# Exploring the IRE1 interactome: from canonical signaling functions to unexpected roles

Simon Le Goupil, Hadrien Laprade, Marc Aubry, Eric Chevet

► To cite this version:

Simon Le Goupil, Hadrien Laprade, Marc Aubry, Eric Chevet. Exploring the IRE1 interactome: from canonical signaling functions to unexpected roles. Journal of Biological Chemistry, 2024, 300 (4), pp.107169. 10.1016/j.jbc.2024.107169. hal-04540346

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# Exploring the IRE1 interactome: From canonical signaling functions to unexpected roles

Received for publication, November 13, 2023, and in revised form, March 4, 2024
https://doi.org/10.1016/j.jbc.2024.107169
Published, Papers in Press, March 15, 2024

Simon Le Goupil<sup>1,2,*</sup> and Eric Chevet<sup>1,2</sup>

From the ¹INSERM U1242, University of Rennes, Rennes, France; ²Centre de Lutte contre le cancer Eugène Marquis, Rennes, France

Reviewed by members of the JBC Editorial Board. Edited by Ronald Wek

The unfolded protein response is a mechanism aiming at restoring endoplasmic reticulum (ER) homeostasis and is likely involved in other adaptive pathways. The unfolded protein response is transduced by three proteins acting as sensors and triggering downstream signaling pathways. Among them, inositol-requiring enzyme 1 alpha (IRE1α) (referred to as IRE1 hereafter), an endoplasmic reticulum-resident type I trans-membrane protein, exerts its function through both kinase and endoribonuclease activities, resulting in both X-box binding protein 1 mRNA splicing and RNA degradation (regulated ire1 dependent decay). An increasing number of studies have re-ported protein-protein interactions as regulators of these signaling mechanisms, and additionally, driving other nonca-nonical functions. In this review, we deliver evolutive and structural insights on IRE1 and further describe how this protein interaction network (interactome) regulates IRE1 signaling abilities or mediates other cellular processes through catalytic-independent mechanisms. Moreover, we focus on newly discovered targets of IRE1 kinase activity and discuss potentially novel IRE1 functions based on the nature of the interactome, thereby identifying new fields to explore regarding this protein's biological roles.

## Endoplasmic reticulum stress and the unfolded protein response

The endoplasmic reticulum (ER) is one of the largest organelles in eukaryotic cells. It was observed for the first time at the end of the 19th century by Charles Garnier and named "ergastoplasm" (1) then defined as ER in 1945 by Porter, Claude, and Fullam (2). Beyond its functions in the regulation of calcium, cells' redox status and lipid homeostasis, the ER has a predominant role in protein synthesis, productive folding, and trafficking (3-6). Indeed, about one-third of the proteome passes through the ER to acquire proper folding before reaching its final destination (7). The ER lumen exhibits a calcium-rich, oxidative environment populated by ER-resident enzymes and foldases (e.g., protein disulfide isomerases (PDIs), chaperones, N-linked glycosylation machinery), which ensure proper protein folding (8, 9). In addition, a protein quality control system is in charge of

either retaining improperly folded proteins in the ER to allow further folding cycles or to promote the degradation of terminally misfolded proteins through ER-associated degradation (ERAD) (10). This process is critical to ensure ER homeostasis, integrating protein synthesis rates, folding, and degradation/export through the secretory pathway (6). Under some circumstances, ER homeostasis can be disrupted when protein folding demand in the ER exceeds the ER capacity to handle productive protein folding. This leads to an accumulation of improperly folded proteins in this compartment, a situation known as ER stress (11). To cope with ER stress, organisms/cells have evolved a strategy termed the unfolded protein response (UPR). The UPR is a signaling pathway triggered upon ER stress mainly to restore ER homeostasis (adaptive UPR). However, if this process fails, the UPR will engage cells on a death path (terminal UPR) (12, 13). Over the years, many studies showed the involvement of UPR signaling in a wide range of diseases such as cancers, metabolic, or degenerative disorders (14, 15).

The UPR is transduced by three ER-resident trans-membrane sensors named respectively the PKR-like endoplasmic reticulum kinase (PERK), the activating transcription factor 6 (ATF6), and the inositol-requiring enzyme 1 (IRE1). PERK is a type 1 transmembrane protein with a luminal sensing domain and a cytosolic kinase catalytic domain. Activated PERK enables the phosphorylation of the eukaryotic translation initiation factor 2A (eIF2α), thereby decreasing overall mRNA translation and therefore the ER protein load (16). However, phosphorylated eIF2α allows bypassing the reading of upstream ORF on translated mRNA by ribosomes to activate the selective synthesis of several proteins, including the ATF4 transcription factor (17). ATF4 targets the transcription of genes whose products are involved in redox homeostasis, amino acid metabolism, or autophagy (18, 19) (Fig. 1). The ATF4-dependent transcription network is also contributing to a negative feedback loop allowing translation to resume (20). Indeed, ATF4 controls the expression of the phosphatase subunit GADD34 which associates with the protein phosphatase 1 catalytic subunit to dephosphorylate eIF2α hence restoring translation (21). Upon accumulation of improperly folded proteins in the ER lumen, the type II ER transmembrane protein ATF6 is translocated in the Golgi apparatus through coat protein complex II transport vesicles, where it is cleaved by the site 1/2 proteases (S1P

<footnote>* For correspondence: Simon Le Goupil, simon.legoupil@univ-rennes1.fr.</footnote>
# Exploring the IRE1 interactome: From canonical signaling functions to unexpected roles

Received for publication, November 13, 2023, and in revised form, March 4, 2024
https://doi.org/10.1016/j.jbc.2024.107169
Published, Papers in Press, March 15, 2024

Simon Le Goupil<sup>1,2,*</sup> and Eric Chevet<sup>1,2</sup>

From the <sup>1</sup>INSERM U1242, University of Rennes, Rennes, France; <sup>2</sup>Centre de Lutte contre le cancer Eugène Marquis, Rennes, France

Reviewed by members of the JBC Editorial Board. Edited by Ronald Wek

The unfolded protein response is a mechanism aiming at restoring endoplasmic reticulum (ER) homeostasis and is likely involved in other adaptive pathways. The unfolded protein response is transduced by three proteins acting as sensors and triggering downstream signaling pathways. Among them, inositol-requiring enzyme 1 alpha (IRE1α) (referred to as IRE1 hereafter), an endoplasmic reticulum-resident type I trans-membrane protein, exerts its function through both kinase and endoribonuclease activities, resulting in both X-box binding protein 1 mRNA splicing and RNA degradation (regulated ire1 dependent decay). An increasing number of studies have re-ported protein-protein interactions as regulators of these signaling mechanisms, and additionally, driving other nonca-nonical functions. In this review, we deliver evolutive and structural insights on IRE1 and further describe how this protein interaction network (interactome) regulates IRE1 signaling abilities or mediates other cellular processes through catalytic-independent mechanisms. Moreover, we focus on newly discovered targets of IRE1 kinase activity and discuss potentially novel IRE1 functions based on the nature of the interactome, thereby identifying new fields to explore regarding this protein's biological roles.

## Endoplasmic reticulum stress and the unfolded protein response

The endoplasmic reticulum (ER) is one of the largest organelles in eukaryotic cells. It was observed for the first time at the end of the 19th century by Charles Garnier and named "ergastoplasm" (1) then defined as ER in 1945 by Porter, Claude, and Fullam (2). Beyond its functions in the regulation of calcium, cells' redox status and lipid homeostasis, the ER has a predominant role in protein synthesis, productive folding, and trafficking (3–6). Indeed, about one-third of the proteome passes through the ER to acquire proper folding before reaching its final destination (7). The ER lumen exhibits a calcium-rich, oxidative environment populated by ER-resident enzymes and foldases (e.g., protein disulfide isomerases (PDIs), chaperones, N-linked glycosylation machinery), which ensure proper protein folding (8, 9). In addition, a protein quality control system is in charge of

either retaining improperly folded proteins in the ER to allow further folding cycles or to promote the degradation of terminally misfolded proteins through ER-associated degradation (ERAD) (10). This process is critical to ensure ER homeostasis, integrating protein synthesis rates, folding, and degradation/export through the secretory pathway (6). Under some circumstances, ER homeostasis can be disrupted when protein folding demand in the ER exceeds the ER capacity to handle productive protein folding. This leads to an accumulation of improperly folded proteins in this compartment, a situation known as ER stress (11). To cope with ER stress, organisms/cells have evolved a strategy termed the unfolded protein response (UPR). The UPR is a signaling pathway triggered upon ER stress mainly to restore ER homeostasis (adaptive UPR). However, if this process fails, the UPR will engage cells on a death path (terminal UPR) (12, 13). Over the years, many studies showed the involvement of UPR signaling in a wide range of diseases such as cancers, metabolic, or degenerative disorders (14, 15).

The UPR is transduced by three ER-resident trans-membrane sensors named respectively the PKR-like endoplasmic reticulum kinase (PERK), the activating transcription factor 6 (ATF6), and the inositol-requiring enzyme 1 (IRE1). PERK is a type 1 transmembrane protein with a luminal sensing domain and a cytosolic kinase catalytic domain. Activated PERK enables the phosphorylation of the eukaryotic translation initiation factor 2A (eIF2α), thereby decreasing overall mRNA translation and therefore the ER protein load (16). However, phosphorylated eIF2α allows bypassing the reading of upstream ORF on translated mRNA by ribosomes to activate the selective synthesis of several proteins, including the ATF4 transcription factor (17). ATF4 targets the transcription of genes whose products are involved in redox homeostasis, amino acid metabolism, or autophagy (18, 19) (Fig. 1). The ATF4-dependent transcription network is also contributing to a negative feedback loop allowing translation to resume (20). Indeed, ATF4 controls the expression of the phosphatase subunit GADD34 which associates with the protein phosphatase 1 catalytic subunit to dephosphorylate eIF2α hence restoring translation (21). Upon accumulation of improperly folded proteins in the ER lumen, the type II ER transmembrane protein ATF6 is translocated in the Golgi apparatus through coat protein complex II transport vesicles, where it is cleaved by the site 1/2 proteases (S1P

<footnote>* For correspondence: Simon Le Goupil, simon.legoupil@univ-rennes1.fr.</footnote>
<figure><img src="image_1.png" /><figcaption>**Figure 1. The unfolded protein response.** Schematic representation of the IRE1, PERK, and ATF6 branches of the UPR. ATF6, activating transcription factor 6; IRE1, inositol-requiring enzyme 1; PERK, PKR-like endoplasmic reticulum kinase.</figcaption></figure>

and S2P) leading to the release of ATF6 cytosolic domain (CD) (ATF6f) in the cytosol (22). ATF6f is a leucine zipper (bZIP) motif containing transcription factor that mostly controls the expression of genes coding for ER chaperones/foldases, thereby leading to an increase in productive protein folding in the ER (23, 24) (Fig. 1).

The most evolutionary conserved stress transducer of the UPR is IRE1, discovered in yeast and mammals in the 90s (25, 26). IRE1 is an ER-resident type 1 transmembrane protein that comprises both BiP-binding (binding-immuno-globulin protein) and misfolded protein binding domain in the ER lumen as well as kinase and endoribonuclease catalytic domains in the cytosol. Upon ER stress, IRE1 oligomerizes and transautophosphorylates, triggering two outcomes managed by the RNase domain (25, 27). First, the nonconventional splicing of X-box binding protein 1 (XBP1) mRNA, where IRE1 removes a 26-nt intron, with further ligation of the remaining 5' and 3' fragments by the tRNA ligase RtcB to form the spliced XBP1 (XBP1s) mRNA (28–30). This splicing event causes a frameshift in the XBP1s transcript, leading to translation of a protein containing a transcription activator domain in the C-terminal end. Hence XBP1s acts as a transcription factor regulating the expression of enzymes involved in lipid biogenesis, of ER

chaperones, and ERAD components (31–34). Second, the RNase activity termed as regulated IRE1dependent decay (RIDD) is responsible for cleaving a pool of RNAs (mRNA, rRNA, miRNA), aiming at decreasing translation or at inducing novel cellular programs (35, 36) (Fig. 1). Importantly, RIDD specificity still remains unclear and might be cell specific. To date, a consensus sequence was identified for RIDD substrates (CUGCAG) within a stem-loop motif (37), but a recent study unveiled targeting of RNAs lacking those criteria and termed this activity as RIDD-lacking endomotif (38). In addition to their signaling, the three UPR sensors exerts a crosstalk between each other as shown by the upregulation of XBP1 expression by both ATF6f and ATF4 (39, 40) or the competitive role of PERK and IRE1 regarding expression of the death receptor 5 (41). Some hypotheses propose a hierarchy among the UPR sensors in human, suggesting that low ER stress is solely resolved by the ATF6 branch, while mild or high ER stress additionally requires IRE1 and PERK branches. Moreover, upon persistent ER stress, it is thought that IRE1 signaling declines in the benefit of PERK, to mediate CHOP-induced apoptosis (42). Numerous studies also point toward scaffolding functions of IRE1 either to regulate its signaling or to mediate other cellular processes (translation, cytoskeleton ...). In this
<figure><img src="image_2.png" /><figcaption>**Figure 1. The unfolded protein response.** Schematic representation of the IRE1, PERK, and ATF6 branches of the UPR. ATF6, activating transcription factor 6; IRE1, inositol-requiring enzyme 1; PERK, PKR-like endoplasmic reticulum kinase.</figcaption></figure>

and S2P) leading to the release of ATF6 cytosolic domain (CD) (ATF6f) in the cytosol (22). ATF6f is a leucine zipper (bZIP) motif containing transcription factor that mostly controls the expression of genes coding for ER chaperones/foldases, thereby leading to an increase in productive protein folding in the ER (23, 24) (Fig. 1).

The most evolutionary conserved stress transducer of the UPR is IRE1, discovered in yeast and mammals in the 90s (25, 26). IRE1 is an ER-resident type 1 transmembrane protein that comprises both BiP-binding (binding-immuno-globulin protein) and misfolded protein binding domain in the ER lumen as well as kinase and endoribonuclease catalytic domains in the cytosol. Upon ER stress, IRE1 oligomerizes and transautophosphorylates, triggering two outcomes managed by the RNase domain (25, 27). First, the nonconventional splicing of X-box binding protein 1 (XBP1) mRNA, where IRE1 removes a 26-nt intron, with further ligation of the remaining 5' and 3' fragments by the tRNA ligase RtcB to form the spliced XBP1 (XBP1s) mRNA (28–30). This splicing event causes a frameshift in the XBP1s transcript, leading to translation of a protein containing a transcription activator domain in the C-terminal end. Hence XBP1s acts as a transcription factor regulating the expression of enzymes involved in lipid biogenesis, of ER

chaperones, and ERAD components (31–34). Second, the RNase activity termed as regulated IRE1dependent decay (RIDD) is responsible for cleaving a pool of RNAs (mRNA, rRNA, miRNA), aiming at decreasing translation or at inducing novel cellular programs (35, 36) (Fig. 1). Importantly, RIDD specificity still remains unclear and might be cell specific. To date, a consensus sequence was identified for RIDD substrates (CUGCAG) within a stem-loop motif (37), but a recent study unveiled targeting of RNAs lacking those criteria and termed this activity as RIDD-lacking endomotif (38). In addition to their signaling, the three UPR sensors exerts a crosstalk between each other as shown by the upregulation of XBP1 expression by both ATF6f and ATF4 (39, 40) or the competitive role of PERK and IRE1 regarding expression of the death receptor 5 (41). Some hypotheses propose a hierarchy among the UPR sensors in human, suggesting that low ER stress is solely resolved by the ATF6 branch, while mild or high ER stress additionally requires IRE1 and PERK branches. Moreover, upon persistent ER stress, it is thought that IRE1 signaling declines in the benefit of PERK, to mediate CHOP-induced apoptosis (42). Numerous studies also point toward scaffolding functions of IRE1 either to regulate its signaling or to mediate other cellular processes (translation, cytoskeleton ...). In this
review, we propose an overview of IRE1 biochemistry and describe the known IRE1 protein--protein interactions (PPis) (Table 1) and their biological outputs.

