Function of ER in reviewing mutated proteins

Function of ER in reviewing mutated proteins

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At least in the case of Cystic Fibrosis it happens that a mutant protein (which could actually function!) is held in the ER because the ER detects it as misfolded. Does this happen in every type of mutant protein?

If so, should I conclude that such genetic disorders are caused because of the total absence of a protein rather than because of a misfolded protein? If not, why would the ER or anything else in the cell allow some mutated proteins to do any job (Like in the case of Sickle cell anaemia, where an amino acid substitution happens)?

How do scientists conclude that a misfolded protein could have done a good enough job (Like in the case of cystic fibrosis)?

Membrane proteins have to go to the ER from where they are transported to the Golgi apparatus. Usually these proteins are translocated into the ER while translation is still going on. There are chaperones (such as calbindin) in ER which help in translocation and also catalyze initial steps of folding. ER is also involved in unfolded protein response. Unless the protein is properly folded it is not transported to Golgi. However, this process is regulated at many steps and I wont add a detailed explanation of the mechanism, in this answer.

To conclude ER is involved in this case because the protein considered is a membrane protein.

Endoplasmic Reticulum

Stress Responses in the Endoplasmic Reticulum

Conditions that flood the ER with excess protein or result in accumulation of misfolded proteins trigger the unfolded protein response (UPR Fig. 20.13 ). Essentially, any condition in which protein import exceeds the protein-folding capacity of the ER triggers the UPR: misfolding of mutant proteins, inhibition of ER glycosylation (eg, by the drug tunicamycin), inhibition of disulfide formation (by reducing agents), or even overproduction of normal proteins. To compensate for these events, this stress-induced signaling pathway upregulates genes that are required to synthesize the entire ER, including its folding machinery. In yeast, the UPR activates more than 300 genes involved with all aspects of ER function, including lipid synthesis, protein translocation, protein folding, glycosylation, and degradation, as well as export to and retrieval from the Golgi apparatus. Developmental programs in metazoans may work through the same genetic controls to determine the abundance of ER in differentiated cells, producing, for example, extensive ER in secretory cells such as plasma, liver, and pancreatic acinar cells.

The UPR depends on three ER transmembrane proteins that sense and respond to stress in the ER: ATF6 (activating transcription factor 6) IRE1 (inositol requiring 1) and PERK/PEK (PKR-like endoplasmic reticulum kinase/pancreatic eIF2a kinase) ( Fig. 20.13 ). Each of these proteins has a different mechanism but all initiate signaling pathways that regulate the production of proteins required for ER function. This allows cells to adjust the capacity of the ER to promote protein folding depending on the demand. Metazoan cells have all three pathways, but yeast have only IRE1.

Under normal conditions the concentration of BiP in the ER lumen exceeds the concentration of unfolded proteins, so free BiP is available to bind to the luminal domains of three ER transmembrane proteins, ATF6, IRE1 and PERK (PKR-like endoplasmic reticulum kinase). These interactions with BiP retain ATF6 in the ER and prevent the dimerization of IRE1 and PERK, keeping them inactive.

If unfolded proteins in the lumen of the ER bind to all of the BiP, IRE1 is free to dimerize. This activates the endoribonuclease activity of the cytoplasmic domain of IRE1, allowing it to remove a small intron from the messenger RNA (mRNA) for XBP1, a bZIP (basic leucine zipper) domain–containing transcription factor (see Fig. 10.14 ). Removing this intron alters the translational reading frame of XBP1 allowing the mRNA to encode a potent transcriptional activator of genes for ER proteins.

Similarly, without free BiP in the ER lumen, PERK dimerizes and phosphorylates eukaryotic translation initiation factor 2 (eIF2). This reduces the frequency of AUG codon recognition and slows the rate of translation initiation on many mRNAs. However, mRNAs for many proteins involved in cell survival and ER functions are preferentially translated under these conditions.

Accumulated unfolded proteins release BiP from ATF6, causing ER transmembrane protein to be transported to the Golgi apparatus, where it is cleaved by S1P and S2P proteases to produce a soluble fragment that is released into the cytoplasm. The fragment moves to the nucleus, where it activates the transcription of target genes, including XBP1.

Aspects of the UPR pathway involving IRE1 and PERK are also important for promoting differentiation in higher eukaryotic cells. For example, IRE1 is activated during B-lymphocyte differentiation into a plasma cell (see Fig. 28.9 ), during which the ER expands fivefold to accommodate the synthesis and secretion of immunoglobulins. Activation of the innate immune response (see Fig. 28.6 ), for example by lipopolysaccharide treatment, also activates IRE1. PERK activity is also required for B-cell differentiation and/or survival.

Endoplasmic reticulum

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Endoplasmic reticulum (ER), in biology, a continuous membrane system that forms a series of flattened sacs within the cytoplasm of eukaryotic cells and serves multiple functions, being important particularly in the synthesis, folding, modification, and transport of proteins . All eukaryotic cells contain an endoplasmic reticulum (ER). In animal cells, the ER usually constitutes more than half of the membranous content of the cell. Differences in certain physical and functional characteristics distinguish the two types of ER, known as rough ER and smooth ER.

What is the endoplasmic reticulum?

  • The endoplasmic reticulum (ER) is a continuous membrane system that forms a series of flattened sacs within the cytoplasm of eukaryotic cells.
  • All eukaryotic cells contain an ER.
  • In animal cells, the ER usually constitutes more than half of the membranous content of the cell.
  • The ER can be classified in two functionally distinct forms: the smooth endoplasmic reticulum (SER) and the rough endoplasmic reticulum (RER).

What is the difference between smooth and rough endoplasmic reticulum?

