Mechanical Properties of Chaperone BiP, the Master Regulator of the Endoplasmic Reticulum

*Hilda M. Alfaro-Valdés, Francesca Burgos-Bravo, Nathalie Casanova-Morales, Diego Quiroga-Roger and Christian A.M. Wilson*

#### **Abstract**

Immunoglobulin heavy-chain-binding protein (BiP protein) is a 75-kDa Hsp70 monomeric ATPase motor that plays broad and crucial roles maintaining proteostasis inside the cell. Its malfunction has been related with the appearance of many and important health problems such as neurodegenerative diseases, cancer, and heart diseases, among others. In particular, it is involved in many endoplasmic reticulum (ER) processes and functions, such as protein synthesis, folding, and assembly, and also it works in the posttranslational mechanism of protein translocation. However, it is unknown what kind of molecular motor BiP works like, since the mechanochemical mechanism that BiP utilizes to perform its work during posttranslational translocation across the ER is not fully understood. One novel approach to study both structural and catalytic properties of BiP considers that the viscoelastic regime behavior of the enzymes (considering them as a spring) and their mechanical properties are correlated with catalysis and ligand binding. Structurally, BiP is formed by two domains, and to establish a correlation between BiP structure and catalysis and how its conformational and viscoelastic changes are coupled to ligand binding, catalysis, and allosterism (information transmitted between the domains), optical tweezers and nano-rheology techniques have been essential in this regard.

**Keywords:** immunoglobulin binding protein (BiP), optical tweezers, nano-rheology, posttranslational translocation, molecular motor

#### **1. Introduction**

The endoplasmic reticulum (ER) is involved in protein synthesis and the folding, assembly, transport, and secretion of nascent proteins [1]. One of the most important functions of the ER involves the quality control (ERQC) of nascent proteins, which is accomplished by ER chaperones [2, 3]. Chaperones are proteins that assist other proteins in the folding process, facilitating correct folding pathways or providing microenvironments in which folding can occur [4]. One of the most important chaperones is BiP protein (immunoglobulin heavy-chain binding protein).

BiP, a monomeric ATPase, has been referred to as the master regulator of the ER because of the broad and crucial roles that play in ER processes and functions [5], such as protein synthesis, folding, assembly, and translocation across the ER [3, 6]. Although BiP is still in early stages of study at a molecular level, some research groups have published findings of great value. These findings suggest that this protein could be a key player in various fields, such as in detection and treatment of serious diseases (neurodegenerative diseases, cancer, and heart diseases, among others) [7, 8]. Until now, most of the previous studies have been focused on the function of BiP with classical biochemical approaches and have not taken into account the mechanical properties of this protein. The role played by force on macromolecular structure and function is a subject of recent intensive research. Mechanical processes are a key component of many biological events. The coupling of mechanics and chemistry is one of the most important features of enzymes, which is highly specific and regulated [9]. Enzymes need to couple their chemical reactions to mechanical motion. In this way, an enzyme can work like a molecular motor using the hydrolysis or binding of ATP, converting this chemical energy to mechanical work. Allosterism and conformational changes are examples of how a chemical event could be transduced to mechanical events regulated by catalysis and ligand binding events based on changes in the elastic properties of domains [10]. Exploring this coupling may contribute to the understanding of the mechanical properties of enzymes, such as the mechanochemical mechanism of BiP. Understanding viscoelasticity is crucial because biological materials show different phenomena such as stiffening or softening upon ligand binding because proteins behave as springs [11, 12]. Due to recent technological progress, it is possible to measure changes in viscoelasticity in the folded state of proteins and we could correlate these changes with functionality. All these new approaches help to solve biological problems based on a mechanical description of molecular mechanisms to obtain a complete view of how the proteins perform their function with high efficiency.

#### **2. The ATP-regulated Hsp70 chaperone BiP is the master regulator of the endoplasmic reticulum**

Approximately, one-third of proteins produced in mammalian cells are folded and assembled in the ER, including secretory, membrane-bound, and some organelle-targeted proteins [13]. In the ER, proteins are translocated into the lumen where they acquire their functional tertiary and quaternary structure [3], and then correctly folded proteins exit the ER and are transported to intracellular organelles and the cell surface. The success of the maturation of a protein in its passage through the secretory pathway is monitored by the ERQC process, which is highly conserved in most eukaryotic organisms [2, 3]. For this, molecular chaperones proteins interact with partially folded or improperly folded polypeptides, facilitating correct folding pathways or providing microenvironments where folding can occur [4]. However, those proteins that fail to fold properly must be translocated back to the cytoplasm and degraded in the proteasomes through a process known as ER-associated degradation (ERAD) [3]. Two main chaperone systems help to fold the proteins in the ER or target them for ERAD if folding fails: lectins such as calnexin/calreticulin, unique to the ER, and the heat shock protein 70 (Hsp70) system, which has many aspects that are common to all Hsp70s. BiP (also known as glucose-regulated protein 78 kDa, HspA5, or Kar2p in yeast) is the only known conventional Hsp70 chaperone in the ER [14, 15].

BiP binds transiently to newly unfolded synthesized proteins translocated posttranslationally into the ER (**Figure 1**). Binding of BiP to the incoming polypeptide

**13**

**Figure 1.**

*Mechanical Properties of Chaperone BiP, the Master Regulator of the Endoplasmic Reticulum*

contributes to efficiency and unidirectionality of transport due to its role as a molecular motor in the posttranslational translocation (will be discussed below). As a molecular chaperone, binding of BiP to hydrophobic patches exposed on nascent unfolded proteins that enter into the ER lumen or incompletely folded nonglycosylated proteins prevents nascent polypeptide chains from folding incorrectly and their interaction with nascent immature secretable proteins synthesized from membrane-bound polysomes. This prevents immature protein denaturation or

*Hsp70 chaperone BiP is a master ER regulator. Under nonstressed conditions (unstressed ER), BiP binds to hydrophobic regions of unfolded polypeptides fully synthesized to favor their posttranslational translocation into the ER lumen. The high substrate binding affinity of BiP to hydrophobic patches is achieved in the ADP-bound state upon the hydrolysis of ATP to ADP. After the translocation, BiP facilitates correct folding of nascent unfolded proteins or incompletely folded proteins nonglycosylated for their subsequently secretion. The proteins that fail to fold properly are targeted for proteasomal degradation in the cytoplasm through the ER-associated degradation (ERAD) pathway. BiP also interacts with the luminal domains of three ER stress sensors: IRE1, PERK, and ATF6 to maintain them in the ER. However, upon accumulation of unfolded/misfolded proteins in the ER lumen (stressed ER), BiP is released from these molecules to interact with unfolded proteins and favor their correct folding. BiP dissociation from these sensors allows their activation that involves: IRE1 dimerization, autophosphorylation, and splicing of Xbp1 and Hac1 mRNA; PERK dimerization, autophosphorylation, and phosphorylation of eIF2α, which lead to the attenuation of protein translation; and ATF6 transportation to the Golgi where it is processed by proteases. The ATF6 cytoplasmic domain obtained after its processing together with Xbp1 and Hac1 is translocated to the nucleus to activate the transcription of UPR-responsive genes.*

Any condition perturbing the correct functioning of the ER, leading to an increase in protein synthesis or to the generation and accumulation of misfolded proteins inside the ER, is known as ER stress [16]. Moreover, misfolded proteins can also aggregate into insoluble higher order structure that has been associated with numerous neurodegenerative human diseases [17]. Adaptation to proteinfolding stress is mediated by the activation of the unfolded protein response (UPR), which has evolved to detect the accumulation of misfolded proteins and activate a cellular response to maintain homeostasis and a normal flux of proteins in the ER, by increasing its folding capacity [18]. In this context, BiP serves as a

degradation and ensures proper folding and its secretion (**Figure 1**).

*DOI: http://dx.doi.org/10.5772/intechopen.82080*

*Mechanical Properties of Chaperone BiP, the Master Regulator of the Endoplasmic Reticulum DOI: http://dx.doi.org/10.5772/intechopen.82080*

#### **Figure 1.**

*Endoplasmic Reticulum*

perform their function with high efficiency.

conventional Hsp70 chaperone in the ER [14, 15].

**endoplasmic reticulum**

BiP, a monomeric ATPase, has been referred to as the master regulator of the ER because of the broad and crucial roles that play in ER processes and functions [5], such as protein synthesis, folding, assembly, and translocation across the ER [3, 6]. Although BiP is still in early stages of study at a molecular level, some research groups have published findings of great value. These findings suggest that this protein could be a key player in various fields, such as in detection and treatment of serious diseases (neurodegenerative diseases, cancer, and heart diseases, among others) [7, 8]. Until now, most of the previous studies have been focused on the function of BiP with classical biochemical approaches and have not taken into account the mechanical properties of this protein. The role played by force on macromolecular structure and function is a subject of recent intensive research. Mechanical processes are a key component of many biological events. The coupling of mechanics and chemistry is one of the most important features of enzymes, which is highly specific and regulated [9]. Enzymes need to couple their chemical reactions to mechanical motion. In this way, an enzyme can work like a molecular motor using the hydrolysis or binding of ATP, converting this chemical energy to mechanical work. Allosterism and conformational changes are examples of how a chemical event could be transduced to mechanical events regulated by catalysis and ligand binding events based on changes in the elastic properties of domains [10]. Exploring this coupling may contribute to the understanding of the mechanical properties of enzymes, such as the mechanochemical mechanism of BiP. Understanding viscoelasticity is crucial because biological materials show different phenomena such as stiffening or softening upon ligand binding because proteins behave as springs [11, 12]. Due to recent technological progress, it is possible to measure changes in viscoelasticity in the folded state of proteins and we could correlate these changes with functionality. All these new approaches help to solve biological problems based on a mechanical description of molecular mechanisms to obtain a complete view of how the proteins

**2. The ATP-regulated Hsp70 chaperone BiP is the master regulator of the** 

Approximately, one-third of proteins produced in mammalian cells are folded

organelle-targeted proteins [13]. In the ER, proteins are translocated into the lumen where they acquire their functional tertiary and quaternary structure [3], and then correctly folded proteins exit the ER and are transported to intracellular organelles and the cell surface. The success of the maturation of a protein in its passage through the secretory pathway is monitored by the ERQC process, which is highly conserved in most eukaryotic organisms [2, 3]. For this, molecular chaperones proteins interact with partially folded or improperly folded polypeptides, facilitating correct folding pathways or providing microenvironments where folding can occur [4]. However, those proteins that fail to fold properly must be translocated back to the cytoplasm and degraded in the proteasomes through a process known as ER-associated degradation (ERAD) [3]. Two main chaperone systems help to fold the proteins in the ER or target them for ERAD if folding fails: lectins such as calnexin/calreticulin, unique to the ER, and the heat shock protein 70 (Hsp70) system, which has many aspects that are common to all Hsp70s. BiP (also known as glucose-regulated protein 78 kDa, HspA5, or Kar2p in yeast) is the only known

BiP binds transiently to newly unfolded synthesized proteins translocated posttranslationally into the ER (**Figure 1**). Binding of BiP to the incoming polypeptide

and assembled in the ER, including secretory, membrane-bound, and some

**12**

*Hsp70 chaperone BiP is a master ER regulator. Under nonstressed conditions (unstressed ER), BiP binds to hydrophobic regions of unfolded polypeptides fully synthesized to favor their posttranslational translocation into the ER lumen. The high substrate binding affinity of BiP to hydrophobic patches is achieved in the ADP-bound state upon the hydrolysis of ATP to ADP. After the translocation, BiP facilitates correct folding of nascent unfolded proteins or incompletely folded proteins nonglycosylated for their subsequently secretion. The proteins that fail to fold properly are targeted for proteasomal degradation in the cytoplasm through the ER-associated degradation (ERAD) pathway. BiP also interacts with the luminal domains of three ER stress sensors: IRE1, PERK, and ATF6 to maintain them in the ER. However, upon accumulation of unfolded/misfolded proteins in the ER lumen (stressed ER), BiP is released from these molecules to interact with unfolded proteins and favor their correct folding. BiP dissociation from these sensors allows their activation that involves: IRE1 dimerization, autophosphorylation, and splicing of Xbp1 and Hac1 mRNA; PERK dimerization, autophosphorylation, and phosphorylation of eIF2α, which lead to the attenuation of protein translation; and ATF6 transportation to the Golgi where it is processed by proteases. The ATF6 cytoplasmic domain obtained after its processing together with Xbp1 and Hac1 is translocated to the nucleus to activate the transcription of UPR-responsive genes.*

contributes to efficiency and unidirectionality of transport due to its role as a molecular motor in the posttranslational translocation (will be discussed below). As a molecular chaperone, binding of BiP to hydrophobic patches exposed on nascent unfolded proteins that enter into the ER lumen or incompletely folded nonglycosylated proteins prevents nascent polypeptide chains from folding incorrectly and their interaction with nascent immature secretable proteins synthesized from membrane-bound polysomes. This prevents immature protein denaturation or degradation and ensures proper folding and its secretion (**Figure 1**).

Any condition perturbing the correct functioning of the ER, leading to an increase in protein synthesis or to the generation and accumulation of misfolded proteins inside the ER, is known as ER stress [16]. Moreover, misfolded proteins can also aggregate into insoluble higher order structure that has been associated with numerous neurodegenerative human diseases [17]. Adaptation to proteinfolding stress is mediated by the activation of the unfolded protein response (UPR), which has evolved to detect the accumulation of misfolded proteins and activate a cellular response to maintain homeostasis and a normal flux of proteins in the ER, by increasing its folding capacity [18]. In this context, BiP serves as a

master UPR regulator and plays essential roles in activating three distinct ER stress sensors: IRE1, PERK, and ATF6 (**Figure 1**). Under nonstressed conditions, BiP binds to IRE1, PERK, and ATF6 by their luminal domains to maintain them in the ER. The accumulation of unfolded/misfolded proteins induces dissociation of BiP from IRE1 and PERK to permit their dimerization, trans-autophosphorylation, and activation [19]. Activated IRE1 initiates mRNA splicing of two transcriptional factors (Xbp1 and Hac1) to generate potent transcriptional activation of UPR target genes. PERK activation involves phosphorylation of the translational elongation factor eLF2 to attenuate protein synthesis. The release of ATF6 favors its transport to Golgi where is cleaved to generate the cytosolic domain of ATF6 that translocate to the nucleus to activate transcription of UPR-responsive genes [20]. Therefore, the activation of these sensors results in the attenuation of translation to reduce the workload of the ER, the transcriptional upregulation of genes encoding ER chaperones to increase the folding capacity of the ER, and the overexpression of the ERAD component to favor the degradation of these unfolded proteins by the proteasome [21, 22]. Thus, BiP participates not only in assisting protein folding, assembly and translocation but also in protein degradation and in the stress adaptability of the ER [1]. One big difference between BiP and lectins is that BiP detects only the unfolded regions of the nascent polypeptide chains, whereas lectins can detect both N-linked glycans of the peptides and unfolded regions [23]. However, it is not yet completely clear how BiP binds to its unfolded substrate because usually peptides are used as substrates instead of complete unfolded proteins. Recently, we developed a new method to study this process by specifically unfolding a complete protein substrate for BiP and measuring in optical tweezers the time that BiP remains bound to its substrate [24]. Previously, a work with DnaK (a close homolog of BiP) shows that it binds and stabilizes also partially folded protein structures [25]. BiP has a crucial role during posttranslational translocation, acting as a molecular motor. Molecular chaperones in the cytoplasm and ER lumen are involved in polypeptide translocation across the ER. Proteins enter the ER by a channel protein complex known as the translocon, discovered in yeast in Randy Schekman's laboratory [26]. In eukaryotic cells, the translocation of proteins is carried out by the Sec61 complex [6, 27]. Sec61 complex consists of three subunits, α, β, and γ, in which the pore to transport the polypeptide chain is created by the α-subunit of Sec61 protein. This complex functions as a passive channel that requires accessory proteins to provide a driving force to facilitate the vectorial translocation of the polypeptide chain through the membrane. Those accessory proteins are molecular motors [28]. Motor enzymes use the energy of nucleotide binding/hydrolysis or product release to generate mechanical work. The two mechanisms of translocation across the ER are co-translational translocation and posttranslational translocation [29]. In the co-translational mechanism, which has been well studied in mammalian systems, the signal sequence at the N-terminus of the nascent polypeptide interacts with the signal recognition protein (SRP) in the cytoplasm, keeping the ribosome attached to the Sec61 complex [6]. In this mechanism, the ribosome acts as an "auxiliary protein," since the driving force for translocation is given by GTP hydrolysis during the elongation of the polypeptide chain [30]. However, the driving force delivered by the ribosome is missing for posttranslationally translocated proteins. In this case, the driving force for polypeptide chain translocation comes from BiP protein [30]. Thus, in posttranslational translocation, after the polypeptides are fully synthesized, cytoplasmic molecular chaperones keep them unfolded to be transported through the Sec61 complex. In this mechanism, the channel partners with another membrane-protein complex, the Sec62/Sec63 complex, and with the lumenal chaperone BiP. However, in spite of the crucial roles of BiP during translocation, it is not fully understood

**15**

mechanism.

**Figure 2.**

*domains closed by red arrows inward.*

*Mechanical Properties of Chaperone BiP, the Master Regulator of the Endoplasmic Reticulum*

if the action of BiP is through an active mechanism of pulling (as a power stroke), mediated by the binding/hydrolysis of ATP, or as a ratchet mechanism (**Figure 2**). In the latter, the polypeptide chain enters the channel passively by Brownian motion, and the BiP protein prevents it from returning to the cytoplasm. The hypothesis of the ratchet mechanism has been supported by employing antibodies against the polypeptide chains passing through the ER lumen [31]. Evidence for the translocation mechanism has been obtained using coarse-grained model simulations [32]. This study suggests that Hsp70 chaperones use an "entropic pulling mechanism," applying a force of about 15pN, and proposes that the Hsp70's would use a combination of ratchet and power stroke mechanisms [33]. Translocation in all eukaryotes is likely to be similar to yeast because of the high identity of amino acids between their channels. The channel interacts with the Sec62/Sec63 complex, with BiP acting as a molecular motor to bias the passive movement of a polypeptide in the Sec61 channel. In bacterial posttranslational translocation, the channel interacts with the cytoplasmic ATPase SecA. SecA moves polypeptides through the SecY channel to the periplasm by a "push and slide" mechanism [34]. Archaea probably use both cotranslational and posttranslational translocation, but it is unknown how posttranslational translocation occurs because these organisms lack SecA, Sec62/Sec63 complex, and BiP [6, 30]. In double membrane system, as in chloroplast, it is mediated by translocon at the outer envelope membrane of chloroplasts (TOC) and translocon at the inner envelope membrane of chloroplasts (TIC), which facility the import of translated proteins with assistant of a TIC associated ATP-driven import motor [35]. However, in mitochondria, the import of preproteins is carried out by translocases called as TOM complex (translocon outer mitochondria membrane) and TIM23 complex (translocon at the inner mitochondrial membrane), where proteins with a hydrophobic sorting signal can be released into the inner membrane and hydrophilic proteins are imported into the matrix by one presequence translocase-associated motor (PAM) in which the force is driven by chaperone mtHsp70 [36]. Therefore, the mHsp70 pulls the presequence by power stroke or Brownian ratchet mechanism to finally translocate the presequence at the mitochondria matrix. This suggests that the mechanism of Hsp70 in the import of preprotein in mitochondria and ER could have similar basic

*Schematic representation of the two mechanisms of BiP in translocation. The figure shows how BiP could be involved in the transport process of the protein into the ER. (A) The ratchet theory is shown in which several BiP molecules would be interacting with the incoming chain, and in this way, the chain will not be returned to the cytoplasm. (B) The theory of power stroke is shown, where BiP binds to the polypeptide chain exerting a force greater than that of the thermal bath. Open domains of BiP are represented by red arrows outward and* 

*DOI: http://dx.doi.org/10.5772/intechopen.82080*

*Mechanical Properties of Chaperone BiP, the Master Regulator of the Endoplasmic Reticulum DOI: http://dx.doi.org/10.5772/intechopen.82080*