## IRE1 across evolution

### IRE1α and IRE1β paralogs

The evolution of the IRE1 gene might have encountered a duplication event, as, unlike in *Saccharomyces cerevisiae*, two IRE1 paralogs exist in mammals, namely IRE1α and IRE1β, respectively encoded by two genes: ERN1 and ERN2,

respectively localized on 17q23.3 and 16p12.2 chromosomal regions and displaying 39% sequence identity (43). Both proteins conserved their structural domains. Whereas IRE1α is ubiquitously expressed, IRE1β is preferentially found in mucosal epithelia (44, 45). Regarding their functions, both IRE1α and IRE1β exerts RIDD activity and IRE1β was found to inhibit mRNA translation by degrading rRNA (43). However, while IRE1α-mediated XBP1 mRNA splicing has been largely described, it remains unclear whether IRE1β is capable of such activity in a physiological setting. Due to nonconserved amino acids in the kinase domain, IRE1β has impaired

<table><caption>**Table 1**

List of described IRE1 interactors</caption><thead><tr><th>Biological functions</th><th>Protein ID</th><th>Protein full name</th><th>Reference</th></tr></thead><tbody><tr><td rowspan="7">IRE1 activation</td><td>Misfolded proteins</td><td>Misfolded proteins</td><td>(85–87)</td></tr><tr><td>AGR2*</td><td>Anterior gradient protein 2</td><td>(51, 95)</td></tr><tr><td>BIP</td><td>Endoplasmic reticulum chaperone BiP</td><td>(88)</td></tr><tr><td>ERDJ4</td><td>DnaJ homolog subfamily B member 9</td><td>(88)</td></tr><tr><td>HSP47</td><td>Serpin family H member 1</td><td>(96)</td></tr><tr><td>PDIA1</td><td>Protein disulfide isomerase A1</td><td>(99)</td></tr><tr><td>PDIA6</td><td>Protein disulfide isomerase A6</td><td>(98)</td></tr><tr><td rowspan="5">IRE1 Oligomerization</td><td>c-ABL</td><td>Tyrosine-protein kinase ABL1</td><td>(104, 105)</td></tr><tr><td>NMIIB</td><td>Nonmuscle myosin heavy chain IIB</td><td>(106)</td></tr><tr><td>RAC</td><td>Ribosome-associated complex</td><td>(103)</td></tr><tr><td>Sec61β</td><td>Protein transport protein Sec61 subunit beta</td><td>(101, 102)</td></tr><tr><td>Sec63</td><td>Translocation protein Sec63 homolog</td><td>(101, 102)</td></tr><tr><td rowspan="7">IRE1 RNase activity</td><td>BAK</td><td>Bcl-2 homologous antagonist/killer</td><td>(109)</td></tr><tr><td>BAX</td><td>Apoptosis regulator BAX</td><td>(109)</td></tr><tr><td>BI-1</td><td>BAX inhibitor 1</td><td>(112, 113)</td></tr><tr><td>BIM</td><td>Bcl-2-like protein 11</td><td>(110)</td></tr><tr><td>c-ABL</td><td>Tyrosine-protein kinase ABL1</td><td>(104)</td></tr><tr><td>NCK</td><td>Cytoplasmic protein NCK1</td><td>(114)</td></tr><tr><td>PUMA</td><td>Bcl-2-binding component 3</td><td>(110)</td></tr><tr><td>RNH1</td><td>Ribonuclease inhibitor</td><td>(111)</td></tr><tr><td rowspan="11">IRE1 Phospho regulation</td><td>AMPK</td><td>AMP-activated protein kinase</td><td>(118, 119)</td></tr><tr><td>Dcr2</td><td>Phosphatase DCR2 (S. cerevisiae)</td><td>(116)</td></tr><tr><td>ERK</td><td>Extracellular signal-regulated kinase</td><td>(120)</td></tr><tr><td>PKA</td><td>Protein kinase A</td><td>(122)</td></tr><tr><td>PP2A</td><td>Protein phosphatase 2A</td><td>(118)</td></tr><tr><td>PP2Ce</td><td>Protein phosphatase 1L</td><td>(117)</td></tr><tr><td>Ptcp2</td><td>Serine/threonine phosphatase of type 2C (S. cerevisiae)</td><td>(26)</td></tr><tr><td>PTP1B</td><td>Tyrosine-protein phosphatase nonreceptor type 1</td><td>(105, 121)</td></tr><tr><td>RACK1</td><td>Receptor for activated C-kinase 1</td><td>(118)</td></tr><tr><td>RPAP2</td><td>RNA polymerase II-associated protein 2</td><td>(41)</td></tr><tr><td>SCP3</td><td>Small carboxy-terminal domain phosphatase 3</td><td>(120)</td></tr><tr><td rowspan="10">PTM and IRE1 degradation</td><td>BIM</td><td>Bcl-2-like protein 11</td><td>(110)</td></tr><tr><td>CHIP</td><td>E3 ubiquitin-protein ligase CHIP</td><td>(129)</td></tr><tr><td>M50</td><td>M50 viral protein</td><td>(128)</td></tr><tr><td>MITOL</td><td>E3 ubiquitin-protein ligase MARCHF5</td><td>(131)</td></tr><tr><td>OS9</td><td>OS9 endoplasmic reticulum lectin</td><td>(127)</td></tr><tr><td>RNF13</td><td>E3 ubiquitin-protein ligase RNF13</td><td>(130)</td></tr><tr><td>SEL1L</td><td>Protein sel-1 homolog 1</td><td>(126, 127)</td></tr><tr><td>SYNV1</td><td>E3 ubiquitin-protein ligase synoviolin</td><td>(126, 127)</td></tr><tr><td>AIP1</td><td>ASK1-interacting protein-1</td><td>(137)</td></tr><tr><td>ASK1</td><td>Apoptotic signaling kinase 1</td><td>(135, 136)</td></tr><tr><td rowspan="5">Cytoskeleton</td><td>JNK</td><td>Mitogen-activated protein kinase 8</td><td>(135)</td></tr><tr><td>MITOL</td><td>E3 ubiquitin-protein ligase MARCHF5</td><td>(131)</td></tr><tr><td>TRAF2</td><td>TNF receptor-associated factor 2</td><td>(135)</td></tr><tr><td>CORO1</td><td>Coronin</td><td>(111)</td></tr><tr><td>FLNA</td><td>Filamin A</td><td>(139)</td></tr><tr><td rowspan="4">SRP and ribosome</td><td>PKCα</td><td>Protein kinase C alpha type</td><td>(139)</td></tr><tr><td>TUBA</td><td>Tubulin A</td><td>(111)</td></tr><tr><td>TUBB</td><td>Tubulin B</td><td>(111)</td></tr><tr><td>RPLs</td><td>Ribosome protein large subunits</td><td>(143)</td></tr><tr><td rowspan="2">Ca<sup>2+</sup> homeostasis</td><td>SRPs</td><td>Signal recognition particles</td><td>(143)</td></tr><tr><td>INSP3R</td><td>Inositol 1,4,5-trisphosphate receptor type 3</td><td>(148)</td></tr><tr><td rowspan="7">IRE1 Kinase target</td><td>STIM1</td><td>Stromal interaction molecule 1</td><td>(149)</td></tr><tr><td>AMPK</td><td>AMP-activated protein kinase</td><td>(119)</td></tr><tr><td>BCL2</td><td>Apoptosis regulator Bcl-2</td><td>(119)</td></tr><tr><td>FMRP</td><td>Fragile X messenger ribonucleoprotein 1</td><td>(150)</td></tr><tr><td>IKK</td><td>IκBα kinase</td><td>(152)</td></tr><tr><td>PUM1</td><td>Pumilio homolog 1</td><td>(151)</td></tr><tr><td>Pumilio</td><td>Pumilio (Drosophila melanogaster)</td><td>(151)</td></tr><tr><td>SPL</td><td>Sphingosine 1-phosphate lyase</td><td>(153)</td></tr></tbody></table>

IRE1, inositol-requiring enzyme 1; PTM, posttranslational modification.

(*) indicates IRE1β specific interactions.
review, we propose an overview of IRE1 biochemistry and describe the known IRE1 protein--protein interactions (PPis) (Table 1) and their biological outputs.

## IRE1 across evolution

### IRE1α and IRE1β paralogs

The evolution of the IRE1 gene might have encountered a duplication event, as, unlike in *Saccharomyces cerevisiae*, two IRE1 paralogs exist in mammals, namely IRE1α and IRE1β, respectively encoded by two genes: ERN1 and ERN2,

respectively localized on 17q23.3 and 16p12.2 chromosomal regions and displaying 39% sequence identity (43). Both proteins conserved their structural domains. Whereas IRE1α is ubiquitously expressed, IRE1β is preferentially found in mucosal epithelia (44, 45). Regarding their functions, both IRE1α and IRE1β exerts RIDD activity and IRE1β was found to inhibit mRNA translation by degrading rRNA (43). However, while IRE1α-mediated XBP1 mRNA splicing has been largely described, it remains unclear whether IRE1β is capable of such activity in a physiological setting. Due to nonconserved amino acids in the kinase domain, IRE1β has impaired

<table><caption>**Table 1**

List of described IRE1 interactors</caption><thead><tr><th>Biological functions</th><th>Protein ID</th><th>Protein full name</th><th>Reference</th></tr></thead><tbody><tr><td rowspan="6">IRE1 activation</td><td>Misfolded proteins</td><td>Misfolded proteins</td><td>(85–87)</td></tr><tr><td>AGR2*</td><td>Anterior gradient protein 2</td><td>(51, 95)</td></tr><tr><td>BIP</td><td>Endoplasmic reticulum chaperone BiP</td><td>(88)</td></tr><tr><td>ERDJ4</td><td>DnaJ homolog subfamily B member 9</td><td>(88)</td></tr><tr><td>HSP47</td><td>Serpin family H member 1</td><td>(96)</td></tr><tr><td>PDIA1</td><td>Protein disulfide isomerase A1</td><td>(99)</td></tr><tr><td rowspan="5">IRE1 Oligomerization</td><td>PDIA6</td><td>Protein disulfide isomerase A6</td><td>(98)</td></tr><tr><td>c-ABL</td><td>Tyrosine-protein kinase ABL1</td><td>(104, 105)</td></tr><tr><td>NMIIB</td><td>Nonmuscle myosin heavy chain IIB</td><td>(106)</td></tr><tr><td>RAC</td><td>Ribosome-associated complex</td><td>(103)</td></tr><tr><td>Sec61β</td><td>Protein transport protein Sec61 subunit beta</td><td>(101, 102)</td></tr><tr><td rowspan="6">IRE1 RNase activity</td><td>Sec63</td><td>Translocation protein Sec63 homolog</td><td>(101, 102)</td></tr><tr><td>BAK</td><td>Bcl-2 homologous antagonist/killer</td><td>(109)</td></tr><tr><td>BAX</td><td>Apoptosis regulator BAX</td><td>(109)</td></tr><tr><td>BI-1</td><td>BAX inhibitor 1</td><td>(112, 113)</td></tr><tr><td>BIM</td><td>Bcl-2-like protein 11</td><td>(110)</td></tr><tr><td>c-ABL</td><td>Tyrosine-protein kinase ABL1</td><td>(104)</td></tr><tr><td rowspan="5">IRE1 Phospho regulation</td><td>NCK</td><td>Cytoplasmic protein NCK1</td><td>(114)</td></tr><tr><td>PUMA</td><td>Bcl-2-binding component 3</td><td>(110)</td></tr><tr><td>RNH1</td><td>Ribonuclease inhibitor</td><td>(111)</td></tr><tr><td>AMPK</td><td>AMP-activated protein kinase</td><td>(118, 119)</td></tr><tr><td>Dcr2</td><td>Phosphatase DCR2 (S. cerevisiae)</td><td>(116)</td></tr><tr><td rowspan="13">PTM and IRE1 degradation</td><td>ERK</td><td>Extracellular signal-regulated kinase</td><td>(120)</td></tr><tr><td>PKA</td><td>Protein kinase A</td><td>(122)</td></tr><tr><td>PP2A</td><td>Protein phosphatase 2A</td><td>(118)</td></tr><tr><td>PP2Ce</td><td>Protein phosphatase 1L</td><td>(117)</td></tr><tr><td>Ptcp2</td><td>Serine/threonine phosphatase of type 2C (S. cerevisiae)</td><td>(26)</td></tr><tr><td>PTP1B</td><td>Tyrosine-protein phosphatase nonreceptor type 1</td><td>(105, 121)</td></tr><tr><td>RACK1</td><td>Receptor for activated C-kinase 1</td><td>(118)</td></tr><tr><td>RPAP2</td><td>RNA polymerase II-associated protein 2</td><td>(41)</td></tr><tr><td>SCP3</td><td>Small carboxy-terminal domain phosphatase 3</td><td>(120)</td></tr><tr><td>BIM</td><td>Bcl-2-like protein 11</td><td>(110)</td></tr><tr><td>CHIP</td><td>E3 ubiquitin-protein ligase CHIP</td><td>(129)</td></tr><tr><td>M50</td><td>M50 viral protein</td><td>(128)</td></tr><tr><td>MITOL</td><td>E3 ubiquitin-protein ligase MARCHF5</td><td>(131)</td></tr><tr><td>OS9</td><td>OS9 endoplasmic reticulum lectin</td><td>(127)</td></tr><tr><td>RNF13</td><td>E3 ubiquitin-protein ligase RNF13</td><td>(130)</td></tr><tr><td>SEL1L</td><td>Protein sel-1 homolog 1</td><td>(126, 127)</td></tr><tr><td>SYNV1</td><td>E3 ubiquitin-protein ligase synoviolin</td><td>(126, 127)</td></tr><tr><td rowspan="6">Apoptosis</td><td>AIP1</td><td>ASK1-interacting protein-1</td><td>(137)</td></tr><tr><td>ASK1</td><td>Apoptotic signaling kinase 1</td><td>(135, 136)</td></tr><tr><td>JNK</td><td>Mitogen-activated protein kinase 8</td><td>(135)</td></tr><tr><td>MITOL</td><td>E3 ubiquitin-protein ligase MARCHF5</td><td>(131)</td></tr><tr><td>TRAF2</td><td>TNF receptor-associated factor 2</td><td>(135)</td></tr><tr><td>CORO1</td><td>Coronin</td><td>(111)</td></tr><tr><td rowspan="5">Cytoskeleton</td><td>FLNA</td><td>Filamin A</td><td>(139)</td></tr><tr><td>PKCα</td><td>Protein kinase C alpha type</td><td>(139)</td></tr><tr><td>TUBA</td><td>Tubulin A</td><td>(111)</td></tr><tr><td>TUBB</td><td>Tubulin B</td><td>(111)</td></tr><tr><td>RPLs</td><td>Ribosome protein large subunits</td><td>(143)</td></tr><tr><td rowspan="2">SRP and ribosome</td><td>SRPs</td><td>Signal recognition particles</td><td>(143)</td></tr><tr><td>INSP3R</td><td>Inositol 1,4,5-trisphosphate receptor type 3</td><td>(148)</td></tr><tr><td rowspan="5">IRE1 Kinase target</td><td>STIM1</td><td>Stromal interaction molecule 1</td><td>(149)</td></tr><tr><td>AMPK</td><td>AMP-activated protein kinase</td><td>(119)</td></tr><tr><td>BCL2</td><td>Apoptosis regulator Bcl-2</td><td>(119)</td></tr><tr><td>FMRP</td><td>Fragile X messenger ribonucleoprotein 1</td><td>(150)</td></tr><tr><td>IKK</td><td>IκBα kinase</td><td>(152)</td></tr><tr><td rowspan="3">Ca<sup>2+</sup> homeostasis</td><td>PUM1</td><td>Pumilio homolog 1</td><td>(151)</td></tr><tr><td>Pumilio</td><td>Pumilio (Drosophila melanogaster)</td><td>(151)</td></tr><tr><td>SPL</td><td>Sphingosine 1-phosphate lyase</td><td>(153)</td></tr></tbody></table>

IRE1, inositol-requiring enzyme 1; PTM, posttranslational modification.

(*) indicates IRE1β specific interactions.
phosphorylation and reduced oligomerization abilities (46–48). Even so, few studies were able to observe a slight XBP1 splicing activity for IRE1β in *in vitro* splicing experiments and in overexpressing cells (49–51). In the latter case, the splicing activity is dependent on cell type and appeared to be reduced in mucosal cell line where IRE1β is usually expressed (51). Concomitantly, IRE1β was found to act as a negative regulator of IRE1α, as its overexpression is able to decrease IRE1α-dependent XBP1 splicing (47). Indeed, IRE1β conserved a capacity to oligomerize with IRE1α and by doing so, acts as a dominant negative by preventing IRE1α functions (47). In this regard, IRE1β is thought to attenuate ER stress upon inflammation induced diseases in mucosal epithelium (44). For the continuation of this review, we will

mainly consider IRE1α (and refer to it as IRE1 unless specified).

### IRE1 across evolution

IRE1 can be divided into three main domains: an N-terminal luminal domain (NLD), a transmembrane domain (TMD), and a CD, the latter comprising both kinase and RNase catalytic activities. These domains are all conserved in metazoan, fungi, and plant orthologs, as shown by the yeast *versus* human IRE1 comparison (Fig. 2A) (52). However, compared to human IRE1, the amino acid sequence differs across the reigns. Indeed, while the BLAST alignment of protein sequences reveals a high degree of conservation in mammals (>90% identity), a low IRE1 sequence conservation is found between human and the other

<figure><img src="image_2.png" /><figcaption>**Figure 2.** Evolutive insights on IRE1. A, schematic representation of yeast (above) and human (below) IRE1 sequence displaying the signal peptide (SP) with the three mains domains: luminal (NLD), transmembrane domain, and the cytosolic domain exerting kinase and RNase activities. B, comparison of IRE1 protein sequence identity and similarity in different species using BLAST. C, GO Manhattan plot using g:Profiler of yeast (up) and human (bottom) IRE1 interactome from curated databases (APID, String and Biogrid). Black arrows highlight terms of interest, and values in brackets represent the number of enriched terms in each Gene Ontology. IRE1, inositol-requiring enzyme 1; NLD, N-terminal luminal domain.</figcaption></figure>
phosphorylation and reduced oligomerization abilities (46–48). Even so, few studies were able to observe a slight XBP1 splicing activity for IRE1β in *in vitro* splicing experiments and in overexpressing cells (49–51). In the latter case, the splicing activity is dependent on cell type and appeared to be reduced in mucosal cell line where IRE1β is usually expressed (51). Concomitantly, IRE1β was found to act as a negative regulator of IRE1α, as its overexpression is able to decrease IRE1α-dependent XBP1 splicing (47). Indeed, IRE1β conserved a capacity to oligomerize with IRE1α and by doing so, acts as a dominant negative by preventing IRE1α functions (47). In this regard, IRE1β is thought to attenuate ER stress upon inflammation induced diseases in mucosal epithelium (44). For the continuation of this review, we will

mainly consider IRE1α (and refer to it as IRE1 unless specified).

### IRE1 across evolution

IRE1 can be divided into three main domains: an N-terminal luminal domain (NLD), a transmembrane domain (TMD), and a CD, the latter comprising both kinase and RNase catalytic activities. These domains are all conserved in metazoan, fungi, and plant orthologs, as shown by the yeast *versus* human IRE1 comparison (Fig. 2A) (52). However, compared to human IRE1, the amino acid sequence differs across the reigns. Indeed, while the BLAST alignment of protein sequences reveals a high degree of conservation in mammals (>90% identity), a low IRE1 sequence conservation is found between human and the other

<figure><img src="image_3.png" /><figcaption>**Figure 2.** Evolutive insights on IRE1. A, schematic representation of yeast (above) and human (below) IRE1 sequence displaying the signal peptide (SP) with the three mains domains: luminal (NLD), transmembrane domain, and the cytosolic domain exerting kinase and RNase activities. B, comparison of IRE1 protein sequence identity and similarity in different species using BLAST. C, GO Manhattan plot using g:Profiler of yeast (up) and human (bottom) IRE1 interactome from curated databases (APID, String and Biogrid). Black arrows highlight terms of interest, and values in brackets represent the number of enriched terms in each Gene Ontology. IRE1, inositol-requiring enzyme 1; NLD, N-terminal luminal domain.</figcaption></figure>
species (approximatively 30% identity according to species) (Fig. 2B) (53). This trend is comparable with the overall similarity of the sequences (>90% in mammals while approximatively 50% in other eukaryotes and <40% in plants) (Fig. 2B). This suggests that both IRE1 kinase and RNase domains share common features and functions across species despite the changes in sequences and implies that the luminal domain diverges more and may show different characteristics which may be linked to (i) IRE1 activation mode in different species and/or (ii) the ability of IRE1 luminal domain to directly and selectively bind to specific motifs which may differ between species. However, IRE1 activity presents significant differences in yeast species. In S. cerevisiae, IRE1 can splice HAC1 mRNA (XBP1 functional homolog in yeast) together with the S. cerevisiae Rlg1p ligase (RtcB homolog in yeast) (27, 54) and to a much lesser extent RIDD (55). Whereas in S. pombe, IRE1 only degrades RNA through RIDD since further analysis found that S. pombe lacks HAC1 (or orthologs) (55, 56). Therefore, one can hypothesize that all eukaryotes may express an IRE1 capable of both splicing and RIDD activities (with different selectivity) and that the occurrence of splicing does not depend on the IRE1 RNase domain but on the expression of the target gene. This suggests an evolutive trait of the IRE1 pathway in managing ER stress, by modifying signaling characteristics and likely to expand its biological functions during evolution. This correlates also with the diversification of the UPR, with the appearance of the PERK and ATF6 arms (57) in higher order organisms (metazoans and chordates).