The ER can be classified in two functionally distinct forms: smooth endoplasmic reticulum (SER) and rough endoplasmic reticulum (RER). The morphological distinction between the two is the presence of protein-synthesizing particles, called ribosomes, attached to the outer surface of the RER. The functions of the SER, a meshwork of fine tubular membrane vesicles, vary considerably from cell to cell, one important role being the synthesis of phospholipids and cholesterol, which are major components of the plasma and internal membranes. The RER is generally a series of connected flattened sacs. It plays a central role in the synthesis and export of proteins and glycoproteins and is best studied in secretory cells specializing in these functions. The many secretory cells in the human body include liver cells secreting serum proteins (e.g., albumin), endocrine cells secreting peptide hormones (e.g., insulin), pancreatic acinar cells secreting digestive enzymes, and cartilage cells secreting collagen.

What is the function of the endoplasmic reticulum?

The endoplasmic reticulum (ER) serves important functions particularly in the synthesis, folding, modification, and transport of proteins. Differences in certain physical and functional characteristics distinguish the two types of ER, known as rough ER (RER) and smooth ER (SER). Ribosomes on RER, which give RER its rough appearance, specialize in the synthesis of proteins that possess a signal sequence that directs them specifically to the ER for processing. Proteins synthesized by the RER have specific final destinations, such as the cell membrane, cell exterior, or the ER itself. SER is involved in the synthesis of lipids, including cholesterol and phospholipids, which are used in the production of new cellular membrane. In cells of the liver, SER contributes to the detoxification of drugs and harmful chemicals. The sarcoplasmic reticulum is a specialized type of SER that regulates calcium ion concentration in the cytoplasm of striated muscle cells.

When was the endoplasmic reticulum discovered?

The ER was first noted in the late 19th century, when studies of stained cells indicated the presence of some type of extensive cytoplasmic structure, then known as the gastroplasm. The electron microscope made possible the study of the morphology of the ER in the 1940s, when it was given its present name.

Rough ER is named for its rough appearance, which is due to the ribosomes attached to its outer (cytoplasmic) surface. Rough ER lies immediately adjacent to the cell nucleus, and its membrane is continuous with the outer membrane of the nuclear envelope. The ribosomes on rough ER specialize in the synthesis of proteins that possess a signal sequence that directs them specifically to the ER for processing. (A number of other proteins in a cell, including those destined for the nucleus and mitochondria, are targeted for synthesis on free ribosomes, or those not attached to the ER membrane see the article ribosome.) Proteins synthesized by the rough ER have specific final destinations. Some proteins, for example, remain within the ER, whereas others are sent to the Golgi apparatus, which lies next to the ER. Proteins secreted from the Golgi apparatus are directed to lysosomes or to the cell membrane still others are destined for secretion to the cell exterior. Proteins targeted for transport to the Golgi apparatus are transferred from ribosomes on rough ER into the rough ER lumen, which serves as the site of protein folding, modification, and assembly.

The proximity of the rough ER to the cell nucleus gives the ER unique control over protein processing. The rough ER is able to rapidly send signals to the nucleus when problems in protein synthesis and folding occur and thereby influences the overall rate of protein translation. When misfolded or unfolded proteins accumulate in the ER lumen, a signaling mechanism known as the unfolded protein response (UPR) is activated. The response is adaptive, such that UPR activation triggers reductions in protein synthesis and enhancements in ER protein-folding capacity and ER-associated protein degradation. If the adaptive response fails, cells are directed to undergo apoptosis (programmed cell death).

Analysis of two mutated vacuolar proteins reveals a degradation pathway in the endoplasmic reticulum or a related compartment of yeast

The fate of a mutant form of each of the two yeast vacuolar enzymes proteinase yscA (PrA) and carboxypeptidase yscY (CPY) has been investigated. Both mutant proteins are rapidly degraded after entering the secretory pathway. Mutant PrA is deleted in 37 amino acids spanning the processing site region of the PrA pro-peptide. The mutant enzyme shows no activity towards maturation of itself or other vacuolar hydrolases, a function of wild-type PrA. Mutant CPY carries an Arg instead of a Gly residue in a highly conserved region, two positions distant from the active-site Ser. In contrast to wild-type CPY, the mutant form was quickly degraded by trypsin in vitro, indicating an altered structure. Using antisera specific for α-1→6 and α-1→3 outer-chain mannose linkages, no Golgi-specific carbohydrate modification could be detected on either mutant protein. Subcellular fractionation studies located both mutant enzymes in the endoplasmic reticulum. Degradation kinetics of both proteins show the same characteristics, indicating similar degradation pathways. The degradation process was shown to be independent of a functional sec18 gene product and takes place before Golgi-specific carbohydrate modifications occur. The proteasome, the major proteolytic activity of the cytoplasm, is not involved in this degradation event. All degradation characteristics of the two mutant proteins are consistent with a degradation process within the endoplasmic reticulum (‘ER degradation’).

Interactions between RAD51 and the BRCA proteins

Despite the apparent dissimilarity in protein sequence and structure, there is considerable evidence that BRCA1 and BRCA2 have common biological functions. BRCA1 and BRCA2 exhibit similar patterns of expression and sub-cellular localisation. They are both expressed in many tissues in a cell-cycle-dependent manner (Bertwistle et al., 1997 Blackshear et al., 1998 Connor et al., 1997b Rajan et al., 1996 Sharan and Bradley, 1997) their levels are highest during S phase, which is suggestive of functions during DNA replication. Both are localised to the nucleus in somatic cells, where they co-exist in characteristic subnuclear foci that redistribute following DNA damage. In meiotic cells, both proteins co-localise to the synaptonemal complexes of developing axial filaments (Chen et al., 1998a).

This pattern of expression and localisation is shared with RAD51, a mammalian homologue of the bacterial protein RecA, which is essential in Escherichia coli for the repair of DNA double-strand breaks (DSBs) by genetic recombination (Kowalczykowski, 2000). Indeed, both BRCA1 and BRCA2 have been reported to bind to RAD51. The BRCA2-RAD51 interaction is direct, in that it can be demonstrated in vitro with recombinant protein fragments (Chen et al., 1998b Wong et al., 1997) and in the yeast two-hybrid system, and appears to be of relatively high stoichiometry. A C-terminal motif spanning residues 3196-3232 of murine BRCA2 has been shown to mediate binding of the protein to the first 98 residues of RAD51 in yeast two-hybrid assays (Sharan et al., 1997). However, an analogous region of human BRCA2, which is 95% identical to the murine sequence, does not bind RAD51 (Wong et al., 1997 Aihara et al., 1999).