**Figure 2.**

*Endoplasmic Reticulum*

master UPR regulator and plays essential roles in activating three distinct ER stress sensors: IRE1, PERK, and ATF6 (**Figure 1**). Under nonstressed conditions, BiP binds to IRE1, PERK, and ATF6 by their luminal domains to maintain them in the ER. The accumulation of unfolded/misfolded proteins induces dissociation of BiP from IRE1 and PERK to permit their dimerization, trans-autophosphorylation, and activation [19]. Activated IRE1 initiates mRNA splicing of two transcriptional factors (Xbp1 and Hac1) to generate potent transcriptional activation of UPR target genes. PERK activation involves phosphorylation of the translational elongation factor eLF2 to attenuate protein synthesis. The release of ATF6 favors its transport to Golgi where is cleaved to generate the cytosolic domain of ATF6 that translocate to the nucleus to activate transcription of UPR-responsive genes [20]. Therefore, the activation of these sensors results in the attenuation of translation to reduce the workload of the ER, the transcriptional upregulation of genes encoding ER chaperones to increase the folding capacity of the ER, and the overexpression of the ERAD component to favor the degradation of these unfolded proteins by the proteasome [21, 22]. Thus, BiP participates not only in assisting protein folding, assembly and translocation but also in protein degradation and in the stress adaptability of the ER [1]. One big difference between BiP and lectins is that BiP detects only the unfolded regions of the nascent polypeptide chains, whereas lectins can detect both N-linked glycans of the peptides and unfolded regions [23]. However, it is not yet completely clear how BiP binds to its unfolded substrate because usually peptides are used as substrates instead of complete unfolded proteins. Recently, we developed a new method to study this process by specifically unfolding a complete protein substrate for BiP and measuring in optical tweezers the time that BiP remains bound to its substrate [24]. Previously, a work with DnaK (a close homolog of BiP) shows that it binds and stabilizes also partially folded protein structures [25]. BiP has a crucial role during posttranslational translocation, acting as a molecular motor. Molecular chaperones in the cytoplasm and ER lumen are involved in polypeptide translocation across the ER. Proteins enter the ER by a channel protein complex known as the translocon, discovered in yeast in Randy Schekman's laboratory [26]. In eukaryotic cells, the translocation of proteins is carried out by the Sec61 complex [6, 27]. Sec61 complex consists of three subunits, α, β, and γ, in which the pore to transport the polypeptide chain is created by the α-subunit of Sec61 protein. This complex functions as a passive channel that requires accessory proteins to provide a driving force to facilitate the vectorial translocation of the polypeptide chain through the membrane. Those accessory proteins are molecular motors [28]. Motor enzymes use the energy of nucleotide binding/hydrolysis or product release to generate mechanical work. The two mechanisms of translocation across the ER are co-translational translocation and posttranslational translocation [29]. In the co-translational mechanism, which has been well studied in mammalian systems, the signal sequence at the N-terminus of the nascent polypeptide interacts with the signal recognition protein (SRP) in the cytoplasm, keeping the ribosome attached to the Sec61 complex [6]. In this mechanism, the ribosome acts as an "auxiliary protein," since the driving force for translocation is given by GTP hydrolysis during the elongation of the polypeptide chain [30]. However, the driving force delivered by the ribosome is missing for posttranslationally translocated proteins. In this case, the driving force for polypeptide chain translocation comes from BiP protein [30]. Thus, in posttranslational translocation, after the polypeptides are fully synthesized, cytoplasmic molecular chaperones keep them unfolded to be transported through the Sec61 complex. In this mechanism, the channel partners with another membrane-protein complex, the Sec62/Sec63 complex, and with the lumenal chaperone BiP. However, in spite of the crucial roles of BiP during translocation, it is not fully understood

**14**

*Schematic representation of the two mechanisms of BiP in translocation. The figure shows how BiP could be involved in the transport process of the protein into the ER. (A) The ratchet theory is shown in which several BiP molecules would be interacting with the incoming chain, and in this way, the chain will not be returned to the cytoplasm. (B) The theory of power stroke is shown, where BiP binds to the polypeptide chain exerting a force greater than that of the thermal bath. Open domains of BiP are represented by red arrows outward and domains closed by red arrows inward.*

if the action of BiP is through an active mechanism of pulling (as a power stroke), mediated by the binding/hydrolysis of ATP, or as a ratchet mechanism (**Figure 2**). In the latter, the polypeptide chain enters the channel passively by Brownian motion, and the BiP protein prevents it from returning to the cytoplasm. The hypothesis of the ratchet mechanism has been supported by employing antibodies against the polypeptide chains passing through the ER lumen [31]. Evidence for the translocation mechanism has been obtained using coarse-grained model simulations [32]. This study suggests that Hsp70 chaperones use an "entropic pulling mechanism," applying a force of about 15pN, and proposes that the Hsp70's would use a combination of ratchet and power stroke mechanisms [33]. Translocation in all eukaryotes is likely to be similar to yeast because of the high identity of amino acids between their channels. The channel interacts with the Sec62/Sec63 complex, with BiP acting as a molecular motor to bias the passive movement of a polypeptide in the Sec61 channel. In bacterial posttranslational translocation, the channel interacts with the cytoplasmic ATPase SecA. SecA moves polypeptides through the SecY channel to the periplasm by a "push and slide" mechanism [34]. Archaea probably use both cotranslational and posttranslational translocation, but it is unknown how posttranslational translocation occurs because these organisms lack SecA, Sec62/Sec63 complex, and BiP [6, 30]. In double membrane system, as in chloroplast, it is mediated by translocon at the outer envelope membrane of chloroplasts (TOC) and translocon at the inner envelope membrane of chloroplasts (TIC), which facility the import of translated proteins with assistant of a TIC associated ATP-driven import motor [35]. However, in mitochondria, the import of preproteins is carried out by translocases called as TOM complex (translocon outer mitochondria membrane) and TIM23 complex (translocon at the inner mitochondrial membrane), where proteins with a hydrophobic sorting signal can be released into the inner membrane and hydrophilic proteins are imported into the matrix by one presequence translocase-associated motor (PAM) in which the force is driven by chaperone mtHsp70 [36]. Therefore, the mHsp70 pulls the presequence by power stroke or Brownian ratchet mechanism to finally translocate the presequence at the mitochondria matrix. This suggests that the mechanism of Hsp70 in the import of preprotein in mitochondria and ER could have similar basic mechanism.

#### **2.1 BiP structure and catalysis**

The effective application of work depends on the elastic properties of a motor based on the softening and stiffening of some domains [37], and it is important to understand how the information is transmitted through domains by BiP. BiP is formed by two domains: a nucleotide-binding domain (NBD), with ATPase activity, connected by a flexible hydrophobic linker to the substrate-binding domain (SBD) (**Figure 3**). The SBD can be further divided into a compact β-sandwich domain harboring, a cleft for substrate binding, and an α-helical domain at its C-terminal end, the so-called "lid" [38]. Many conformational changes, such as the open and close movement of the lid and the variation in the distance between the SBD and NBD, have been associated with the ATPase cycle of BiP in the ER. Once BiP binds K<sup>+</sup> and ATP, its NBD and SBD come into close proximity to each other and the lid of the SBD opens, which results in a form that binds substrates with low affinity [3]. Also, a number of BiP cofactors have been discovered that assist in controlling the substrate-binding cycle and its localization within the ER. Nucleotide exchange factors (NEFs) help in the transition from the ADP to the ATP bound state for BiP, catalyzing the release of substrate. Hsp70 hydrolysis of ATP to ADP is accelerated by Hsp40 family members or so-called J domain proteins. The J-domain binds to Hsp70 and stimulates its ATPase activity [39]. In addition to controlling the localization and activity of Hsp70's, J-domain proteins may also bind the substrate themselves and help with the initial delivery of the substrate to Hsp70 chaperone. In the mammalian ER, there are seven J-domain proteins (ERdj1–7) that assist with the diverse functions of BiP [40]. After the Mg2+-dependent hydrolysis of ATP, BiP enters a state with low on and off rates for substrates [3]. For elongated/peptide substrates, the lid closes over the bound substrate; for globular substrates, there are direct interactions between the lid and the substrate, but the lid may not close completely [3]. The SBD and NBD become farther apart upon substrate binding and ATP hydrolysis, which is less pronounced for globular substrates. ADP must be exchanged for ATP in order to release the substrate and make BiP available for another round of client binding. Ca2+ increases the affinity for ADP, whereas NEFs Grp170 and Sil1 facilitate the nucleotide exchange reaction [3]. Conformational changes in murine BiP during ATPase cycle have been determined by Förster Resonance Energy Transfer (FRET) at the single molecule level, showing that NBD and SBD come into close

#### **Figure 3.**

*Structure of ATP-bound BiP in the open conformation. BiP has two domains, NBD (light green) and SBD (dark green). The latter has a subdomain that acts as an α helix lid that covers the binding pocket for polypeptides formed by β sheets. In the ATP-bound BiP conformation, the lid is open. This structure corresponds to protein data bank number: 5E84 and was drawn as a ribbon diagram, using PyMOL molecular visualization system.*

**17**

*Mechanical Properties of Chaperone BiP, the Master Regulator of the Endoplasmic Reticulum*

contact with a mean distance 58–75 Å [41]. Additionally, by using NMR residual dipolar coupling, spin labeling, and dynamics methods, it has been determined in DnaK that the NBD and SBD are loosely linked and can move in cones of 35° with respect to each other [42]. Moreover, the distance between the base and the lid of the SBD domain in Hsp70 has been calculated to be 77 Å by means of FRET [43]. Also, there is a crystal structure of human BiP bound to ATP that shows similar distances [44]. The conformational changes and movements of BiP are not independent for each domain because an important communication and coupling

Most HSP70 do their work coupling the Mg2+-dependent hydrolisis of ATP to large conformational changes, involving movements of its structural domains (SBD and NBD) and the interdomain linker. So, HSP70 protein function rely on a dynamic ATP dependent cycle in which several conformations are visited, with different substrate binding affinities in them [45–47]. For example in DnaK, ATP binding favors a compact, domain-docked, and linker-bound conformation, which has low ATPase activity [3, 45]. Substrate binding to this state stabilizes a transient domain-undocked conformation, with a linker-bound state, that has high ATPase activity and fast substrate binding and release kinetics but low substrate affinity. Then, when ATP is hydrolyzed to ADP, it is favored a domainundocked conformation, linker-unbound state, which has high substrate affinity but very slow substrate binding and release kinetics [45–48]. Recently, X-ray structures of ATP-bound DnaK [49] and human BiP [44] have shown that both proteins have big structural similarity, but their functional activity (and between different Hsp70s) varies significantly between them [3]. On this ground, considering the fact that in spite of the structural similarity between different Hsp70, they have different functional activity; it was suggested that an important feature that should modulate Hsp70 function was its allosteric communication between both structural domains, mediated by the interdomain linker [44, 50]. Basically, the allosteric mechanism transmits information on the nucleotide state from NBD to SBD and on the substrate occupancy state from SBD back to NBD [51]. At the beginning, three different ideas explaining how interdomain communication occurs have been suggested. In the *E. coli* Hsp70 DnaK, bound nucleotide is sensed by residues in NBD that are closer to the bound ATP, in particular, a proline residue (Pro143) and a surface-exposed arginine (Arg151), and the communication with the SBD domain is thought to be via this proline, which can likely undergo cis/trans isomerization [52]. Replacement of the arginine completely disrupted the mutual allosteric communication between ATPase domain and substrate binding domain. Moreover, arginine had been shown to be an important residue in allosteric communication in other proteins [53]. Replacement of the proline destabilized the allosteric communication, increasing the rate of spontaneous transition between ATP-like and ADP-like states. Interestingly, all residues of the proposed DnaK sensor-relay system are conserved in BiP [3, 54]. In addition to this putative proline-focused sensor-relay system, threonine in position 37 (Thr37) in NBD plays a particularly important role as a nucleotide sensor in a hamster BiP [55], likely due to a direct interaction of its hydroxyl group with the γ-phosphate oxygen of bound ATP. Once this position was mutated, no more conformational change occurred, while nucleotide binding and hydrolysis were not affected [3]. The third known communication path between NBD and SBD occurs through the conserved hydrophobic linker, which connects both domains.

*DOI: http://dx.doi.org/10.5772/intechopen.82080*

exists between them.

**2.2 BiP allosteric mechanism**

*Mechanical Properties of Chaperone BiP, the Master Regulator of the Endoplasmic Reticulum DOI: http://dx.doi.org/10.5772/intechopen.82080*

contact with a mean distance 58–75 Å [41]. Additionally, by using NMR residual dipolar coupling, spin labeling, and dynamics methods, it has been determined in DnaK that the NBD and SBD are loosely linked and can move in cones of 35° with respect to each other [42]. Moreover, the distance between the base and the lid of the SBD domain in Hsp70 has been calculated to be 77 Å by means of FRET [43]. Also, there is a crystal structure of human BiP bound to ATP that shows similar distances [44]. The conformational changes and movements of BiP are not independent for each domain because an important communication and coupling exists between them.

#### **2.2 BiP allosteric mechanism**

*Endoplasmic Reticulum*

ER. Once BiP binds K<sup>+</sup>

**2.1 BiP structure and catalysis**

The effective application of work depends on the elastic properties of a motor based on the softening and stiffening of some domains [37], and it is important to understand how the information is transmitted through domains by BiP. BiP is formed by two domains: a nucleotide-binding domain (NBD), with ATPase activity, connected by a flexible hydrophobic linker to the substrate-binding domain (SBD) (**Figure 3**). The SBD can be further divided into a compact β-sandwich domain harboring, a cleft for substrate binding, and an α-helical domain at its C-terminal end, the so-called "lid" [38]. Many conformational changes, such as the open and close movement of the lid and the variation in the distance between the SBD and NBD, have been associated with the ATPase cycle of BiP in the

each other and the lid of the SBD opens, which results in a form that binds substrates with low affinity [3]. Also, a number of BiP cofactors have been discovered that assist in controlling the substrate-binding cycle and its localization within the ER. Nucleotide exchange factors (NEFs) help in the transition from the ADP to the ATP bound state for BiP, catalyzing the release of substrate. Hsp70 hydrolysis of ATP to ADP is accelerated by Hsp40 family members or so-called J domain proteins. The J-domain binds to Hsp70 and stimulates its ATPase activity [39]. In addition to controlling the localization and activity of Hsp70's, J-domain proteins may also bind the substrate themselves and help with the initial delivery of the substrate to Hsp70 chaperone. In the mammalian ER, there are seven J-domain proteins (ERdj1–7) that assist with the diverse functions of BiP [40]. After the Mg2+-dependent hydrolysis of ATP, BiP enters a state with low on and off rates for substrates [3]. For elongated/peptide substrates, the lid closes over the bound substrate; for globular substrates, there are direct interactions between the lid and the substrate, but the lid may not close completely [3]. The SBD and NBD become farther apart upon substrate binding and ATP hydrolysis, which is less pronounced for globular substrates. ADP must be exchanged for ATP in order to release the substrate and make BiP available for another round of client binding. Ca2+ increases the affinity for ADP, whereas NEFs Grp170 and Sil1 facilitate the nucleotide exchange reaction [3]. Conformational changes in murine BiP during ATPase cycle have been determined by Förster Resonance Energy Transfer (FRET) at the single molecule level, showing that NBD and SBD come into close

*Structure of ATP-bound BiP in the open conformation. BiP has two domains, NBD (light green) and SBD (dark green). The latter has a subdomain that acts as an α helix lid that covers the binding pocket for polypeptides formed by β sheets. In the ATP-bound BiP conformation, the lid is open. This structure corresponds to protein data bank number: 5E84 and was drawn as a ribbon diagram, using PyMOL molecular* 

and ATP, its NBD and SBD come into close proximity to

**16**

**Figure 3.**

*visualization system.*

Most HSP70 do their work coupling the Mg2+-dependent hydrolisis of ATP to large conformational changes, involving movements of its structural domains (SBD and NBD) and the interdomain linker. So, HSP70 protein function rely on a dynamic ATP dependent cycle in which several conformations are visited, with different substrate binding affinities in them [45–47]. For example in DnaK, ATP binding favors a compact, domain-docked, and linker-bound conformation, which has low ATPase activity [3, 45]. Substrate binding to this state stabilizes a transient domain-undocked conformation, with a linker-bound state, that has high ATPase activity and fast substrate binding and release kinetics but low substrate affinity. Then, when ATP is hydrolyzed to ADP, it is favored a domainundocked conformation, linker-unbound state, which has high substrate affinity but very slow substrate binding and release kinetics [45–48]. Recently, X-ray structures of ATP-bound DnaK [49] and human BiP [44] have shown that both proteins have big structural similarity, but their functional activity (and between different Hsp70s) varies significantly between them [3]. On this ground, considering the fact that in spite of the structural similarity between different Hsp70, they have different functional activity; it was suggested that an important feature that should modulate Hsp70 function was its allosteric communication between both structural domains, mediated by the interdomain linker [44, 50]. Basically, the allosteric mechanism transmits information on the nucleotide state from NBD to SBD and on the substrate occupancy state from SBD back to NBD [51]. At the beginning, three different ideas explaining how interdomain communication occurs have been suggested. In the *E. coli* Hsp70 DnaK, bound nucleotide is sensed by residues in NBD that are closer to the bound ATP, in particular, a proline residue (Pro143) and a surface-exposed arginine (Arg151), and the communication with the SBD domain is thought to be via this proline, which can likely undergo cis/trans isomerization [52]. Replacement of the arginine completely disrupted the mutual allosteric communication between ATPase domain and substrate binding domain. Moreover, arginine had been shown to be an important residue in allosteric communication in other proteins [53]. Replacement of the proline destabilized the allosteric communication, increasing the rate of spontaneous transition between ATP-like and ADP-like states. Interestingly, all residues of the proposed DnaK sensor-relay system are conserved in BiP [3, 54]. In addition to this putative proline-focused sensor-relay system, threonine in position 37 (Thr37) in NBD plays a particularly important role as a nucleotide sensor in a hamster BiP [55], likely due to a direct interaction of its hydroxyl group with the γ-phosphate oxygen of bound ATP. Once this position was mutated, no more conformational change occurred, while nucleotide binding and hydrolysis were not affected [3]. The third known communication path between NBD and SBD occurs through the conserved hydrophobic linker, which connects both domains.

Upon ATP binding, the linker binds to a cleft in NBD, which is important in transmitting the nucleotide state of NBD to SBD and increases ATP hydrolysis of the NBD once bound to the cleft. Basically, it has been suggested that allostery results from an energetic tug-of-war between domain conformations and formation of two orthogonal interfaces: between the NBD and SBD and between the helical lid and the β subdomain of the SBD [46]. More recently, "soft" amino acid substitutions have been performed in BiP, trying to affect the allosteric communication between SBD and NBD, uncoupling the substrate-binding site with the NBD-SBD interdomain interface. In particular, Val461 was mutated to Phe; Ile526 to Val; Ile437 to Val; and Ile538 to Val. It has been reported that in the presence of ATP, all these "soft" mutations affected the equilibrium between the domain-docked and domain-undocked conformations, suggesting that this residue enables allosteric control of BiP conformational ensemble [45].

Moreover, allosterism in BiP has been studied at the single molecule level with optical tweezer manipulation [24]. The results showed that BiP binds reversibly to the unfolded state of MJ0366 (substrate protein), preventing its refolding, and that this effect depends on both the type and concentration of nucleotides. Additionally, more clues about BiP allosteric mechanism have arisen from BiP ensemble measurements performed with nano-rheological experimental setup, which will be explain later.

Finally, it has been studied how the posttranslational modification of BiP by AMPylation onto Thr518 [56] could affect BiP conformational cycle, modulating in this way the allosteric mechanism of BiP. The results showed that effectively, this modification shifted BiP conformational equilibrium toward the domain-docked conformation in the presence of ATP, stabilizing the domain docking in the absence of ATP and demonstrating posttranslational fine tuning of BiP conformational equilibrium [45].

As a general overview, BiP allosteric mechanism has a high level of complexity, as it has different layers of control. From a structural point of view, there are residues that exert the communication between the SBD and NBD domains, and other residues that are involved in stabilizing conformational ensembles of BiP that affect allosteric communication. Moreover, changes in the mechanical properties of BiP are also involved in the allosteric mechanism regulation, as it has been demonstrated with the nano-rheological studies. Finally, posttranslational modifications also play a role in this chaperone function, as their importance in shuffling conformational ensembles, involved in this ATP and Mg2+ dependent cycle, has been demonstrated.