### Comparison of IRE1 interactomes across evolution

To analyze the diversity of IRE1 PPis across evolution, we collected IRE1 interactome data from S. cerevisiae and *Homo sapiens*, the most documented species on public repositories (agile protein interactomes dataserver, String and Biogrid) (58–60). Interestingly, agile protein interactomes dataserver, the most complete database reports 300,000 interactions among the 5800 proteins studied in yeast while in human, 700,000 interactions are distributed among 14,500 proteins. Regarding IRE1, analysis of those repositories documents a total of 101 interactions for human against 121 for yeast. Gene Ontology analysis carried out through g:Profiler revealed a different pattern of enrichment between human and yeast IRE1 human interactome. Indeed, every ontology (molecular function, biological process, and cellular component) displays much more terms in human than in yeast (Fig. 2C) (61). Regarding the content of these ontologies, while in yeast IRE1 interactors are mainly linked to stress-related processes such as protein folding or ERAD, human IRE1 interactors are connected with many more diverse processes in addition to the UPR, from signaling pathways (with the mitogen-activated protein kinase (MAPK) signaling), trafficking (coat protein complex II) to apoptosis or ubiquitination (Fig. 2C). We analyzed the number of terms enriched between the interactomes of a significant number of *H. sapiens* and *S. cerevisiae* orthologs and found that the richer and denser functional enrichment observed in the human IRE1 interactome is significantly different (p value = 0.0014, not shown) than the mean of interactome differences of other

orthologs. Altogether, it suggests that IRE1 is part of a specific subset of proteins, which gains new functions across evolution through their protein interaction network.

## IRE1 structural insights

### IRE1 luminal domain

A first crystal structure of the conserved core region of IRE1 NLD was obtained in S. cerevisiae in 2005 (62). It shows a dimer harboring a putative protein/protein interface (Fig. 3A). This structure describes that the IRE1 NLD is able to dimerize through both sides of the NLD and suggests that IRE1 could form high order oligomers necessary for its activation. This crystalized dimer also shows a “groove” structure between the two monomers similar to the major histocompatibility peptide-binding complexes (MHCs) (62). Interestingly, mutations in the NLD contact sites were shown to abolish IRE1 activation (63). In comparison, human IRE1 NLD dimer structure, obtained by crystallography, reveals a triangle shape of each monomer, composed of three β-sheets that appear different from the yeast NLD. This structural difference could be explained by the length of this domain in both species (Fig. 2A) and by the previously discussed poor conservation of the sequence as shown by the amino acid percentage identity between human and yeast (Fig. 2B). Upon dimerization, one face of a triangle interacts with another similar structure in an antiparallel manner, creating an MHC-like motif (Fig. 3B). This motif presents an interface able to form hydrophobic and hydrogen bonds, the last especially through the K121 amino acid (64). The following MHC-like structure is comparable to that of yeast, but the closed groove suggests a different peptide binding capacity in sensing unfolded protein (64). Of note, cancer-associated somatic mutations in the luminal domain of human IRE1 were reported to affect IRE1 signaling (65).

### IRE1 transmembrane domain

The IRE1 TMD consists in a single amino acid stretch exhibiting a juxta-membrane amphipathic helix (AH) containing polar residues facing the lumen and a transmembrane helix (66) (Fig. 3, C and D). Interestingly, analysis of the TMD from different species revealed a conservation of these AH and transmembrane helix from mammals to plants (66). In yeast, a study identified a link between IRE1 activation and lipid sensing properties regulated by IRE1 AH (66). It was found that the AH could sense lipid composition of the ER membrane and that mutation in this domain led to impaired HAC1 mRNA splicing (66). They further discussed a model in which IRE1 AH could locally compress the membrane, allowing stable formation of oligomers (66). Another study showed that lipid unbalance induces UPR and that IRE1 counteracts membrane stress-induced cell death by reprogramming protein homeostasis and maintains membrane functions without affecting its composition (67). The same team recently reported that spliced HAC1 regulates the expression of different genes according to either lipid bilayer stress or unfolded protein stress (68). In human, IRE1 TMD has also been documented for lipid-sensing properties with IRE1-detecting lipid
species (approximatively 30% identity according to species) (Fig. 2B) (53). This trend is comparable with the overall similarity of the sequences (>90% in mammals while approximatively 50% in other eukaryotes and <40% in plants) (Fig. 2B). This suggests that both IRE1 kinase and RNase domains share common features and functions across species despite the changes in sequences and implies that the luminal domain diverges more and may show different characteristics which may be linked to (i) IRE1 activation mode in different species and/or (ii) the ability of IRE1 luminal domain to directly and selectively bind to specific motifs which may differ between species. However, IRE1 activity presents significant differences in yeast species. In S. cerevisiae, IRE1 can splice HAC1 mRNA (XBP1 functional homolog in yeast) together with the S. cerevisiae Rlg1p ligase (RtcB homolog in yeast) (27, 54) and to a much lesser extent RIDD (55). Whereas in S. pombe, IRE1 only degrades RNA through RIDD since further analysis found that S. pombe lacks HAC1 (or orthologs) (55, 56). Therefore, one can hypothesize that all eukaryotes may express an IRE1 capable of both splicing and RIDD activities (with different selectivity) and that the occurrence of splicing does not depend on the IRE1 RNase domain but on the expression of the target gene. This suggests an evolutive trait of the IRE1 pathway in managing ER stress, by modifying signaling characteristics and likely to expand its biological functions during evolution. This correlates also with the diversification of the UPR, with the appearance of the PERK and ATF6 arms (57) in higher order organisms (metazoans and chordates).

### Comparison of IRE1 interactomes across evolution

To analyze the diversity of IRE1 PPis across evolution, we collected IRE1 interactome data from S. cerevisiae and *Homo sapiens*, the most documented species on public repositories (agile protein interactomes dataserver, String and Biogrid) (58–60). Interestingly, agile protein interactomes dataserver, the most complete database reports 300,000 interactions among the 5800 proteins studied in yeast while in human, 700,000 interactions are distributed among 14,500 proteins. Regarding IRE1, analysis of those repositories documents a total of 101 interactions for human against 121 for yeast. Gene Ontology analysis carried out through g:Profiler revealed a different pattern of enrichment between human and yeast IRE1 human interactome. Indeed, every ontology (molecular function, biological process, and cellular component) displays much more terms in human than in yeast (Fig. 2C) (61). Regarding the content of these ontologies, while in yeast IRE1 interactors are mainly linked to stress-related processes such as protein folding or ERAD, human IRE1 interactors are connected with many more diverse processes in addition to the UPR, from signaling pathways (with the mitogen-activated protein kinase (MAPK) signaling), trafficking (coat protein complex II) to apoptosis or ubiquitination (Fig. 2C). We analyzed the number of terms enriched between the interactomes of a significant number of *H. sapiens* and *S. cerevisiae* orthologs and found that the richer and denser functional enrichment observed in the human IRE1 interactome is significantly different (p value = 0.0014, not shown) than the mean of interactome differences of other

orthologs. Altogether, it suggests that IRE1 is part of a specific subset of proteins, which gains new functions across evolution through their protein interaction network.

## IRE1 structural insights

### IRE1 luminal domain

A first crystal structure of the conserved core region of IRE1 NLD was obtained in S. cerevisiae in 2005 (62). It shows a dimer harboring a putative protein/protein interface (Fig. 3A). This structure describes that the IRE1 NLD is able to dimerize through both sides of the NLD and suggests that IRE1 could form high order oligomers necessary for its activation. This crystalized dimer also shows a “groove” structure between the two monomers similar to the major histocompatibility peptide-binding complexes (MHCs) (62). Interestingly, mutations in the NLD contact sites were shown to abolish IRE1 activation (63). In comparison, human IRE1 NLD dimer structure, obtained by crystallography, reveals a triangle shape of each monomer, composed of three β-sheets that appear different from the yeast NLD. This structural difference could be explained by the length of this domain in both species (Fig. 2A) and by the previously discussed poor conservation of the sequence as shown by the amino acid percentage identity between human and yeast (Fig. 2B). Upon dimerization, one face of a triangle interacts with another similar structure in an antiparallel manner, creating an MHC-like motif (Fig. 3B). This motif presents an interface able to form hydrophobic and hydrogen bonds, the last especially through the K121 amino acid (64). The following MHC-like structure is comparable to that of yeast, but the closed groove suggests a different peptide binding capacity in sensing unfolded protein (64). Of note, cancer-associated somatic mutations in the luminal domain of human IRE1 were reported to affect IRE1 signaling (65).

### IRE1 transmembrane domain

The IRE1 TMD consists in a single amino acid stretch exhibiting a juxta-membrane amphipathic helix (AH) containing polar residues facing the lumen and a transmembrane helix (66) (Fig. 3, C and D). Interestingly, analysis of the TMD from different species revealed a conservation of these AH and transmembrane helix from mammals to plants (66). In yeast, a study identified a link between IRE1 activation and lipid sensing properties regulated by IRE1 AH (66). It was found that the AH could sense lipid composition of the ER membrane and that mutation in this domain led to impaired HAC1 mRNA splicing (66). They further discussed a model in which IRE1 AH could locally compress the membrane, allowing stable formation of oligomers (66). Another study showed that lipid unbalance induces UPR and that IRE1 counteracts membrane stress-induced cell death by reprogramming protein homeostasis and maintains membrane functions without affecting its composition (67). The same team recently reported that spliced HAC1 regulates the expression of different genes according to either lipid bilayer stress or unfolded protein stress (68). In human, IRE1 TMD has also been documented for lipid-sensing properties with IRE1-detecting lipid
<figure><img src="image_3.png" /><figcaption>B</figcaption></figure>

<figure><img src="image_4.png" /><figcaption>C</figcaption></figure>

<figure><img src="image_5.png" /><figcaption>D</figcaption></figure>

<figure><img src="image_6.png" /><figcaption>E</figcaption></figure>

<figure><img src="image_7.png" /><figcaption>F</figcaption></figure>

<figure><img src="image_8.png" /><figcaption>G</figcaption></figure>

<figure><img src="image_9.png" /></figure>

**Figure 3.** Comparative structure of IRE1 in *Homo sapiens* and *Saccharomyces cerevisiae*. Dimer structures of luminal domain of (A) yeast (PDB: 2BE1) and (B) human (PDB: 2HZ6) IRE1. Each color represents a monomer. Helical wheel schematic representation of IRE1 yeast (C) and (D) human amphipathic helix from the transmembrane domain. The arrow represents the hydrophobic moment. E, crystal structure of dimeric IRE1 cytosolic domain of yeast (PDB: 2RIO). F, crystal structures of human dimeric IRE1 cytosolic domain with back-to-back (PDB: 6W3C, left) and face-to-face (PDB: 3P23, right) conformations. G, crystal structure of yeast IRE1 cytosolic oligomer (PDB: 3FBV). Each color represents a monomer, and the black arrow illustrates the progressive 52° rotation between each dimer. IRE1, inositol-requiring enzyme 1.

bilayer stress to trigger UPR activation (69, 70). Moreover, by using fluorescence complementation assays, Cho and colleagues showed in cells that lipid saturation of the ER membrane with palmitate induces IRE1 (lacking NLD) dimerization through the TMD, by a mechanism involving W457 and S450

residues (71). In addition, the IRE1 TMD has also been proposed to be involved in the interaction with the membrane-embedded translocon component Sec61β. Indeed, alteration of IRE1 TMD by replacing it with calnexin TMD abolished its interaction with the translocon (72).

<figure><img src="image_10.png" /></figure>
<figure><img src="image_4.png" /><figcaption>B</figcaption></figure>

<figure><img src="image_5.png" /><figcaption>C</figcaption></figure>

<figure><img src="image_6.png" /><figcaption>D</figcaption></figure>

<figure><img src="image_7.png" /><figcaption>E</figcaption></figure>

<figure><img src="image_8.png" /><figcaption>F</figcaption></figure>

<figure><img src="image_9.png" /><figcaption>G</figcaption></figure>

<figure><img src="image_10.png" /></figure>

**Figure 3.** Comparative structure of IRE1 in *Homo sapiens* and *Saccharomyces cerevisiae*. Dimer structures of luminal domain of (A) yeast (PDB: 2BE1) and (B) human (PDB: 2HZ6) IRE1. Each color represents a monomer. Helical wheel schematic representation of IRE1 yeast (C) and (D) human amphipathic helix from the transmembrane domain. The arrow represents the hydrophobic moment. E, crystal structure of dimeric IRE1 cytosolic domain of yeast (PDB: 2RIO). F, crystal structures of human dimeric IRE1 cytosolic domain with back-to-back (PDB: 6W3C, left) and face-to-face (PDB: 3P23, right) conformations. G, crystal structure of yeast IRE1 cytosolic oligomer (PDB: 3FBV). Each color represents a monomer, and the black arrow illustrates the progressive 52° rotation between each dimer. IRE1, inositol-requiring enzyme 1.

bilayer stress to trigger UPR activation (69, 70). Moreover, by using fluorescence complementation assays, Cho and colleagues showed in cells that lipid saturation of the ER membrane with palmitate induces IRE1 (lacking NLD) dimerization through the TMD, by a mechanism involving W457 and S450

residues (71). In addition, the IRE1 TMD has also been proposed to be involved in the interaction with the membrane-embedded translocon component Sec61β. Indeed, alteration of IRE1 TMD by replacing it with calnexin TMD abolished its interaction with the translocon (72).
IRE1 CD

The CD of IRE1 is the signaling region of IRE1. As such, it
shows two main domains exerting, respectively a serine/thre-
onine kinase activity and an RNase activity. The former leads
to IRE1 transautophoshorylation upon oligomerization,
thereby leading to the activation of the latter. IRE1 RNase
induces the noncanonical splicing of HAC1 mRNA in yeast
and of XBP1 mRNA in human, with further ligation by the
Rlg1 (yeast) or RtcB (human) RNA ligases, respectively. This
process allows for the synthesis of the spliced form of the
XBP1s/Hac1¹p transcription factors, thereby promoting the
expression of UPR target genes (28, 73, 74). IRE1 cytosolic
regions presents two main fragments: a kinase domain (yeast
674–980; human 571–832), and a RNase domain (yeast
983–1115; human 835–963) (Fig. 2A). Structural evidence in
yeast showed that the phosphorylated IRE1 kinase/RNase
domain is able to dimerize in its active form (75). This dimer is
arranged in a back-to-back orientation, meaning that the ki-
nase domain of each monomer is not in contact with their
target, thus suggesting that the phosphorylation occurs by
contact with another dimer (Fig. 3E). This activating IRE1
transautophosphorylation is further supported by the ability of
the CD to form higher order oligomers (76). The phosphory-
lation occurring on S840, S841, and T844 in yeast IRE1 leads
to the formation of both intramolecular and intermolecular
(with another IRE1) salt bridges promoting the self-activation
of the RNase domain (76). In contrast, hyperphosphorylation
of a 28 amino acid loop (residues 864–892) containing seven
serine and two threonine residues may participate in the
deactivation of IRE1 by promoting its disassembly, a regulatory
mechanism to adapt to a sustained ER stress (77). In human,
the crystal structure of the IRE1 CD was found to resemble
that of the yeast IRE1, with a back-to-back orientation bringing
the RNase domains in close proximity, compatible with the
cleavage of known substrates (78, 79) (Fig. 3F, left). This would
be relevant regarding the modest conservation of sequence
(~60% similarity) (Fig. 2B). Although, another structure was
obtained after IRE1 dephosphorylation, called the face-to-face
conformation (Fig. 3F, right), where the RNase domains of two
monomers have minimal interactions. This last orientation
brings another possibility regarding the trans-
autophosphorylation, where the kinase domain of a monomer
is in close contact with the activation loop of a monomer to
achieve the reciprocal phosphorylation at S724, S726, and S729
(80, 81). Despite the lake of observation of this crystal struc-
ture in yeast, the interface between the two kinase domains is
conserved, suggesting that such conformation exists for all
IRE1 homologs. These two states are part of a model in which
IRE1 first acquires the face-to-face conformation and then
proceeds to the transphosphorylation of the activation loop,
leading to reorganization in a back-to-back conformation that
brings two RNase domains in proximity, thus promoting IRE1
activity. Additionally, an oligomeric structure of yeast IRE1
CDs was obtained by crystallography, displaying fourteen
monomers assembled in seven back-to-back dimers exerting a
52 degree rotation between each dimer, therefore confirming
the ability of the CD to form high order oligomer (Fig. 3G)

(76). Nevertheless, this structure does not take into consider-
ation the constraints imposed by the membrane insertion of
IRE1. A foci structure containing five IRE1 dimers was further
observed in yeast by microscopy. Altogether, it shows that
IRE1 monomers gather into larger signaling platform at the ER
(63). To allow such a complex, the IRE1 linker sequence
positioned between the kinase/RNase and the TMDs might be
of importance by providing flexibility to the protein. In
agreement with this, an *in vitro* experiment evaluating the
oligomerization capacity of the cytosolic portion of IRE1
showed that deletion of the linker region prevents formation of
oligomers (76). To date, there is no available crystal structure
of a full-length IRE1 in any species, thereby preventing the
elucidation of the exact conformation of IRE1 active form,
since its oligomerization and activation may also be governed
(at least in part) by the luminal domain.