The interaction of RAD51 with human BRCA2 is mediated primarily, if not exclusively, by the eight BRC repeats (Wong et al., 1997). Each repeat, with the exception of BRC5 and BRC6, can bind individually to RAD51 in two-hybrid assays, as well as in vitro when expressed as a GST fusion protein. Their relative RAD51-binding capacities vary considerably (Chen et al., 1999), BRC4 being about four times as active as BRC1 in two-hybrid assays. PCR mutagenesis defines a binding consensus of about 30 residues present in both BRC1 and BRC4. Interestingly, despite the conservation of this core motif, different residues appear to be important in BRC1 and BRC4 for RAD51 binding. This, and the large difference in the affinity for RAD51, indicates that the mode of interaction of BRC1 or BRC4 with RAD51 could be quite distinct.

A region encompassing residues 758-1064 in BRCA1 was first reported to be involved in its interaction with RAD51 (Scully et al., 1997). It remains unclear, however, whether the two proteins can bind directly. Co-immunoprecipitation from cell extracts reveals an interaction of low stoichiometry, which has not yet been demonstrated in yeast two-hybrid assays or in vitro with recombinant proteins.

BRCA1 and BRCA2 co-localise in mitotic and meiotic cells (Chen et al., 1998a) and physically associate with one another through a region in BRCA1 (residues 1314-1863) distinct from that reported to bind to RAD51. Again, the interaction may not be direct, and it appears to involve a small fraction (perhaps 2-5%) of the total cellular pool of each protein. Moreover, recent efforts to characterise the protein complex associated with BRCA1 by biochemical purification and mass spectroscopy do not report the presence of appreciable amounts of either RAD51 or BRCA2 (Wang et al., 2000).

Thus, of the reported physical interactions between BRCA1, BRCA2 and RAD51, the BRCA2-RAD51 interaction appears to be the best established. Tantalising though existing evidence may be, it remains to be firmly demonstrated that BRCA1 functions together with BRCA2 and RAD51 in a multi-molecular complex, and the functional significance of their reported interactions is yet to be rigorously defined.

ER-associated degradation: Protein quality control and beyond

A. Ruggiano and O. Foresti contributed equally to this paper.

Annamaria Ruggiano, Ombretta Foresti, Pedro Carvalho ER-associated degradation: Protein quality control and beyond. J Cell Biol 17 March 2014 204 (6): 869–879. doi:

Even with the assistance of many cellular factors, a significant fraction of newly synthesized proteins ends up misfolded. Cells evolved protein quality control systems to ensure that these potentially toxic species are detected and eliminated. The best characterized of these pathways, the ER-associated protein degradation (ERAD), monitors the folding of membrane and secretory proteins whose biogenesis takes place in the endoplasmic reticulum (ER). There is also increasing evidence that ERAD controls other ER-related functions through regulated degradation of certain folded ER proteins, further highlighting the role of ERAD in cellular homeostasis.

Newly synthesized membrane and secreted proteins enter the ER in an unfolded state through a protein-conducting channel named the translocon (Rapoport, 2007). In the ER, a myriad of chaperones and modifying enzymes assist their membrane integration and folding. In many cases folding involves post-translational modifications, such as glycosylation or disulfide bond formation (Braakman and Hebert, 2013). At this stage many proteins are also assembled into multisubunit complexes with defined stoichiometries. As newly synthesized proteins reach a native conformation, they leave the ER to perform their function elsewhere either along the secretory pathway or outside of the cell.

Despite all the resources dedicated to protein folding, a significant fraction of newly synthesized polypeptides entering the ER fails to acquire a native conformation (Hartl and Hayer-Hartl, 2009). The degree of misfolding of these proteins varies considerably and can have several causes such as mutations, substoichiometric amounts of a binding partner, or merely a shortage of chaperone availability. In most cases, the misfolded molecules are retained in the ER and eventually become substrates of the ER-associated protein degradation (ERAD), a collection of quality-control mechanisms that clears the ER from these potentially harmful species. Inactivation of ERAD results in the accumulation of misfolded proteins in the lumen and membrane of the ER, a condition known as ER stress that is common to several diseases (Walter and Ron, 2011). For this reason, ERAD plays a key role in ER homeostasis across eukaryotes. Genetic ablation of a number of ERAD components leads to embryonic lethality in mice, also highlighting the importance of this process in cellular and organismal homeostasis (Yagishita et al., 2005 Francisco et al., 2010 Eura et al., 2012). Whether this essential function of ERAD during mouse development is due to its role in the degradation of misfolded proteins remains to be determined.

Certain folded, perfectly active proteins are also targeted by ERAD. However, their degradation is highly regulated and only occurs in the presence of a specific signal. The best-characterized regulated substrate is the 3-hydroxy-3-methylglutaryl acetyl-coenzyme-A reductase (HMGR), a key enzyme in sterol biosynthesis (Gil et al., 1985 Hampton et al., 1996 Bays et al., 2001a Song et al., 2005). Both in yeast and in mammals, HMGR degradation by ERAD is part of a feedback inhibition system critical for sterol homeostasis. Interestingly, another enzyme of the sterol biosynthetic pathway, squalene monooxygenase (Erg1 in yeast and SQLE in mammals), was recently identified as a regulated ERAD substrate (Foresti et al., 2013). The degradation of Erg1/SQLE by ERAD is again part of a feedback inhibition system to prevent the accumulation of intermediate sterol metabolites, which are toxic for cells (Foresti et al., 2013). Recent evidence shows that regulation of the synthesis of sterols and other sterol-derived metabolites by ERAD is also present in plants (Doblas et al., 2013 Pollier et al., 2013). This evolutionarily conserved role of ERAD in sterol regulation might have been one of its primordial functions.