#### **2.3 Mechanical aspects**

Considering that translocation through the ER is a crucial process to maintain homeostasis inside the cell, it is essential to have a mechanistic understanding of the role that BiP has in translocation to maintain proteostasis. Therefore, classical biochemical assays, or ensemble studies, have been conducted to study each of these processes without taking into account the measurement of forces and changes in elasticity. Single molecule level studies, called *in singulo* studies, have become very relevant during recent years. These studies have become the gold standard to study biomolecular mechanisms because of their advantages when it comes to obtaining specific information about biological phenomena, and it also permits the application of force in molecules [57]. *In singulo* studies are very direct approaches, following the behavior of an individual molecule in time, thus making it possible to obtain not just the average behavior of many molecules, but rather the

**19**

*Mechanical Properties of Chaperone BiP, the Master Regulator of the Endoplasmic Reticulum*

whole distribution and individual behaviors of a population that may not be homogeneous. It is possible to study a single biomolecule by visualizing it or manipulating it, with the most common approaches being single molecule fluorescence and single molecule force spectroscopy [9]. With force spectroscopy, it is possible to mechanically manipulate and apply forces to molecules in a highly specific manner [58]. This technique lets us measure the mechanical stability of particular domains, instead of the whole protein, thus allowing us to determine the energetic coupling between one domain and the other. Techniques such as atomic force microscopy (AFM), magnetic tweezers, and optical tweezers allow the direct application of mechanical forces to biological macromolecules and let us study the conformational changes [9, 59]. One example of single molecule studies with BiP has been the analysis of the conformational cycle of BiP achieved by single molecule and ensemble FRET measurements. In this study, the authors determined that nucleotide binding resulted in concerted domain movements of BiP. Conformational transitions of the lid domain allowed BiP to discriminate between peptide and protein substrates [41]. Also, we recently developed a method to measure how BiP binds to its substrate using optical tweezers [24]. Without single molecule approaches, it is very difficult to learn about how BiP binds to its substrate, since the substrate of BiP is an unfolded peptide, and if we unfold the substrate, we may also unfold BiP. However, by optical tweezers manipulation, we can specifically unfold the substrate without affecting BiP. Another study, by directly pulling DnaK using optical tweezers, the authors were able to study the mechanics and the order of events of unfolding of each domain of this Hsp70 [60, 61]. This study shows that DnaK has more than two mechanical intermediates in each domain. All the single molecule techniques that exert force on the protein are not able to measure small changes in distance at subnanometer resolution at low forces (below 1–5 pN), and so it is difficult to correlate the elastic properties of the folded protein with ligand binding. A new technique called nano-rheology developed in Giovanni Zocchi's laboratory at the University of California at Los Angeles (UCLA) allows measurement of elasticity in folded proteins [62]. Nano-rheology is a traditional rheology experiment, in which an oscillatory force is directly applied to the protein and where average deformation is measured [63]. This technique exploits sub-Angstrom resolution to study the mechanical properties of the folded state of proteins by applying low force to the proteins in bulk [12]. The universal mechanical property of the folded state is the viscoelastic behavior, meaning, when a protein is subjected to a force, it can behave as an elastic or viscous material, getting stiffer or softer (flexible). Then, stiff and soft here refer to both elastic and viscous mechanical responses; the two are coupled because the structure is viscoelastic [63]. This behavior is relevant for the large conformational changes of protein which often accompany substrate binding in proteins [12, 64]. Using this technique (**Figure 4**), we studied the mechanical properties of BiP, considering the viscoelastic behavior upon ligand binding. We observed that the folded state of the protein behaves like a viscoelastic material, getting softer when it binds nucleotides but stiffer when it binds peptide substrate. The explanation for this mechanical behavior is related to the ATPase cycle of BiP. As shown **Figure 4B**, when the protein is in the presence of ATP, the protein is softer state because the lid is more flexible and the NBD and SBD domains are closer [50, 65]. Also, the protein is in softer state in the presence of ADP, but the structural reason is different. The hydrophobic linker is more flexible, and the domains seem to be in a dynamic distance distribution [3, 6]. Finally, the protein is more rigid or stiffer in the presence of the HTFPAVL peptide substrate. The structural reason is because the lid is close [6]. Additionally, it was observed in presence of peptide the dissociation constant (KD) for ADP decreased

*DOI: http://dx.doi.org/10.5772/intechopen.82080*

#### *Mechanical Properties of Chaperone BiP, the Master Regulator of the Endoplasmic Reticulum DOI: http://dx.doi.org/10.5772/intechopen.82080*

whole distribution and individual behaviors of a population that may not be homogeneous. It is possible to study a single biomolecule by visualizing it or manipulating it, with the most common approaches being single molecule fluorescence and single molecule force spectroscopy [9]. With force spectroscopy, it is possible to mechanically manipulate and apply forces to molecules in a highly specific manner [58]. This technique lets us measure the mechanical stability of particular domains, instead of the whole protein, thus allowing us to determine the energetic coupling between one domain and the other. Techniques such as atomic force microscopy (AFM), magnetic tweezers, and optical tweezers allow the direct application of mechanical forces to biological macromolecules and let us study the conformational changes [9, 59]. One example of single molecule studies with BiP has been the analysis of the conformational cycle of BiP achieved by single molecule and ensemble FRET measurements. In this study, the authors determined that nucleotide binding resulted in concerted domain movements of BiP. Conformational transitions of the lid domain allowed BiP to discriminate between peptide and protein substrates [41]. Also, we recently developed a method to measure how BiP binds to its substrate using optical tweezers [24]. Without single molecule approaches, it is very difficult to learn about how BiP binds to its substrate, since the substrate of BiP is an unfolded peptide, and if we unfold the substrate, we may also unfold BiP. However, by optical tweezers manipulation, we can specifically unfold the substrate without affecting BiP. Another study, by directly pulling DnaK using optical tweezers, the authors were able to study the mechanics and the order of events of unfolding of each domain of this Hsp70 [60, 61]. This study shows that DnaK has more than two mechanical intermediates in each domain. All the single molecule techniques that exert force on the protein are not able to measure small changes in distance at subnanometer resolution at low forces (below 1–5 pN), and so it is difficult to correlate the elastic properties of the folded protein with ligand binding. A new technique called nano-rheology developed in Giovanni Zocchi's laboratory at the University of California at Los Angeles (UCLA) allows measurement of elasticity in folded proteins [62]. Nano-rheology is a traditional rheology experiment, in which an oscillatory force is directly applied to the protein and where average deformation is measured [63]. This technique exploits sub-Angstrom resolution to study the mechanical properties of the folded state of proteins by applying low force to the proteins in bulk [12]. The universal mechanical property of the folded state is the viscoelastic behavior, meaning, when a protein is subjected to a force, it can behave as an elastic or viscous material, getting stiffer or softer (flexible). Then, stiff and soft here refer to both elastic and viscous mechanical responses; the two are coupled because the structure is viscoelastic [63]. This behavior is relevant for the large conformational changes of protein which often accompany substrate binding in proteins [12, 64]. Using this technique (**Figure 4**), we studied the mechanical properties of BiP, considering the viscoelastic behavior upon ligand binding. We observed that the folded state of the protein behaves like a viscoelastic material, getting softer when it binds nucleotides but stiffer when it binds peptide substrate. The explanation for this mechanical behavior is related to the ATPase cycle of BiP. As shown **Figure 4B**, when the protein is in the presence of ATP, the protein is softer state because the lid is more flexible and the NBD and SBD domains are closer [50, 65]. Also, the protein is in softer state in the presence of ADP, but the structural reason is different. The hydrophobic linker is more flexible, and the domains seem to be in a dynamic distance distribution [3, 6]. Finally, the protein is more rigid or stiffer in the presence of the HTFPAVL peptide substrate. The structural reason is because the lid is close [6]. Additionally, it was observed in presence of peptide the dissociation constant (KD) for ADP decreased

*Endoplasmic Reticulum*

later.

equilibrium [45].

demonstrated.

**2.3 Mechanical aspects**

control of BiP conformational ensemble [45].

Upon ATP binding, the linker binds to a cleft in NBD, which is important in transmitting the nucleotide state of NBD to SBD and increases ATP hydrolysis of the NBD once bound to the cleft. Basically, it has been suggested that allostery results from an energetic tug-of-war between domain conformations and formation of two orthogonal interfaces: between the NBD and SBD and between the helical lid and the β subdomain of the SBD [46]. More recently, "soft" amino acid substitutions have been performed in BiP, trying to affect the allosteric communication between SBD and NBD, uncoupling the substrate-binding site with the NBD-SBD interdomain interface. In particular, Val461 was mutated to Phe; Ile526 to Val; Ile437 to Val; and Ile538 to Val. It has been reported that in the presence of ATP, all these "soft" mutations affected the equilibrium between the domain-docked and domain-undocked conformations, suggesting that this residue enables allosteric

Moreover, allosterism in BiP has been studied at the single molecule level with optical tweezer manipulation [24]. The results showed that BiP binds reversibly to the unfolded state of MJ0366 (substrate protein), preventing its refolding, and that this effect depends on both the type and concentration of nucleotides. Additionally, more clues about BiP allosteric mechanism have arisen from BiP ensemble measurements performed with nano-rheological experimental setup, which will be explain

Finally, it has been studied how the posttranslational modification of BiP by AMPylation onto Thr518 [56] could affect BiP conformational cycle, modulating in this way the allosteric mechanism of BiP. The results showed that effectively, this modification shifted BiP conformational equilibrium toward the domain-docked conformation in the presence of ATP, stabilizing the domain docking in the absence of ATP and demonstrating posttranslational fine tuning of BiP conformational

As a general overview, BiP allosteric mechanism has a high level of complexity, as it has different layers of control. From a structural point of view, there are residues that exert the communication between the SBD and NBD domains, and other residues that are involved in stabilizing conformational ensembles of BiP that affect allosteric communication. Moreover, changes in the mechanical properties of BiP are also involved in the allosteric mechanism regulation, as it has been demonstrated with the nano-rheological studies. Finally, posttranslational modifications also play a role in this chaperone function, as their importance in shuffling conformational ensembles, involved in this ATP and Mg2+ dependent cycle, has been

Considering that translocation through the ER is a crucial process to maintain homeostasis inside the cell, it is essential to have a mechanistic understanding of the role that BiP has in translocation to maintain proteostasis. Therefore, classical biochemical assays, or ensemble studies, have been conducted to study each of these processes without taking into account the measurement of forces and changes in elasticity. Single molecule level studies, called *in singulo* studies, have become very relevant during recent years. These studies have become the gold standard to study biomolecular mechanisms because of their advantages when it comes to obtaining specific information about biological phenomena, and it also permits the application of force in molecules [57]. *In singulo* studies are very direct approaches, following the behavior of an individual molecule in time, thus making it possible to obtain not just the average behavior of many molecules, but rather the

**18**

#### **Figure 4.**

*Mechanical aspects of BiP. (A) BiP nano-rheology setup shows the flow chamber with BiP attached to both gold surfaces, the parallel plates capacitor geometry used for mechanical excitation, and the evanescent wave scattering optics used for read out. BiP was directly tethered between a gold film surface evaporated on a glass slide and 20 nm diameter GNPs, constituting the lower part of a thick flow chamber. BiP attachment proceeds via two exposed cys residues at positions 166 and 518, located in NBD and SBD, respectively. GNPs are covered with ssDNAs on the surface to negatively charge them. (B) Model for mechanical response of BiP in the presence of different ligands in ATPase cycle. The illustration shows BiP unbound state. In ATPase cycle of BiP, the structure is softer in two cases: first, in the presence of ATP, the lid is more flexible and the domains are closer leading to an important rigidity decrease. Second, in the presence of ADP, the domains are separated by the linker elongation. SBD seems to be in a dynamic distance distribution with a general trend toward domain separation. Finally, the structure is stiffer in the presence of peptide because the lid of BiP is closed, then generating a compact state.*

1000 times, demonstrating that peptide binding dramatically increases the affinity for ADP which evidences the allosteric coupling between SBD and NBD domains [66].

#### **3. Conclusion**

Changes in the conformational state and viscoelastic properties of BiP triggered by ATP binding and/or hydrolysis are essential for allosteric communication between its domains (NBD and SBD), as these could supply the mechanical work to move the peptides through the Sec61 channel, with BiP behaving as a molecular motor. It is still not completely known how BiP applied the force in the peptide that is translocating or if it just uses the water bath. Taking into account the important role of BiP in proteostasis and diseases, an in-depth study of the functioning of the mechanics of BiP with new technology has major relevance to future research and development in science, biomedicine, and health, as well as in technological developments in biotechnology and even education, thus opening up new investigative directions of great potential and impact for science worldwide.

#### **Acknowledgements**

This work was supported by the Fondo Nacional de Desarrollo Científico y Tecnológico (FONDECYT), project 1,181,361 and PCI PII20150073. We thank all member of the Biochemistry laboratory, Universidad de Chile.

**21**

**Author details**

Hilda M. Alfaro-Valdés†

University of Chile, Chile

Diego Quiroga-Roger†

provided the original work is properly cited.

\*Address all correspondence to: yitowilson@gmail.com

†All authors contributed equally to this book chapter

© 2018 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/ by/3.0), which permits unrestricted use, distribution, and reproduction in any medium,

, Francesca Burgos-Bravo†

and Christian A.M. Wilson\*†

Biochemistry and Molecular Biology Department, Chemistry and Pharmaceutical Sciences Faculty, Mechanobiology and Single Molecule Biochemistry Laboratory,

, Nathalie Casanova-Morales†

,

*Mechanical Properties of Chaperone BiP, the Master Regulator of the Endoplasmic Reticulum*

*DOI: http://dx.doi.org/10.5772/intechopen.82080*

#### **Conflict of interest**

None conflict of interest.

*Mechanical Properties of Chaperone BiP, the Master Regulator of the Endoplasmic Reticulum DOI: http://dx.doi.org/10.5772/intechopen.82080*

### **Author details**

*Endoplasmic Reticulum*

domains [66].

**Figure 4.**

**3. Conclusion**

worldwide.

**Acknowledgements**

**Conflict of interest**

None conflict of interest.

1000 times, demonstrating that peptide binding dramatically increases the affinity for ADP which evidences the allosteric coupling between SBD and NBD

*Mechanical aspects of BiP. (A) BiP nano-rheology setup shows the flow chamber with BiP attached to both gold surfaces, the parallel plates capacitor geometry used for mechanical excitation, and the evanescent wave scattering optics used for read out. BiP was directly tethered between a gold film surface evaporated on a glass slide and 20 nm diameter GNPs, constituting the lower part of a thick flow chamber. BiP attachment proceeds via two exposed cys residues at positions 166 and 518, located in NBD and SBD, respectively. GNPs are covered with ssDNAs on the surface to negatively charge them. (B) Model for mechanical response of BiP in the presence of different ligands in ATPase cycle. The illustration shows BiP unbound state. In ATPase cycle of BiP, the structure is softer in two cases: first, in the presence of ATP, the lid is more flexible and the domains are closer leading to an important rigidity decrease. Second, in the presence of ADP, the domains are separated by the linker elongation. SBD seems to be in a dynamic distance distribution with a general trend toward domain separation. Finally, the structure is stiffer in the presence of peptide because the lid of BiP is closed, then generating a compact state.*

Changes in the conformational state and viscoelastic properties of BiP triggered by ATP binding and/or hydrolysis are essential for allosteric communication between its domains (NBD and SBD), as these could supply the mechanical work to move the peptides through the Sec61 channel, with BiP behaving as a molecular motor. It is still not completely known how BiP applied the force in the peptide that is translocating or if it just uses the water bath. Taking into account the important role of BiP in proteostasis and diseases, an in-depth study of the functioning of the mechanics of BiP with new technology has major relevance to future research and development in science, biomedicine, and health, as well as in technological developments in biotechnology and even education, thus opening up new investigative directions of great potential and impact for science

This work was supported by the Fondo Nacional de Desarrollo Científico y Tecnológico (FONDECYT), project 1,181,361 and PCI PII20150073. We thank all

member of the Biochemistry laboratory, Universidad de Chile.

**20**

Hilda M. Alfaro-Valdés† , Francesca Burgos-Bravo† , Nathalie Casanova-Morales† , Diego Quiroga-Roger† and Christian A.M. Wilson\*† Biochemistry and Molecular Biology Department, Chemistry and Pharmaceutical Sciences Faculty, Mechanobiology and Single Molecule Biochemistry Laboratory, University of Chile, Chile

\*Address all correspondence to: yitowilson@gmail.com

†All authors contributed equally to this book chapter

© 2018 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/ by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

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2017;**114**(23):6040-6045

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[62] Alavi Z, Ariyaratne A, Zocchi G. Nano-rheology measurements reveal that the hydration layer of enzymes partially controls conformational dynamics. Applied Physics Letters.

[63] Casanova-Morales N, Alavi Z, Wilson CA, Zocchi G. Identifying chaotropic and kosmotropic agents by nanorheology. The Journal of Physical Chemistry B. 2018;**122**(14):3754-3759

[64] Huang X, Peng X, Wang Y, Wang Y, Shin DM, El-Sayed MA, et al. A Reexamination of active and passive tumor targeting by using rod-shaped gold nanocrystals and covalently conjugated peptide ligands. ACS Nano.

[65] Rist W, Graf C, Bukau B, Mayer MP. Amide hydrogen exchange reveals conformational changes in Hsp70 chaperones important for allosteric regulation. Journal of Biological Chemistry. 2006;**281**(24):16493-16501

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intramolecular vibrational energy flow in proteins reveals functionally important residues. The Journal of Physical Chemistry Letters.

[54] Wisniewska M, Karlberg T, Lehtiö L, Johansson I, Kotenyova T, Moche M, et al. Crystal structures of the ATPase domains of four human Hsp70 isoforms: HSPA1L/Hsp70-hom, HSPA2/Hsp70- 2, HSPA6/Hsp70B', and HSPA5/BiP/ GRP78. PLoS One. 2010;**5**(1):e8625

[55] Wei J, Gaut JR, Hendershot LM. In vitro dissociation of BiP-peptide complexes requires a conformational change in BiP after ATP binding but does not require ATP hydrolysis. Journal of Biological Chemistry. 1995;**270**(44):26677-26682

[56] Preissler S, Rato C, Chen R, Antrobus R, Ding S, Fearnley IM, et al. AMPylation matches BiP activity to client protein load in the endoplasmic reticulum. eLife. 2015;**4**:e12621

[57] Bustamante C. In singulo biochemistry: When less is more. Annual Review of Biochemistry.

[58] Shank EA, Cecconi C, Dill JW, Marqusee S, Bustamante C. The folding cooperativity of a protein is controlled by its chain topology. Nature.

[59] Neuman KC, Nagy A. Singlemolecule force spectroscopy: Optical tweezers, magnetic tweezers and atomic force microscopy. Nature Methods.

[60] Bauer D, Merz DR, Pelz B, Theisen KE, Yacyshyn G, Mokranjac D, et al. Nucleotides regulate the mechanical hierarchy between subdomains of the nucleotide binding domain of the Hsp70 chaperone DnaK. Proceedings of the National Academy of Sciences.

2015;**112**(33):10389-10394

2008;**77**(1):45-50

2010;**465**:637

2008;**5**:491

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intramolecular vibrational energy flow in proteins reveals functionally important residues. The Journal of Physical Chemistry Letters. 2011;**2**(16):2073-2078

*Endoplasmic Reticulum*

2014;**113**(19):198101

[36] Wiedemann N, Pfanner N. Mitochondrial machineries for protein import and assembly. Annual Review of [45] Wieteska L, Shahidi S, Zhuravleva A. Allosteric fine-tuning of the conformational equilibrium poises the chaperone BiP for post-translational regulation. eLife. 2017;**6**:e29430

[46] Zhuravleva A, Gierasch LM. Substrate-binding domain

conformational dynamics mediate Hsp70 allostery. Proceedings of the National Academy of Sciences.

[47] McCarty JS, Buchberger A, Reinstein J, Bukau B. The role of ATP in the functional cycle of the DnaK chaperone system. Journal of Molecular

Biology. 1995;**249**(1):126-137

[48] Zhuravleva A, Clerico Eugenia M, Gierasch Lila M. An Interdomain energetic tug-of-war creates the allosterically active state in Hsp70 molecular chaperones. Cell. 2012;**151**(6):1296-1307

[49] Kityk R, Kopp J, Sinning I, Mayer MP. Structure and dynamics of the ATP-bound open conformation of Hsp70 chaperones. Molecular Cell.

[50] Swain JF, Dinler G, Sivendran R, Montgomery DL, Stotz M, Gierasch LM. Hsp70 chaperone ligands control domain association via an allosteric mechanism mediated by the interdomain linker. Molecular Cell. 2007;**26**(1):27-39

[51] Banerjee R, Jayaraj GG, Peter JJ, Kumar V, Mapa K. Monitoring conformational heterogeneity of the lid of DnaK substrate-binding domain during its chaperone cycle. The FEBS Journal. 2016;**283**(15):2853-2868

[52] Vogel M, Bukau B, Mayer MP. Allosteric regulation of Hsp70 chaperones by a proline switch. Molecular Cell. 2006;**21**(3):359-367

[53] Martínez L, Figueira ACM, Webb P, Polikarpov I, Skaf MS. Mapping the

2012;**48**(6):863-874

2015;**2015**:06692

Biochemistry. 2017;**86**:685-714

[37] Ariyaratne A, Wu C, Tseng CY, Zocchi G. Dissipative dynamics of enzymes. Physical Review Letters.

[38] Zhu X, Zhao X, Burkholder WF, Gragerov A, Ogata CM, Gottesman ME, et al. Structural analysis of substrate binding by the molecular chaperone DnaK. Science. 1996;**272**(5268):1606

[39] Braakman I, Hebert DN. Protein folding in the endoplasmic reticulum. Cold Spring Harbor Perspectives in

[40] Otero JH, Lizák B, Hendershot LM. Life and death of a BiP substrate. Seminars in Cell & Developmental Biology. 2010;**21**(5):472-478

[41] Marcinowski M, Höller M, Feige MJ,

[42] Bertelsen EB, Chang L, Gestwicki

[43] Mapa K, Sikor M, Kudryavtsev V, Waegemann K, Kalinin S, Seidel CAM, et al. The conformational dynamics of the mitochondrial Hsp70 chaperone. Molecular Cell. 2010;**38**(1):89-100

[44] Yang J, Nune M, Zong Y, Zhou L, Liu Q. Close and allosteric opening of the polypeptide-binding site in a human Hsp70 chaperone BiP. Structure.

JE, Zuiderweg ER. Solution conformation of wild-type *E. coli* Hsp70 (DnaK) chaperone complexed with ADP and substrate. Proceedings of the National Academy of Sciences.

2009;**106**(21):8471-8476

2015;**23**(12):2191-2203

Baerend D, Lamb DC, Buchner J. Substrate discrimination of the chaperone BiP by autonomous and cochaperone-regulated conformational transitions. Nature Structural & Molecular Biology. 2011;**18**:150

Biology. 2013;**5**(5):a013201

**24**

[54] Wisniewska M, Karlberg T, Lehtiö L, Johansson I, Kotenyova T, Moche M, et al. Crystal structures of the ATPase domains of four human Hsp70 isoforms: HSPA1L/Hsp70-hom, HSPA2/Hsp70- 2, HSPA6/Hsp70B', and HSPA5/BiP/ GRP78. PLoS One. 2010;**5**(1):e8625

[55] Wei J, Gaut JR, Hendershot LM. In vitro dissociation of BiP-peptide complexes requires a conformational change in BiP after ATP binding but does not require ATP hydrolysis. Journal of Biological Chemistry. 1995;**270**(44):26677-26682

[56] Preissler S, Rato C, Chen R, Antrobus R, Ding S, Fearnley IM, et al. AMPylation matches BiP activity to client protein load in the endoplasmic reticulum. eLife. 2015;**4**:e12621

[57] Bustamante C. In singulo biochemistry: When less is more. Annual Review of Biochemistry. 2008;**77**(1):45-50

[58] Shank EA, Cecconi C, Dill JW, Marqusee S, Bustamante C. The folding cooperativity of a protein is controlled by its chain topology. Nature. 2010;**465**:637

[59] Neuman KC, Nagy A. Singlemolecule force spectroscopy: Optical tweezers, magnetic tweezers and atomic force microscopy. Nature Methods. 2008;**5**:491

[60] Bauer D, Merz DR, Pelz B, Theisen KE, Yacyshyn G, Mokranjac D, et al. Nucleotides regulate the mechanical hierarchy between subdomains of the nucleotide binding domain of the Hsp70 chaperone DnaK. Proceedings of the National Academy of Sciences. 2015;**112**(33):10389-10394

[61] Mandal SS, Merz DR, Buchsteiner M, Dima RI, Rief M, Žoldák G. Nanomechanics of the substrate binding domain of Hsp70 determine its allosteric ATP-induced conformational change. Proceedings of the National Academy of Sciences. 2017;**114**(23):6040-6045

[62] Alavi Z, Ariyaratne A, Zocchi G. Nano-rheology measurements reveal that the hydration layer of enzymes partially controls conformational dynamics. Applied Physics Letters. 2015;**106**(20):203702

[63] Casanova-Morales N, Alavi Z, Wilson CA, Zocchi G. Identifying chaotropic and kosmotropic agents by nanorheology. The Journal of Physical Chemistry B. 2018;**122**(14):3754-3759

[64] Huang X, Peng X, Wang Y, Wang Y, Shin DM, El-Sayed MA, et al. A Reexamination of active and passive tumor targeting by using rod-shaped gold nanocrystals and covalently conjugated peptide ligands. ACS Nano. 2010;**4**(10):5887-5896

[65] Rist W, Graf C, Bukau B, Mayer MP. Amide hydrogen exchange reveals conformational changes in Hsp70 chaperones important for allosteric regulation. Journal of Biological Chemistry. 2006;**281**(24):16493-16501

[66] https://onlinelibrary.wiley.com/doi/ full/10.1002/pro.3432

Chapter 3

Abstract

1. Introduction

27

processes occur in the ER [1–4].