While IRE1 activation might be subjected to different modes
that remain debated (see Activation in ER lumen below), it is
possible that the changes in sequence and structure between
yeast and human IRE1 (Figs. 2A and 3, A and B) lead to
modulation of the signaling properties. Especially, as the
luminal domain is the most divergent between species, it can
confer a variability in binding misfolded proteins or other
partners, leading to the complexification and fine-tuning of this
signaling. This would be concomitant with the appearance of
other UPR sensors in metazoans. Nonetheless, one must notice
that despite the three UPR sensors are expressed in Caeno-
rhabditis elegans, the IRE1-XBP1s axis was shown to represent
the main path to cope with ER stress in this organism (82).
Additionally, while the CD retains a similar structure between
species, the changes in sequence and length of this domain
(~50 amino acids more in the yeast IRE1 compared to human)
may modify the oligomerization kinetic as yeast IRE1 was
observed to form higher order oligomer, which were not
observed in human so far. Those differences also emerge as a
parameter regulating the binding and response to small mol-
ecules. Indeed, quercetin, a flavonoid, was first found to bind to
yeast IRE1 in a small pocket formed by kinase and RNase
domain residues (amino acids 838–844 and 1036–1040, called
Q-site) (83). This compound induces a modification of yeast
IRE1 nucleotide-binding site and further activates IRE1. Later,
an O-glycoside quercetin derivative, the quercitrin, was char-
acterized to bind to another binding site, located at the dimer
interface region of the human RNase domain (amino acids
830–834). This molecule leads to an opposite effect, by inhib-
iting IRE1 (84). Altogether, it suggests that IRE1 from yeast and
human respond differently to external stimuli.

Protein-protein interactions regulating IRE1 signaling:
the IRE1 signalosome

Protein-protein interactions

Activation in ER lumen

IRE1 luminal domain has mainly been described as
responsible for sensing misfolded proteins. Yet these sensing
mechanisms are not fully understood and three models are
currently proposed. On the one hand, the "direct association"
IRE1 CD

The CD of IRE1 is the signaling region of IRE1. As such, it
shows two main domains exerting, respectively a serine/thre-
onine kinase activity and an RNase activity. The former leads
to IRE1 transautophoshorylation upon oligomerization,
thereby leading to the activation of the latter. IRE1 RNase
induces the noncanonical splicing of HAC1 mRNA in yeast
and of XBP1 mRNA in human, with further ligation by the
Rlg1 (yeast) or RtcB (human) RNA ligases, respectively. This
process allows for the synthesis of the spliced form of the
XBP1s/Hac1¹p transcription factors, thereby promoting the
expression of UPR target genes (28, 73, 74). IRE1 cytosolic
regions presents two main fragments: a kinase domain (yeast
674–980; human 571–832), and a RNase domain (yeast
983–1115; human 835–963) (Fig. 2A). Structural evidence in
yeast showed that the phosphorylated IRE1 kinase/RNase
domain is able to dimerize in its active form (75). This dimer is
arranged in a back-to-back orientation, meaning that the ki-
nase domain of each monomer is not in contact with their
target, thus suggesting that the phosphorylation occurs by
contact with another dimer (Fig. 3E). This activating IRE1
transautophosphorylation is further supported by the ability of
the CD to form higher order oligomers (76). The phosphory-
lation occurring on S840, S841, and T844 in yeast IRE1 leads
to the formation of both intramolecular and intermolecular
(with another IRE1) salt bridges promoting the self-activation
of the RNase domain (76). In contrast, hyperphosphorylation
of a 28 amino acid loop (residues 864–892) containing seven
serine and two threonine residues may participate in the
deactivation of IRE1 by promoting its disassembly, a regulatory
mechanism to adapt to a sustained ER stress (77). In human,
the crystal structure of the IRE1 CD was found to resemble
that of the yeast IRE1, with a back-to-back orientation bringing
the RNase domains in close proximity, compatible with the
cleavage of known substrates (78, 79) (Fig. 3F, left). This would
be relevant regarding the modest conservation of sequence
(~60% similarity) (Fig. 2B). Although, another structure was
obtained after IRE1 dephosphorylation, called the face-to-face
conformation (Fig. 3F, right), where the RNase domains of two
monomers have minimal interactions. This last orientation
brings another possibility regarding the trans-
autophosphorylation, where the kinase domain of a monomer
is in close contact with the activation loop of a monomer to
achieve the reciprocal phosphorylation at S724, S726, and S729
(80, 81). Despite the lake of observation of this crystal struc-
ture in yeast, the interface between the two kinase domains is
conserved, suggesting that such conformation exists for all
IRE1 homologs. These two states are part of a model in which
IRE1 first acquires the face-to-face conformation and then
proceeds to the transphosphorylation of the activation loop,
leading to reorganization in a back-to-back conformation that
brings two RNase domains in proximity, thus promoting IRE1
activity. Additionally, an oligomeric structure of yeast IRE1
CDs was obtained by crystallography, displaying fourteen
monomers assembled in seven back-to-back dimers exerting a
52 degree rotation between each dimer, therefore confirming
the ability of the CD to form high order oligomer (Fig. 3G)

(76). Nevertheless, this structure does not take into consider-
ation the constraints imposed by the membrane insertion of
IRE1. A foci structure containing five IRE1 dimers was further
observed in yeast by microscopy. Altogether, it shows that
IRE1 monomers gather into larger signaling platform at the ER
(63). To allow such a complex, the IRE1 linker sequence
positioned between the kinase/RNase and the TMDs might be
of importance by providing flexibility to the protein. In
agreement with this, an *in vitro* experiment evaluating the
oligomerization capacity of the cytosolic portion of IRE1
showed that deletion of the linker region prevents formation of
oligomers (76). To date, there is no available crystal structure
of a full-length IRE1 in any species, thereby preventing the
elucidation of the exact conformation of IRE1 active form,
since its oligomerization and activation may also be governed
(at least in part) by the luminal domain.

While IRE1 activation might be subjected to different modes
that remain debated (see Activation in ER lumen below), it is
possible that the changes in sequence and structure between
yeast and human IRE1 (Figs. 2A and 3, A and B) lead to
modulation of the signaling properties. Especially, as the
luminal domain is the most divergent between species, it can
confer a variability in binding misfolded proteins or other
partners, leading to the complexification and fine-tuning of this
signaling. This would be concomitant with the appearance of
other UPR sensors in metazoans. Nonetheless, one must notice
that despite the three UPR sensors are expressed in Caeno-
rhabditis elegans, the IRE1-XBP1s axis was shown to represent
the main path to cope with ER stress in this organism (82).
Additionally, while the CD retains a similar structure between
species, the changes in sequence and length of this domain
(~50 amino acids more in the yeast IRE1 compared to human)
may modify the oligomerization kinetic as yeast IRE1 was
observed to form higher order oligomer, which were not
observed in human so far. Those differences also emerge as a
parameter regulating the binding and response to small mol-
ecules. Indeed, quercetin, a flavonoid, was first found to bind to
yeast IRE1 in a small pocket formed by kinase and RNase
domain residues (amino acids 838–844 and 1036–1040, called
Q-site) (83). This compound induces a modification of yeast
IRE1 nucleotide-binding site and further activates IRE1. Later,
an O-glycoside quercetin derivative, the quercitrin, was char-
acterized to bind to another binding site, located at the dimer
interface region of the human RNase domain (amino acids
830–834). This molecule leads to an opposite effect, by inhib-
iting IRE1 (84). Altogether, it suggests that IRE1 from yeast and
human respond differently to external stimuli.

Protein-protein interactions regulating IRE1 signaling:
the IRE1 signalosome

Protein-protein interactions

Activation in ER lumen

IRE1 luminal domain has mainly been described as
responsible for sensing misfolded proteins. Yet these sensing
mechanisms are not fully understood and three models are
currently proposed. On the one hand, the "direct association"
model where peptides exposed at the surface of improperly
folded proteins bind directly to IRE1 NLD, promoting its
oligomerization and further activation. This model was first
thought to occur in yeast but not human IRE1 (85, 86), how-
ever, a recent study described this mechanism in human where
the direct binding induces a conformational change and leads
to IRE1 oligomerization (87). On the other hand, the
"competition models" presents IRE1 as a signal transducer
(instead of direct sensor), where the chaperone BiP interacts
with IRE1 NLD to repress IRE1 signaling. Here, two mecha-
nisms were proposed and are still a matter of debate in the
field. One sustains that in the absence of improperly folded
proteins, the ER chaperone ERD4 binds IRE1 NLD and re-
cruits BiP substrate binding domain to IRE1 NLD, stabilizing
its monomeric form and inhibiting its oligomerization (88, 89).
The other mechanism involves direct binding of BiP nucleo-
tide binding domain to IRE1 NLD, where misfolded proteins
bind BiP substrate binding domain, triggering an allosteric
conformational change, releasing BiP from IRE1 and enabling
its oligomerization (90–92). In both cases, upon presence of
misfolded proteins, BiP is titrated away from IRE1, thus
allowing its oligomerization and further activation. Such
mechanisms do not seem to exist in S. cerevisiae (with the BiP
ortholog kar2) (93, 94). Interestingly, a similar mechanism
seems to occur for IREβ with the ER chaperone AGR2, where
the last binds to IREβ luminal domain to favorize the mono-
meric state (51, 95).

In addition to those models, other PPis in the ER lumen
were shown to modulate IRE1 activation. Immunoprecipi-
tation coupled to mass spectrometry experiments, followed
by biochemical approaches confirmation, showed that the ER
chaperone HSP47 interacts with IRE1 NLD with high affinity
upon ER stress, competing with BiP binding and leading to
IRE1 oligomerization or stabilization of the oligomeric state
(96). At last, oxidation of C148 in IRE1 luminal domain al-
lows the protein disulfide isomerase PDIA6 to form disulfide
bonds with oligomeric IRE1 and converting it into mono-
meric entities therefore limiting its signaling (97, 98). ER-
stress induced phosphorylation of another protein disulfide
isomerase, PDIA1, was also reported to interact with IRE1
NLD, leading to an attenuation of excessive IRE1
activation (99).

Oligomerization

IRE1 oligomerization is necessary to ensure signaling. IRE1
can dimerize and possibly form high order oligomers
composed of several dimers. However, it remains unclear how
the degree of oligomerization defines IRE1 signaling outputs.
While IRE1 tetramers are thought to preferentially catalyze
XBP1 splicing, IRE1 dimers tend to be more prone to mediate
RIDD (55, 76). Using single molecule tracking and HaloTag,
IRE1 was shown to mainly exist in an inactive back-to-back
dimer (100). Upon ER stress, IRE1 dimers form transient
higher order oligomers, allowing the reciprocal dimers to be
positioned in a "face-to-face like" configuration. Therefore, it is
interesting to focus on interactions able to fine tune IRE1

oligomerization state. In this regard, studies reported that the
translocon subunit Sec61β interacts with IRE1 TMD, forming
a complex with another translocon subunit Sec63, preventing
IRE1 oligomerization and further activation by recruiting BiP
(72, 101, 102). Additionally, it was shown that the ribosome-
associated complex (RAC) chaperone is involved in the regu-
lation of IRE1 oligomerization state. Observation of IRE1
cluster foci using microscopy revealed that RAC promotes
IRE1 high order oligomerization to enhance XBP1 mRNA
splicing but not RIDD (103). However, the physical interaction
between RAC and IRE1 needs further confirmation. The
Abelson tyrosine-protein kinase 1 (c-ABL) was found to
interact with IRE1 CD upon ER stress. Independently of its
kinase activity, c-ABL acts as a scaffold protein and promotes
IRE1 oligomerization and further hyperactivation, leading to
RIDD and terminal UPR (104). Moreover, c-ABL modulates
the IRE1 signaling cascade by phosphorylating the tRNA ligase
RtcB at least on three tyrosine residues (306, 316, 475), with
Y306 phosphorylation reducing RtcB interaction with IRE1
and subsequently XBP1 mRNA splicing (105). One might also
hypothesize that dissociation of the IRE1-RtcB complex by c-
ABL might leads to the formation of IRE1 clusters able to
perform RIDD. Altogether, it confers a role of switch to c-ABL,
able to fine-tune IRE1 signaling through different mechanisms.
At last, a link between IRE1 signaling and cytoskeleton was
unveiled. The nonmuscle myosin IIB (NMIIB) interacts with
IRE1 during ER stress. While it seems that NMIIB is not
directly involved in IRE1 phosphorylation and dimerization,
NMIIB drives IRE1 foci formation, promoting splicing of XBP1
mRNA (106). Interestingly, this mechanism was found to
require the ATPase activity of NMIIB as well as a functional
actin network, suggesting that the actomyosin contractility of
NMIIB is required for IRE1α signaling (106).

RNase activity

Regarding interactions directly regulating the RNase activ-
ity, the previously described relationship between IRE1 and
c-ABL was also found to act as a genotoxic stress sensor, by
promoting RIDD activation and further cleavage of PPP2R1A
and RuvB-like AAA ATPase 1 mRNA, two proteins involved in
dephosphorylation of DNA damage proteins (107). The
apoptosis regulator BAX (BAX) and Bcl-2 Homologous
Antagonist/Killer (BAK), two proapoptotic proteins (108),
were shown by coimmunoprecipitation to bind IRE1 (prefer-
entially in its active form) upon high ER stress and were found
to promote XBP1 mRNA splicing (109). Similarly, other PPis
were found to enhance IRE1 signaling. This is illustrated by the
association of IRE1 with the Bcl-2 family members Bcl-2-like
protein 11 (BIM) and PUMA, leading to sustained XBP1
mRNA splicing upon ER stress (110).

Conversely, some PPis were shown to impair IRE1 RNase
activity. An IRE1 immunoprecipitation coupled to mass
spectrometry experiment identified RNH1, a ribonuclease in-
hibitor, to be present in complex with IRE1. This result was
confirmed using both immunoprecipitation and proximity
ligation. Surprisingly, ER stress increases this interaction
model where peptides exposed at the surface of improperly
folded proteins bind directly to IRE1 NLD, promoting its
oligomerization and further activation. This model was first
thought to occur in yeast but not human IRE1 (85, 86), how-
ever, a recent study described this mechanism in human where
the direct binding induces a conformational change and leads
to IRE1 oligomerization (87). On the other hand, the
"competition models" presents IRE1 as a signal transducer
(instead of direct sensor), where the chaperone BiP interacts
with IRE1 NLD to repress IRE1 signaling. Here, two mecha-
nisms were proposed and are still a matter of debate in the
field. One sustains that in the absence of improperly folded
proteins, the ER chaperone ERD4 binds IRE1 NLD and re-
cruits BiP substrate binding domain to IRE1 NLD, stabilizing
its monomeric form and inhibiting its oligomerization (88, 89).
The other mechanism involves direct binding of BiP nucleo-
tide binding domain to IRE1 NLD, where misfolded proteins
bind BiP substrate binding domain, triggering an allosteric
conformational change, releasing BiP from IRE1 and enabling
its oligomerization (90–92). In both cases, upon presence of
misfolded proteins, BiP is titrated away from IRE1, thus
allowing its oligomerization and further activation. Such
mechanisms do not seem to exist in S. cerevisiae (with the BiP
ortholog kar2) (93, 94). Interestingly, a similar mechanism
seems to occur for IREβ with the ER chaperone AGR2, where
the last binds to IREβ luminal domain to favorize the mono-
meric state (51, 95).

In addition to those models, other PPis in the ER lumen
were shown to modulate IRE1 activation. Immunoprecipi-
tation coupled to mass spectrometry experiments, followed
by biochemical approaches confirmation, showed that the ER
chaperone HSP47 interacts with IRE1 NLD with high affinity
upon ER stress, competing with BiP binding and leading to
IRE1 oligomerization or stabilization of the oligomeric state
(96). At last, oxidation of C148 in IRE1 luminal domain al-
lows the protein disulfide isomerase PDIA6 to form disulfide
bonds with oligomeric IRE1 and converting it into mono-
meric entities therefore limiting its signaling (97, 98). ER-
stress induced phosphorylation of another protein disulfide
isomerase, PDIA1, was also reported to interact with IRE1
NLD, leading to an attenuation of excessive IRE1
activation (99).

Oligomerization

IRE1 oligomerization is necessary to ensure signaling. IRE1
can dimerize and possibly form high order oligomers
composed of several dimers. However, it remains unclear how
the degree of oligomerization defines IRE1 signaling outputs.
While IRE1 tetramers are thought to preferentially catalyze
XBP1 splicing, IRE1 dimers tend to be more prone to mediate
RIDD (55, 76). Using single molecule tracking and HaloTag,
IRE1 was shown to mainly exist in an inactive back-to-back
dimer (100). Upon ER stress, IRE1 dimers form transient
higher order oligomers, allowing the reciprocal dimers to be
positioned in a "face-to-face like" configuration. Therefore, it is
interesting to focus on interactions able to fine tune IRE1

oligomerization state. In this regard, studies reported that the
translocon subunit Sec61β interacts with IRE1 TMD, forming
a complex with another translocon subunit Sec63, preventing
IRE1 oligomerization and further activation by recruiting BiP
(72, 101, 102). Additionally, it was shown that the ribosome-
associated complex (RAC) chaperone is involved in the regu-
lation of IRE1 oligomerization state. Observation of IRE1
cluster foci using microscopy revealed that RAC promotes
IRE1 high order oligomerization to enhance XBP1 mRNA
splicing but not RIDD (103). However, the physical interaction
between RAC and IRE1 needs further confirmation. The
Abelson tyrosine-protein kinase 1 (c-ABL) was found to
interact with IRE1 CD upon ER stress. Independently of its
kinase activity, c-ABL acts as a scaffold protein and promotes
IRE1 oligomerization and further hyperactivation, leading to
RIDD and terminal UPR (104). Moreover, c-ABL modulates
the IRE1 signaling cascade by phosphorylating the tRNA ligase
RtcB at least on three tyrosine residues (306, 316, 475), with
Y306 phosphorylation reducing RtcB interaction with IRE1
and subsequently XBP1 mRNA splicing (105). One might also
hypothesize that dissociation of the IRE1-RtcB complex by c-
ABL might leads to the formation of IRE1 clusters able to
perform RIDD. Altogether, it confers a role of switch to c-ABL,
able to fine-tune IRE1 signaling through different mechanisms.
At last, a link between IRE1 signaling and cytoskeleton was
unveiled. The nonmuscle myosin IIB (NMIIB) interacts with
IRE1 during ER stress. While it seems that NMIIB is not
directly involved in IRE1 phosphorylation and dimerization,
NMIIB drives IRE1 foci formation, promoting splicing of XBP1
mRNA (106). Interestingly, this mechanism was found to
require the ATPase activity of NMIIB as well as a functional
actin network, suggesting that the actomyosin contractility of
NMIIB is required for IRE1α signaling (106).