The ERAD machinery is also exploited by certain viruses to degrade host proteins thereby escaping immune surveillance. Well characterized examples are the degradation of newly synthesized major histocompatibility complex class I (MHC I) heavy chain (Wiertz et al., 1996a) or CD4 molecules by the human cytomegalovirus or the immunodeficiency virus (HIV Fujita et al., 1997 Schubert et al., 1998), respectively. Moreover, some bacterial toxins, such as cholera, and viruses, like simian virus 40 (SV40), travel to the ER retrogradely through the secretory pathway. At the ER these toxins and viruses exploit ERAD components to reach the cytosol, where ultimately they will act (Tsai et al., 2001 Schelhaas et al., 2007 Bernardi et al., 2008).

Finally, ERAD components are also involved in the turnover of several soluble proteins in the cytoplasm and the nucleus of cells (Swanson et al., 2001 Ravid et al., 2006 Yamasaki et al., 2007). Most of these cases, however, involve only a subset of the ERAD steps and components. In sum, although a complete repertoire of substrates is not available, it is clear that misfolded proteins are not the exclusive clients of ERAD.

ERAD, linking ER quality control to cytoplasmic protein degradation

The earliest evidence for protein quality control at the ER came from observations that unassembled subunits of the T cell receptor were rapidly degraded in the cells (Lippincott-Schwartz et al., 1988). This degradation occurred independently of lysosomal proteases, leading to the proposal that the ER itself would house some uncharacterized proteolytic activity toward misfolded proteins. Then a landmark study in yeast showed that the degradation of a short-lived misfolded ER membrane protein was blocked in cells lacking Ubc6, a component of the ubiquitin conjugation machinery (Sommer and Jentsch, 1993). The ubiquitin system mediates the covalent attachment of ubiquitin, a small 76–amino acid protein, to target proteins in the cytoplasm by the sequential action of activating (E1), conjugating (E2), and ligase (E3) enzymes (Pickart, 2001). Ubiquitin-modified proteins are then recognized and degraded by the proteasome. The involvement of the ubiquitin–proteasome system in ER protein quality control was confirmed by studies on the degradation of mutant and wild-type cystic fibrosis transmembrane conductance regulator (CFTR), a large membrane protein with a complicated folding process (Jensen et al., 1995 Ward et al., 1995). Inhibition of proteasome function led to accumulation of CFTR molecules, and interestingly, a significant fraction of these was detected as ubiquitin conjugates (Jensen et al., 1995 Ward et al., 1995). Soon after, it became clear that a similar mechanism could also account for the degradation of luminal misfolded proteins such as CPY*, a mutant version of the yeast vacuolar carboxypeptidase Y and a prototype ERAD substrate (Hiller et al., 1996). Together, these papers demonstrated that aberrant proteins in the lumen and membrane of the ER are degraded in the cytoplasm where the components of the ubiquitin–proteasome system reside.

Ubiquitin ligase complexes: The hubs in ERAD

Subsequent genetic and biochemical studies, primarily in budding yeast but also in mammalian cells, identified many ERAD components and led to a general understanding of the organization of the pathway. An important realization was that the “one-size-fits-all” model does not apply to ERAD and that this pathway encompasses multiple branches with distinct specificity for different classes of misfolded proteins (Taxis et al., 2003 Vashist and Ng, 2004 Carvalho et al., 2006 Bernasconi et al., 2010 Christianson et al., 2012). However, irrespective of the branch, the same sequence of events leads to the degradation of all ERAD substrates (Fig. 1 A). The first step is the recognition of a substrate in the crowded ER environment. Then the substrate is transported across the ER lipid bilayer back into the cytoplasm, a step known as retrotranslocation. On the cytosolic side of the ER membrane, the substrate is ubiquitinated by a membrane-associated ubiquitin ligase (or E3 ligase). Subsequently, the ubiquitinated substrate is extracted from the membrane in an ATP-dependent manner and released in the cytoplasm for degradation by the proteasome. The execution of these steps is coordinated by a membrane-embedded protein complex named after the E3 ligase at its core. The canonical E3 ligases involved in ERAD are themselves multispanning membrane proteins, in which the RING (really interesting new gene) domain responsible for the ligase activity is in the cytoplasm. These E3 ligase complexes are best characterized in yeast (Fig. 1 B and Table 1) where Doa10 (Swanson et al., 2001) and Hrd1 (Bordallo et al., 1998 Bays et al., 2001a) assemble into the Doa10 and the Hrd1 complexes, respectively, each responsible for the degradation of a class of ERAD substrates (Carvalho et al., 2006).

Based on the analysis of a few model substrates, the E3 ligase complex specificity appears to be determined by the location of the misfolded lesion on a substrate relative to the ER membrane: proteins with misfolded domains in the cytoplasmic side of the membrane (ERAD-C substrates) are degraded via the Doa10 complex proteins with luminal (ERAD-L substrates) or intramembrane (ERAD-M substrates) misfolded domains are targeted to the Hrd1 complex (Fig. 1 B and Table 1 Taxis et al., 2003 Vashist and Ng, 2004 Carvalho et al., 2006). Factors involved in substrate recognition are unique to the E3 ligase complex and likely determine the substrate specificity of each ERAD branch. On the other hand, the components that act at late steps of ERAD, such as the Cdc48 ATPase complex (p97 in mammals) required for membrane extraction of ubiquitinated substrates (Bays et al., 2001b Ye et al., 2001 Jarosch et al., 2002 Rabinovich et al., 2002), are common to both E3 ligase complexes.

In mammalian cells the best-studied E3 ligases are Hrd1 and Gp78 (Table 1). They are both homologous to yeast Hrd1 but assemble distinct E3 ligase complexes that preferably target different substrates (Schulze et al., 2005 Mueller et al., 2008 Bernasconi et al., 2010 Christianson et al., 2012 Burr et al., 2013). Several more E3 ligases have been implicated in ERAD in mammalian cells (such as Rma1/Rnf5, Trc8, Rfp2, Rnf170, and Rnf185) but these are still poorly characterized. Only few substrates are known for each ligase and a preference for particular ERAD substrate classes has been difficult to infer (Claessen et al., 2012).

How are ERAD substrates recognized?

Recognition of misfolded proteins.