Endoplasmic Reticulum-

Associated Degradation (ERAD)

Burcu Erbaykent Tepedelen and Petek Ballar Kirmizibayrak

The newly synthesized proteins are kept in the endoplasmic reticulum (ER) until their maturation is completed. The accurate protein folding is vital for homeostasis, but this process is error-prone since it is chemically complicated. Aberrant folding may result in aggregates having a toxic gain of function or may lead to a loss of protein function; therefore, protein misfolding can lead to several pathologies. The ER protein quality control mechanism monitors the fidelity of protein folding. Those proteins that fail to fold or assemble properly are subjected to degradation via a process known as ER-associated degradation (ERAD). Besides clearing proteins having folding problems, ERAD is also known to regulate the levels of some physiological proteins including 3-hydroxy-3-methylglutarylcoenzymeA reductase (HMGR) catalyzing the rate-limiting step of cholesterol biosynthesis. ERAD is a complex, multistep process starting with the recognition and

targeting of substrates, followed by ubiquitination, retrotranslocation and

Keywords: endoplasmic reticulum-associated degradation (ERAD), protein misfolding, ubiquitin-mediated degradation, proteasomal degradation

The endoplasmic reticulum (ER) is an extensive network of flattened, membrane-enclosed tubes or sacs that extends throughout the cytosol [1]. ER has important roles in many biochemical processes required for cell survival and normal cellular functions. ER regulates these cellular processes through proteins that are localized in its complex network structures [1–3]. In addition to protein synthesis, significant cellular activities such as protein transport and folding, lipid and steroid synthesis, carbohydrate metabolism, calcium storage and protein quality control

Approximately one-third of all newly synthesized proteins are targeted to the ER

and traffic to other organelles of secretory pathway, plasma membrane or the extracellular space [5]. Protein translocation to the ER occurs through Sec61 complex [6, 7]. As synchronized with translocation, protein is exposed to the ER's oxidizing and calcium-rich environment, which is suitable for protein folding and

together with its tight regulation will be discussed.

proteasomal degradation. A large number of ERAD factors functioning in different molecular machineries increases the complexity of mammalian ERAD. ERAD is fundamental for human health and there is increasing evidence linking ERAD with various diseases. Here, the different modules/machineries of the ERAD process

#### Chapter 3

## Endoplasmic Reticulum-Associated Degradation (ERAD)

Burcu Erbaykent Tepedelen and Petek Ballar Kirmizibayrak

#### Abstract

The newly synthesized proteins are kept in the endoplasmic reticulum (ER) until their maturation is completed. The accurate protein folding is vital for homeostasis, but this process is error-prone since it is chemically complicated. Aberrant folding may result in aggregates having a toxic gain of function or may lead to a loss of protein function; therefore, protein misfolding can lead to several pathologies. The ER protein quality control mechanism monitors the fidelity of protein folding. Those proteins that fail to fold or assemble properly are subjected to degradation via a process known as ER-associated degradation (ERAD). Besides clearing proteins having folding problems, ERAD is also known to regulate the levels of some physiological proteins including 3-hydroxy-3-methylglutarylcoenzymeA reductase (HMGR) catalyzing the rate-limiting step of cholesterol biosynthesis. ERAD is a complex, multistep process starting with the recognition and targeting of substrates, followed by ubiquitination, retrotranslocation and proteasomal degradation. A large number of ERAD factors functioning in different molecular machineries increases the complexity of mammalian ERAD. ERAD is fundamental for human health and there is increasing evidence linking ERAD with various diseases. Here, the different modules/machineries of the ERAD process together with its tight regulation will be discussed.

Keywords: endoplasmic reticulum-associated degradation (ERAD), protein misfolding, ubiquitin-mediated degradation, proteasomal degradation

#### 1. Introduction

The endoplasmic reticulum (ER) is an extensive network of flattened, membrane-enclosed tubes or sacs that extends throughout the cytosol [1]. ER has important roles in many biochemical processes required for cell survival and normal cellular functions. ER regulates these cellular processes through proteins that are localized in its complex network structures [1–3]. In addition to protein synthesis, significant cellular activities such as protein transport and folding, lipid and steroid synthesis, carbohydrate metabolism, calcium storage and protein quality control processes occur in the ER [1–4].

Approximately one-third of all newly synthesized proteins are targeted to the ER and traffic to other organelles of secretory pathway, plasma membrane or the extracellular space [5]. Protein translocation to the ER occurs through Sec61 complex [6, 7]. As synchronized with translocation, protein is exposed to the ER's oxidizing and calcium-rich environment, which is suitable for protein folding and

co- and post-translational modifications such as glycosylation, disulfide bond formation and glycosylphosphatidylinositol (GPI) anchoring [8]. During this folding process, many proteins such as lectin-type molecular chaperones (e.g., calnexin (CNX) or calreticulin (CLR)), HSP70-like chaperone BiP) and enzymes like protein disulfide isomerases (PDI) work in association with each other [4, 9, 10]. Conformational maturation and folding of the proteins in the ER are instantly controlled through the added N-glycan groups to decide whether the proteins are directed to distant compartments via the secretory pathway or included in the refolding cycle [11, 12].

by binding seconder messenger inositol 1,4,5-triphosphate (IP3); type II

metastasis suppressor KAI1 levels [22, 25–28].

Endoplasmic Reticulum-Associated Degradation (ERAD)

DOI: http://dx.doi.org/10.5772/intechopen.82043

cytoplasm [31].

during ERAD pathway [12, 19–21].

viral infection and albinism [4, 33].

2. Molecular mechanisms of ERAD

place in the ER.

29

patterns of the ERAD will be summarized and presented.

2.1 Protein folding process and recognition of misfolded proteins

iodothyronine deiodinase, an ER-localized enzyme converting thyroxin (T4) to the biologically active hormone triiodothyronine (T3) and GABA neurotransmitter receptor responsible for the reduction of neuronal excitability and the tumor

Some viruses hijack the ERAD system through encoding effectors by serving as adaptors that redirect correctly folded molecules towards degradation. US2 and US11, the human cytomegalovirus gene products, induce degradation of major histocompatibility complex (MHC) class I heavy chain, which enables virusinfected cell to avoid detection by the immune system [29]. Similarly, Vpu is a glycoprotein encoded in the human immunodeficiency virus (HIV-1) genome and binds and targets newly synthesized CD4 for degradation [30], allowing them to evade immunosurveillance. Moreover, toxins like diphtheria, cholera and ricin enter the cell by endocytosis and move to the ER. They use the ERAD system to escape from the ER lumen and gain access to their enzymatic substrates in the

ERAD is a highly complicated and regulated mechanism in which the diversity and combination of components change according to the protein to be destroyed [19–21, 32]. Maturation-defective proteins are removed from the ER lumen or lipid

proteasome. The ubiquitin system is an integral part of the ERAD and is composed of factors necessary for the recruitment, processing and binding of ubiquitin chains to substrates [24]. In other words, ERAD is composed of steps that include substrate

proteasomal degradation. Several key molecules such as E1, E2, and E3 enzymes responsible for ubiquitin transfer, channel components responsible for retrotranslocation, chaperones and cofactor proteins function in a synchronized manner

This critical role of ERAD in the regulation of cell homeostasis is an evident that ERAD disorders will have important effects on cell survival. Furthermore, it has been shown that aberrations in ERAD function play a role in the pathology of nearly 70 diseases such as cystic fibrosis, α1-antitrypsin (AAT) insufficiency, diabetes, neurodegenerative diseases (Parkinson, Alzheimer's and Huntington's diseases),

In this section, the knowledge related to the basic mechanism and regulation

About 30% of the total proteins and all transmembrane proteins of the cell are synthesized in the ER, which acts as a portal for entry into the secretory pathway via the Sec61 channel [7–8]. As being translocated, the N terminal hydrophobic signal sequence of newly synthesized protein is cleaved by a peptidase complex [34]. Co- and post-translational modifications such as disulfide bond formation, initial steps of N-glycosylation, and glycophosphatidylinositol (GPI) anchorage take

The oxidizing environment of ER assists the formation of disulfide bonds, which

stabilizes tertiary protein structure and facilitates protein assembly. During the folding process, disulfide bonds are formed through the oxidation of pairs of free thiols on cysteine residues by protein disulfide isomerases (PDIs). PDIs act as

bilayer by retrotranslocation through the ERAD pathway and degraded by

selection, modification with ubiquitin chain, retrotranslocation and 26S

The folding process is not completely accurate. In mammals, 30% of all newly synthesized proteins are estimated to be incorrectly folded [13]. However, genetic mutations, errors in transcription and translation, toxic compounds and cellular stresses such as defects in cellular redox regulation due to hypoxia, oxidants and reducing agents that interact with disulfide bonds in the ER lumen, glucose starvation and abnormalities in calcium regulation lead to a significant increase in the ratio of incorrectly folded proteins [4, 11, 14]. Adequate removal of these unwanted proteins is crucial for protecting cells from proteotoxicity caused by the formation of protein aggregates through the re-opening of hydrophobic residues as well as by unfolded or misfolded proteins that may compete with their properly folded counterparts for substrate binding or for complex formation with partners. Even though the primary damage of these unwanted proteins is restricted to the cell they reside, the damage gets wider if it is a secretory protein [11]. Therefore, there is a robust control via "Protein Quality Control Mechanisms" for the removal of defective proteins in living cells, and thus, only properly folded proteins are allowed to exit from ER lumen to the secretory pathway [11, 15–18]. When the folding process fails, the terminal mannose residues from the core glycan chain are gradually removed, allowing the proteins to be recognized by mannose-specific lectins and defective proteins are transferred to the 26S proteasome for degradation through the protein quality control mechanism called "ER-associated degradation (ERAD)" [19–21].

In addition to misfolding proteins, ERAD also targets some proteins that might fold into their native structures under the right conditions and also orphan subunits of oligomeric complexes. The chloride channel protein CFTR (cystic fibrosis transmembrane conductance regulator) is the best example, where it is targeted to ERAD as a consequence of its complex and inefficient folding pathway. The low folding efficiency is further decreased upon mutation as seen in CFTRΔF508. CFTRΔF508 is the most common mutation found in cystic fibrosis patients, can fold and function in plasma membrane; thus, degradation of CFTR via ERAD is obtrusive. ERAD also functions in supporting the correct stoichiometry of multimeric protein complexes by degrading components that are produced in excess of the limiting monomer [22]. For example, the unassembled subunits of T cell receptor-like TCRα and CD3δ are also well-known ERAD substrates [23]. These proteins contain charged residues in the intramembrane sections promoting the assembly of complexes. However, when oligomerization is not proper, these residues might initiate degradation via recruiting specific ERAD factors [23].

ERAD also functions in cell homeostasis by regulating the endogenous levels of many enzymes and signal molecules especially those localized to the ER membrane or plasma membrane under physiological conditions [24]. For instance, ERAD plays a homeostatic role in the regulation of HMG-CoA reductase (HMGR), which is the key enzyme of cholesterol metabolism; apolipoprotein B, an essential secreted protein member of triacylglycerol-rich lipoproteins responsible for the export of lipids, triglycerides and cholesterol; hepatic cytochrome P450 enzyme 3A4 metabolizing endo- and xenobiotics; IP3 receptor, an ER-localized protein allowing Ca2+ release

co- and post-translational modifications such as glycosylation, disulfide bond formation and glycosylphosphatidylinositol (GPI) anchoring [8]. During this folding process, many proteins such as lectin-type molecular chaperones (e.g., calnexin (CNX) or calreticulin (CLR)), HSP70-like chaperone BiP) and enzymes like protein disulfide isomerases (PDI) work in association with each other [4, 9, 10]. Conformational maturation and folding of the proteins in the ER are instantly controlled through the added N-glycan groups to decide whether the proteins are directed to distant compartments via the secretory pathway or included in the refolding cycle

The folding process is not completely accurate. In mammals, 30% of all newly synthesized proteins are estimated to be incorrectly folded [13]. However, genetic mutations, errors in transcription and translation, toxic compounds and cellular stresses such as defects in cellular redox regulation due to hypoxia, oxidants and reducing agents that interact with disulfide bonds in the ER lumen, glucose starvation and abnormalities in calcium regulation lead to a significant increase in the ratio of incorrectly folded proteins [4, 11, 14]. Adequate removal of these unwanted proteins is crucial for protecting cells from proteotoxicity caused by the formation of protein aggregates through the re-opening of hydrophobic residues as well as by unfolded or misfolded proteins that may compete with their properly folded counterparts for substrate binding or for complex formation with partners. Even though the primary damage of these unwanted proteins is restricted to the cell they reside, the damage gets wider if it is a secretory protein [11]. Therefore, there is a robust control via "Protein Quality Control Mechanisms" for the removal of defective proteins in living cells, and thus, only properly folded proteins are allowed to exit from ER lumen to the secretory pathway [11, 15–18]. When the folding process fails, the terminal mannose residues from the core glycan chain are gradually removed, allowing the proteins to be recognized by mannose-specific lectins and defective proteins are transferred to the 26S proteasome for degradation through the protein quality control mechanism called "ER-associated degradation (ERAD)"

In addition to misfolding proteins, ERAD also targets some proteins that might fold into their native structures under the right conditions and also orphan subunits of oligomeric complexes. The chloride channel protein CFTR (cystic fibrosis transmembrane conductance regulator) is the best example, where it is targeted to ERAD as a consequence of its complex and inefficient folding pathway. The low folding efficiency is further decreased upon mutation as seen in CFTRΔF508. CFTRΔF508 is the most common mutation found in cystic fibrosis patients, can fold and function in plasma membrane; thus, degradation of CFTR via ERAD is obtrusive. ERAD also functions in supporting the correct stoichiometry of multimeric protein complexes by degrading components that are produced in excess of the limiting monomer [22]. For example, the unassembled subunits of T cell receptor-like TCRα and CD3δ are also well-known ERAD substrates [23]. These proteins contain charged residues in the intramembrane sections promoting the assembly of complexes. However, when oligomerization is not proper, these residues might initiate degra-

ERAD also functions in cell homeostasis by regulating the endogenous levels of many enzymes and signal molecules especially those localized to the ER membrane or plasma membrane under physiological conditions [24]. For instance, ERAD plays a homeostatic role in the regulation of HMG-CoA reductase (HMGR), which is the key enzyme of cholesterol metabolism; apolipoprotein B, an essential secreted protein member of triacylglycerol-rich lipoproteins responsible for the export of lipids, triglycerides and cholesterol; hepatic cytochrome P450 enzyme 3A4 metabolizing endo- and xenobiotics; IP3 receptor, an ER-localized protein allowing Ca2+ release

dation via recruiting specific ERAD factors [23].

[11, 12].

Endoplasmic Reticulum

[19–21].

28

by binding seconder messenger inositol 1,4,5-triphosphate (IP3); type II iodothyronine deiodinase, an ER-localized enzyme converting thyroxin (T4) to the biologically active hormone triiodothyronine (T3) and GABA neurotransmitter receptor responsible for the reduction of neuronal excitability and the tumor metastasis suppressor KAI1 levels [22, 25–28].

Some viruses hijack the ERAD system through encoding effectors by serving as adaptors that redirect correctly folded molecules towards degradation. US2 and US11, the human cytomegalovirus gene products, induce degradation of major histocompatibility complex (MHC) class I heavy chain, which enables virusinfected cell to avoid detection by the immune system [29]. Similarly, Vpu is a glycoprotein encoded in the human immunodeficiency virus (HIV-1) genome and binds and targets newly synthesized CD4 for degradation [30], allowing them to evade immunosurveillance. Moreover, toxins like diphtheria, cholera and ricin enter the cell by endocytosis and move to the ER. They use the ERAD system to escape from the ER lumen and gain access to their enzymatic substrates in the cytoplasm [31].

ERAD is a highly complicated and regulated mechanism in which the diversity and combination of components change according to the protein to be destroyed [19–21, 32]. Maturation-defective proteins are removed from the ER lumen or lipid bilayer by retrotranslocation through the ERAD pathway and degraded by proteasome. The ubiquitin system is an integral part of the ERAD and is composed of factors necessary for the recruitment, processing and binding of ubiquitin chains to substrates [24]. In other words, ERAD is composed of steps that include substrate selection, modification with ubiquitin chain, retrotranslocation and 26S proteasomal degradation. Several key molecules such as E1, E2, and E3 enzymes responsible for ubiquitin transfer, channel components responsible for retrotranslocation, chaperones and cofactor proteins function in a synchronized manner during ERAD pathway [12, 19–21].

This critical role of ERAD in the regulation of cell homeostasis is an evident that ERAD disorders will have important effects on cell survival. Furthermore, it has been shown that aberrations in ERAD function play a role in the pathology of nearly 70 diseases such as cystic fibrosis, α1-antitrypsin (AAT) insufficiency, diabetes, neurodegenerative diseases (Parkinson, Alzheimer's and Huntington's diseases), viral infection and albinism [4, 33].

In this section, the knowledge related to the basic mechanism and regulation patterns of the ERAD will be summarized and presented.

#### 2. Molecular mechanisms of ERAD

#### 2.1 Protein folding process and recognition of misfolded proteins

About 30% of the total proteins and all transmembrane proteins of the cell are synthesized in the ER, which acts as a portal for entry into the secretory pathway via the Sec61 channel [7–8]. As being translocated, the N terminal hydrophobic signal sequence of newly synthesized protein is cleaved by a peptidase complex [34]. Co- and post-translational modifications such as disulfide bond formation, initial steps of N-glycosylation, and glycophosphatidylinositol (GPI) anchorage take place in the ER.

The oxidizing environment of ER assists the formation of disulfide bonds, which stabilizes tertiary protein structure and facilitates protein assembly. During the folding process, disulfide bonds are formed through the oxidation of pairs of free thiols on cysteine residues by protein disulfide isomerases (PDIs). PDIs act as

cycles, and after initial oxidation, disulfide bonds are sometimes isomerized by PDI and ERp57, which is a thiol oxidoreductase, in order to stabilize the correct folding of protein [35]. Conversely, the reduction of disulfide bonds of misfolded proteins is necessary for retrotranslocation step of ERAD. Indeed, PDI enables the retrotranslocation of the simian virüs-40 (SV-40) and cholera toxin [36, 37]. ERdj5, an ER oxidoreductase, reduces disulfide bonds and interacts with EDEM (ERdegradation enhancing mannosidase-like protein) and also accelerates the step of retrotranslocation of SV-40 [37]. ERDJ5 also regulates the degradation of diseasecausing α1-antitrypsin variant (null Hong Kong) [38].

misfolded lesion (ER lumen, ER membrane and cytoplasm) cytosolic or luminal chaperones function in the recognition and targeting for the degradation [44].

of ERMan1 [49]. It appeared that EDEMs also play an important role in

proteins. In summary, EDEMs are directly or indirectly involved in

CNX, receives substrates from CNX cycle and facilitates ERAD substrate

mannose-trimmed proteins for ERAD (Figure 1).

Endoplasmic Reticulum-Associated Degradation (ERAD)

DOI: http://dx.doi.org/10.5772/intechopen.82043

Protein quality control and targeting misfolding proteins to the ERAD.

Figure 1.