RNase activity

Regarding interactions directly regulating the RNase activ-
ity, the previously described relationship between IRE1 and
c-ABL was also found to act as a genotoxic stress sensor, by
promoting RIDD activation and further cleavage of PPP2R1A
and RuvB-like AAA ATPase 1 mRNA, two proteins involved in
dephosphorylation of DNA damage proteins (107). The
apoptosis regulator BAX (BAX) and Bcl-2 Homologous
Antagonist/Killer (BAK), two proapoptotic proteins (108),
were shown by coimmunoprecipitation to bind IRE1 (prefer-
entially in its active form) upon high ER stress and were found
to promote XBP1 mRNA splicing (109). Similarly, other PPis
were found to enhance IRE1 signaling. This is illustrated by the
association of IRE1 with the Bcl-2 family members Bcl-2-like
protein 11 (BIM) and PUMA, leading to sustained XBP1
mRNA splicing upon ER stress (110).

Conversely, some PPis were shown to impair IRE1 RNase
activity. An IRE1 immunoprecipitation coupled to mass
spectrometry experiment identified RNH1, a ribonuclease in-
hibitor, to be present in complex with IRE1. This result was
confirmed using both immunoprecipitation and proximity
ligation. Surprisingly, ER stress increases this interaction
between IRE1 and RNH1, thereby reducing XBP1 mRNA
splicing (111). This counterintuitive outcome might indicate
that IRE1–RNH1 interaction acts as a regulatory mechanism,
where RNH1 decreases IRE1 activity upon prolonged ER
stress. The BAX inhibitor-1 (BI-1) was found to compete with
BAX and BAK for IRE1. BI-1 interacts with the IRE1 C-ter-
minal region and reduces IRE1 RNase activity (112, 113).
Under basal conditions, the interaction of the SH2/SH3-
containing adaptor protein 1 with IRE1 carboxy-terminal re-
gion was shown to repress IRE1, a mechanism proposed to
avoid a signaling leakage (114).

Protein modification

Phosphorylation-dependent regulations of IRE1 signaling

Evidences unveiled a role of IRE1 phosphorylation in determining its signaling outcomes (115). As such, IRE1 faces phosphorylation-dephosphorylation cycles relying on PPis. These regulating events were studied and revealed interesting mechanisms. Regarding the downregulation of IRE1 signaling through dephosphorylation, two yeast phosphatases—Ptc2p serine/threonine phosphatase of type 2C and Dcr2—were found to interact with phosphorylated IRE1 and further de-phosphorylate it to downregulate the splicing of HAC1 mRNA (26, 116). Interestingly, it was found that Dcr2 specifically interacts with IRE1 when phosphorylated at S840 and S841 residues (116). This finding was extended to human thanks to another study describing IRE1 interaction with the ER membrane phosphatase PP2Ce, the human homolog of the S. cerevisiae Ptcp2 (117). Here, PP2Ce dephosphorylates IRE1 S724 upon ER stress, acting as a negative regulator of IRE1 signaling. Of note, both IRE1α and IRE1β were shown to partner with PP2Ce, unveiling a probable common regulatory mechanism (117). Coimmunoprecipitation assays found that the scaffold protein RACK1 interacts with IRE1 linker domain (between the kinase and the TMD) (118, 119). In basal condition, RACK1 recruits the phosphatase PP2A, which de-phosphorylate IRE1 (120). A recent article uncovered through screening different serine-threonine phosphatases that the small carboxy-terminal domain phosphatase 3 was able to interact directly with IRE1 and further dephosphorylate the protein (120). We previously mentioned the crosstalk between the UPR sensors. In this regard, a study investigated the role of the PERK pathway in PPi-dependent attenuation of IRE1 signaling. Upon acute ER stress, PERK-prolonged activation leads to the phosphatase RPAP2 to interact and de-phosphorylate IRE1, thereby decreasing its signaling (41). Another mechanism involving the protein tyrosine phosphatase PTP1B was shown to regulate both catalytic and scaffolding functions of IRE1 (105, 121). Here, PTP1B deficiency or catalytically inactive impairs XBP1 splicing and c-Jun-N-terminal-kinase (JNK) activation (121), while further investigations revealed that PTP1B dephosphorylates RtcB, thus enhancing its association with IRE1 for XBP1 mRNA splicing (105). However, to date, direct interaction between IRE1 and PTP1B requires confirmation.

Conversely, other proteins were found to phosphorylate
IRE1. Interestingly, the previously mentioned IRE1-RACK1
complex was found to play another role. Upon increased ER
stress, RACK1 recruits the AMP-activated protein kinase,
which phosphorylates IRE1 (118, 119). Hence, RACK1 acts as a
platform regulating IRE1 signaling according to ER stress
through the phosphorylation status. Glucagon-stimulated PKA
was reported to phosphorylate IRE1, indicating a role in
glucose metabolism and suggesting an ER-lumen-indepen-
dent activation (122). At last, a mechanism by which KRas
mutated tumors elaborate tumor resistance to KRas inhibitors
was shown to involve ERK1/2-mediated phosphorylation of
IRE1, thus preventing its degradation by the ERAD machinery
(120). This led us to further document the role of post-
translational modifications (PTMs) of IRE1 and its
degradation.

PTMs and IRE1 degradation

N-glycosylation is a PTM occurring during synthesis and maturation of ER proteins, participating in their folding and functionality (8, 123). IRE1 is N-glycosylated at N176 (124, 125) but there is no clear evidence mentioning a role for this PTM in regulating IRE1 activity. Despite that, few studies reported that mutations preventing this PTM do not impair IRE1 folding nor oligomerization properties (124, 125). Also, N-glycosylation disruption causes ER stress, and some might hypothesize that it could be a sensor-initiating IRE1 cascade.

Alternatively, ERAD was shown to control IRE1 expression and activity, as shown by the SYVN1/SEL1L-dependent ubiquitination of IRE1, promoting its degradation (126, 127). In this case, IRE1 is sent to ERAD in basal conditions, where both BiP and the lectin protein OS9 interacts with IRE1 in the lumen, recruiting the SYNV1/SEL1L complex and subsequently allowing IRE1 cytosolic ubiquitination by the E3 ligase SYNV1 (127). OS9 being a lectin, we suggest that it recognizes IRE1 through its N-glycan chain, therefore conferring the N-glycan a role in IRE1 stability. Cytomegalovirus M50 was found to take advantage of this ERAD-dependent IRE1 degradation, where the M50 viral protein interacts and tethers with both IRE1 and SEL1L to promote IRE1 degradation (128). IRE1 was also found to be a substrate of the E3 ubiquitin-protein ligase CHIP (also involved in ERAD). Authors proposed a dual role for CHIP-dependent IRE1 ubiquitination during ER stress: first, ubiquitination of IRE1 K545, enhancing IRE1 phosphorylation and second, of IRE1 K828, facilitating recruitment of TNF receptor-associated factor 2 (TRAF2) for JNK pathway activation (129). This is supported by another study, where the E3 ligase RNF13 interacts with IRE1, exerting enhanced XBP1 splicing and JNK activation (130). On another note, a recent study identified IRE1 to be a substrate of the mitochondria ubiquitin ligase MARCHF5 (MITOL), which ubiquitinylate IRE1 K481 with a K63 ubiquitin chain at mitochondria-ER contact sites (131). This ubiquitination was proposed to disrupt BIM stabilizing interaction with IRE1 oligomers, thus inhibiting IRE1 signaling (110, 131).
between IRE1 and RNH1, thereby reducing XBP1 mRNA
splicing (111). This counterintuitive outcome might indicate
that IRE1-RNH1 interaction acts as a regulatory mechanism,
where RNH1 decreases IRE1 activity upon prolonged ER
stress. The BAX inhibitor-1 (BI-1) was found to compete with
BAX and BAK for IRE1. BI-1 interacts with the IRE1 C-ter-
minal region and reduces IRE1 RNase activity (112, 113).
Under basal conditions, the interaction of the SH2/SH3-
containing adaptor protein 1 with IRE1 carboxy-terminal re-
gion was shown to repress IRE1, a mechanism proposed to
avoid a signaling leakage (114).

Protein modification

Phosphorylation-dependent regulations of IRE1 signaling

Evidences unveiled a role of IRE1 phosphorylation in determining its signaling outcomes (115). As such, IRE1 faces phosphorylation-dephosphorylation cycles relying on PPis. These regulating events were studied and revealed interesting mechanisms. Regarding the downregulation of IRE1 signaling through dephosphorylation, two yeast phosphatases—Ptc2p serine/threonine phosphatase of type 2C and Dcr2—were found to interact with phosphorylated IRE1 and further de-phosphorylate it to downregulate the splicing of HAC1 mRNA (26, 116). Interestingly, it was found that Dcr2 specifically interacts with IRE1 when phosphorylated at S840 and S841 residues (116). This finding was extended to human thanks to another study describing IRE1 interaction with the ER membrane phosphatase PP2Ce, the human homolog of the S. cerevisiae Ptcp2 (117). Here, PP2Ce dephosphorylates IRE1 S724 upon ER stress, acting as a negative regulator of IRE1 signaling. Of note, both IRE1α and IRE1β were shown to partner with PP2Ce, unveiling a probable common regulatory mechanism (117). Coimmunoprecipitation assays found that the scaffold protein RACK1 interacts with IRE1 linker domain (between the kinase and the TMD) (118, 119). In basal condition, RACK1 recruits the phosphatase PP2A, which de-phosphorylate IRE1 (120). A recent article uncovered through screening different serine-threonine phosphatases that the small carboxy-terminal domain phosphatase 3 was able to interact directly with IRE1 and further dephosphorylate the protein (120). We previously mentioned the crosstalk between the UPR sensors. In this regard, a study investigated the role of the PERK pathway in PPi-dependent attenuation of IRE1 signaling. Upon acute ER stress, PERK-prolonged activation leads to the phosphatase RPAP2 to interact and de-phosphorylate IRE1, thereby decreasing its signaling (41). Another mechanism involving the protein tyrosine phosphatase PTP1B was shown to regulate both catalytic and scaffolding functions of IRE1 (105, 121). Here, PTP1B deficiency or catalytically inactive impairs XBP1 splicing and c-Jun-N-terminal-kinase (JNK) activation (121), while further investigations revealed that PTP1B dephosphorylates RtcB, thus enhancing its association with IRE1 for XBP1 mRNA splicing (105). However, to date, direct interaction between IRE1 and PTP1B requires confirmation.

Conversely, other proteins were found to phosphorylate
IRE1. Interestingly, the previously mentioned IRE1-RACK1
complex was found to play another role. Upon increased ER
stress, RACK1 recruits the AMP-activated protein kinase,
which phosphorylates IRE1 (118, 119). Hence, RACK1 acts as a
platform regulating IRE1 signaling according to ER stress
through the phosphorylation status. Glucagon-stimulated PKA
was reported to phosphorylate IRE1, indicating a role in
glucose metabolism and suggesting an ER-lumen-indepen-
dent activation (122). At last, a mechanism by which KRas
mutated tumors elaborate tumor resistance to KRas inhibitors
was shown to involve ERK1/2-mediated phosphorylation of
IRE1, thus preventing its degradation by the ERAD machinery
(120). This led us to further document the role of post-
translational modifications (PTMs) of IRE1 and its
degradation.

PTMs and IRE1 degradation

N-glycosylation is a PTM occurring during synthesis and maturation of ER proteins, participating in their folding and functionality (8, 123). IRE1 is N-glycosylated at N176 (124, 125) but there is no clear evidence mentioning a role for this PTM in regulating IRE1 activity. Despite that, few studies reported that mutations preventing this PTM do not impair IRE1 folding nor oligomerization properties (124, 125). Also, N-glycosylation disruption causes ER stress, and some might hypothesize that it could be a sensor-initiating IRE1 cascade.

Alternatively, ERAD was shown to control IRE1 expression and activity, as shown by the SYVN1/SEL1L-dependent ubiquitination of IRE1, promoting its degradation (126, 127). In this case, IRE1 is sent to ERAD in basal conditions, where both BiP and the lectin protein OS9 interacts with IRE1 in the lumen, recruiting the SYNV1/SEL1L complex and subsequently allowing IRE1 cytosolic ubiquitination by the E3 ligase SYNV1 (127). OS9 being a lectin, we suggest that it recognizes IRE1 through its N-glycan chain, therefore conferring the N-glycan a role in IRE1 stability. Cytomegalovirus M50 was found to take advantage of this ERAD-dependent IRE1 degradation, where the M50 viral protein interacts and tethers with both IRE1 and SEL1L to promote IRE1 degradation (128). IRE1 was also found to be a substrate of the E3 ubiquitin-protein ligase CHIP (also involved in ERAD). Authors proposed a dual role for CHIP-dependent IRE1 ubiquitination during ER stress: first, ubiquitination of IRE1 K545, enhancing IRE1 phosphorylation and second, of IRE1 K828, facilitating recruitment of TNF receptor-associated factor 2 (TRAF2) for JNK pathway activation (129). This is supported by another study, where the E3 ligase RNF13 interacts with IRE1, exerting enhanced XBP1 splicing and JNK activation (130). On another note, a recent study identified IRE1 to be a substrate of the mitochondria ubiquitin ligase MARCHF5 (MITOL), which ubiquitinylate IRE1 K481 with a K63 ubiquitin chain at mitochondria-ER contact sites (131). This ubiquitination was proposed to disrupt BIM stabilizing interaction with IRE1 oligomers, thus inhibiting IRE1 signaling (110, 131).
# IRE1 PPis associated with other functions/outcomes

## Protein-protein interactions

### Apoptosis

IRE1 signaling has been connected to the activation of the apoptosis cascade. Indeed, upon prolonged ER stress, IRE1 RIDD activity cleaves nonselectively several prosurvival miRNAs. Among them, miR-17, a miRNA known to repress TXNIP expression, subsequently triggers caspase 1 and 2/interleukin-1β-dependent apoptosis (36, 132). Of note another report showed that ER stress did not lead to the activation of Caspase 2 to initiate apoptosis (133). Several other studies have also connected IRE1 PPis with cell death, occurring during terminal UPR (134). This is linked to the activation of the proapoptotic JNK pathway, which first involves the interaction of IRE1 with TRAF2 and the further recruitment ASK1, also known as apoptotic signaling kinase 1 (135, 136). ASK1 may then phosphorylate JNK, thereby initiating the JNK pathway and leading to apoptosis. A new actor in the TRAF2/ASK1/JNK axis was also identified as the ASK1-interacting-protein 1 (AIP1), which, upon ER stress, is recruited on IRE1 prior to TRAF2 and further promotes JNK pathway activation (137). Additionally, this study presented evidence that AIP1 could enhance IRE1 dimerization (137). We previously mentioned IRE1 interactions with the Bcl-2 family member BIM and PUMA. These interactions occur through the BH3 domain of these proteins, which is necessary for their canonical proapoptotic signaling (109). It was shown that knock out of BIM and PUMA reduces both IRE1 RIDD and JNK activation (110). Therefore, BIM and PUMA might promote apoptosis by sustaining cleavage of prosurvival miRNAs by RIDD and activation of the JNK pathway. Accordingly, the other members of the Bcl-2 family BAX and BAK, also containing BH3 domains, may act in a similar manner by interacting with IRE1 (109). Therefore, it confers a noncanonical role of these BH3 domain proteins in inducing cell death. Conversely, we also hypothesize on this model that the previously described protein BI-1 can inhibit cell death by competing with BAX and BAK interactions with IRE1. More recently, another IRE1 PPi was reported to exert antiapoptotic functions, involving the previously described interaction between IRE1 and MITOL, shown to inhibit JNK phosphorylation (131).

### Cytoskeleton

In the past years, few studies reported a crosstalk between UPR and cytoskeleton as mentioned with NMIIB or through the interaction of the actin-binding protein FLNA with PERK (106, 138). Interestingly, a mechanism involving both IRE1 and FLNA showed that IRE1 promotes cell migration through FLNA phosphorylation. In this study, IRE1 associates with FLNA and acts as a recruiting platform for the kinase PKCα, enabling the phosphorylation of FLNA, which in turns drives actin cytoskeleton remodeling and cell migration (139). This interaction was shown to participate in migration of cells infected by toxoplasma, where the parasite engages the

IRE1-FLNA axis (140). Additionally, several high-throughput interactome screenings identified other cytoskeletal proteins such as CORO1 proteins or tubulin α and β, however, they require further validation (111, 139). IRE1 interacting with cytoskeleton proteins may also participate in membrane contact sites between ER and plasma membrane (ER-PM contact sites). This complex is known to regulate $Ca^{2+}$ and lipid homeostasis, two processes associated with IRE1 signaling (141, 142).

### Translation machinery: signal recognition particle and ribosome

High translational load can result in nonefficient protein folding in the ER by overwhelming the folding capacity of this compartment, hence, triggering ER stress. Given the proximity of IRE1 with the protein synthesis machinery, a crosstalk has been discovered between IRE1 signaling and mRNA translation. Cross-linking experiments revealed interactions between IRE1 and translating ribosomes at the cytosolic ER surface, through the ribosomal subunits RPL9, RPL10A, RPS14, and RPS24 (143). In the same study, immunoprecipitation analyses showed IRE1 interaction with proteins from the signal recognition particle (SRP) (143). These results correlate with studies describing that XBP1 mRNA begins translation before being brought to the ER membrane by the SRP pathway to undergo IRE1-dependent splicing (144–146). Moreover, we can hypothesize a role for IRE1 in monitoring the newly synthesized polypeptides entering the ER during the cotranslational translocation, as a way to avoid protein accumulation into the ER. Of note, those interactions with SRP and ribosomal proteins did not modify the catalytic functions of IRE1, thus suggesting a catalytic-independent regulation of the UPR which remains to be further investigated.

### Calcium homeostasis

The ER is an intracellular organelle known to be involved in the maintenance of $Ca^{2+}$ homeostasis. It was suggested that IRE1 could play a role in regulating $Ca^{2+}$ metabolism independently of its enzymatic activities (147, 148). Indeed, IRE1 CD was shown to interact with the inositol-1,4,5-trisphosphate receptors. This PPi allows mitochondria to localize close to the ER in the mitochondria-associated ER membrane, another membrane contact site, and further promote calcium transfer to support mitochondrial respiration (148). This is supported by recent findings describing the interaction between IRE1 and the stromal interaction molecule 1 (STIM1) in the ER lumen (149). This interaction connects the UPR and $Ca^{2+}$ metabolism, where STIM1 promotes XBP1 mRNA splicing upon ER $Ca^{2+}$ store depletion and IRE1 promotes STIM1-dependent ER-PM contact sites, triggering the store operated $Ca^{2+}$ entry pathway.