The commitment to degradation by ERAD occurs at the level of substrate recognition therefore, this step needs to be tightly controlled. Inefficient detection of misfolded proteins leads to their accumulation, ultimately affecting cell function (Travers et al., 2000 Jonikas et al., 2009). On the other hand, overactive ERAD would likely have its cost, with the degradation of significant amounts of folding intermediates. For this reason, substrate recognition by ERAD has to be finely balanced. This is a complex task considering the ER environment, in which a complete spectrum of protein species coexist, from newly synthesized unfolded molecules to fully folded proteins.

Folding intermediates and terminally misfolded proteins share structural similarities, for example the exposure of hydrophobic patches that are normally hidden once proteins acquire the native structure. These molecules are kept in a soluble state by binding to chaperones such as the ER Hsp70 (Kar2 in yeast and BiP in mammals), which are essential for the folding of newly synthesized polypeptides as well as for the disposal of misfolded proteins. However, chaperones on their own do not appear to determine the fate of their clients. Instead, recognition factors that are part of the E3 ligase complexes play a major role in ERAD substrate selection. For example, the recognition of ERAD-L substrates requires the luminal components of the Hrd1 complex Hrd3, Kar2 and, in the case of glycosylated substrates, the lectin Yos9 (Plemper et al., 1997, 1999 Bhamidipati et al., 2005 Kim et al., 2005 Szathmary et al., 2005 Carvalho et al., 2006 Denic et al., 2006 Gauss et al., 2006a). The yeast derlin Der1, a membrane protein of the Hrd1 complex, might also be involved in ERAD-L substrate recognition (Gauss et al., 2006b Stanley et al., 2011). Moreover, certain ERAD-M substrates appear to be recognized directly by the E3 ligase Hrd1 (Sato et al., 2009). However, the features recognized on the misfolded proteins by these ERAD factors are largely unknown.

An informative exception is the recognition of luminal misfolded N-linked glycoproteins in Saccharomyces cerevisiae (Fig. 2 A). As they enter the ER lumen, proteins are often modified at asparagine residues (in the context of the N-X-S/T sequence) with a well-defined, branched glycan moiety made up of three glucose, nine mannose, and two N-acetylglucosamine residues, Glc3–Man9–GlcNAc2 (Fig. 2 A Braakman and Hebert, 2013). This N-linked glycan is subsequently trimmed by several enzymes. Early glycan-processing enzymes such as glucosidases lead to the binding of lectins that facilitate the folding of the newly synthesized proteins. In contrast, late acting enzymes, such as the mannosidase Htm1, trigger the binding of a different lectin that engages the protein in ERAD (Jakob et al., 2001 Quan et al., 2008 Clerc et al., 2009). This difference in the kinetics of the glycan-trimming enzymes provides an opportunity for newly synthesized proteins to acquire the native conformation and traffic beyond the ER (Fig. 2 A). A long ER residency, indicative of folding problems, results in the processing of the misfolded glycoproteins by Htm1, which generates a biochemical mark (α1,6-linked mannose) decoded by the lectin Yos9, an ERAD substrate recognition factor (Quan et al., 2008 Clerc et al., 2009).

Both yeast Htm1 and its mammalian counterpart EDEM are in complex with oxidoreductases (Pdi1 in yeast, Erdj5 in mammals), required for the stability of Htm1 and also for reducing disulfide bonds in misfolded proteins, which might affect subsequent ERAD steps (Ushioda et al., 2008 Clerc et al., 2009). The binding of Yos9 to the α1,6-linked mannose is not sufficient to trigger the degradation of the misfolded protein. This processed glycan must be located in an unstructured polypeptide segment that is bound by Hrd3 (Xie et al., 2009). The delivery of substrates to Hrd3 might be facilitated by the luminal chaperone Kar2 that is essential for ERAD (Plemper et al., 1997 Denic et al., 2006 Xie et al., 2009). The dual recognition of a specific N-linked glycan by Yos9 and an unstructured segment by Hrd3 likely enhances the stringency of ERAD substrate recognition, perhaps by a kinetic proofreading mechanism (Fig. 2 A Denic et al., 2006).

The recognition mechanism of misfolded luminal N-linked glycoproteins is likely similar in mammalian cells because the yeast components are largely conserved in higher eukaryotes (Table 1). However, the situation might be more complicated. For example, OS-9 and XTP3-B, the mammalian homologues of Yos9, use their glycan-binding domain not only to interact with glycans in the misfolded protein but also with Sel1, the mammalian version of Hrd3, which is itself a glycoprotein (Christianson et al., 2008). Whether the interactions with Sel1 and the substrates occur sequentially or correspond to different pools of OS-9/XTP3-B is unclear. In either case, using the same domain to interact with a component and a substrate offers OS-9/XTP3-B an additional mechanism to regulate recognition of N-glycosylated ERAD substrates.

Recent work in yeast suggests that a different type of glycosylation, O-mannosylation, plays an important role in removing certain luminal proteins from futile folding cycles and thus favoring their degradation by ERAD after prolonged residency in the ER (Goder and Melero, 2011 Xu et al., 2013). The enzymes involved in protein O-mannosylation physically associate with ERAD machinery, but how O-mannosylated proteins are captured by the ERAD components is still unclear (Goder and Melero, 2011). Therefore, O-mannosylation is another appealing mechanism for generating an irreversible biochemical mark on proteins displaying folding problems.

A common feature between ERAD substrate recognition by N-glycan trimming and O-mannosylating enzymes is that both appear to be slow processes, requiring substrates to stay for relatively prolonged periods in the ER. Whether other mechanisms involved in recognition of misfolded proteins by ERAD also require a lag period in the ER is not known. Nevertheless, it is curious that newly synthesized proteins were shown to be protected from degradation for a period of time, even under conditions that favor their misfolding (Vabulas and Hartl, 2005). Therefore, recognition of misfolded proteins might have evolved as an intrinsically slow process, perhaps to spare some folding intermediates from prematurely engaging in ERAD.

Adaptor-mediated substrate recognition.