31

demannosylation of substrates [50]. EDEM1 also prevents reglycosylation and promotes retrotranslocation and degradation of some ERAD substrates [51]. On the other hand, while mannosidase homology domain (MHD) of Htm1p is necessary for substrate binding, mammalian EDEM1 binds misfolded proteins independent of MHD domain, and therefore, EDEM1 substrate binding may not require mannose trimming or even glycosylation [52]. Thus, in addition to N-linked oligosaccharide moieties of glycoproteins, EDEM1 can recognize the folding lesions of misfolded

demannosylation of glycoproteins and/or serve as receptors that bind and target

Truncation of terminal mannose from branch C exposes α terminal α1,6-bonded mannose residues functioning as a recognition signal for ERAD lectins such as OS9 (Yos9 in yeast) and XTP3-B (Figure 2). Through their mannose-6-phosphate receptor homology (MRH) domain, both proteins primarily recognize α1,6-linked mannose j. Additionally, OS-9 also recognizes α1,6-linked mannose e and c [53]. Several reports suggest that factors (EDEMs, OS9 and XTP3-B) required for substrate recognition and targeting reside within supramolecular complexes and/or interact with important ERAD regulators [54]. For example, EDEM1 interacts with

It is possible to study substrate recognition during ERAD using model misfolded proteins. It is clear that de-mannosylation is required for degradation of misfolded glycoproteins since inhibition of this mannose trimming stabilizes misfolded glycoproteins in the ER [45]. Overexpression of ERMan1 accelerates the degradation of N-glycosylated proteins [39, 46]. The resulting Man8-GlcNAc2 containing glycoprotein after this trimming becomes a substrate for EDEM1 (ER-degradation enhancing mannosidase-like protein 1, Htm1p in yeast)—a mannosidase-related lectin in the ER. It was further proposed that misfolded glycoproteins interact with ERManI and EDEM1 for their ERAD, and lectin-carbohydrate interaction found to be crucial for EDEM substrate recognition [47]. Although ERMan1 was suggested to be a biological timer initiating the ERAD of misfolded proteins [48], recent studies revealed that mannosidases are not solely responsible for intensive demannosylation during ERAD, especially under non-basal conditions. Under ER stress (unfolded protein response active) conditions, the transcriptional elevation of EDEM1 enhances the ERAD efficiency by suppressing proteolytic downregulation

Folding is aided by molecular chaperones shepherding against misfolding and unfolding. Chaperone-like glycans bind to N-glycans playing a crucial role in protein folding and degradation. It is apparent that N-glycosylation, quality control of protein folding and ERAD are functionally linked. After entering to the ER, a large majority of the newly synthesized polypeptide chain are being N-linked glycosylated. The oligosaccharyltransferase enzyme recognizes the Asn-X-Ser/Thr consensus sequence in the most of the nascent protein molecule and covalently integrates a high mannose containing core glycan groups (Glc3Man9GlcNAc2) from dolichol localized on the ER membrane to the protein [39]. Due to the very short half-life of triglycosylated form of protein-bound oligosaccharide, glycan processing starts immediately after the transfer of precursor glycan groups through glucosidase enzymes. Following cleavage of two of three glucose residues, the nascent protein could interact with quality control lectins like CNX and CLR. This interaction is preserved until cleavage of remaining glucose residue. After releasing the glycoprotein from CNX/CLR cycle, final glucose is also trimmed creating unglycosylated substrate. This compromises the interaction of substrate with the lectin chaperones. At this stage, if protein is properly folded, it could exit the ER for their final destination. However, if glycoprotein is still unfolded, it is retained in the ER and reglucosylated by UDP-glucose:glycoprotein glucosyltransferase and rebound with CNX and CLR giving protein more time for proper folding [40, 41]. It is not yet understood the mechanisms involved in the termination of reglycosylation/deglycosylation cycles. However, it is clear that, if the polypeptide chain cannot reach its mature form after repeated folding attempts, terminal mannose residues from the core glycan chain are gradually removed by ER α1,2 mannosidase I (ERMan1). ERMan1 produces Man8GlcNAc2 isomer by removing a mannose residue from the middle branch of N-glycans. By this trimming, glycoprotein becomes poorer substrates for reglycosylation and exit from the CNX cycle [11].

The hydrophobic patches of properly folded proteins are usually buried within the interior of soluble proteins. However, those patches could be exposed in misfolded proteins. If a protein has exposed hydrophobic surfaces, BiP binds to it in order to hide these aggregation-prone surfaces for proper folding attempts by preventing aggregation. However, if folding does not succeed or delayed, extended chaperone-misfolding protein interaction serve for a sophisticated process where protein is transferred to other chaperones and/or to the ERAD process [27, 42].

It is well accepted that the first step of ERAD is selection of misfolded proteins by chaperones. As early as 1999, it was found that yeast ERAD substrates strikingly differed in their requirement for the ER-luminal Hsp70, BiP [43]. The degradation of soluble substrates such as pαF and a mutant form of the vacuolar protease carboxypeptidase Y\* (CPY\*) were dependent on BiP, while degradation of transmembrane proteins Pdr5\*p, Ste6-166p, Sec61-2p and Hmg2p occurred in a BiPindependent manner. In 2004, it has been shown that substrates with cytosolic domain such as Ste6-166p were degraded BiP-independently, while proteins with luminal defects required BiP, suggesting that depending on the topology of

#### Endoplasmic Reticulum-Associated Degradation (ERAD) DOI: http://dx.doi.org/10.5772/intechopen.82043

cycles, and after initial oxidation, disulfide bonds are sometimes isomerized by PDI and ERp57, which is a thiol oxidoreductase, in order to stabilize the correct folding of protein [35]. Conversely, the reduction of disulfide bonds of misfolded proteins

retrotranslocation of the simian virüs-40 (SV-40) and cholera toxin [36, 37]. ERdj5, an ER oxidoreductase, reduces disulfide bonds and interacts with EDEM (ERdegradation enhancing mannosidase-like protein) and also accelerates the step of retrotranslocation of SV-40 [37]. ERDJ5 also regulates the degradation of disease-

Folding is aided by molecular chaperones shepherding against misfolding and unfolding. Chaperone-like glycans bind to N-glycans playing a crucial role in protein folding and degradation. It is apparent that N-glycosylation, quality control of protein folding and ERAD are functionally linked. After entering to the ER, a large

glycosylated. The oligosaccharyltransferase enzyme recognizes the Asn-X-Ser/Thr consensus sequence in the most of the nascent protein molecule and covalently integrates a high mannose containing core glycan groups (Glc3Man9GlcNAc2) from dolichol localized on the ER membrane to the protein [39]. Due to the very short

processing starts immediately after the transfer of precursor glycan groups through glucosidase enzymes. Following cleavage of two of three glucose residues, the nascent protein could interact with quality control lectins like CNX and CLR. This interaction is preserved until cleavage of remaining glucose residue. After releasing the glycoprotein from CNX/CLR cycle, final glucose is also trimmed creating unglycosylated substrate. This compromises the interaction of substrate with the lectin chaperones. At this stage, if protein is properly folded, it could exit the ER for their final destination. However, if glycoprotein is still unfolded, it is retained in the ER and reglucosylated by UDP-glucose:glycoprotein glucosyltransferase and rebound with CNX and CLR giving protein more time for proper folding [40, 41]. It

reglycosylation/deglycosylation cycles. However, it is clear that, if the polypeptide chain cannot reach its mature form after repeated folding attempts, terminal mannose residues from the core glycan chain are gradually removed by ER α1,2 mannosidase I (ERMan1). ERMan1 produces Man8GlcNAc2 isomer by removing a

The hydrophobic patches of properly folded proteins are usually buried within

misfolded proteins. If a protein has exposed hydrophobic surfaces, BiP binds to it in order to hide these aggregation-prone surfaces for proper folding attempts by preventing aggregation. However, if folding does not succeed or delayed, extended chaperone-misfolding protein interaction serve for a sophisticated process where protein is transferred to other chaperones and/or to the ERAD process [27, 42]. It is well accepted that the first step of ERAD is selection of misfolded proteins by chaperones. As early as 1999, it was found that yeast ERAD substrates strikingly differed in their requirement for the ER-luminal Hsp70, BiP [43]. The degradation of soluble substrates such as pαF and a mutant form of the vacuolar protease carboxypeptidase Y\* (CPY\*) were dependent on BiP, while degradation of transmembrane proteins Pdr5\*p, Ste6-166p, Sec61-2p and Hmg2p occurred in a BiPindependent manner. In 2004, it has been shown that substrates with cytosolic domain such as Ste6-166p were degraded BiP-independently, while proteins with luminal defects required BiP, suggesting that depending on the topology of

is necessary for retrotranslocation step of ERAD. Indeed, PDI enables the

majority of the newly synthesized polypeptide chain are being N-linked

half-life of triglycosylated form of protein-bound oligosaccharide, glycan

is not yet understood the mechanisms involved in the termination of

mannose residue from the middle branch of N-glycans. By this trimming, glycoprotein becomes poorer substrates for reglycosylation and exit from the

the interior of soluble proteins. However, those patches could be exposed in

CNX cycle [11].

30

causing α1-antitrypsin variant (null Hong Kong) [38].

Endoplasmic Reticulum

misfolded lesion (ER lumen, ER membrane and cytoplasm) cytosolic or luminal chaperones function in the recognition and targeting for the degradation [44].

It is possible to study substrate recognition during ERAD using model misfolded proteins. It is clear that de-mannosylation is required for degradation of misfolded glycoproteins since inhibition of this mannose trimming stabilizes misfolded glycoproteins in the ER [45]. Overexpression of ERMan1 accelerates the degradation of N-glycosylated proteins [39, 46]. The resulting Man8-GlcNAc2 containing glycoprotein after this trimming becomes a substrate for EDEM1 (ER-degradation enhancing mannosidase-like protein 1, Htm1p in yeast)—a mannosidase-related lectin in the ER. It was further proposed that misfolded glycoproteins interact with ERManI and EDEM1 for their ERAD, and lectin-carbohydrate interaction found to be crucial for EDEM substrate recognition [47]. Although ERMan1 was suggested to be a biological timer initiating the ERAD of misfolded proteins [48], recent studies revealed that mannosidases are not solely responsible for intensive demannosylation during ERAD, especially under non-basal conditions. Under ER stress (unfolded protein response active) conditions, the transcriptional elevation of EDEM1 enhances the ERAD efficiency by suppressing proteolytic downregulation of ERMan1 [49]. It appeared that EDEMs also play an important role in demannosylation of substrates [50]. EDEM1 also prevents reglycosylation and promotes retrotranslocation and degradation of some ERAD substrates [51]. On the other hand, while mannosidase homology domain (MHD) of Htm1p is necessary for substrate binding, mammalian EDEM1 binds misfolded proteins independent of MHD domain, and therefore, EDEM1 substrate binding may not require mannose trimming or even glycosylation [52]. Thus, in addition to N-linked oligosaccharide moieties of glycoproteins, EDEM1 can recognize the folding lesions of misfolded proteins. In summary, EDEMs are directly or indirectly involved in demannosylation of glycoproteins and/or serve as receptors that bind and target mannose-trimmed proteins for ERAD (Figure 1).

Truncation of terminal mannose from branch C exposes α terminal α1,6-bonded mannose residues functioning as a recognition signal for ERAD lectins such as OS9 (Yos9 in yeast) and XTP3-B (Figure 2). Through their mannose-6-phosphate receptor homology (MRH) domain, both proteins primarily recognize α1,6-linked mannose j. Additionally, OS-9 also recognizes α1,6-linked mannose e and c [53].

Several reports suggest that factors (EDEMs, OS9 and XTP3-B) required for substrate recognition and targeting reside within supramolecular complexes and/or interact with important ERAD regulators [54]. For example, EDEM1 interacts with CNX, receives substrates from CNX cycle and facilitates ERAD substrate

Figure 1. Protein quality control and targeting misfolding proteins to the ERAD.

substrates [64]. HERP interacts with Derlin1, and the partially oxidized Ig-K-LC is transferred from BiP to the HERP-Derlin1-Hrd1 complex and subsequently directed to proteasomal degradation [65]. Besides BiP, ERdj5 as disulfide reductase is also indicated to be important for ERAD of non-glycosylated proteins [63]. The nonglycosylated substrates captured by BiP are transferred to ERdj5 for the cleavage of disulfide bonds. Then, these substrates are transferred to SEL1L by the help of BiP for retrotranslocation [63]. Besides BiP, both OS9 and XTP3-B have been implicated

Ubiquitin is a 76 amino acid polypeptide encoded on multiple genes. It is ubiquitously expressed in all eukaryotic cells and highly conserved from yeast to human. Ubiquitin can be covalently conjugated to other proteins as monomers or as chains through a complex, highly regulated process called ubiquitination. Although there are reports for evidence of Ser- and Thr-linked ubiquitination, ubiquitin chain is generally attached on the Lys residue on misfolded protein. Lys-6, -11, -27, -29, 33,


the fate and function of the proteins. The most well-established role of

localization, stability and activity of target proteins [9].

ubiquitination (mono/poly) and the linkages of ubiquitin chains affect the fate,

Ubiquitination has a regulatory role in almost all cellular processes by altering

ubiquitination is targeting proteins for degradation by the 26S proteasome, and the most efficient way of targeting proteins to the proteasome is by tagging them with chains of ubiquitin [66]. This targeting requires modification of proteins with chains of four or more ubiquitins attached through lysine 48 (K48) and the specific recognition of these chains by the 19S cap of the 26S proteasome [67]. Mainly Lys-48 but rarely Lys-11-based polyubiquitin chains are reported to bind onto ERAD

Ubiquitination regulates several critical cellular functions, often by mediating the selective degradation of important regulatory proteins. Antigen presentation, inflammatory response induction and cell cycle progression are few examples. As expected, malfunctioning of ubiquitin-dependent proteolysis has implications for cancer and several inherited diseases, such as Angelman syndrome, Parkinson's

The role of ubiquitination, however, is not limited to proteasomal targeting. The

type of residue that the chain is built is critical for the fate of the ubiquitinated protein. Monoubiquitination has effects in protein trafficking, including endocytosis and lysosomal targeting. Polyubiquitin chains conjugated through K48 or other lysines (often K63) also have effect on proteasome-independent mechanisms, such as DNA repair, regulation of transcription factor activity and protein kinase

Ubiquitination is a multi-enzyme process. Three enzymes are involved: E1 ubiquitin activating enzyme, E2-ubiquitin conjugating enzyme and E3-ubiquitin ligase. During ubiquitination, E1 forms a thiol-ester bond between its active cysteine and C-terminal glycine of ubiquitin in an adenosine triphosphate (ATP) dependent manner. Ubiquitin on E1 is now activated and transferred to the active cysteine of E2 by a trans-thiolation reaction. E3 binds both to E2 and substrate and facilitates the formation of an isopeptide linkage between C-terminal glycine of ubiquitin and an internal lysine residue on substrate. Ubiquitin modification is

Today only 2 E1 enzymes and 35 E2 enzymes have been identified in mammals, but there are approximately 100 E3 in yeast and at least 600 in humans [71, 72]. E3s

dynamic and could be removed by deubiquitination enzymes (DUBs).

in the ERAD of non-glycosylated proteins [12].

Endoplasmic Reticulum-Associated Degradation (ERAD)

DOI: http://dx.doi.org/10.5772/intechopen.82043

2.2 Ubiquitination

substrates [68].

activation [70].

33

disease and Alzheimer's disease [69].

Figure 2. Schematic representation of ERAD using the Hrd1 complex as model.

degradation such as NHK-α1-antitrypsin mutant [55–57]. EDEM1 also associates with the components of ER retrotranslocation machinery. It is suggested that EDEM1 binds misfolded proteins and uses its MHD domain to target aberrant proteins to the ER-resident glycoprotein SEL1L protein of the Hrd1-SEL1L ubiquitin ligase complex [58]. SEL1L scaffolds several luminal substrate recognition factors and links them to Hrd1. OS9 and XTP3-B also associate with Hrd1-SEL1L complex, which also includes BiP and GRP94 [59, 60]. Furthermore, XTP3-B is proposed to link BiP with Hrd1 complex [60]. According to a hypothesis, these three chaperones (EDEM1, OS9 and XTP3-B) function as oligomers, where one monomer interacts with substrate and another with Hrd1-SEL1L complex [61]. Additionally, EDEM1 also interacts with Derlins, a transmembrane protein, which is a candidate for translocon [62]; furthermore, Derlin2 is shown to enhance the interaction of EDEM1 with a cytosolic AAA-ATPase p97, which couples ATP hydrolysis to the retrotranslocation of misfolded proteins [50].

It is clear that substrate recognition step of ERAD is a complicated mechanism, in which several different enzymes and chaperones having distinct but concerted roles in the ERAD are involved. Moreover, depending on substrates, the number and features of involved proteins vary. For example, concerted roles of EDEM, ERdj5 and BiP in the degradation of misfolded proteins have been suggested [63]. After exiting CNX-CLR cycle, EDEM1 further trims the Man8-GlcNAc2 glycan structure and ERdj5 reduces disulfate bonds. Concomitantly, ERdj5 activates BiP's ATPase activity. ADP-bound BiP binds to the misfolded protein and holds it in a retrotranslocation component form until it transfers to the retrotranslocation complex [63].

ERAD is also involved in the quality control of non-glycosylated proteins, which is independent of lectin-like proteins. Immunoglobulin light chain (Ig-K-LC), a non-glycosylated ERAD substrate, is degraded in a BiP-dependent manner. Okuda-Shimizu and Hendershot have characterized an ERAD pathway for this nonglycosylated BiP substrate [64] and different protein interaction dynamics seen to play a role in this process. Ig-K-LC has two intramolecular disulfide bonds, and its fully oxidized form does not have ability to pass from the ER to the cytoplasm. BiP interacts with only partially oxidized form of the Ig, preventing the full oxidation of Ig-K-LC and thereby facilitating its release from the ER [64]. Furthermore, a transmembrane UBL domain-containing protein, homoCys-responsive ER-resident protein (HERP), has been implicated as a receptor for non-glycosylated BiP

substrates [64]. HERP interacts with Derlin1, and the partially oxidized Ig-K-LC is transferred from BiP to the HERP-Derlin1-Hrd1 complex and subsequently directed to proteasomal degradation [65]. Besides BiP, ERdj5 as disulfide reductase is also indicated to be important for ERAD of non-glycosylated proteins [63]. The nonglycosylated substrates captured by BiP are transferred to ERdj5 for the cleavage of disulfide bonds. Then, these substrates are transferred to SEL1L by the help of BiP for retrotranslocation [63]. Besides BiP, both OS9 and XTP3-B have been implicated in the ERAD of non-glycosylated proteins [12].

#### 2.2 Ubiquitination

degradation such as NHK-α1-antitrypsin mutant [55–57]. EDEM1 also associates with the components of ER retrotranslocation machinery. It is suggested that EDEM1 binds misfolded proteins and uses its MHD domain to target aberrant proteins to the ER-resident glycoprotein SEL1L protein of the Hrd1-SEL1L ubiquitin ligase complex [58]. SEL1L scaffolds several luminal substrate recognition factors and links them to Hrd1. OS9 and XTP3-B also associate with Hrd1-SEL1L complex, which also includes BiP and GRP94 [59, 60]. Furthermore, XTP3-B is proposed to link BiP with Hrd1 complex [60]. According to a hypothesis, these three chaperones (EDEM1, OS9 and XTP3-B) function as oligomers, where one monomer interacts with substrate and another with Hrd1-SEL1L complex [61]. Additionally, EDEM1 also interacts with Derlins, a transmembrane protein, which is a candidate for translocon [62]; furthermore, Derlin2 is shown to enhance the interaction of EDEM1 with a cytosolic AAA-ATPase p97, which couples ATP hydrolysis to the

It is clear that substrate recognition step of ERAD is a complicated mechanism, in which several different enzymes and chaperones having distinct but concerted roles in the ERAD are involved. Moreover, depending on substrates, the number and features of involved proteins vary. For example, concerted roles of EDEM, ERdj5 and BiP in the degradation of misfolded proteins have been suggested [63]. After exiting CNX-CLR cycle, EDEM1 further trims the Man8-GlcNAc2 glycan structure and ERdj5 reduces disulfate bonds. Concomitantly, ERdj5 activates BiP's ATPase activity. ADP-bound BiP binds to the misfolded protein and holds it in a retrotranslocation component form until it transfers to the retrotranslocation com-

ERAD is also involved in the quality control of non-glycosylated proteins, which is independent of lectin-like proteins. Immunoglobulin light chain (Ig-K-LC), a non-glycosylated ERAD substrate, is degraded in a BiP-dependent manner. Okuda-Shimizu and Hendershot have characterized an ERAD pathway for this nonglycosylated BiP substrate [64] and different protein interaction dynamics seen to play a role in this process. Ig-K-LC has two intramolecular disulfide bonds, and its fully oxidized form does not have ability to pass from the ER to the cytoplasm. BiP interacts with only partially oxidized form of the Ig, preventing the full oxidation of Ig-K-LC and thereby facilitating its release from the ER [64]. Furthermore, a transmembrane UBL domain-containing protein, homoCys-responsive ER-resident protein (HERP), has been implicated as a receptor for non-glycosylated BiP

retrotranslocation of misfolded proteins [50].

Schematic representation of ERAD using the Hrd1 complex as model.

plex [63].

32

Figure 2.

Endoplasmic Reticulum

Ubiquitin is a 76 amino acid polypeptide encoded on multiple genes. It is ubiquitously expressed in all eukaryotic cells and highly conserved from yeast to human. Ubiquitin can be covalently conjugated to other proteins as monomers or as chains through a complex, highly regulated process called ubiquitination. Although there are reports for evidence of Ser- and Thr-linked ubiquitination, ubiquitin chain is generally attached on the Lys residue on misfolded protein. Lys-6, -11, -27, -29, 33, -48 and -63 are the residues used for ubiquitin linkage. Both the type of ubiquitination (mono/poly) and the linkages of ubiquitin chains affect the fate, localization, stability and activity of target proteins [9].

Ubiquitination has a regulatory role in almost all cellular processes by altering the fate and function of the proteins. The most well-established role of ubiquitination is targeting proteins for degradation by the 26S proteasome, and the most efficient way of targeting proteins to the proteasome is by tagging them with chains of ubiquitin [66]. This targeting requires modification of proteins with chains of four or more ubiquitins attached through lysine 48 (K48) and the specific recognition of these chains by the 19S cap of the 26S proteasome [67]. Mainly Lys-48 but rarely Lys-11-based polyubiquitin chains are reported to bind onto ERAD substrates [68].

Ubiquitination regulates several critical cellular functions, often by mediating the selective degradation of important regulatory proteins. Antigen presentation, inflammatory response induction and cell cycle progression are few examples. As expected, malfunctioning of ubiquitin-dependent proteolysis has implications for cancer and several inherited diseases, such as Angelman syndrome, Parkinson's disease and Alzheimer's disease [69].