### Direct IRE1 kinase-dependent phosphorylation

For many years, the only known substrate of the IRE1 kinase domain was IRE1 itself. However, few recent publications reported novel targets for IRE1 kinase activity. The previously described IRE1-RACK1 axis was shown to participate in BCL-2 phosphorylation by IRE1 at S70, a phosphorylation site
# IRE1 PPis associated with other functions/outcomes

## Protein-protein interactions

### Apoptosis

IRE1 signaling has been connected to the activation of the apoptosis cascade. Indeed, upon prolonged ER stress, IRE1 RIDD activity cleaves nonselectively several prosurvival miRNAs. Among them, miR-17, a miRNA known to repress TXNIP expression, subsequently triggers caspase 1 and 2/interleukin-1β-dependent apoptosis (36, 132). Of note another report showed that ER stress did not lead to the activation of Caspase 2 to initiate apoptosis (133). Several other studies have also connected IRE1 PPis with cell death, occurring during terminal UPR (134). This is linked to the activation of the proapoptotic JNK pathway, which first involves the interaction of IRE1 with TRAF2 and the further recruitment ASK1, also known as apoptotic signaling kinase 1 (135, 136). ASK1 may then phosphorylate JNK, thereby initiating the JNK pathway and leading to apoptosis. A new actor in the TRAF2/ASK1/JNK axis was also identified as the ASK1-interacting-protein 1 (AIP1), which, upon ER stress, is recruited on IRE1 prior to TRAF2 and further promotes JNK pathway activation (137). Additionally, this study presented evidence that AIP1 could enhance IRE1 dimerization (137). We previously mentioned IRE1 interactions with the Bcl-2 family member BIM and PUMA. These interactions occur through the BH3 domain of these proteins, which is necessary for their canonical proapoptotic signaling (109). It was shown that knock out of BIM and PUMA reduces both IRE1 RIDD and JNK activation (110). Therefore, BIM and PUMA might promote apoptosis by sustaining cleavage of prosurvival miRNAs by RIDD and activation of the JNK pathway. Accordingly, the other members of the Bcl-2 family BAX and BAK, also containing BH3 domains, may act in a similar manner by interacting with IRE1 (109). Therefore, it confers a noncanonical role of these BH3 domain proteins in inducing cell death. Conversely, we also hypothesize on this model that the previously described protein BI-1 can inhibit cell death by competing with BAX and BAK interactions with IRE1. More recently, another IRE1 PPi was reported to exert antiapoptotic functions, involving the previously described interaction between IRE1 and MITOL, shown to inhibit JNK phosphorylation (131).

### Cytoskeleton

In the past years, few studies reported a crosstalk between UPR and cytoskeleton as mentioned with NMIIB or through the interaction of the actin-binding protein FLNA with PERK (106, 138). Interestingly, a mechanism involving both IRE1 and FLNA showed that IRE1 promotes cell migration through FLNA phosphorylation. In this study, IRE1 associates with FLNA and acts as a recruiting platform for the kinase PKCα, enabling the phosphorylation of FLNA, which in turns drives actin cytoskeleton remodeling and cell migration (139). This interaction was shown to participate in migration of cells infected by toxoplasma, where the parasite engages the

IRE1-FLNA axis (140). Additionally, several high-throughput interactome screenings identified other cytoskeletal proteins such as CORO1 proteins or tubulin α and β, however, they require further validation (111, 139). IRE1 interacting with cytoskeleton proteins may also participate in membrane contact sites between ER and plasma membrane (ER-PM contact sites). This complex is known to regulate $Ca^{2+}$ and lipid homeostasis, two processes associated with IRE1 signaling (141, 142).

### Translation machinery: signal recognition particle and ribosome

High translational load can result in nonefficient protein folding in the ER by overwhelming the folding capacity of this compartment, hence, triggering ER stress. Given the proximity of IRE1 with the protein synthesis machinery, a crosstalk has been discovered between IRE1 signaling and mRNA translation. Cross-linking experiments revealed interactions between IRE1 and translating ribosomes at the cytosolic ER surface, through the ribosomal subunits RPL9, RPL10A, RPS14, and RPS24 (143). In the same study, immunoprecipitation analyses showed IRE1 interaction with proteins from the signal recognition particle (SRP) (143). These results correlate with studies describing that XBP1 mRNA begins translation before being brought to the ER membrane by the SRP pathway to undergo IRE1-dependent splicing (144–146). Moreover, we can hypothesize a role for IRE1 in monitoring the newly synthesized polypeptides entering the ER during the cotranslational translocation, as a way to avoid protein accumulation into the ER. Of note, those interactions with SRP and ribosomal proteins did not modify the catalytic functions of IRE1, thus suggesting a catalytic-independent regulation of the UPR which remains to be further investigated.

### Calcium homeostasis

The ER is an intracellular organelle known to be involved in the maintenance of $Ca^{2+}$ homeostasis. It was suggested that IRE1 could play a role in regulating $Ca^{2+}$ metabolism independently of its enzymatic activities (147, 148). Indeed, IRE1 CD was shown to interact with the inositol-1,4,5-trisphosphate receptors. This PPi allows mitochondria to localize close to the ER in the mitochondria-associated ER membrane, another membrane contact site, and further promote calcium transfer to support mitochondrial respiration (148). This is supported by recent findings describing the interaction between IRE1 and the stromal interaction molecule 1 (STIM1) in the ER lumen (149). This interaction connects the UPR and $Ca^{2+}$ metabolism, where STIM1 promotes XBP1 mRNA splicing upon ER $Ca^{2+}$ store depletion and IRE1 promotes STIM1-dependent ER-PM contact sites, triggering the store operated $Ca^{2+}$ entry pathway.

### Direct IRE1 kinase-dependent phosphorylation

For many years, the only known substrate of the IRE1 kinase domain was IRE1 itself. However, few recent publications reported novel targets for IRE1 kinase activity. The previously described IRE1-RACK1 axis was shown to participate in BCL-2 phosphorylation by IRE1 at S70, a phosphorylation site
known to exert antiapoptotic functions (119). Liu and colleagues also showed that IRE1 interacts and promotes AMP-activated protein kinase phosphorylation exerting prosurvival signaling, nevertheless the direct/indirect modification remains unclear (119). The fragile X mental retardation protein (FMRP) was also identified as a new IRE1 kinase substrate, possibly regulating cholesterol transport and efferocytosis (150). In addition, pumilio, a RNA-binding protein in *drosophila*, was shown to be phosphorylated by IRE1, allowing its binding to XBP1 mRNA and protecting it against IRE1 RIDD activity (151). PUM1, the human homolog of pumilio, has also been shown to be phosphorylated by IRE1 (151). Another study has reported an interaction between IRE1 and

the IκBα kinase complex (IKK). The use of a kinase defective IRE1 mutant (K599A) revealed a reduced phosphorylation of IκBα kinase, rescued by transfection with a WT IRE1 construct (152). This links IRE1 with the NF-κB pathway, involved in many cellular processes such as inflammation or apoptosis. At last, the sphingosine 1-phosphate lyase (SPL) was shown to be phosphorylated by IRE1, inhibiting its enzymatic activity and resulting in activation of a mitochondrial-dependent UPR (153). Even though the experiments performed were mostly *in vitro*, these discoveries open a new field in the exploration of IRE1 kinase-dependent signaling and raise questions about potential implication of IRE1 in processes not limited to the UPR.

<figure><img src="image_11.png" /><figcaption>**Figure 4.** Summary of the current IRE1 interactome. Clusters of IRE1 interactors regulating IRE1 signaling (A) or exerting noncanonical functions (B). C, integrated view of IRE1 interactions mediating its signaling and other functions. (*) indicates IRE1β-specific interactions. AIP1, ASK1-interacting protein 1; FMRP, fragile X mental retardation protein; IKK, IκBα kinase complex; IRE1, inositol-requiring enzyme 1; MITOL, mitochondria ubiquitin ligase MARCHF5; SPL, sphingosine 1-phosphate lyase</figcaption></figure>
known to exert antiapoptotic functions (119). Liu and colleagues also showed that IRE1 interacts and promotes AMP-activated protein kinase phosphorylation exerting prosurvival signaling, nevertheless the direct/indirect modification remains unclear (119). The fragile X mental retardation protein (FMRP) was also identified as a new IRE1 kinase substrate, possibly regulating cholesterol transport and efferocytosis (150). In addition, pumilio, a RNA-binding protein in *drosophila*, was shown to be phosphorylated by IRE1, allowing its binding to XBP1 mRNA and protecting it against IRE1 RIDD activity (151). PUM1, the human homolog of pumilio, has also been shown to be phosphorylated by IRE1 (151). Another study has reported an interaction between IRE1 and

the IκBα kinase complex (IKK). The use of a kinase defective IRE1 mutant (K599A) revealed a reduced phosphorylation of IκBα kinase, rescued by transfection with a WT IRE1 construct (152). This links IRE1 with the NF-κB pathway, involved in many cellular processes such as inflammation or apoptosis. At last, the sphingosine 1-phosphate lyase (SPL) was shown to be phosphorylated by IRE1, inhibiting its enzymatic activity and resulting in activation of a mitochondrial-dependent UPR (153). Even though the experiments performed were mostly *in vitro*, these discoveries open a new field in the exploration of IRE1 kinase-dependent signaling and raise questions about potential implication of IRE1 in processes not limited to the UPR.

<figure><img src="image_11.png" /><figcaption>**Figure 4.** Summary of the current IRE1 interactome. Clusters of IRE1 interactors regulating IRE1 signaling (A) or exerting noncanonical functions (B). C, integrated view of IRE1 interactions mediating its signaling and other functions. (*) indicates IRE1β-specific interactions. AIP1, ASK1-interacting protein 1; FMRP, fragile X mental retardation protein; IKK, IκBα kinase complex; IRE1, inositol-requiring enzyme 1; MITOL, mitochondria ubiquitin ligase MARCHF5; SPL, sphingosine 1-phosphate lyase</figcaption></figure>
Concluding remarks and outstanding questions

IRE1 first appeared as a transducer of the UPR, in order to
restore a balance in ER homeostasis and help cells adapt to
stress. The evolutive interactome unveils possible new roles of
IRE1 in mammals. IRE1 is connected with immunity, inflam-
mation, and metabolism and is therefore involved in disease
progression (154, 155). While most of the studies focus on
IRE1 RNase activity, understanding the dynamic of IRE1 in-
teractions may also provide useful insights on disease devel-
opment and potential therapies. This review summarizes the
known IRE1 interactors and classifies them according to their
role in regulating IRE1 signaling (Fig. 4A) or exerting nonca-
nonical (RNase independent) functions (Fig. 4B). While a
majority of the interactors were characterized to regulate IRE1
canonical signaling, an increasing number of evidences tend to
attribute to IRE1 a wide range of diverse functions (Fig. 4C).

IRE1 signaling can therefore be regulated by interactors at
every step of its activation. We term this protein interaction
network regulating IRE1 signaling functions as the IRE1 sig-
nalosome. The latter can be modulated by specific signals;
hence, deciphering how the regulation of the IRE1 cascade
exerts different cellular outcomes might represent an
appealing avenue to better understand IRE1 signaling. We
found recent studies describing new targets of the IRE1 kinase
activity. Protein phosphorylation is a crucial mechanism in
transducing signals and regulating cellular functions (156).
Considering IRE1 as a regular kinase opens a new field of in-
vestigations. Studying the sequence and modeling the complex
of the known IRE1-dependent phosphorylation could provide
some insights on potential other targets. Lastly, IRE1 is known
for its role in regulating the translational load through RIDD in
the cytosol. However, some studies reported a possible local-
ization of IRE1 in the nucleus (143, 157–159) and confirming
this would require further investigations. Considering the
continuity between ER membrane and nuclear envelope, we
can hypothesize a nonconventional localization of IRE1, where
it would unveil a new landscape of regulators and regulate
gene expression independently from RIDD.

Acknowledgments—We thank Dr Xavier Guillory for his assistance regarding IRE1 structural representations.

Author contributions—S. L. G., H. L., M. A., and E. C. investigation; S. L. G., H. L., M. A., and E. C. conceptualization; S. L. G., H. L., M. A., and E. C. writing-original draft; S. L. G., H. L., M. A., and E. C. writing-review and editing.

Funding and additional information—This work was funded by grants from Institut National du Cancer (INCa, PLBio) and Fondation pour la Recherche Médicale (FRM; EQU202403018041), INSERM (International Research Project – TUPRIC) to E. C. S. L. G. was funded by a PhD scholarship from the French Ministry of research and H. L. was funded by a postdoctoral fellowship from Région Bretagne.

Conflict of interest—E. C. is the founder of Thabor Therapeutics (https://www.thabor-tx.com/). The other authors declare that they have no conflicts of interest with the contents of this article.

Abbreviations—The abbreviations used are: AH, amphipathic helix; ASK1, apoptotic signaling kinase 1; ATF, activating transcription factor; BI-1, BAX inhibitor-1; CD, cytosolic domain; ER, endoplasmic reticulum; ERAD, ER-associated degradation; IRE1, inositol-requiring enzyme 1; JNK, c-Jun-N-terminal-kinase; MHC, major histocompatibility peptide-binding complex; NLD, N-terminal luminal domain; NM IIB, nonmuscle myosin IIB; PDI, protein disulfide isomerase; PERK, PKR-like endoplasmic reticulum kinase; PTM, posttranslational modification; RAC, ribosome-associated complex; RIDD, regulated IRE1-dependent decay; RNH, ribonuclease inhibitor; SRP, signal recognition particle; STIM1, stromal interaction molecule 1; UPR, unfolded protein response; XBP1, X-box binding protein 1.

References

1. Garnier, C. (1897) *Les filaments basaux des cellules glandulaires*. Note préliminaire., Bibliographie Anatomique [Paris]: [Berger-Levrault]

2. Porter, K. R., Claude, A., and Fullam, E. F. (1945) A study of tissue culture cells by electron microscopy. *J. Exp. Med.* **81**, 233–246

3. Ashby, M. C., and Tepikin, A. V. (2001) ER calcium and the functions of intracellular organelles. *Semin. Cell Dev. Biol.* **12**, 11–17

4. Fagone, P., and Jackowski, S. (2009) Membrane phospholipid synthesis and endoplasmic reticulum function. *J. Lipid Res.* **50**, S311–S316

5. Braakman, I., and Hebert, D. N. (2013) Protein folding in the endoplasmic reticulum. *Cold Spring Harb. Perspect. Biol.* **5**, a013201

6. Almanza, A., Carlesso, A., Chintha, C., Creedican, S., Doultsinos, D., Leuzzi, B., *et al.* (2019) Endoplasmic reticulum stress signalling - from basic mechanisms to clinical applications. *FEBS J.* **286**, 241–278

7. Ghaemmaghami, S., Huh, W. K., Bower, K., Howson, R. W., Belle, A., Dephoure, N., *et al.* (2003) Global analysis of protein expression in yeast. *Nature* **425**, 737–741

8. Hanson, S. R., Culyba, E. K., Hsu, T. L., Wong, C. H., Kelly, J. W., and Powers, E. T. (2009) The core trisaccharide of an N-linked glycoprotein intrinsically accelerates folding and enhances stability. *Proc. Natl. Acad. Sci. U. S. A.* **106**, 3131–3136

9. Ellgaard, L., and Helenius, A. (2003) Quality control in the endoplasmic reticulum. *Nat. Rev. Mol. Cell Biol.* **4**, 181–191

10. Olzmann, J. A., Kopito, R. R., and Christianson, J. C. (2013) The mammalian endoplasmic reticulum-associated degradation system. *Cold Spring Harb. Perspect. Biol.* **5**, a013185

11. Schröder, M., and Kaufman, R. J. (2005) The mammalian unfolded protein response. *Annu. Rev. Biochem.* **74**, 739–789

12. Shore, G. C., Papa, F. R., and Oakes, S. A. (2011) Signaling cell death from the endoplasmic reticulum stress response. *Curr. Opin. Cell Biol.* **23**, 143–149

13. McGrath, E. P., Centonze, F. G., Chevet, E., Avril, T., and Lafont, E. (2021) Death sentence: the tale of a fallen endoplasmic reticulum. *Biochim. Biophys. Acta Mol. Cell Res.* **1868**, 119001

14. Hughes, D., and Mallucci, G. R. (2019) The unfolded protein response in neurodegenerative disorders – therapeutic modulation of the PERK pathway. *FEBS J.* **286**, 342–355

15. Madden, E., Logue, S. E., Healy, S. J., Manie, S., and Samali, A. (2019) The role of the unfolded protein response in cancer progression: from oncogenesis to chemoresistance. *Biol. Cell* **111**, 1–17

16. Hetz, C., Zhang, K., and Kaufman, R. J. (2020) Mechanisms, regulation and functions of the unfolded protein response. *Nat. Rev. Mol. Cell Biol.* **21**, 421–438

17. Vattem, K. M., and Wek, R. C. (2004) Reinitiation involving upstream ORFs regulates ATF4 mRNA translation in mammalian cells. *Proc. Natl. Acad. Sci. U. S. A.* **101**, 11269–11274

18. B'chir, W., Maurin, A. C., Carraro, V., Averous, J., Jousse, C., Muranishi, Y., et al. (2013) The eIF2α/ATF4 pathway is essential for stress-induced autophagy gene expression. *Nucleic Acids Res.* **41**, 7683–7699
Concluding remarks and outstanding questions

IRE1 first appeared as a transducer of the UPR, in order to
restore a balance in ER homeostasis and help cells adapt to
stress. The evolutive interactome unveils possible new roles of
IRE1 in mammals. IRE1 is connected with immunity, inflam-
mation, and metabolism and is therefore involved in disease
progression (154, 155). While most of the studies focus on
IRE1 RNase activity, understanding the dynamic of IRE1 in-
teractions may also provide useful insights on disease devel-
opment and potential therapies. This review summarizes the
known IRE1 interactors and classifies them according to their
role in regulating IRE1 signaling (Fig. 4A) or exerting nonca-
nonical (RNase independent) functions (Fig. 4B). While a
majority of the interactors were characterized to regulate IRE1
canonical signaling, an increasing number of evidences tend to
attribute to IRE1 a wide range of diverse functions (Fig. 4C).