The degradation of specific folded proteins by ERAD is mediated by the same general machinery, but the recognition of these substrates involves distinct factors. A simple mechanism to target a native protein to ERAD is by a substrate-specific adaptor. For example, the human cytomegalovirus encodes ER membrane adaptor proteins, US2 and US11, which bind independently to newly synthesized MHC I molecules and deliver them to ERAD components for degradation. As a consequence, infected cells display less MHC I complex at their surface and escape detection by the immune system (Wiertz et al., 1996a). Despite this common outcome, the two viral proteins interact differently with ERAD components. US11 uses its transmembrane domain to recruit MHC I into a complex which contains Derlin-1 as well as Sel1L, the p97 ATPase complex and its membrane adaptor UBXD8 (Lilley and Ploegh, 2004 Ye et al., 2004 Mueller et al., 2008). Intriguingly, the E3 ligase required for US11-mediated MHC I degradation is not known and both Hrd1 and Gp78 E3 ligases, which are found in complex with Derlin-1, appear to be dispensable. US2, on the other hand, delivers its substrate to the ERAD ligase complex containing the E3 Trc8 and SPP, the signal peptide peptidase (Fig. 2 B Loureiro et al., 2006 Stagg et al., 2009). The precise function of SPP in ERAD is not known and it is not even clear whether it involves its proteolytic activity. Despite these differences, both US2 and US11 act as adaptors to deliver a specific ERAD substrate, MHC I heavy chain, to the E3 ligase complexes that promote its degradation.

A similar mechanism targets CD4 for degradation in cells expressing the HIV-encoded ER membrane protein Vpu. In this case, Vpu works not only as the substrate adaptor for CD4 but also as a scaffold to recruit a cytosolic E3 ligase complex, SCF βTrCP , required for CD4 ubiquitination (Fujita et al., 1997 Schubert et al., 1998). In vitro reconstitution of Vpu-mediated CD4 ubiquitination revealed that the specificity of adaptor-mediated substrate selection can be further increased at the level of substrate ubiquitination, which can be counteracted by the activity of de-ubiquitinating enzymes (Zhang et al., 2013). The balance between these activities helps discriminating small differences in adaptor–substrate affinity. Whether this mechanism aids the selection of other ERAD substrates is not known yet.

The strategy of using a substrate-specific adaptor is not exclusive to viral encoded proteins. Both in Drosophila and in mammalian cells, the derlin-related iRhom proteins function as adaptors in the ERAD-mediated degradation of EGFR ligands as they traffic through the ER (Zettl et al., 2011). Elegant genetic experiments in flies showed that this mode of regulated ERAD was important to control sleeping behavior that requires EGFR signaling (Zettl et al., 2011). It is likely that more substrate-specific adaptors will be identified as our knowledge of the mechanisms of regulated ERAD expands.

Signal-mediated substrate recognition.

A substrate-specific adaptor also functions in the regulated ERAD of HMGR, a key enzyme of the sterol biosynthetic pathway. In this case the adaptor, either Insig-1 or Insig-2, does not bind constitutively to HMGR (Song et al., 2005). Instead, the interaction only occurs in the presence of 24,25-dihydrolanosterol, an intermediate metabolite in sterol biosynthesis. Under low sterols levels HMGR is a stable protein, actively producing sterol precursors (Fig. 2 C). On the other hand, high sterol synthesis leads to a rise in 24,25-dihydrolanosterol concentration, which favors the binding of HMGR to one of the Insig proteins, its delivery to an E3 ligase complex, and consequently its degradation by the proteasome (Fig. 2 C). Whereas Gp78 and the Trc8 were originally implicated in HMGR regulated ERAD, recent data suggest that additional E3 ligases might also be involved (Song et al., 2005 Lee et al., 2010 Jo et al., 2011 Tsai et al., 2012). Degradation of HMGR by ERAD results in reduced flux through the sterol biosynthetic pathway and reestablishment of membrane lipid homeostasis.

Interestingly, Insig-1 (but not Insig-2) is itself subjected to reciprocal sterol-regulated ERAD (Lee et al., 2006). Depletion of cellular sterols stimulates Insig-1 ubiquitination by the E3 Gp78 ligase complex. Conversely, if sterol levels are high Insig-1 binds to SCAP, another key ER membrane protein required for sterol homeostasis, leading to a much longer Insig-1 half-life. These data illustrate the complex interplay between the opposing effects of sterols on the stability of the HMGR enzyme and one of its adaptors for regulated degradation by ERAD.

In yeast, sterol homeostasis also involves negative feedback of an HMGR homologue, Hmg2 (Hampton et al., 1996). Like in mammals, degradation of yeast Hmg2 is controlled by the Insig-like proteins Nsg1 and Nsg2 and requires the E3 ligase Hrd1 (Bays et al., 2001a Gardner et al., 2001 Flury et al., 2005). In fact, Hrd1 was originally identified in a genetic screen for mutants defective in HMGR degradation (HRD genes Hampton et al., 1996). The binding of Hmg2 to Nsg1 is also modulated by sterol levels (Theesfeld and Hampton, 2013). However, in contrast to mammalian cells, the binding of Nsg1 promotes Hmg2 stability, indicating that the recognition of this substrate is mechanistically different in the two systems (Flury et al., 2005 Theesfeld and Hampton, 2013). Based on limited proteolysis experiments, it has been proposed that Nsg1 and Nsg2 work as Hmg2-specific chaperones and that in their absence Hmg2 presents sufficient conformation instability to engage in ERAD as a misfolded protein (Flury et al., 2005 Shearer and Hampton, 2005). This degradation is further accelerated by high concentrations of an early sterol-intermediate metabolite, geranylgeranyl pyrophosphate (Theesfeld and Hampton, 2013). Therefore, a change in the affinity to a binding partner is another strategy to target a protein for ERAD in a signal-dependent manner.