The role of ubiquitination, however, is not limited to proteasomal targeting. The type of residue that the chain is built is critical for the fate of the ubiquitinated protein. Monoubiquitination has effects in protein trafficking, including endocytosis and lysosomal targeting. Polyubiquitin chains conjugated through K48 or other lysines (often K63) also have effect on proteasome-independent mechanisms, such as DNA repair, regulation of transcription factor activity and protein kinase activation [70].

Ubiquitination is a multi-enzyme process. Three enzymes are involved: E1 ubiquitin activating enzyme, E2-ubiquitin conjugating enzyme and E3-ubiquitin ligase. During ubiquitination, E1 forms a thiol-ester bond between its active cysteine and C-terminal glycine of ubiquitin in an adenosine triphosphate (ATP) dependent manner. Ubiquitin on E1 is now activated and transferred to the active cysteine of E2 by a trans-thiolation reaction. E3 binds both to E2 and substrate and facilitates the formation of an isopeptide linkage between C-terminal glycine of ubiquitin and an internal lysine residue on substrate. Ubiquitin modification is dynamic and could be removed by deubiquitination enzymes (DUBs).

Today only 2 E1 enzymes and 35 E2 enzymes have been identified in mammals, but there are approximately 100 E3 in yeast and at least 600 in humans [71, 72]. E3s catalyzing the transfer of active ubiquitin moieties on the substrate are responsible for substrate specificity. There are two large families of E3s: (1) HECT [homologous to E6-associated protein (E6AP) C-terminus] domain E3s and (2) RING [really interesting new gene] domain E3s. HECT domain E3s share a 350-residue region harboring a strictly conserved cysteine residue that forms an essential thiol-ester intermediate during catalysis. That is why ubiquitin is transferred to the active-site cysteine of the HECT domain followed by transfer to substrate or to a substratebound multi-ubiquitin chain. The RING finger defines the largest family of E3s. RING fingers range from 40 to 100 amino acids and are defined by eight conserved cysteine and histidine residues that coordinate two zinc ions stabilizing a characteristic cross-braced conformation. For RING E3s, current evidence indicates that ubiquitin is transferred directly from E2 to substrate [69, 70].

variety of different partners allowing its communication with proteins on both sites of ER membrane. gp78 uses a VIM (VCP-interacting motif) segment to bind p97/ VCP [77] and CUE domain recruiting a multiprotein complex composed of Bag6

After initial E3-mediated ubiquitin attachment, ubiquitin chain extension ("polyubiquitination") occurs by the covalent modification of additional ubiquitin monomers onto a Lys residue in a previously linked ubiquitin. This forms an extended isopeptide-linked polyubiquitin chain. In some selected cases, the cooperative extension of a polyubiquitin chain is by the E4s, ubiquitin chain extension

The ERAD substrates must be retrotranslocated to the cytosol for proteasomal degradation and the cytoplasmic AAA+ ATPase p97 (VCP or Cdc48p in yeast) is the main retrotranslocation protein providing the mechanical force required for removal of proteins from the ER. It is an essential protein having many roles in diverse biological processes, such as endoplasmic reticulum-associated degradation (ERAD), homotypic membrane fusion, transcriptional control, cell cycle regulation, autophagy, endosomal sorting and regulating protein degradation at the outer

p97/VCP has a multidomain structure including N domain, D1 weak ATPase, D2 major ATPase and C domain [86–88]. p97/VCP functions as a homohexamer and D1 domain is responsible for oligomerization independent of nucleotide binding. The change in the conformation of hexameric ring by ATP hydrolysis is persistent with

The diversity in cellular functions of p97/VCP is dictated by the variety of its partner proteins that interact with its N domain. p97/VCP associates with several E3s like Hrd1 and gp78, DUBs like ataxin3 and YOD1 and ERAD accessory factors such as UbxD2 and VIMP. Moreover, many p97/VCP interacting proteins (Ufd1-Npl4 dimer, gp78 etc.) bind directly to ubiquitin. p97/VCP functions as a segregase using the energy from ATP hydrolysis to segregate ubiquitinated proteins from large immobile complexes of ER to the cytosol. This cytosolic protein is recruited to the ER membrane through its interaction with membrane-embedded ERAD components. There are at least seven different ERAD members that could interact with p97/VCP via certain motifs such as VIM motif (gp78 and SVIP), UBX domains (UBXD2 and UBXD8), SHP boxes (Derlin1 and Derlin2) and uncharacterized cytosolic regions of

Hrd1 and VIMP that found to have p97/VCP-binding motif [12, 42, 90].

Retrotranslocation is tightly coupled with both ubiquitination and proteasomal degradation. In most cases, inhibiting ubiquitination prevents both degradation and retrotranslocation. The interaction of p97/VCP/CDC48p with its cofactor Ufd1- Npl4 dimer enhances its affinity to ubiquitin (Figure 2). However, it has been also suggested that Hrd1-mediated ERAD requires well-established retrotranslocation machinery, the p97/VCP–Ufd1–Npl4 complex, whereas the gp78 pathway needs

Many deubiquitinating enzymes (DUBs) in mammalian cells, including Ataxin3, USP13, USP25 and YOD1, are also implicated in the ERAD through physical interaction with ERAD core machinery [72, 91, 92]. Several studies revealed that p97/ VCP interacts with DUBs. However, the function of DUBs in the ERAD is still not fully characterized. Otu1p (yeast homolog of YOD1) binds to CDC48p and trims the polyubiquitin chain, resulting oligoubiquitin chains with up to 10 ubiquitin molecules. It has been further suggested that releasing substrates from CDC48p requires DUBs [93]. Consistently, catalytically inactive YOD1 inhibits retrotranslocation of

2.3 Retrotranslocation and shuttling substrates to the proteasome

and its cofactors [79].

enzymes, that facilitate ERAD [80–82].

Endoplasmic Reticulum-Associated Degradation (ERAD)

DOI: http://dx.doi.org/10.5772/intechopen.82043

mitochondrial membrane [83–85].

only p97/VCP and Npl4 [75].

35

its function in retrotranslocation [88, 89].

Ubiquitination step marks ERAD substrates for proteasomal degradation. In yeast, Doa10p and Hrd1p ligases are mainly responsible for ubiquitination of ERAD substrates, but additional E3s shown to contribute to the ERAD under special circumstances [9]. Depending on the topology of misfolded lesion, factors required for ERAD vary. In yeast, three ERAD pathways have been proposed. ERAD-C, ERAD-L and ERAD-M target proteins with lesions in the cytoplasmic, luminal and membrane domains, respectively [44, 73, 74]. ERAD-L substrates use the Hrd1p ubiquitin ligase complex containing Hrd1p, Hrd3p, Usa1p, Der1p, and Yos9p, whereas ERAD-M substrates use Hrd1p and Hrd3p, only in some cases Usa1p [68]. Hrd3p is specifically important for structural integrity of Hrd1p complex. Hrd3p stabilizes Hrd1p, and when it is absent, Hrd1p is auto-ubiquitinated and rapidly degraded. Hrd3p and its mammalian homolog SEL1L also function as an adaptor bridging substrate recognition, ubiquitination and retrotranslocation in Hrd1 mediated ERAD. On the other hand, ERAD-C substrates interact with the Doa10p ubiquitin ligase complex. These three pathways have been identified only in yeast and mammalian has more complicated machinery. Even in yeast, some membrane proteins require both Doa10p and Hrd1p E3s; thus, these pathways could overlap [42].

Although Hrd1p and Doa10p are conserved evolutionary (mammalian homologs: Hrd1 and TEB4, respectively), the number of ERAD E3s in mammals is highly expanded. Besides Hrd1 and TEB4, gp78, RNF5/RMA1, RNF170, RNF185, Trc8, RNF103, RFP2, Fbx2, Fbx6, Parkin, CHIP and UBE4a are other characterized ERAD E3s [9, 27]. Hrd1 and gp78, both homologues to yeast Hrd1p, are the most studied ERAD E3 indicated for degradation of several substrates, some of which are associated with the quality of disease-related proteins. HMG-CoA reductase, apolipoprotein B, cytochrome P450 CYP3A4, CFTRΔF508, z-variant antitrypsin, CD3δ and KAI1 are shown to be degraded via gp78-mediated ERAD, whereas studies have been suggested that Hrd1 is important for the ERAD of GABAb receptor, Nrf2, Pael-receptor mutant tyrosinase, z-variant antitrypsin and gp78 [22, 75–78]. Only a couple of substrates are known for other E3 ligases. It is also interesting that multiple E3s often function in the degradation on same substrate either in parallel or in tandem.

As Hrd1p in yeast, Hrd1 in mammals functions in a multi-protein complex. While it is complex with EDEM1, Derlins, OS9, XTP-3B and SEL1-L have been linked with degradation of glycosylated substrates (Figure 2), and another Hrd1 complex utilizing BiP, HERP and Derlin1 functions in the degradation of nonglycosylated substrates. Other ERAD factors have also been shown to interact with Hrd1 including UBXD2 and UBXD8 that interact with p97/VCP and recently identified chaperones such as ubiquilin and BAG6. Similarly, gp78, the second major mammalian ERAD E3 enzyme, functions in multiprotein complex in conjunction with E2 enzyme UBE2G2. Besides its diversity on substrate specificity, gp78 also has Endoplasmic Reticulum-Associated Degradation (ERAD) DOI: http://dx.doi.org/10.5772/intechopen.82043

catalyzing the transfer of active ubiquitin moieties on the substrate are responsible for substrate specificity. There are two large families of E3s: (1) HECT [homologous to E6-associated protein (E6AP) C-terminus] domain E3s and (2) RING [really interesting new gene] domain E3s. HECT domain E3s share a 350-residue region harboring a strictly conserved cysteine residue that forms an essential thiol-ester intermediate during catalysis. That is why ubiquitin is transferred to the active-site cysteine of the HECT domain followed by transfer to substrate or to a substratebound multi-ubiquitin chain. The RING finger defines the largest family of E3s. RING fingers range from 40 to 100 amino acids and are defined by eight conserved cysteine and histidine residues that coordinate two zinc ions stabilizing a characteristic cross-braced conformation. For RING E3s, current evidence indicates that

Ubiquitination step marks ERAD substrates for proteasomal degradation. In yeast, Doa10p and Hrd1p ligases are mainly responsible for ubiquitination of ERAD substrates, but additional E3s shown to contribute to the ERAD under special circumstances [9]. Depending on the topology of misfolded lesion, factors required for ERAD vary. In yeast, three ERAD pathways have been proposed. ERAD-C, ERAD-L and ERAD-M target proteins with lesions in the cytoplasmic, luminal and membrane domains, respectively [44, 73, 74]. ERAD-L substrates use the Hrd1p ubiquitin ligase complex containing Hrd1p, Hrd3p, Usa1p, Der1p, and Yos9p, whereas ERAD-M substrates use Hrd1p and Hrd3p, only in some cases Usa1p [68]. Hrd3p is specifically important for structural integrity of Hrd1p complex. Hrd3p stabilizes Hrd1p, and when it is absent, Hrd1p is auto-ubiquitinated and rapidly degraded. Hrd3p and its mammalian homolog SEL1L also function as an adaptor bridging substrate recognition, ubiquitination and retrotranslocation in Hrd1 mediated ERAD. On the other hand, ERAD-C substrates interact with the Doa10p ubiquitin ligase complex. These three pathways have been identified only in yeast

ubiquitin is transferred directly from E2 to substrate [69, 70].

and mammalian has more complicated machinery. Even in yeast, some

could overlap [42].

Endoplasmic Reticulum

in tandem.

34

membrane proteins require both Doa10p and Hrd1p E3s; thus, these pathways

Although Hrd1p and Doa10p are conserved evolutionary (mammalian homologs: Hrd1 and TEB4, respectively), the number of ERAD E3s in mammals is highly expanded. Besides Hrd1 and TEB4, gp78, RNF5/RMA1, RNF170, RNF185, Trc8, RNF103, RFP2, Fbx2, Fbx6, Parkin, CHIP and UBE4a are other characterized ERAD E3s [9, 27]. Hrd1 and gp78, both homologues to yeast Hrd1p, are the most studied ERAD E3 indicated for degradation of several substrates, some of which are associated with the quality of disease-related proteins. HMG-CoA reductase, apolipoprotein B, cytochrome P450 CYP3A4, CFTRΔF508, z-variant antitrypsin, CD3δ and KAI1 are shown to be degraded via gp78-mediated ERAD, whereas studies have been suggested that Hrd1 is important for the ERAD of GABAb receptor, Nrf2, Pael-receptor mutant tyrosinase, z-variant antitrypsin and gp78 [22, 75–78]. Only a couple of substrates are known for other E3 ligases. It is also interesting that multiple E3s often function in the degradation on same substrate either in parallel or

As Hrd1p in yeast, Hrd1 in mammals functions in a multi-protein complex. While it is complex with EDEM1, Derlins, OS9, XTP-3B and SEL1-L have been linked with degradation of glycosylated substrates (Figure 2), and another Hrd1 complex utilizing BiP, HERP and Derlin1 functions in the degradation of nonglycosylated substrates. Other ERAD factors have also been shown to interact with Hrd1 including UBXD2 and UBXD8 that interact with p97/VCP and recently identified chaperones such as ubiquilin and BAG6. Similarly, gp78, the second major mammalian ERAD E3 enzyme, functions in multiprotein complex in conjunction with E2 enzyme UBE2G2. Besides its diversity on substrate specificity, gp78 also has variety of different partners allowing its communication with proteins on both sites of ER membrane. gp78 uses a VIM (VCP-interacting motif) segment to bind p97/ VCP [77] and CUE domain recruiting a multiprotein complex composed of Bag6 and its cofactors [79].

After initial E3-mediated ubiquitin attachment, ubiquitin chain extension ("polyubiquitination") occurs by the covalent modification of additional ubiquitin monomers onto a Lys residue in a previously linked ubiquitin. This forms an extended isopeptide-linked polyubiquitin chain. In some selected cases, the cooperative extension of a polyubiquitin chain is by the E4s, ubiquitin chain extension enzymes, that facilitate ERAD [80–82].

#### 2.3 Retrotranslocation and shuttling substrates to the proteasome

The ERAD substrates must be retrotranslocated to the cytosol for proteasomal degradation and the cytoplasmic AAA+ ATPase p97 (VCP or Cdc48p in yeast) is the main retrotranslocation protein providing the mechanical force required for removal of proteins from the ER. It is an essential protein having many roles in diverse biological processes, such as endoplasmic reticulum-associated degradation (ERAD), homotypic membrane fusion, transcriptional control, cell cycle regulation, autophagy, endosomal sorting and regulating protein degradation at the outer mitochondrial membrane [83–85].

p97/VCP has a multidomain structure including N domain, D1 weak ATPase, D2 major ATPase and C domain [86–88]. p97/VCP functions as a homohexamer and D1 domain is responsible for oligomerization independent of nucleotide binding. The change in the conformation of hexameric ring by ATP hydrolysis is persistent with its function in retrotranslocation [88, 89].

The diversity in cellular functions of p97/VCP is dictated by the variety of its partner proteins that interact with its N domain. p97/VCP associates with several E3s like Hrd1 and gp78, DUBs like ataxin3 and YOD1 and ERAD accessory factors such as UbxD2 and VIMP. Moreover, many p97/VCP interacting proteins (Ufd1-Npl4 dimer, gp78 etc.) bind directly to ubiquitin. p97/VCP functions as a segregase using the energy from ATP hydrolysis to segregate ubiquitinated proteins from large immobile complexes of ER to the cytosol. This cytosolic protein is recruited to the ER membrane through its interaction with membrane-embedded ERAD components. There are at least seven different ERAD members that could interact with p97/VCP via certain motifs such as VIM motif (gp78 and SVIP), UBX domains (UBXD2 and UBXD8), SHP boxes (Derlin1 and Derlin2) and uncharacterized cytosolic regions of Hrd1 and VIMP that found to have p97/VCP-binding motif [12, 42, 90].

Retrotranslocation is tightly coupled with both ubiquitination and proteasomal degradation. In most cases, inhibiting ubiquitination prevents both degradation and retrotranslocation. The interaction of p97/VCP/CDC48p with its cofactor Ufd1- Npl4 dimer enhances its affinity to ubiquitin (Figure 2). However, it has been also suggested that Hrd1-mediated ERAD requires well-established retrotranslocation machinery, the p97/VCP–Ufd1–Npl4 complex, whereas the gp78 pathway needs only p97/VCP and Npl4 [75].

Many deubiquitinating enzymes (DUBs) in mammalian cells, including Ataxin3, USP13, USP25 and YOD1, are also implicated in the ERAD through physical interaction with ERAD core machinery [72, 91, 92]. Several studies revealed that p97/ VCP interacts with DUBs. However, the function of DUBs in the ERAD is still not fully characterized. Otu1p (yeast homolog of YOD1) binds to CDC48p and trims the polyubiquitin chain, resulting oligoubiquitin chains with up to 10 ubiquitin molecules. It has been further suggested that releasing substrates from CDC48p requires DUBs [93]. Consistently, catalytically inactive YOD1 inhibits retrotranslocation of

ERAD substrates [91]. In conclusion, many p97-associated DUBs serve as positive regulators of ERAD.

or misfolding proteins in the ER lumen triggers "ER stress" by decreasing free chaperone levels [105]. In response to this cellular stress, the pathway known as the "Unfolded Protein Response (UPR)" is activated and results in specific cellular functions classified as adaptation, alarm and apoptosis [4]. Three transmembrane proteins with luminal domains that sense the changes in the ER environment function as UPR sensor proteins are inositol requiring enzyme-1 (IRE1), activating transcription factor 6 (ATF6) and protein kinase RNA-like endoplasmic reticulum kinase (PERK). PERK is a serine/threonine kinase, and IRE1 possesses both kinase and endoribonuclease domains [27, 50]. These sensors initiate signal transduction by sensing the presence of unfolded proteins in the ER lumen and thus control the UPR pathway [15, 18, 106]. All these transmembrane proteins interact with BIP under basal conditions. However, when unfolded proteins are present, BIP dissociates from the UPR sensor proteins. After dissociation of BIP, PERK and IRE1 dimerize and become activated by auto-phosphorylation, whereas ATF6 become translocated to the Golgi and proteolytically cleaved [27, 50]. Activated PERK phosphorylates translation factor eIF2α attenuating protein synthesis to limit protein load. IRE1 activates XBP-1 that enhances transcription of ERAD factors [27, 50]. On the other pathway, ATF6 upregulates many genes that encode ERresident chaperones and folding assistants like BIP, CNX, CLR and PDI. To summarize, with the induction of UPR in the cell, the overall translation is inhibited for several hours primarily to slow down the entry of newly synthesized proteins to the ER, the amount of chaperones and ER protein folding capacity is increased for proper folding of accumulated unfolded proteins, and thus, the normal ER function and homeostasis are protected [4, 107]. UPR also enhances ERAD capacity by upregulating some of the ERAD genes to ensure that defective proteins are

Endoplasmic Reticulum-Associated Degradation (ERAD)

DOI: http://dx.doi.org/10.5772/intechopen.82043

degraded when the folding attempts fail [21–23]. EDEM proteins, Hrd1, SVIP, OS9 and gp78, are only some of the targets of the ER stress-induced Ire1/Xbp1 pathway [62, 108–111]. If the cellular stress is consistently increasing, UPR induces cell death

It has been suggested that large or prolonged variations such as change in Ca2+ or

redox homeostasis, exposure to pathogens and large-scale accumulation of misfolded proteins may induce UPR to adapt ERAD activity. However, smaller or

more transient fluctuations on ER load may be overcome rapidly by posttranslational pathways that control stability, localization and assembly of ERAD components [23]. For example, reversible ADP ribosylation adapts BIP response for short-term fluctuations [113]. Reversible palmitoylation changes the sub-organelle distribution of CNX [114, 115]. Moreover, many ERAD factors/enhancers, including EDEM1, ER Man1, HERP, OS9, SEL1L and gp78, have fast turnover. This is important since when protein misfolding crisis is over, ERAD activity should rapidly turn back to the basal levels. Many ERAD factors then rapidly degraded via a process called ERAD tuning [23]. ERAD tuning does not require signal transduction from the ER to the nucleus [23]. Hrd1 was suggested to be a central regulator of ERAD tuning. It has been shown that Hrd1 ubiquitinates gp78 E3 enzyme and enhances its degradation, which in turn causes inhibition of gp78-mediated ERAD. Very recently, Hrd1 was also found to regulate the stability of OS9 [116]. Hrd1 also undergoes auto-ubiquitination to induce its own proteasomal degradation [117]. Another homeostatic control mechanism, in which ERAD activity itself is regulated post-translationally and independent of UPR, is degradation of EDEM1, OS9 and SEL1L by the E2 enzyme UBC6e, a component of Hrd1 supramolecular

Another type of ERAD regulation occurs via substrate-specific adaptor, as reported for HMGR. The adaptor proteins, Insig1 or Insig2, bind to HMGR only in the presence of 24,25-dihydrolanosterol, an intermediate molecule in sterol

mechanisms such as apoptosis or autophagy [4, 14, 112].

complex [118].

