell-contact regulation and altered responses to autocrine motility factor correlate with increased malignancy in prostate cancer cells,” International Journal of Cancer, vol. 63, no. 1, pp. 100–105, 1995.

T. Otto, W. Birchmeier, U. Schmidt et al., “Inverse relation of E-cadherin and autocrine motility factor receptor expression as a prognostic factor in patients with bladder carcinomas,” Cancer Research, vol. 54, no. 12, pp. 3120–3123, 1994.

T. Otto, A. Bex, U. Schmidt, A. Raz, and H. Rubben, “Improved prognosis assessment for patients with bladder carcinoma,” The American Journal of Pathology, vol. 150, no. 6, pp. 1919–1923, 1997.

K. Kawanishi, Y. Doki, H. Shiozaki et al., “Correlation between loss of E-cadherin expression and overexpression of autocrine motility factor receptor in association with progression of human gastric cancers,” American Journal of Clinical Pathology, vol. 113, no. 2, pp. 266–274, 2000.

Y. C. Tsai, A. Mendoza, J. M. Mariano et al., “The ubiquitin ligase gp78 promotes sarcoma metastasis by targeting KAI1 for degradation,” Nature Medicine, vol. 13, no. 12, pp. 1504–1509, 2007.

B. Joshi, L. Li, and I. R. Nabi, “A role for KAI1 in promotion of cell proliferation and mammary gland hyperplasia by the gp78 ubiquitin ligase,” The Journal of Biological Chemistry, vol. 285, no. 12, pp. 8830–8839, 2010.

L. Wang, G. Hou, L. Xue, J. Li, P. Wei, and P. Xu, “Autocrine motility factor receptor signaling pathway promotes cell invasion via activation of ROCK-2 in esophageal squamous cell cancer cells,” Cancer Investigation, vol. 28, no. 10, pp. 993–1003, 2010.

M. Kajiro, R. Hirota, Y. Nakajima et al., “The ubiquitin ligase CHIP acts as an upstream regulator of oncogenic pathways,” Nature Cell Biology, vol. 11, no. 3, pp. 312–319, 2009.