Squalene monooxygenase (SQLE), another enzyme of the sterol biosynthetic pathway, is also targeted by regulated ERAD both in yeast and in mammals (Foresti et al., 2013). SQLE degradation requires the yeast Doa10 E3 ligase complex or its mammalian homologue Teb4, indicating that two independent branches of ERAD control distinct steps in sterol biosynthesis (Foresti et al., 2013). Although the mechanism for the recognition of SQLE by ERAD machinery is still not known, it is clear that Insigs are dispensable for this recognition, both in mammals and in yeast (Gill et al., 2011 Foresti et al., 2013). In mammals, the N-terminal domain of SQLE is necessary and sufficient for cholesterol-dependent degradation (Gill et al., 2011). Whether this domain binds directly to cholesterol or interacts with an ERAD-specific adaptor is not known. The mechanism for SQLE recognition by ERAD in yeast is likely to be different because this N-terminal domain is only conserved among certain animals.

Based on these few examples, it is clear that signal-mediated ERAD depends on the ability of cells to sense the concentration of some lipids in their membranes and on specific adaptors to selectively degrade key enzymes. Our knowledge on the mechanisms by which other classes of regulated ERAD substrates are recognized will grow as more of these are identified.

Shipping out the trash: Substrate retrotranslocation and cytoplasmic events in ERAD

After being selected, ERAD substrates are retrotranslocated across the ER membrane back into the cytoplasm. In the case of misfolded luminal proteins the complete polypeptide needs to be retrotranslocated, whereas for membrane substrates this step requires the transport of only certain domains. As a consequence, ubiquitination of luminal substrates only occurs at late stages of retrotranslocation, once a portion of the substrate has been exposed to the cytoplasm. In contrast, ubiquitination of most, but not all (Burr et al., 2013), membrane substrates is coupled to their retrotranslocation.

In analogy to the transport of newly synthesized proteins into the ER or mitochondria, it has been postulated that retrotranslocation occurs through a protein-conducting channel. However, the identity of the retrotranslocation channel has been at the center of an intense debate that is almost as old as the research in this field. Over the years, several channel candidates have been proposed but it has been difficult to gather definitive evidence in support of any of them. The Sec61 translocon used for protein import into the ER was the first proposed retrotranslocation channel (Wiertz et al., 1996b). Sec61 was found to interact with ERAD substrates both in yeast and in mammalian cells as well as with the yeast proteasome (Wiertz et al., 1996b Kalies et al., 2005 Scott and Schekman, 2008). However, the significance of these associations is not clear. Recent work showed that proteins engaging the Sec61 translocon aberrantly or persistently in their way into the ER become substrates of the Hrd1 ligase complex, which might explain the interaction between Sec61 and some ERAD substrates (Rubenstein et al., 2012). In addition, certain yeast sec61 mutants displayed defects in degrading model ERAD substrates, even under conditions in which general “forward” translocation appeared not to be dramatically affected (Pilon et al., 1997 Plemper et al., 1997 Willer et al., 2008). It remains to be determined whether this phenotype is caused by a specific impairment in retrotranslocation.

The E3 ligase complexes interact with ERAD substrates immediately before and after their retrotranslocation, indicating this step occurs in their immediate vicinity. Therefore, multispanning membrane proteins within the E3 ligase complexes have also been seen as good candidates to mediate retrotranslocation (Ye et al., 2001 Lilley and Ploegh, 2004 Kreft et al., 2006 Horn et al., 2009 Carvalho et al., 2010 Mehnert et al., 2014). These include the E3 ligases themselves as well as proteins of the Derlin family (Der1 in yeast), which are essential for the degradation of all luminal ERAD substrates but whose function has remained elusive. In vitro experiments using mammalian-derived microsomes loaded with the yeast ERAD substrate mutant pro-α-factor indicate that Derlin might be involved in retrotranslocation (Wahlman et al., 2007). In this simplified system, in which synthesis and retrotranslocation were uncoupled, the substrate cross-linked to Derlin but not to Sec61. Moreover, its retrotranslocation was blocked by anti-Derlin antibodies whereas antibodies directed to Sec61 had no effect (Wahlman et al., 2007). It should be noted that mutant pro-α-factor is a noncanonical substrate because its retrotranslocation does not require ubiquitination. Interactions between the yeast Der1 and the prototype ERAD-L substrate CPY* were also detected by site-specific photocrosslinking in yeast (Carvalho et al., 2010 Stanley et al., 2011 Mehnert et al., 2014). These cross-links were seen even in cells lacking the substrate recognition factors Hrd3 and Yos9, and were increased if conserved polar residues in the membrane domain of Der1 were mutated. An interpretation of these results is that Der1 mediates the transfer of substrates from the recognition factors Hrd3/Yos9 to the Hrd1 ligase inside the membrane (Mehnert et al., 2014).

A compelling piece of evidence for a function during substrate retrotranslocation was reported for the E3 ligase Hrd1 (Fig. 3 Carvalho et al., 2010). Although commonly working in the context of the Hrd1 complex, overexpressed Hrd1 can mediate the degradation of ERAD-L substrates even in the absence of its membrane partners Hrd3, Der1, and Usa1 (Plemper et al., 1999 Denic et al., 2006 Carvalho et al., 2010). Under these conditions Hrd1 selectivity for misfolded proteins is lost, suggesting that the other subunits of the complex are critical to control Hrd1 activity and substrate specificity (Denic et al., 2006). Hrd1 interacts with a sizeable region of a modified version of CPY* during the early stages of retrotranslocation, as assayed by site-specific photocrosslinking (Carvalho et al., 2010). Importantly, the interaction likely occurs inside of the ER bilayer because it is lost if substrate recognition is blocked or if the transmembrane segments of Hrd1 are mutated. Hrd1 contains only six transmembrane segments therefore, ERAD-L substrate retrotranslocation requires Hrd1 oligomerization, which normally is facilitated by Usa1 but can occur spontaneously upon Hrd1 overexpression (Horn et al., 2009 Carvalho et al., 2010). All these data make a strong case for a direct role of Hrd1 in the retrotranslocation of ERAD-L substrates, but the possibility that it works with some partner(s), such as Der1, Sec61, or other unknown factors cannot be excluded at this point. Moreover, it is not known whether this is a unique feature of Hrd1 or whether the membrane domains of other E3 ligases also participate in the retrotranslocation of other classes of ERAD substrates.