37

Several putative retrotranslocation channels have been proposed such as the Sec61 complex, members of Derlin family and polytopic E3s such as Hrd1 and gp78. Sec61 is one of the proposed channel protein mutants, which prevented degradation of some ERAD substrates in yeast [94, 95]. Cholera toxin also translocates from ER by utilizing Sec61 [96]. On the other hand, retrotranslocation of some other ERAD substrates has been suggested to depend on Derlins [97, 98], a family of polytopic transmembrane ER proteins linked to some ERAD substrates. Moreover, Derlin1 recruits p97/VCP [99], a key protein of retrotranslocation, which provides energy for the process. Derlin1 also interacts with some E3s like Hrd1, gp78 and RNF5 forming large complexes on the ER membrane [9]. Recently, Hrd1 ubiquitin ligase has been suggested to be the top candidate for retrotranslocation channel [9]. Autoubiquitination of Hrd1p in its RING finger domain triggers conformational change allowing the misfolded luminal domain of a substrate to move across the membrane. Thus, it was suggested that Hrd1 forms an ubiquitin-gated proteinconducting channel [33]. It has also been suggested that proteins might exit the ER via the formation of lipid droplets or lipid droplets serve as an intermediate step for substrates en route to the proteasome [100]. However, studies in yeast suggested that lipid droplet formation is dispensable for ERAD-L and ERAD-M [101].

Once retrotranslocated from ER to the cytosol, ERAD substrates should be rapidly targeted to the proteasome for degradation in order to avoid accumulation of aggregates in the cytosol. Consistently, proteasomal inhibition also stabilizes ERAD substrates in the ER lumen. For the coupling of retrotranslocation with degradation, ubiquitinated substrates must be recognized by cytosolic proteins functioning as ubiquitin receptors. Ubiquitin-binding domain containing proteins has ability to shuttle ubiquitinated proteins from retrotranslocation complex at the ER membrane to the proteasome since these proteins interact both with proteasome and p97/VCP. Indeed, it has been suggested that p97/VCP bridges the ER to the proteasome by forming a complex with mHR23B (homolog of yeast Rad23p)- PNGase [102] (Figure 2). In yeast, the substrates are probably transferred from CDC48p to the proteasome indirectly via ubiquitin- and proteasome-binding domains containing shuttling factors Rad23p and Dsk2p [103, 104]. Recently, Bag6/ Bat3/Scythe has been characterized as a novel chaperone system with regulatory functions in protein degradation [79]. The chaperone holdase activity of this system keeps some retrotranslocated substrates in a soluble state for proteasome degradation. Bag6, also a partner protein of gp78 E3 enzyme, interacts with proteasome, and proteins like ubiquilin that known as proteasome adaptor proteins suggesting Bag6 might act between p97/VCP and proteasome to hand substrates off from retrotranslocation machinery to the proteasome.

#### 3. Regulation of ERAD

Regulation of ERAD in normal and pathological conditions is also of great importance since hyper-ERAD may cause in loss-of-function phenotypes upon unnecessary degradation of folding intermediates as seen in CFTR and hypo-ERAD may result in gain-of-function phenotypes upon accumulation and/or aggregation of misfolded and unassembled proteins. Several studies suggested different regulation paths for ERAD activity via ubiquitin ligases and their dynamic ERAD complexes, UPR and endogenous ERAD inhibitors.

It is thought that ERAD functions at relatively low levels under basal conditions, but under proteotoxic stress its activity is enhanced. Accumulation of the unfolding

#### Endoplasmic Reticulum-Associated Degradation (ERAD) DOI: http://dx.doi.org/10.5772/intechopen.82043

ERAD substrates [91]. In conclusion, many p97-associated DUBs serve as positive

Several putative retrotranslocation channels have been proposed such as the Sec61 complex, members of Derlin family and polytopic E3s such as Hrd1 and gp78. Sec61 is one of the proposed channel protein mutants, which prevented degradation of some ERAD substrates in yeast [94, 95]. Cholera toxin also translocates from ER by utilizing Sec61 [96]. On the other hand, retrotranslocation of some other ERAD substrates has been suggested to depend on Derlins [97, 98], a family of polytopic transmembrane ER proteins linked to some ERAD substrates. Moreover, Derlin1 recruits p97/VCP [99], a key protein of retrotranslocation, which provides energy for the process. Derlin1 also interacts with some E3s like Hrd1, gp78 and RNF5 forming large complexes on the ER membrane [9]. Recently, Hrd1 ubiquitin ligase has been suggested to be the top candidate for retrotranslocation channel [9]. Autoubiquitination of Hrd1p in its RING finger domain triggers conformational change allowing the misfolded luminal domain of a substrate to move across the membrane. Thus, it was suggested that Hrd1 forms an ubiquitin-gated protein-

conducting channel [33]. It has also been suggested that proteins might exit the ER via the formation of lipid droplets or lipid droplets serve as an intermediate step for substrates en route to the proteasome [100]. However, studies in yeast suggested that lipid droplet formation is dispensable for ERAD-L and ERAD-M [101]. Once retrotranslocated from ER to the cytosol, ERAD substrates should be rapidly targeted to the proteasome for degradation in order to avoid accumulation of aggregates in the cytosol. Consistently, proteasomal inhibition also stabilizes ERAD substrates in the ER lumen. For the coupling of retrotranslocation with degradation, ubiquitinated substrates must be recognized by cytosolic proteins functioning as ubiquitin receptors. Ubiquitin-binding domain containing proteins has ability to shuttle ubiquitinated proteins from retrotranslocation complex at the ER membrane to the proteasome since these proteins interact both with proteasome and p97/VCP. Indeed, it has been suggested that p97/VCP bridges the ER to the proteasome by forming a complex with mHR23B (homolog of yeast Rad23p)- PNGase [102] (Figure 2). In yeast, the substrates are probably transferred from CDC48p to the proteasome indirectly via ubiquitin- and proteasome-binding domains containing shuttling factors Rad23p and Dsk2p [103, 104]. Recently, Bag6/ Bat3/Scythe has been characterized as a novel chaperone system with regulatory functions in protein degradation [79]. The chaperone holdase activity of this system keeps some retrotranslocated substrates in a soluble state for proteasome degradation. Bag6, also a partner protein of gp78 E3 enzyme, interacts with proteasome, and proteins like ubiquilin that known as proteasome adaptor proteins suggesting Bag6 might act between p97/VCP and proteasome to hand substrates off from

Regulation of ERAD in normal and pathological conditions is also of great importance since hyper-ERAD may cause in loss-of-function phenotypes upon unnecessary degradation of folding intermediates as seen in CFTR and hypo-ERAD may result in gain-of-function phenotypes upon accumulation and/or aggregation of misfolded and unassembled proteins. Several studies suggested different regulation paths for ERAD activity via ubiquitin ligases and their dynamic ERAD com-

It is thought that ERAD functions at relatively low levels under basal conditions, but under proteotoxic stress its activity is enhanced. Accumulation of the unfolding

retrotranslocation machinery to the proteasome.

plexes, UPR and endogenous ERAD inhibitors.

3. Regulation of ERAD

36

regulators of ERAD.

Endoplasmic Reticulum

or misfolding proteins in the ER lumen triggers "ER stress" by decreasing free chaperone levels [105]. In response to this cellular stress, the pathway known as the "Unfolded Protein Response (UPR)" is activated and results in specific cellular functions classified as adaptation, alarm and apoptosis [4]. Three transmembrane proteins with luminal domains that sense the changes in the ER environment function as UPR sensor proteins are inositol requiring enzyme-1 (IRE1), activating transcription factor 6 (ATF6) and protein kinase RNA-like endoplasmic reticulum kinase (PERK). PERK is a serine/threonine kinase, and IRE1 possesses both kinase and endoribonuclease domains [27, 50]. These sensors initiate signal transduction by sensing the presence of unfolded proteins in the ER lumen and thus control the UPR pathway [15, 18, 106]. All these transmembrane proteins interact with BIP under basal conditions. However, when unfolded proteins are present, BIP dissociates from the UPR sensor proteins. After dissociation of BIP, PERK and IRE1 dimerize and become activated by auto-phosphorylation, whereas ATF6 become translocated to the Golgi and proteolytically cleaved [27, 50]. Activated PERK phosphorylates translation factor eIF2α attenuating protein synthesis to limit protein load. IRE1 activates XBP-1 that enhances transcription of ERAD factors [27, 50]. On the other pathway, ATF6 upregulates many genes that encode ERresident chaperones and folding assistants like BIP, CNX, CLR and PDI. To summarize, with the induction of UPR in the cell, the overall translation is inhibited for several hours primarily to slow down the entry of newly synthesized proteins to the ER, the amount of chaperones and ER protein folding capacity is increased for proper folding of accumulated unfolded proteins, and thus, the normal ER function and homeostasis are protected [4, 107]. UPR also enhances ERAD capacity by upregulating some of the ERAD genes to ensure that defective proteins are degraded when the folding attempts fail [21–23]. EDEM proteins, Hrd1, SVIP, OS9 and gp78, are only some of the targets of the ER stress-induced Ire1/Xbp1 pathway [62, 108–111]. If the cellular stress is consistently increasing, UPR induces cell death mechanisms such as apoptosis or autophagy [4, 14, 112].

It has been suggested that large or prolonged variations such as change in Ca2+ or redox homeostasis, exposure to pathogens and large-scale accumulation of misfolded proteins may induce UPR to adapt ERAD activity. However, smaller or more transient fluctuations on ER load may be overcome rapidly by posttranslational pathways that control stability, localization and assembly of ERAD components [23]. For example, reversible ADP ribosylation adapts BIP response for short-term fluctuations [113]. Reversible palmitoylation changes the sub-organelle distribution of CNX [114, 115]. Moreover, many ERAD factors/enhancers, including EDEM1, ER Man1, HERP, OS9, SEL1L and gp78, have fast turnover. This is important since when protein misfolding crisis is over, ERAD activity should rapidly turn back to the basal levels. Many ERAD factors then rapidly degraded via a process called ERAD tuning [23]. ERAD tuning does not require signal transduction from the ER to the nucleus [23]. Hrd1 was suggested to be a central regulator of ERAD tuning. It has been shown that Hrd1 ubiquitinates gp78 E3 enzyme and enhances its degradation, which in turn causes inhibition of gp78-mediated ERAD. Very recently, Hrd1 was also found to regulate the stability of OS9 [116]. Hrd1 also undergoes auto-ubiquitination to induce its own proteasomal degradation [117]. Another homeostatic control mechanism, in which ERAD activity itself is regulated post-translationally and independent of UPR, is degradation of EDEM1, OS9 and SEL1L by the E2 enzyme UBC6e, a component of Hrd1 supramolecular complex [118].

Another type of ERAD regulation occurs via substrate-specific adaptor, as reported for HMGR. The adaptor proteins, Insig1 or Insig2, bind to HMGR only in the presence of 24,25-dihydrolanosterol, an intermediate molecule in sterol

biosynthesis. Under low sterol levels, HMGR is stable; however, when sterol levels are high, Insig-HMGR interaction become favored, leading delivery of HMGR to E3 complex following by its proteasomal degradation [119]. Likewise, ERAD-mediated degradations of apolipoprotein and IP3R are initiated when lipid levels are low and calcium levels are high, respectively [23].

References

[1] Alberts B, Johnson A, Lewis J, Morgan D, Raff R, Roberts M, et al. Membrane transport of small molecules

Endoplasmic Reticulum-Associated Degradation (ERAD)

DOI: http://dx.doi.org/10.5772/intechopen.82043

1800:172-180. DOI: 10.1016/j.

[9] Christianson JC, Ye Y. Cleaning up in the endoplasmic reticulum: Ubiquitin in charge. Nature Structural & Molecular Biology. 2014;21:325-335. DOI: 10.1038/

[10] Araki K, Nagata K. Protein folding and quality control in the ER. Cold Spring Harbor Perspectives in Biology.

[11] Romisch K. Endoplasmic reticulumassociated degradation. Annual Review of Cell and Developmental Biology. 2005;21:435-456. DOI: 10.1146/annurev.

2011;3:a007526. DOI: 10.1101/

cshperspect.a007526

cellbio.21.012704.133250

[12] Olzmann JA, Kopito RR, Christianson JC. The mammalian endoplasmic reticulum-associated degradation system. Cold Spring Harbor Perspectives in Medicine. 2013;5. DOI:

10.1101/cshperspect.a013185

newly synthesized proteins by proteasomes. Nature. 2000;404: 770-774. DOI: 10.1038/35008096

59:335-344. DOI: 10.1016/j.

[15] Nishikawa S, Brodsky JL, Nakatsukasa K. Roles of molecular chaperones in endoplasmic reticulum (ER) quality control and ER-associated degradation (ERAD). Journal of Biochemistry. 2005;137:551-555. DOI:

molcel.2015.06.010

10.1093/jb/mvi068

[13] Schubert U, Anton LC, Gibbs J, Norbury CC, Yewdell JW, Bennink JR. Rapid degradation of a large fraction of

[14] Morito D, Nagata K. Pathogenic hijacking of ER-associated degradation: Is ERAD flexible? Molecular Cell. 2015;

[16] Nyathi Y, Wilkinson BM, Pool MR.

Co-translational targeting and

bbagen.2009.07.029

nsmb.2793

and the electrical properties of membranes: The endoplasmic reticulum. In: Richte ML, editor. Molecular Biology of The Cell. 6th ed. Garland Science, Taylor&Francis

Group. 2015. pp. 669-691

s00018-015-2052-6

Annual Review of Cell and

Discovery. 2008;7:1013-1030

cellbio.21.012704.133214

embor.7400832

39

[2] Schwarz DS, Blower MD. The endoplasmic reticulum: Structure, function and response to cellular signaling. Cellular and Molecular Life Sciences. 2016;73:79-94. DOI: 10.1007/

[3] Lee MCS, Miller EA, Goldberg J, Orci L, Schekman R. Bi-directional protein transport between the ER and Golgi.

Developmental Biology. 2004;20:87-123

[4] Kim I, Xu WJ, Reed JC. Cell death and endoplasmic reticulum stress: Disease relevance and therapeutic opportunities. Nature Reviews. Drug

[5] Ghaemmaghami S, Huh WK, Bower K, Howson RW, Belle A, Dephoure N, et al. Global analysis of protein expression in yeast. Nature. 2003;425: 737-741. DOI: 10.1038/nature02046

[6] Osborne AR, Rapoport TA, van den Berg B. Protein translocation by the Sec61/SecY channel. Annual Review of Cell and Developmental Biology. 2005; 21:529-550. DOI: 10.1146/annurev.

[7] Robson A, Collinson I. The structure of the Sec complex and the problem of protein translocation. EMBO Reports. 2006;7:1099-1103. DOI: 10.1038/sj.

[8] Yoshida Y, Tanaka K. Lectin-like ERAD players in ER and cytosol. Biochimica et Biophysica Acta. 2010;

DUBs are also proposed as factors that regulate ERAD. As explained above, several DUBs have been reported to interact with p97/VCP and function as positive regulators of retrotranslocation. Additionally, some DUBs are linked with the regulation of E3 enzyme stability. For example, USP19, an ER-anchored DUB, rescues HRD1 from proteasomal degradation and thereby regulates HRD1 stability [120]. Similarly, USP19 enhances the stability and activity of another E3 MARCH6 [121].

SVIP (small VCP interacting protein), a VCP-interaction motif (VIM) containing protein, is the first identified endogen ERAD inhibitor. SVIP interacts with p97/VCP and Derlin1 and inhibits the ubiquitination and degradation of gp78 dependent ERAD substrates [111]. Another endogen ERAD inhibitor is SAKS1. SAKS1 binds to the polyubiquitin chain of the substrate and p97/VCP and attenuates the ERAD process [122].

ERAD activity can also be controlled by hormonal regulation. Glucocorticoids have been suggested to ameliorate ER stress by promoting correct folding of secreted proteins and enhancing removal of misfolded proteins from the ER probably through induction of UPR. Recently, androgen-mediated regulation of ERAD has been reported. Androgen treatment upregulated the expression of Os9, p97/ VCP, Ufd1, Npl4, Hrd1 and gp78, but downregulated ERAD inhibitor SVIP, which in turn enhanced the proteolytic activity of ERAD in androgen-sensitive prostate cancer cells [123]. Furthermore, the regulation of ERAD by androgen is mediated via AR and is partially or fully independent on the androgen-mediated induction of IRE1α branch [123].

#### Conflict of interest

The authors declare no conflict of interest.

#### Author details

Burcu Erbaykent Tepedelen<sup>1</sup> and Petek Ballar Kirmizibayrak<sup>2</sup>

1 Faculty of Science, Molecular Biology and Genetics, Uludag University, Bursa, Turkey

2 Biochemistry Department, Faculty of Pharmacy, Ege University, Izmir, Turkey

\*Address all correspondence to: petek.ballar@ege.edu.tr

© 2019 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/ by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Endoplasmic Reticulum-Associated Degradation (ERAD) DOI: http://dx.doi.org/10.5772/intechopen.82043

#### References

biosynthesis. Under low sterol levels, HMGR is stable; however, when sterol levels are high, Insig-HMGR interaction become favored, leading delivery of HMGR to E3 complex following by its proteasomal degradation [119]. Likewise, ERAD-mediated degradations of apolipoprotein and IP3R are initiated when lipid levels are low and

DUBs are also proposed as factors that regulate ERAD. As explained above, several DUBs have been reported to interact with p97/VCP and function as positive regulators of retrotranslocation. Additionally, some DUBs are linked with the regulation of E3 enzyme stability. For example, USP19, an ER-anchored DUB, rescues HRD1 from proteasomal degradation and thereby regulates HRD1 stability [120]. Similarly, USP19 enhances the stability and activity of another E3 MARCH6 [121]. SVIP (small VCP interacting protein), a VCP-interaction motif (VIM) containing protein, is the first identified endogen ERAD inhibitor. SVIP interacts with p97/VCP and Derlin1 and inhibits the ubiquitination and degradation of gp78 dependent ERAD substrates [111]. Another endogen ERAD inhibitor is SAKS1. SAKS1 binds to the polyubiquitin chain of the substrate and p97/VCP and attenu-

ERAD activity can also be controlled by hormonal regulation. Glucocorticoids

have been suggested to ameliorate ER stress by promoting correct folding of secreted proteins and enhancing removal of misfolded proteins from the ER probably through induction of UPR. Recently, androgen-mediated regulation of ERAD has been reported. Androgen treatment upregulated the expression of Os9, p97/ VCP, Ufd1, Npl4, Hrd1 and gp78, but downregulated ERAD inhibitor SVIP, which in turn enhanced the proteolytic activity of ERAD in androgen-sensitive prostate cancer cells [123]. Furthermore, the regulation of ERAD by androgen is mediated via AR and is partially or fully independent on the androgen-mediated induction of

calcium levels are high, respectively [23].

Endoplasmic Reticulum

ates the ERAD process [122].

IRE1α branch [123].

Conflict of interest

Author details

Turkey

38

The authors declare no conflict of interest.

Burcu Erbaykent Tepedelen<sup>1</sup> and Petek Ballar Kirmizibayrak<sup>2</sup>

\*Address all correspondence to: petek.ballar@ege.edu.tr

provided the original work is properly cited.

1 Faculty of Science, Molecular Biology and Genetics, Uludag University, Bursa,

2 Biochemistry Department, Faculty of Pharmacy, Ege University, Izmir, Turkey

© 2019 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/ by/3.0), which permits unrestricted use, distribution, and reproduction in any medium,

[1] Alberts B, Johnson A, Lewis J, Morgan D, Raff R, Roberts M, et al. Membrane transport of small molecules and the electrical properties of membranes: The endoplasmic reticulum. In: Richte ML, editor. Molecular Biology of The Cell. 6th ed. Garland Science, Taylor&Francis Group. 2015. pp. 669-691

[2] Schwarz DS, Blower MD. The endoplasmic reticulum: Structure, function and response to cellular signaling. Cellular and Molecular Life Sciences. 2016;73:79-94. DOI: 10.1007/ s00018-015-2052-6

[3] Lee MCS, Miller EA, Goldberg J, Orci L, Schekman R. Bi-directional protein transport between the ER and Golgi. Annual Review of Cell and Developmental Biology. 2004;20:87-123

[4] Kim I, Xu WJ, Reed JC. Cell death and endoplasmic reticulum stress: Disease relevance and therapeutic opportunities. Nature Reviews. Drug Discovery. 2008;7:1013-1030

[5] Ghaemmaghami S, Huh WK, Bower K, Howson RW, Belle A, Dephoure N, et al. Global analysis of protein expression in yeast. Nature. 2003;425: 737-741. DOI: 10.1038/nature02046

[6] Osborne AR, Rapoport TA, van den Berg B. Protein translocation by the Sec61/SecY channel. Annual Review of Cell and Developmental Biology. 2005; 21:529-550. DOI: 10.1146/annurev. cellbio.21.012704.133214

[7] Robson A, Collinson I. The structure of the Sec complex and the problem of protein translocation. EMBO Reports. 2006;7:1099-1103. DOI: 10.1038/sj. embor.7400832

[8] Yoshida Y, Tanaka K. Lectin-like ERAD players in ER and cytosol. Biochimica et Biophysica Acta. 2010; 1800:172-180. DOI: 10.1016/j. bbagen.2009.07.029

[9] Christianson JC, Ye Y. Cleaning up in the endoplasmic reticulum: Ubiquitin in charge. Nature Structural & Molecular Biology. 2014;21:325-335. DOI: 10.1038/ nsmb.2793

[10] Araki K, Nagata K. Protein folding and quality control in the ER. Cold Spring Harbor Perspectives in Biology. 2011;3:a007526. DOI: 10.1101/ cshperspect.a007526

[11] Romisch K. Endoplasmic reticulumassociated degradation. Annual Review of Cell and Developmental Biology. 2005;21:435-456. DOI: 10.1146/annurev. cellbio.21.012704.133250

[12] Olzmann JA, Kopito RR, Christianson JC. The mammalian endoplasmic reticulum-associated degradation system. Cold Spring Harbor Perspectives in Medicine. 2013;5. DOI: 10.1101/cshperspect.a013185

[13] Schubert U, Anton LC, Gibbs J, Norbury CC, Yewdell JW, Bennink JR. Rapid degradation of a large fraction of newly synthesized proteins by proteasomes. Nature. 2000;404: 770-774. DOI: 10.1038/35008096

[14] Morito D, Nagata K. Pathogenic hijacking of ER-associated degradation: Is ERAD flexible? Molecular Cell. 2015; 59:335-344. DOI: 10.1016/j. molcel.2015.06.010

[15] Nishikawa S, Brodsky JL, Nakatsukasa K. Roles of molecular chaperones in endoplasmic reticulum (ER) quality control and ER-associated degradation (ERAD). Journal of Biochemistry. 2005;137:551-555. DOI: 10.1093/jb/mvi068

[16] Nyathi Y, Wilkinson BM, Pool MR. Co-translational targeting and

translocation of proteins to the endoplasmic reticulum. Biochimica et Biophysica Acta. 2013;1833:2392-2402. DOI: 10.1016/j.bbamcr.2013.02.021

[17] Brandizzi F, Barlowe C. Organization of the ER-Golgi interface for membrane traffic control. Nature Reviews Molecular Cell Biology. 2013; 14:382-392. DOI: 10.1038/nrm3588

[18] Rao RV, Ellerby HM, Bredesen DE. Coupling endoplasmic reticulum stress to the cell death program. Cell Death and Differentiation. 2004;11:372-380. DOI: 10.1038/sj.cdd.4401378

[19] Hegde RS, Ploegh HL. Quality and quantity control at the endoplasmic reticulum. Current Opinion in Cell Biology. 2010;22:437-446. DOI: 10.1016/j.ceb.2010.05.005

[20] Iwawaki T, Oikawa D. The role of the unfolded protein response in diabetes mellitus. Seminars in Immunopathology. 2013;35:333-350. DOI: 10.1007/s00281-013-0369-5

[21] Scheper W, Hoozemans JJ. The unfolded protein response in neurodegenerative diseases: A neuropathological perspective. Acta Neuropathologica. 2015;130:315-331. DOI: 10.1007/s00401-015-1462-8

[22] Printsev I, Curiel D, Carraway KL 3rd. Membrane protein quantity control at the endoplasmic reticulum. The Journal of Membrane Biology. 2017;250: 379-392. DOI: 10.1007/s00232-016- 9931-0

[23] Merulla J, Fasana E, Solda T, Molinari M. Specificity and regulation of the endoplasmic reticulum-associated degradation machinery. Traffic. 2013; 14:767-777. DOI: 10.1111/tra.12068

[24] Hampton RY, Gardner RG, Rine J. Role of 26S proteasome and HRD genes in the degradation of 3-hydroxy-3-methylglutaryl-CoA reductase, an

integral endoplasmic reticulum membrane protein. Molecular Biology of the Cell. 1996;7:2029-2044

[32] Duan XH, Zhou YB, Teng X, Tang CS, Qi YF. Endoplasmic reticulum stress-mediated apoptosis is activated in vascular calcification. Biochemical and Biophysical Research Communications. 2009;387:694-699. DOI: 10.1016/j.