IRE1 signaling can therefore be regulated by interactors at
every step of its activation. We term this protein interaction
network regulating IRE1 signaling functions as the IRE1 sig-
nalosome. The latter can be modulated by specific signals;
hence, deciphering how the regulation of the IRE1 cascade
exerts different cellular outcomes might represent an
appealing avenue to better understand IRE1 signaling. We
found recent studies describing new targets of the IRE1 kinase
activity. Protein phosphorylation is a crucial mechanism in
transducing signals and regulating cellular functions (156).
Considering IRE1 as a regular kinase opens a new field of in-
vestigations. Studying the sequence and modeling the complex
of the known IRE1-dependent phosphorylation could provide
some insights on potential other targets. Lastly, IRE1 is known
for its role in regulating the translational load through RIDD in
the cytosol. However, some studies reported a possible local-
ization of IRE1 in the nucleus (143, 157–159) and confirming
this would require further investigations. Considering the
continuity between ER membrane and nuclear envelope, we
can hypothesize a nonconventional localization of IRE1, where
it would unveil a new landscape of regulators and regulate
gene expression independently from RIDD.

Acknowledgments—We thank Dr Xavier Guillory for his assistance regarding IRE1 structural representations.

Author contributions—S. L. G., H. L., M. A., and E. C. investigation; S. L. G., H. L., M. A., and E. C. conceptualization; S. L. G., H. L., M. A., and E. C. writing-original draft; S. L. G., H. L., M. A., and E. C. writing-review and editing.

Funding and additional information—This work was funded by grants from Institut National du Cancer (INCa, PLBio) and Fondation pour la Recherche Médicale (FRM; EQU202403018041), INSERM (International Research Project – TUPRIC) to E. C. S. L. G. was funded by a PhD scholarship from the French Ministry of research and H. L. was funded by a postdoctoral fellowship from Région Bretagne.

Conflict of interest—E. C. is the founder of Thabor Therapeutics (https://www.thabor-tx.com/). The other authors declare that they have no conflicts of interest with the contents of this article.

*Abbreviations*—The abbreviations used are: AH, amphipathic helix; ASK1, apoptotic signaling kinase 1; ATF, activating transcription factor; BI-1, BAX inhibitor-1; CD, cytosolic domain; ER, endoplasmic reticulum; ERAD, ER-associated degradation; IRE1, inositol-requiring enzyme 1; JNK, c-Jun-N-terminal-kinase; MHC, major histocompatibility peptide-binding complex; NLD, N-terminal luminal domain; NM IIB, nonmuscle myosin IIB; PDI, protein disulfide isomerase; PERK, PKR-like endoplasmic reticulum kinase; PTM, posttranslational modification; RAC, ribosome-associated complex; RIDD, regulated IRE1-dependent decay; RNH, ribonuclease inhibitor; SRP, signal recognition particle; STIM1, stromal interaction molecule 1; UPR, unfolded protein response; XBP1, X-box binding protein 1.

References

1. Garnier, C. (1897) *Les filaments basaux des cellules glandulaires*. Note préliminaire., Bibliographie Anatomique [Paris]: [Berger-Levrault]

2. Porter, K. R., Claude, A., and Fullam, E. F. (1945) A study of tissue culture cells by electron microscopy. *J. Exp. Med.* **81**, 233–246

3. Ashby, M. C., and Tepikin, A. V. (2001) ER calcium and the functions of intracellular organelles. *Semin. Cell Dev. Biol.* **12**, 11–17

4. Fagone, P., and Jackowski, S. (2009) Membrane phospholipid synthesis and endoplasmic reticulum function. *J. Lipid Res.* **50**, S311–S316

5. Braakman, I., and Hebert, D. N. (2013) Protein folding in the endoplasmic reticulum. *Cold Spring Harb. Perspect. Biol.* **5**, a013201

6. Almanza, A., Carlesso, A., Chintha, C., Creedican, S., Doultsinos, D., Leuzzi, B., *et al.* (2019) Endoplasmic reticulum stress signalling - from basic mechanisms to clinical applications. *FEBS J.* **286**, 241–278

7. Ghaemmaghami, S., Huh, W. K., Bower, K., Howson, R. W., Belle, A., Dephoure, N., *et al.* (2003) Global analysis of protein expression in yeast. *Nature* **425**, 737–741

8. Hanson, S. R., Culyba, E. K., Hsu, T. L., Wong, C. H., Kelly, J. W., and Powers, E. T. (2009) The core trisaccharide of an N-linked glycoprotein intrinsically accelerates folding and enhances stability. *Proc. Natl. Acad. Sci. U. S. A.* **106**, 3131–3136

9. Ellgaard, L., and Helenius, A. (2003) Quality control in the endoplasmic reticulum. *Nat. Rev. Mol. Cell Biol.* **4**, 181–191

10. Olzmann, J. A., Kopito, R. R., and Christianson, J. C. (2013) The mammalian endoplasmic reticulum-associated degradation system. *Cold Spring Harb. Perspect. Biol.* **5**, a013185

11. Schröder, M., and Kaufman, R. J. (2005) The mammalian unfolded protein response. *Annu. Rev. Biochem.* **74**, 739–789

12. Shore, G. C., Papa, F. R., and Oakes, S. A. (2011) Signaling cell death from the endoplasmic reticulum stress response. *Curr. Opin. Cell Biol.* **23**, 143–149

13. McGrath, E. P., Centonze, F. G., Chevet, E., Avril, T., and Lafont, E. (2021) Death sentence: the tale of a fallen endoplasmic reticulum. *Biochim. Biophys. Acta Mol. Cell Res.* **1868**, 119001

14. Hughes, D., and Mallucci, G. R. (2019) The unfolded protein response in neurodegenerative disorders – therapeutic modulation of the PERK pathway. *FEBS J.* **286**, 342–355

15. Madden, E., Logue, S. E., Healy, S. J., Manie, S., and Samali, A. (2019) The role of the unfolded protein response in cancer progression: from oncogenesis to chemoresistance. *Biol. Cell* **111**, 1–17

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20. Novoa, I., Zhang, Y., Zeng, H., Jungreis, R., Harding, H. P., and Ron, D. (2003) Stress-induced gene expression requires programmed recovery from translational repression. *EMBO J.* **22**, 1180–1187

21. Novoa, I., Zeng, H., Harding, H. P., and Ron, D. (2001) Feedback inhibition of the unfolded protein response by GADD34-mediated dephosphorylation of eIF2α. *J. Cell Biol.* **153**, 1011–1022

22. Haze, K., Yoshida, H., Yanagi, H., Yura, T., and Mori, K. (1999) Mammalian transcription factor ATF6 is synthesized as a trans-membrane protein and activated by proteolysis in response to endoplasmic reticulum stress. *Mol. Biol. Cell* **10**, 3787–3799

23. Wu, J., Rutkowski, D. T., Dubois, M., Swathirajan, J., Saunders, T., Wang, J., et al. (2007) ATF6α Optimizes long-term endoplasmic reticulum function to protect cells from chronic stress. *Dev. Cell* **13**, 351–364

24. Shoulders, M. D., Ryno, L. M., Genereux, J. C., Moresco, J. J., Tu, P. G., Wu, C., et al. (2013) Stress-independent activation of XBP1s and/or ATF6 reveals three functionally diverse ER proteostasis environments. *Cell Rep.* **3**, 1279–1292

25. Mori, K., Ma, W., Gething, M. J., and Sambrook, J. (1993) A transmembrane protein with a cdc2+/CDC28-related kinase activity is required for signaling from the ER to the nucleus. *Cell* **74**, 743–756

26. Welihinda, A. A., Tirasophon, W., Green, S. R., and Kaufman, R. J. (1998) Protein serine/threonine phosphatase Ptc2p negatively regulates the unfolded-protein response by dephosphorylating Ire1p kinase. *Mol. Cell. Biol.* **18**, 1967–1977

27. Cox, J. S., Shamu, C. E., and Walter, P. (1993) Transcriptional induction of genes encoding endoplasmic reticulum resident proteins requires a transmembrane protein kinase. *Cell* **73**, 1197–1206

28. Jurkin, J., Henkel, T., Nielsen, A. F., Minnich, M., Popow, J., Kaufmann, T., et al. (2014) The mammalian tRNA ligase complex mediates splicing of XBP1 mRNA and controls antibody secretion in plasma cells. *EMBO J.* **33**, 2922–2936

29. Kosmaczewski, S. G., Edwards, T. J., Han, S. M., Eckwahl, M. J., Meyer, B. I., Peach, S., et al. (2014) The RtcB RNA ligase is an essential component of the metazoan unfolded protein response. *EMBO Rep.* **15**, 1278–1285

30. Lu, Y., Liang, F. X., and Wang, X. (2014) A synthetic biology approach identifies the mammalian UPR RNA ligase RtcB. *Mol. Cell* **55**, 758–770

31. Sidrauski, C., and Walter, P. (1997) The transmembrane kinase Ire1p is a site-specific Endonuclease that initiates mRNA splicing in the unfolded protein response. *Cell* **90**, 1031–1039

32. Lee, A. H., Iwakoshi, N. N., and Glimcher, L. H. (2003) XBP-1 regulates a subset of endoplasmic reticulum resident chaperone genes in the unfolded protein response. *Mol. Cell. Biol.* **23**, 7448–7459

33. Lee, A. H., Scapa, E. F., Cohen, D. E., and Glimcher, L. H. (2008) Regulation of hepatic lipogenesis by the transcription factor XBP1. *Science* **320**, 1492–1496

34. Yamamoto, K., Suzuki, N., Wada, T., Okada, T., Yoshida, H., Kaufman, R. J., et al. (2008) Human HRD1 promoter Carries a functional unfolded protein response element to which XBP1 but not ATF6 directly binds. *J. Biochem.* **144**, 477–486

35. Hollien, J., and Weissman, J. S. (2006) Decay of endoplasmic reticulum-localized mRNAs during the unfolded protein response. *Science* **313**, 104–107

36. Upton, J. P., Wang, L., Han, D., Wang, E. S., Huskey, N. E., Lim, L., et al. (2012) IRE1α cleaves select microRNAs during ER stress to Derepress translation of Proapoptotic caspase-2. *Science* **338**, 818–822

37. Oikawa, D., Tokuda, M., Hosoda, A., and Iwawaki, T. (2010) Identification of a consensus element recognized and cleaved by IRE1α. *Nucleic Acids Res.* **38**, 6265–6273

38. Le Thomas, A., Ferri, E., Marsters, S., Harnoss, J. M., Lawrence, D. A., Zuazo-Gaztelu, I., et al. (2021) Decoding non-canonical mRNA decay by the endoplasmic-reticulum stress sensor IRE1α. *Nat Commun* **12**, 7310

39. Yoshida, H., Matsui, T., Yamamoto, A., Okada, T., and Mori, K. (2001) XBP1 mRNA is induced by ATF6 and spliced by IRE1 in response to ER stress to Produce a highly active transcription factor. *Cell* **107**, 881–891

40. Tsuru, A., Imai, Y., Saito, M., and Kohno, K. (2016) Novel mechanism of enhancing IRE1α-XBP1 signalling via the PERK-ATF4 pathway. *Sci. Rep.* **6**, 24217

41. Chang, T. K., Lawrence, D. A., Lu, M., Tan, J., Harnoss, J. M., Marsters, S. A., et al. (2018) Coordination between two branches of the unfolded protein response determines apoptotic cell fate. *Mol. Cell* **71**, 629–636.e5

42. Lin, J. H., Li, H., Yasumura, D., Cohen, H. R., Zhang, C., Panning, B., et al. (2007) IRE1 signaling affects cell fate during the unfolded protein response. *Science* **318**, 944–949

43. Iwawaki, T., Hosoda, A., Okuda, T., Kamigori, Y., Nomura-Furuwatari, C., Kimata, Y., et al. (2001) Translational control by the ER trans-membrane kinase/ribonuclease IRE1 under ER stress. *Nat. Cell Biol.* **3**, 158–164

44. Bertolotti, A., Wang, X., Novoa, I., Jungreis, R., Schlessinger, K., Cho, J. H., et al. (2001) Increased sensitivity to dextran sodium sulfate colitis in IRE1β-deficient mice. *J. Clin. Invest.* **107**, 585–593

45. Martino, M. B., Jones, L., Brighton, B., Ehre, C., Abdulah, L., Davis, C. W., et al. (2013) The ER stress transducer IRE1β is required for airway epithelial mucin production. *Mucosal Immunol.* **6**, 639–654

46. Imagawa, Y., Hosoda, A., Sasaki, S., Tsuru, A., and Kohno, K. (2008) RNase domains determine the functional difference between IRE1α and IRE1β. *FEBS Lett.* **582**, 656–660

47. Grey, M. J., Cloots, E., Simpson, M. S., LeDuc, N., Serebrenik, Y. V., De Luca, H., et al. (2020) IRE1β negatively regulates IRE1α signaling in response to endoplasmic reticulum stress. *J. Cell Biol.* **219**, e201904048

48. Cloots, E., Simpson, M. S., De Nolf, C., Lencer, W. I., Janssens, S., and Grey, M. J. (2021) Evolution and function of the epithelial cell-specific ER stress sensor IRE1β. *Mucosal Immunol.* **14**, 1235–1246

49. Calfon, M., Zeng, H., Urano, F., Till, J. H., Hubbard, S. R., Harding, H. P., et al. (2002) IRE1 couples endoplasmic reticulum load to secretory capacity by processing the XBP-1 mRNA. *Nature* **415**, 92–96

50. Feldman, H. C., Vidadala, V. N., Potter, Z. E., Papa, F. R., Backes, B. J., and Maly, D. J. (2019) Development of a chemical Toolset for studying the Paralog-specific function of IRE1. *ACS Chem. Biol.* **14**, 2595–2605

51. Cloots, E., Guilbert, P., Provost, M., Neidhardt, L., Van de Velde, E., Fayazpour, F., et al. (2023) Activation of goblet-cell stress sensor IRE1β is controlled by the mucin chaperone AGR2. *EMBO J.* **43**, 1–24

52. Zhang, L., Zhang, C., and Wang, A. (2016) Divergence and conservation of the major UPR branch IRE1-bZIP signaling pathway across eukaryotes. *Sci. Rep.* **6**, 27362

53. Altschul, S. F., Gish, W., Miller, W., Myers, E. W., and Lipman, D. J. (1990) Basic local alignment search tool. *J. Mol. Biol.* **215**, 403–410

54. Cox, J. S., and Walter, P. (1996) A novel mechanism for regulating activity of a transcription factor that controls the unfolded protein response. *Cell* **87**, 391–404

55. Tam, A. B., Koong, A. C., and Niwa, M. (2014) Ire1 has Distinct catalytic mechanisms for XBP1/HAC1 splicing and RIDD. *Cell Rep.* **9**, 850–858

56. Kimmig, P., Diaz, M., Zheng, J., Williams, C. C., Lang, A., Aragón, T., et al. (2012) The unfolded protein response in fission yeast modulates stability of select mRNAs to maintain protein homeostasis. *Elife.* **1**, e00048

57. Hollien, J. (2013) Evolution of the unfolded protein response. *Biochim. Biophys. Acta* **1833**, 2458–2463

58. Alonso-López, D., Campos-Laborie, F. J., Gutiérrez, M. A., Lambourne, L., Calderwood, M. A., Vidal, M., et al. (2019) APID database: redefining protein-protein interaction experimental evidences and binary interactomes. *Database (Oxford)* **2019**, baz005

59. Szklarczyk, D., Gable, A. L., Lyon, D., Junge, A., Wyder, S., Huerta-Cepas, J., et al. (2019) STRING v11: protein-protein association networks with increased coverage, supporting functional discovery in genome-wide experimental datasets. *Nucleic Acids Res.* **47**, D607–D613

60. Oughtred, R., Rust, J., Chang, C., Breitkreutz, B. J., Stark, C., Willems, A., et al. (2021) The BioGRID database: a comprehensive biomedical resource of curated protein, genetic, and chemical interactions. *Protein Sci.* **30**, 187–200

61. Kolberg, L., Raudvere, U., Kuzmin, I., Adler, P., Vilo, J., and Peterson, H. (2023) g:Profiler—interoperable web service for functional enrichment
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20. Novoa, I., Zhang, Y., Zeng, H., Jungreis, R., Harding, H. P., and Ron, D. (2003) Stress-induced gene expression requires programmed recovery from translational repression. *EMBO J.* **22**, 1180–1187

21. Novoa, I., Zeng, H., Harding, H. P., and Ron, D. (2001) Feedback inhibition of the unfolded protein response by GADD34-mediated dephosphorylation of eIF2α. *J. Cell Biol.* **153**, 1011–1022

22. Haze, K., Yoshida, H., Yanagi, H., Yura, T., and Mori, K. (1999) Mammalian transcription factor ATF6 is synthesized as a trans-membrane protein and activated by proteolysis in response to endoplasmic reticulum stress. *Mol. Biol. Cell* **10**, 3787–3799

23. Wu, J., Rutkowski, D. T., Dubois, M., Swathirajan, J., Saunders, T., Wang, J., et al. (2007) ATF6α Optimizes long-term endoplasmic reticulum function to protect cells from chronic stress. *Dev. Cell* **13**, 351–364

24. Shoulders, M. D., Ryno, L. M., Genereux, J. C., Moresco, J. J., Tu, P. G., Wu, C., et al. (2013) Stress-independent activation of XBP1s and/or ATF6 reveals three functionally diverse ER proteostasis environments. *Cell Rep.* **3**, 1279–1292

25. Mori, K., Ma, W., Gething, M. J., and Sambrook, J. (1993) A transmembrane protein with a cdc2+/CDC28-related kinase activity is required for signaling from the ER to the nucleus. *Cell* **74**, 743–756

26. Welihinda, A. A., Tirasophon, W., Green, S. R., and Kaufman, R. J. (1998) Protein serine/threonine phosphatase Ptc2p negatively regulates the unfolded-protein response by dephosphorylating Ire1p kinase. *Mol. Cell. Biol.* **18**, 1967–1977

27. Cox, J. S., Shamu, C. E., and Walter, P. (1993) Transcriptional induction of genes encoding endoplasmic reticulum resident proteins requires a transmembrane protein kinase. *Cell* **73**, 1197–1206

28. Jurkin, J., Henkel, T., Nielsen, A. F., Minnich, M., Popow, J., Kaufmann, T., et al. (2014) The mammalian tRNA ligase complex mediates splicing of XBP1 mRNA and controls antibody secretion in plasma cells. *EMBO J.* **33**, 2922–2936