DOI: http://dx.doi.org/10.5772/intechopen.82043

Endoplasmic Reticulum-Associated Degradation (ERAD)

DOI: 10.1146/annurev.biochem.

[40] Caramelo JJ, Castro OA, Alonso LG, De Prat-Gay G, Parodi AJ. UDP-Glc: glycoprotein glucosyltransferase recognizes structured and solvent accessible hydrophobic patches in molten globule-like folding intermediates. Proceedings of the National Academy of Sciences of the United States of America. 2003;100: 86-91. DOI: 10.1073/pnas.262661199

[41] Taylor SC, Thibault P, Tessier DC, Bergeron JJ, Thomas DY. Glycopeptide specificity of the secretory protein folding sensor UDP-glucose

glycoprotein:glucosyltransferase. EMBO Reports. 2003;4:405-411. DOI: 10.1038/

[42] Vembar SS, Brodsky JL. One step at

associated degradation. Nature Reviews. Molecular Cell Biology. 2008;9:944-957.

[43] Brodsky JL, Werner ED, Dubas ME, Goeckeler JL, Kruse KB, McCracken AA.

degradation demonstrates that protein export and import are mechanistically distinct. The Journal of Biological Chemistry. 1999;274:3453-3460

a time: Endoplasmic reticulum-

The requirement for molecular chaperones during endoplasmic reticulum-associated protein

[44] Vashist S, Ng DT. Misfolded proteins are sorted by a sequential checkpoint mechanism of ER quality control. The Journal of Cell Biology. 2004;165:41-52. DOI: 10.1083/

[45] Jakob CA, Burda P, Roth J, Aebi M. Degradation of misfolded endoplasmic

Saccharomyces cerevisiae is determined by a specific oligosaccharide structure. The Journal of Cell Biology. 1998;142:

reticulum glycoproteins in

jcb.200309132

1223-1233

73.011303.073752

sj.embor.embor797

DOI: 10.1038/nrm2546

[33] Baldridge RD, Rapoport TA. Autoubiquitination of the Hrd1 ligase triggers protein retrotranslocation in ERAD. Cell. 2016;166:394-407. DOI:

[34] Stroud RM, Walter P. Signal sequence recognition and protein targeting. Current Opinion in Structural

[35] Ellgaard L, Ruddock LW. The human protein disulphide isomerase family: Substrate interactions and functional properties. EMBO Reports.

2005;6:28-32. DOI: 10.1038/sj.

[37] Schelhaas M, Malmstrom J, Pelkmans L, Haugstetter J, Ellgaard L, Grunewald K, et al. Simian Virus 40 depends on ER protein folding and quality control factors for entry into host cells. Cell. 2007;131:516-529. DOI:

10.1016/j.cell.2007.09.038

1443-1456

41

[38] Gillece P, Luz JM, Lennarz WJ, de La Cruz FJ, Romisch K. Export of a cysteine-free misfolded secretory protein from the endoplasmic reticulum for degradation requires interaction with protein disulfide isomerase. The Journal of Cell Biology. 1999;147:

[39] Helenius A, Aebi M. Roles of N-linked glycans in the endoplasmic

reticulum. Annual Review of Biochemistry. 2004;73:1019-1049.

[36] Tsai B, Rodighiero C, Lencer WI, Rapoport TA. Protein disulfide isomerase acts as a redox-dependent chaperone to unfold cholera toxin. Cell.

10.1016/j.cell.2016.05.048

Biology. 1999;9:754-759

embor.7400311

2001;104:937-948

bbrc.2009.07.085

[25] Tsai YC, Mendoza A, Mariano JM, Zhou M, Kostova Z, Chen B, et al. The ubiquitin ligase gp78 promotes sarcoma metastasis by targeting KAI1 for degradation. Nature Medicine. 2007;13: 1504-1509. DOI: 10.1038/nm1686

[26] Zhang H, Hoff H, Sell C. Insulin-like growth factor I-mediated degradation of insulin receptor substrate-1 is inhibited by epidermal growth factor in prostate epithelial cells. The Journal of Biological Chemistry. 2000;275:22558-22562. DOI: 10.1074/jbc.M000412200

[27] Guerriero CJ, Brodsky JL. The delicate balance between secreted protein folding and endoplasmic reticulum-associated degradation in human physiology. Physiological Reviews. 2012;92:537-576

[28] Kim S-M, Wang Y, Nabavi N, Liu Y, Correia MA. Hepatic cytochromes P450: Structural degrons and barcodes, posttranslational modifications and cellular adapters in the ERAD-endgame. Drug Metabolism Reviews. 2016;48: 405-433

[29] Loureiro J, Ploegh HL. Antigen presentation and the ubiquitinproteasome system in host-pathogen interactions. Advances in Immunology. 2006;92:225-305. DOI: 10.1016/ S0065-2776(06)92006-9

[30] Margottin F, Bour SP, Durand H, Selig L, Benichou S, Richard V, et al. A novel human WD protein, h-beta TrCp, that interacts with HIV-1 Vpu connects CD4 to the ER degradation pathway through an F-box motif. Molecular Cell. 1998;1:565

[31] Jarosch E, Lenk U, Sommer T. Endoplasmic reticulum-associated protein degradation. International Review of Cytology. 2003;223:39-81 Endoplasmic Reticulum-Associated Degradation (ERAD) DOI: http://dx.doi.org/10.5772/intechopen.82043

[32] Duan XH, Zhou YB, Teng X, Tang CS, Qi YF. Endoplasmic reticulum stress-mediated apoptosis is activated in vascular calcification. Biochemical and Biophysical Research Communications. 2009;387:694-699. DOI: 10.1016/j. bbrc.2009.07.085

translocation of proteins to the endoplasmic reticulum. Biochimica et Biophysica Acta. 2013;1833:2392-2402. DOI: 10.1016/j.bbamcr.2013.02.021

Endoplasmic Reticulum

[17] Brandizzi F, Barlowe C.

DOI: 10.1038/sj.cdd.4401378

Organization of the ER-Golgi interface for membrane traffic control. Nature Reviews Molecular Cell Biology. 2013; 14:382-392. DOI: 10.1038/nrm3588

integral endoplasmic reticulum membrane protein. Molecular Biology

of the Cell. 1996;7:2029-2044

10.1074/jbc.M000412200

405-433

[27] Guerriero CJ, Brodsky JL. The delicate balance between secreted protein folding and endoplasmic reticulum-associated degradation in human physiology. Physiological Reviews. 2012;92:537-576

[28] Kim S-M, Wang Y, Nabavi N, Liu Y, Correia MA. Hepatic cytochromes P450: Structural degrons and barcodes, posttranslational modifications and cellular adapters in the ERAD-endgame. Drug Metabolism Reviews. 2016;48:

[29] Loureiro J, Ploegh HL. Antigen presentation and the ubiquitinproteasome system in host-pathogen interactions. Advances in Immunology.

2006;92:225-305. DOI: 10.1016/ S0065-2776(06)92006-9

[30] Margottin F, Bour SP, Durand H, Selig L, Benichou S, Richard V, et al. A novel human WD protein, h-beta TrCp, that interacts with HIV-1 Vpu connects CD4 to the ER degradation pathway through an F-box motif. Molecular Cell. 1998;1:565

[31] Jarosch E, Lenk U, Sommer T. Endoplasmic reticulum-associated protein degradation. International Review of Cytology. 2003;223:39-81

[25] Tsai YC, Mendoza A, Mariano JM, Zhou M, Kostova Z, Chen B, et al. The ubiquitin ligase gp78 promotes sarcoma metastasis by targeting KAI1 for degradation. Nature Medicine. 2007;13: 1504-1509. DOI: 10.1038/nm1686

[26] Zhang H, Hoff H, Sell C. Insulin-like growth factor I-mediated degradation of insulin receptor substrate-1 is inhibited by epidermal growth factor in prostate epithelial cells. The Journal of Biological Chemistry. 2000;275:22558-22562. DOI:

[18] Rao RV, Ellerby HM, Bredesen DE. Coupling endoplasmic reticulum stress to the cell death program. Cell Death and Differentiation. 2004;11:372-380.

[19] Hegde RS, Ploegh HL. Quality and quantity control at the endoplasmic reticulum. Current Opinion in Cell Biology. 2010;22:437-446. DOI: 10.1016/j.ceb.2010.05.005

[20] Iwawaki T, Oikawa D. The role of the unfolded protein response in diabetes mellitus. Seminars in Immunopathology. 2013;35:333-350. DOI: 10.1007/s00281-013-0369-5

[21] Scheper W, Hoozemans JJ. The unfolded protein response in neurodegenerative diseases: A neuropathological perspective. Acta Neuropathologica. 2015;130:315-331. DOI: 10.1007/s00401-015-1462-8

[22] Printsev I, Curiel D, Carraway KL 3rd. Membrane protein quantity control at the endoplasmic reticulum. The Journal of Membrane Biology. 2017;250: 379-392. DOI: 10.1007/s00232-016-

[23] Merulla J, Fasana E, Solda T,

Molinari M. Specificity and regulation of the endoplasmic reticulum-associated degradation machinery. Traffic. 2013; 14:767-777. DOI: 10.1111/tra.12068

[24] Hampton RY, Gardner RG, Rine J. Role of 26S proteasome and HRD genes in the degradation of 3-hydroxy-3-methylglutaryl-CoA reductase, an

9931-0

40

[33] Baldridge RD, Rapoport TA. Autoubiquitination of the Hrd1 ligase triggers protein retrotranslocation in ERAD. Cell. 2016;166:394-407. DOI: 10.1016/j.cell.2016.05.048

[34] Stroud RM, Walter P. Signal sequence recognition and protein targeting. Current Opinion in Structural Biology. 1999;9:754-759

[35] Ellgaard L, Ruddock LW. The human protein disulphide isomerase family: Substrate interactions and functional properties. EMBO Reports. 2005;6:28-32. DOI: 10.1038/sj. embor.7400311

[36] Tsai B, Rodighiero C, Lencer WI, Rapoport TA. Protein disulfide isomerase acts as a redox-dependent chaperone to unfold cholera toxin. Cell. 2001;104:937-948

[37] Schelhaas M, Malmstrom J, Pelkmans L, Haugstetter J, Ellgaard L, Grunewald K, et al. Simian Virus 40 depends on ER protein folding and quality control factors for entry into host cells. Cell. 2007;131:516-529. DOI: 10.1016/j.cell.2007.09.038

[38] Gillece P, Luz JM, Lennarz WJ, de La Cruz FJ, Romisch K. Export of a cysteine-free misfolded secretory protein from the endoplasmic reticulum for degradation requires interaction with protein disulfide isomerase. The Journal of Cell Biology. 1999;147: 1443-1456

[39] Helenius A, Aebi M. Roles of N-linked glycans in the endoplasmic reticulum. Annual Review of Biochemistry. 2004;73:1019-1049.

DOI: 10.1146/annurev.biochem. 73.011303.073752

[40] Caramelo JJ, Castro OA, Alonso LG, De Prat-Gay G, Parodi AJ. UDP-Glc: glycoprotein glucosyltransferase recognizes structured and solvent accessible hydrophobic patches in molten globule-like folding intermediates. Proceedings of the National Academy of Sciences of the United States of America. 2003;100: 86-91. DOI: 10.1073/pnas.262661199

[41] Taylor SC, Thibault P, Tessier DC, Bergeron JJ, Thomas DY. Glycopeptide specificity of the secretory protein folding sensor UDP-glucose glycoprotein:glucosyltransferase. EMBO Reports. 2003;4:405-411. DOI: 10.1038/ sj.embor.embor797

[42] Vembar SS, Brodsky JL. One step at a time: Endoplasmic reticulumassociated degradation. Nature Reviews. Molecular Cell Biology. 2008;9:944-957. DOI: 10.1038/nrm2546

[43] Brodsky JL, Werner ED, Dubas ME, Goeckeler JL, Kruse KB, McCracken AA. The requirement for molecular chaperones during endoplasmic reticulum-associated protein degradation demonstrates that protein export and import are mechanistically distinct. The Journal of Biological Chemistry. 1999;274:3453-3460

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cell.2007.11.023

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bies.201200159

44

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DOI: http://dx.doi.org/10.5772/intechopen.82043

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[123] Erzurumlu Y, Ballar P. Androgen mediated regulation of endoplasmic reticulum-associated degradation and its effects on prostate cancer. Scientific Reports. 2017;7:40719. DOI: 10.1038/

10.3390/ijms17111829

10.1074/jbc.M110.158030

srep40719

47

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endoplasmic reticulum: A target for new anticancer drugs. In Vivo. 2007;21:

[113] Chambers JE, Petrova K, Tomba G,

[114] Lynes EM, Bui M, Yap MC, Benson MD, Schneider B, Ellgaard L, et al. Palmitoylated TMX and calnexin target

membrane. The EMBO Journal. 2012;31:

[115] Lakkaraju AK, Abrami L, Lemmin T, Blaskovic S, Kunz B, Kihara A, et al.

component of the ribosome–translocon complex. The EMBO Journal. 2012;31:

[116] Ye Y, Baek S-H, Ye Y, Zhang T. Proteomic characterization of

endogenous substrates of mammalian ubiquitin ligase Hrd1. Cell & Bioscience.

[117] Nadav E, Shmueli A, Barr H, Gonen H, Ciechanover A, Reiss Y. A novel mammalian endoplasmic reticulum ubiquitin ligase homologous to the yeast Hrd1. Biochemical and Biophysical Research Communications.

[118] Hagiwara M, Ling J, Koenig PA, Ploegh HL. Posttranscriptional regulation of glycoprotein quality control in the endoplasmic reticulum is controlled by the E2 Ub-conjugating enzyme UBC6e. Molecular Cell. 2016;

[119] Song BL, Sever N, DeBose-Boyd RA. Gp78, a membrane-anchored ubiquitin ligase, associates with Insig-1

ubiquitination to degradation of HMG CoA reductase. Molecular Cell. 2005;19:

63:753-767. DOI: 10.1016/j.

and couples sterol-regulated

829-840. DOI: 10.1016/j. molcel.2005.08.009

molcel.2016.07.014

ribosylation adapts an ER chaperone response to short-term fluctuations in unfolded protein load. The Journal of Cell Biology. 2012;198:371-385

Vendruscolo M, Ron D. ADP

to the mitochondria-associated

Palmitoylated calnexin is a key

457-470

1823-1835

2018;8:46

2003;303:91-97

[106] Ogata M, Hino SI, Saito A, Morikawa K, Kondo S, Kanemoto S, et al. Autophagy is activated for cell survival after endoplasmic reticulum stress. Molecular and Cellular Biology. 2006;26:9220-9231. DOI: 10.1128/

[107] Brodsky JL. Cleaning up: ERassociated degradation to the rescue. Cell. 2012;151:1163-1167. DOI: 10.1016/j.

[108] Olivari S, Galli C, Alanen H, Ruddock L, Molinari M. A novel stressinduced EDEM variant regulating endoplasmic reticulum-associated glycoprotein degradation. The Journal of Biological Chemistry. 2005;280:

[109] Yoshida H, Matsui T, Hosokawa N, Kaufman RJ, Nagata K, Mori K. A time-

mammalian unfolded protein response. Developmental Cell. 2003;4:265-271

[110] Alcock F, Swanton E. Mammalian OS-9 is upregulated in response to endoplasmic reticulum stress and facilitates ubiquitination of misfolded glycoproteins. Journal of Molecular Biology. 2009;385:1032-1042

[111] Ballar P, Zhong Y, Nagahama M,

Identification of SVIP as an endogenous inhibitor of endoplasmic reticulumassociated degradation. The Journal of Biological Chemistry. 2007;282: 33908-33914. DOI: 10.1074/jbc.

[112] Storm M, Sheng X, Arnoldussen YJ, Saatcioglu F. Prostate cancer and the unfolded protein response. Oncotarget. 2016;7:54051-54066. DOI: 10.18632/

Tagaya M, Shen Y, Fang S.

M704446200

oncotarget.9912

46

dependent phase shift in the

215-226

Endoplasmic Reticulum

Mcb.01453-06

cell.2012.11.012

2424-2428

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[122] LaLonde DP, Bretscher A. The UBX protein SAKS1 negatively regulates endoplasmic reticulum-associated degradation and p97-dependent degradation. The Journal of Biological Chemistry. 2011;286:4892-4901. DOI: 10.1074/jbc.M110.158030

[123] Erzurumlu Y, Ballar P. Androgen mediated regulation of endoplasmic reticulum-associated degradation and its effects on prostate cancer. Scientific Reports. 2017;7:40719. DOI: 10.1038/ srep40719

**49**

**Chapter 4**

**Abstract**

Autophagy

*and Han-Jung Chae*

and inhibition signaling network.

**1. Endoplasmic reticulum**

**Keywords:** ER stress, autophagy, calcium, lysosome

Endoplasmic Reticulum Stress and

In eukaryotic cells, the aggregation of the endoplasmic reticulum (ER)-mediated unfolded or misfolded proteins leads to disruption of the ER homeostasis, which can trigger ER stress. To restore the ER homeostasis, the ER stress activates the intracellular signaling cascade from the ER to the nucleus, referred to as the unfolded protein response (UPR). Autophagy primitively portrayed as an evolutionarily conserved process is involved in cellular homeostasis by facilitating the lysosomal degradation pathway for the recycling and elimination of intracellular defective macromolecules and organelles. Autophagy is tightly regulated by the protective mechanism of UPR. The UPR and autophagy are interlinked, which indicates that the ER stress can not only induce autophagy but also suppress it. Here, we discuss the molecular mechanism of ER stress and autophagy and their induction

The endoplasmic reticulum (ER) is a central membrane-bound organelle constructed from a dynamic network of tubules involved in cellular processes such as protein synthesis, gluconeogenesis, lipid synthesis and processing, and calcium storage and release in the cell and contributes to the generation of autophagosomes and peroxisomes [1]. The extension of ER morphology depends on the cell's activity and lineage; it is organized into subcompartments of different shapes, such as cisternae and tubules. ER appears as two main interconnected compartments, namely, the smooth endoplasmic reticulum (SER) and the rough endoplasmic reticulum (RER), which are abundant in different proportions in different cell lineages [2]. RER is less tubular than the SER, which forms an interrelated network of subdomains of ER; the RER is illuminated with ribosomes on their membranes, which are absent in the SER. RER has appeared in all cells and its density is higher, similar to that of the Golgi apparatus and nucleus because in all cells the nascent polypeptide is cotranslationally inserted into their membranes from the ER membrane. However, SER is present in only certain cell types, such as the liver cells, steroid-synthesizing cells, neurons, and muscle cells. SER is involved in the generation of steroid hormones within the adrenal cortex and endocrine glands and acts as a center for detoxification and protein transportation [3, 4]. A remarkable number of proteins are Ca2+ dependent and need a completely oxidizing environment [5]. In the lumen, the

*Mohammad Fazlul Kabir, Hyung-Ryong Kim* 

#### **Chapter 4**