29. Kosmaczewski, S. G., Edwards, T. J., Han, S. M., Eckwahl, M. J., Meyer, B. I., Peach, S., et al. (2014) The RtcB RNA ligase is an essential component of the metazoan unfolded protein response. *EMBO Rep.* **15**, 1278–1285

30. Lu, Y., Liang, F. X., and Wang, X. (2014) A synthetic biology approach identifies the mammalian UPR RNA ligase RtcB. *Mol. Cell* **55**, 758–770

31. Sidrauski, C., and Walter, P. (1997) The transmembrane kinase Ire1p is a site-specific Endonuclease that initiates mRNA splicing in the unfolded protein response. *Cell* **90**, 1031–1039

32. Lee, A. H., Iwakoshi, N. N., and Glimcher, L. H. (2003) XBP-1 regulates a subset of endoplasmic reticulum resident chaperone genes in the unfolded protein response. *Mol. Cell. Biol.* **23**, 7448–7459

33. Lee, A. H., Scapa, E. F., Cohen, D. E., and Glimcher, L. H. (2008) Regulation of hepatic lipogenesis by the transcription factor XBP1. *Science* **320**, 1492–1496

34. Yamamoto, K., Suzuki, N., Wada, T., Okada, T., Yoshida, H., Kaufman, R. J., et al. (2008) Human HRD1 promoter Carries a functional unfolded protein response element to which XBP1 but not ATF6 directly binds. *J. Biochem.* **144**, 477–486

35. Hollien, J., and Weissman, J. S. (2006) Decay of endoplasmic reticulum-localized mRNAs during the unfolded protein response. *Science* **313**, 104–107

36. Upton, J. P., Wang, L., Han, D., Wang, E. S., Huskey, N. E., Lim, L., et al. (2012) IRE1α cleaves select microRNAs during ER stress to Derepress translation of Proapoptotic caspase-2. *Science* **338**, 818–822

37. Oikawa, D., Tokuda, M., Hosoda, A., and Iwawaki, T. (2010) Identification of a consensus element recognized and cleaved by IRE1α. *Nucleic Acids Res.* **38**, 6265–6273

38. Le Thomas, A., Ferri, E., Marsters, S., Harnoss, J. M., Lawrence, D. A., Zuazo-Gaztelu, I., et al. (2021) Decoding non-canonical mRNA decay by the endoplasmic-reticulum stress sensor IRE1α. *Nat Commun* **12**, 7310

39. Yoshida, H., Matsui, T., Yamamoto, A., Okada, T., and Mori, K. (2001) XBP1 mRNA is induced by ATF6 and spliced by IRE1 in response to ER stress to Produce a highly active transcription factor. *Cell* **107**, 881–891

40. Tsuru, A., Imai, Y., Saito, M., and Kohno, K. (2016) Novel mechanism of enhancing IRE1α-XBP1 signalling via the PERK-ATF4 pathway. *Sci. Rep.* **6**, 24217

41. Chang, T. K., Lawrence, D. A., Lu, M., Tan, J., Harnoss, J. M., Marsters, S. A., et al. (2018) Coordination between two branches of the unfolded protein response determines apoptotic cell fate. *Mol. Cell* **71**, 629–636.e5

42. Lin, J. H., Li, H., Yasumura, D., Cohen, H. R., Zhang, C., Panning, B., et al. (2007) IRE1 signaling affects cell fate during the unfolded protein response. *Science* **318**, 944–949

43. Iwawaki, T., Hosoda, A., Okuda, T., Kamigori, Y., Nomura-Furuwatari, C., Kimata, Y., et al. (2001) Translational control by the ER trans-membrane kinase/ribonuclease IRE1 under ER stress. *Nat. Cell Biol.* **3**, 158–164

44. Bertolotti, A., Wang, X., Novoa, I., Jungreis, R., Schlessinger, K., Cho, J. H., et al. (2001) Increased sensitivity to dextran sodium sulfate colitis in IRE1β-deficient mice. *J. Clin. Invest.* **107**, 585–593

45. Martino, M. B., Jones, L., Brighton, B., Ehre, C., Abdulah, L., Davis, C. W., et al. (2013) The ER stress transducer IRE1β is required for airway epithelial mucin production. *Mucosal Immunol.* **6**, 639–654

46. Imagawa, Y., Hosoda, A., Sasaki, S., Tsuru, A., and Kohno, K. (2008) RNase domains determine the functional difference between IRE1α and IRE1β. *FEBS Lett.* **582**, 656–660

47. Grey, M. J., Cloots, E., Simpson, M. S., LeDuc, N., Serebrenik, Y. V., De Luca, H., et al. (2020) IRE1β negatively regulates IRE1α signaling in response to endoplasmic reticulum stress. *J. Cell Biol.* **219**, e201904048

48. Cloots, E., Simpson, M. S., De Nolf, C., Lencer, W. I., Janssens, S., and Grey, M. J. (2021) Evolution and function of the epithelial cell-specific ER stress sensor IRE1β. *Mucosal Immunol.* **14**, 1235–1246

49. Calfon, M., Zeng, H., Urano, F., Till, J. H., Hubbard, S. R., Harding, H. P., et al. (2002) IRE1 couples endoplasmic reticulum load to secretory capacity by processing the XBP-1 mRNA. *Nature* **415**, 92–96

50. Feldman, H. C., Vidadala, V. N., Potter, Z. E., Papa, F. R., Backes, B. J., and Maly, D. J. (2019) Development of a chemical Toolset for studying the Paralog-specific function of IRE1. *ACS Chem. Biol.* **14**, 2595–2605

51. Cloots, E., Guilbert, P., Provost, M., Neidhardt, L., Van de Velde, E., Fayazpour, F., et al. (2023) Activation of goblet-cell stress sensor IRE1β is controlled by the mucin chaperone AGR2. *EMBO J.* **43**, 1–24

52. Zhang, L., Zhang, C., and Wang, A. (2016) Divergence and conservation of the major UPR branch IRE1-bZIP signaling pathway across eukaryotes. *Sci. Rep.* **6**, 27362

53. Altschul, S. F., Gish, W., Miller, W., Myers, E. W., and Lipman, D. J. (1990) Basic local alignment search tool. *J. Mol. Biol.* **215**, 403–410

54. Cox, J. S., and Walter, P. (1996) A novel mechanism for regulating activity of a transcription factor that controls the unfolded protein response. *Cell* **87**, 391–404

55. Tam, A. B., Koong, A. C., and Niwa, M. (2014) Ire1 has Distinct catalytic mechanisms for XBP1/HAC1 splicing and RIDD. *Cell Rep.* **9**, 850–858

56. Kimmig, P., Diaz, M., Zheng, J., Williams, C. C., Lang, A., Aragón, T., et al. (2012) The unfolded protein response in fission yeast modulates stability of select mRNAs to maintain protein homeostasis. *Elife.* **1**, e00048

57. Hollien, J. (2013) Evolution of the unfolded protein response. *Biochim. Biophys. Acta* **1833**, 2458–2463

58. Alonso-López, D., Campos-Laborie, F. J., Gutiérrez, M. A., Lambourne, L., Calderwood, M. A., Vidal, M., et al. (2019) APID database: redefining protein-protein interaction experimental evidences and binary interactomes. *Database (Oxford)* **2019**, baz005

59. Szklarczyk, D., Gable, A. L., Lyon, D., Junge, A., Wyder, S., Huerta-Cepas, J., et al. (2019) STRING v11: protein-protein association networks with increased coverage, supporting functional discovery in genome-wide experimental datasets. *Nucleic Acids Res.* **47**, D607–D613

60. Oughtred, R., Rust, J., Chang, C., Breitkreutz, B. J., Stark, C., Willems, A., et al. (2021) The BioGRID database: a comprehensive biomedical resource of curated protein, genetic, and chemical interactions. *Protein Sci.* **30**, 187–200

61. Kolberg, L., Raudvere, U., Kuzmin, I., Adler, P., Vilo, J., and Peterson, H. (2023) g:Profiler—interoperable web service for functional enrichment
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141. Zaman, M. F., Nenadic, A., Radojičić, A., Rosado, A., and Beh, C. T. (2020) Sticking with it: ER-PM membrane contact sites as a Coordinating Nexus for regulating lipids and proteins at the cell Cortex. *Front. Cell Dev. Biol.* **8**, 675

142. Li, C., Qian, T., He, R., Wan, C., Liu, Y., and Yu, H. (2021) Endoplasmic reticulum–plasma membrane contact sites: regulators, mechanisms, and physiological functions. *Front. Cell Dev. Biol.* **9**, 627700

143. Acosta-Alvear, D., Karagöz, G. E., Fröhlich, F., Li, H., Walther, T. C., and Walter, P. (2018) The unfolded protein response and endoplasmic reticulum protein targeting machineries converge on the stress sensor IRE1. *Elife* **7**, e43036

144. Yanagitani, K., Imagawa, Y., Iwawaki, T., Hosoda, A., Saito, M., Kimata, Y., et al. (2009) Cotranslational targeting of XBP1 protein to the membrane promotes Cytoplasmic splicing of its Own mRNA. *Mol. Cell* **34**, 191–200

145. Yanagitani, K., Kimata, Y., Kadokura, H., and Kohno, K. (2011) Translational pausing ensures membrane targeting and Cytoplasmic splicing of XBP1u mRNA. *Science* **331**, 586–589

146. Kanda, S., Yanagitani, K., Yokota, Y., Esaki, Y., and Kohno, K. (2016) Autonomous translational pausing is required for XBP1u mRNA recruitment to the ER via the SRP pathway. *Proc. Natl. Acad. Sci. U. S. A.* **113**, E5886–E5895

147. Son, S. M., Byun, J., Roh, S. E., Kim, S. J., and Mook-Jung, I. (2014) Reduced IRE1α mediates apoptotic cell death by disrupting calcium homeostasis via the InsP3 receptor. *Cell Death Dis.* **5**, e1188

148. Carreras-Sureda, A., Jaña, F., Urra, H., Durand, S., Mortenson, D. E., Sagredo, A., et al. (2019) Non-canonical function of IRE1α determines mitochondria-associated endoplasmic reticulum composition to control calcium transfer and bioenergetics. *Nat. Cell Biol.* **21**, 755–767

149. Carreras-Sureda, A., Zhang, X., Laubry, L., Brunetti, J., Koenig, S., Wang, X., et al. (2023) The ER stress sensor IRE1 interacts with STIM1 to promote store-operated calcium entry, T cell activation, and muscular differentiation. *Cell Rep.* **42**, 113540

150. Yildirim, Z., Baboo, S., Hamid, S. M., Dogan, A. E., Tufanli, O., Robi-chaud, S., et al. (2022) Intercepting IRE1 kinase-FMRP signaling prevents atherosclerosis progression. *EMBO Mol. Med.* **14**, e15344

151. Cairrão, F., Santos, C. C., Le Thomas, A., Marsters, S., Ashkenazi, A., and Domingos, P. M. (2022) Pumilio protects Xbp1 mRNA from regulated Ire1-dependent decay. *Nat. Commun.* **13**, 1587

152. Tam, A. B., Mercado, E. L., Hoffmann, A., and Niwa, M. (2012) ER stress activates NF-κB by integrating functions of basal IKK activity, IRE1 and PERK. *PLoS One* **7**, e45078

153. Yildirim, A. D., Citir, M., Dogan, A. E., Veli, Z., Yildirim, Z., Tufanli, O., et al. (2022) ER stress-induced sphingosine-1-phosphate lyase phosphorylation potentiates the mitochondrial unfolded protein response. *J. Lipid Res.* **63**, 100279

154. Huang, S., Xing, Y., and Liu, Y. (2019) Emerging roles for the ER stress sensor IRE1α in metabolic regulation and disease. *J. Biol. Chem.* **294**, 18726–18741

155. Dong, H., Adams, N. M., Xu, Y., Cao, J., Allan, D. S. J., Carlyle, J. R., et al. (2019) The IRE1 endoplasmic reticulum stress sensor activates natural killer cell immunity in part by regulating c-Myc. *Nat. Immunol.* **20**, 865–878

156. Ardito, F., Giuliani, M., Perrone, D., Troiano, G., and Lo Muzio, L. (2017) The crucial role of protein phosphorylation in cell signaling and its use as targeted therapy (Review). *Int. J. Mol. Med.* **40**, 271–280

157. Niwa, M., Sidrauski, C., Kaufman, R. J., and Walter, P. (1999) A role for Presenilin-1 in nuclear accumulation of Ire1 fragments and induction of the mammalian unfolded protein response. *Cell* **99**, 691–702

158. Lee, K., Tirasophon, W., Shen, X., Michalak, M., Prywes, R., Okada, T., et al. (2002) IRE1-mediated unconventional mRNA splicing and S2P-mediated ATF6 cleavage merge to regulate XBP1 in signaling the unfolded protein response. *Genes Dev.* **16**, 452–466

159. Goffin, L., Vodala, S., Fraser, C., Ryan, J., Timms, M., Meusburger, S., et al. (2006) The unfolded protein response transducer Ire1p Contains a nuclear localization sequence recognized by Multiple β Importins. *Mol. Biol. Cell* **17**, 5309–5323
protein response to enhance migration and Dissemination of infected Host cells. *mBio* **11**, e00915–e00920

141. Zaman, M. F., Nenadic, A., Radojičić, A., Rosado, A., and Beh, C. T. (2020) Sticking with it: ER-PM membrane contact sites as a Coordinating Nexus for regulating lipids and proteins at the cell Cortex. *Front. Cell Dev. Biol.* **8**, 675

142. Li, C., Qian, T., He, R., Wan, C., Liu, Y., and Yu, H. (2021) Endoplasmic reticulum–plasma membrane contact sites: regulators, mechanisms, and physiological functions. *Front. Cell Dev. Biol.* **9**, 627700

143. Acosta-Alvear, D., Karagöz, G. E., Fröhlich, F., Li, H., Walther, T. C., and Walter, P. (2018) The unfolded protein response and endoplasmic reticulum protein targeting machineries converge on the stress sensor IRE1. *Elife* **7**, e43036

144. Yanagitani, K., Imagawa, Y., Iwawaki, T., Hosoda, A., Saito, M., Kimata, Y., et al. (2009) Cotranslational targeting of XBP1 protein to the membrane promotes Cytoplasmic splicing of its Own mRNA. *Mol. Cell* **34**, 191–200

145. Yanagitani, K., Kimata, Y., Kadokura, H., and Kohno, K. (2011) Translational pausing ensures membrane targeting and Cytoplasmic splicing of XBP1u mRNA. *Science* **331**, 586–589

146. Kanda, S., Yanagitani, K., Yokota, Y., Esaki, Y., and Kohno, K. (2016) Autonomous translational pausing is required for XBP1u mRNA recruitment to the ER via the SRP pathway. *Proc. Natl. Acad. Sci. U. S. A.* **113**, E5886–E5895

147. Son, S. M., Byun, J., Roh, S. E., Kim, S. J., and Mook-Jung, I. (2014) Reduced IRE1α mediates apoptotic cell death by disrupting calcium homeostasis via the InsP3 receptor. *Cell Death Dis.* **5**, e1188

148. Carreras-Sureda, A., Jaña, F., Urra, H., Durand, S., Mortenson, D. E., Sagredo, A., et al. (2019) Non-canonical function of IRE1α determines mitochondria-associated endoplasmic reticulum composition to control calcium transfer and bioenergetics. *Nat. Cell Biol.* **21**, 755–767

149. Carreras-Sureda, A., Zhang, X., Laubry, L., Brunetti, J., Koenig, S., Wang, X., et al. (2023) The ER stress sensor IRE1 interacts with STIM1 to promote store-operated calcium entry, T cell activation, and muscular differentiation. *Cell Rep.* **42**, 113540

150. Yildirim, Z., Baboo, S., Hamid, S. M., Dogan, A. E., Tufanli, O., Robi-chaud, S., et al. (2022) Intercepting IRE1 kinase-FMRP signaling prevents atherosclerosis progression. *EMBO Mol. Med.* **14**, e15344

151. Cairrão, F., Santos, C. C., Le Thomas, A., Marsters, S., Ashkenazi, A., and Domingos, P. M. (2022) Pumilio protects Xbp1 mRNA from regulated Ire1-dependent decay. *Nat. Commun.* **13**, 1587

152. Tam, A. B., Mercado, E. L., Hoffmann, A., and Niwa, M. (2012) ER stress activates NF-κB by integrating functions of basal IKK activity, IRE1 and PERK. *PLoS One* **7**, e45078

153. Yildirim, A. D., Citir, M., Dogan, A. E., Veli, Z., Yildirim, Z., Tufanli, O., et al. (2022) ER stress-induced sphingosine-1-phosphate lyase phosphorylation potentiates the mitochondrial unfolded protein response. *J. Lipid Res.* **63**, 100279

154. Huang, S., Xing, Y., and Liu, Y. (2019) Emerging roles for the ER stress sensor IRE1α in metabolic regulation and disease. *J. Biol. Chem.* **294**, 18726–18741

155. Dong, H., Adams, N. M., Xu, Y., Cao, J., Allan, D. S. J., Carlyle, J. R., et al. (2019) The IRE1 endoplasmic reticulum stress sensor activates natural killer cell immunity in part by regulating c-Myc. *Nat. Immunol.* **20**, 865–878

156. Ardito, F., Giuliani, M., Perrone, D., Troiano, G., and Lo Muzio, L. (2017) The crucial role of protein phosphorylation in cell signaling and its use as targeted therapy (Review). *Int. J. Mol. Med.* **40**, 271–280

157. Niwa, M., Sidrauski, C., Kaufman, R. J., and Walter, P. (1999) A role for Presenilin-1 in nuclear accumulation of Ire1 fragments and induction of the mammalian unfolded protein response. *Cell* **99**, 691–702

158. Lee, K., Tirasophon, W., Shen, X., Michalak, M., Prywes, R., Okada, T., et al. (2002) IRE1-mediated unconventional mRNA splicing and S2P-mediated ATF6 cleavage merge to regulate XBP1 in signaling the unfolded protein response. *Genes Dev.* **16**, 452–466

159. Goffin, L., Vodala, S., Fraser, C., Ryan, J., Timms, M., Meusburger, S., et al. (2006) The unfolded protein response transducer Ire1p Contains a nuclear localization sequence recognized by Multiple β Importins. *Mol. Biol. Cell* **17**, 5309–5323