In most of these cross-linking experiments the retrotranslocation of CPY* was dampened by fusing it to a very tightly folded domain, indicating that ERAD-L substrates need to be unfolded before this step (Bhamidipati et al., 2005 Carvalho et al., 2010). Whether unfolding is also a prerequisite for the retrotranslocation of other classes of ERAD substrates is not settled yet (Fiebiger et al., 2002 Tirosh et al., 2003).

At the cytoplasmic side of the ER membrane, substrates are ubiquitinated, a modification that allows their recognition by the Cdc48/p97 ATPase complex composed of a homohexamer of Cdc48/p97 and by the cofactors Ufd1 and Npl4 (Bays et al., 2001b Ye et al., 2001, 2003 Jarosch et al., 2002 Rabinovich et al., 2002). The recruitment of this ATPase complex to the ER membrane is facilitated by ubiquitin regulatory X (UBX) domain–containing proteins, Ubx2 in yeast and UBXD8 in mammals (Neuber et al., 2005 Schuberth and Buchberger, 2005 Mueller et al., 2008). The ATPase activity of the Cdc48/p97 complex provides the driving force to move ubiquitinated substrates out of the membrane into the cytosol (Ye et al., 2003). Although the role of Cdc48/p97 in this process is well established, the mechanism that couples ATP hydrolysis to membrane extraction of the substrate is still not understood. In addition to the Cdc48/p97 complex, the ATPase subunits of the proteasome regulatory particle were also shown to play a role in retrotranslocation of some ERAD substrates (Lipson et al., 2008). The driving force for the retrotranslocation of the few non-ubiquitinated substrates, like the cholera toxin or pro-α-factor, is not known (Kothe et al., 2005 Moore et al., 2013).

The Cdc48/p97 complex also serves as a platform for other ubiquitin-modifying enzymes such as de-ubiquitinating enzymes (DUBs Rumpf and Jentsch, 2006 Jentsch and Rumpf, 2007 Ernst et al., 2009). Although the role of some of these Cdc48/p97-binding factors is not clear yet, it was shown that interfering with the DUBs YOD1 and ataxin-3 affected substrate retrotranslocation (Wang et al., 2006 Ernst et al., 2009). The requirement for DUB activity during retrotranslocation suggests that the process involves cycles of ubiquitination and de-ubiquitination. In some cases these cycles might be important to increase the specificity of substrate recognition, as was shown for the Vpu-mediated degradation of CD4 (Zhang et al., 2013). Interestingly, the retrotranslocation of some noncanonical substrates like cholera toxin, which are not ubiquitinated, is also affected by manipulation of both E3 ligase and DUB activities (Hassink et al., 2006 Bernardi et al., 2013). These results suggest that retrotranslocation requires the ubiquitination of some factors other than the substrates. Future studies should test some obvious candidates such as the components of the E3 ligase complexes themselves.

There is some recent evidence that the yeast Cdc48 complex might be acting also much earlier, during the initial stages of retrotranslocation of an ERAD-L substrate, before it is exposed to the cytoplasm (Fig. 3). Using a cross-linking strategy, it was shown that the interaction between Hrd1 and an early retrotranslocation intermediate was lost in mutants of the Cdc48 complex (Carvalho et al., 2010). Interestingly, a similar defect was detected in Hrd1 mutants impaired for E3 ligase activity (Carvalho et al., 2010). Based on these observations it was proposed that in early stages of ERAD-L substrate retrotranslocation, Hrd1 induces the ubiquitination of an ERAD component, perhaps Hrd1 itself. This ubiquitination signals the recruitment of the Cdc48 complex, which upon ATP hydrolysis would induce changes in the conformation or in the oligomeric status of Hrd1, resulting in substrate retrotranslocation. This model is consistent with the well-established role of Cdc48/p97 in the disassembly and remodeling of protein complexes (Jentsch and Rumpf, 2007). Once the substrate emerges on the cytosolic side and is ubiquitinated by Hrd1, the ATPase complex binds to it and promotes the final stages of its retrotranslocation. Although many aspects of this model still wait for experimental support, it would provide a unifying role for the Cdc48/p97 ATPase as the driving force for substrate retrotranslocation in ERAD (Fig. 3).

After being released from the membrane, substrates are kept soluble and transferred to the proteasome by cytosolic chaperones such as the BAG6 complex (Claessen and Ploegh, 2011 Wang et al., 2011 Xu et al., 2012) or shuttle factors like Rad23 and Dsk2 (Medicherla et al., 2004). The long journey of ERAD substrates ends with their degradation by the proteasome.

Conclusions and future perspectives

In recent years there has been tremendous progress in understanding ERAD. The identification of most of the components involved in this process and how these are pieced together and organized in the different ERAD branches were important achievements. A major challenge for the future is to reveal the mechanistic aspects of the pathway. Such developments should, for example, help in discerning the basis by which misfolded proteins are recognized in each of the different ERAD branches.

The mechanisms of substrate retrotranslocation and the roles played by the different components, such as the E3 ligases and the Cdc48/p97 complex, will certainly be another interesting area to follow. However, progress on these topics might require the development of new approaches, such as in vitro systems with purified components recapitulating individual ERAD steps.

In early days, ERAD was perceived as a process mainly dedicated to ER protein quality control. The picture now emerging places ERAD closer to other ubiquitination systems in the cytoplasm and nucleus, which control the turnover of specific proteins to achieve a certain physiological state. Therefore, another major challenge for the coming years is the detailed characterization of the roles of ERAD beyond quality control. Although the central role of ERAD in sterol homeostasis is unequivocal, it will be important to clarify whether ERAD has a more general role in the regulated degradation of folded ER proteins and in that way modulates other ER-related functions. A systematic and rigorous identification of regulated ERAD substrates should help in addressing these issues. Uncovering the intersections of ERAD with other cellular pathways will provide important insights into the mechanisms of ER and cellular homeostasis.


Hepatitis C virus plays a significant role in the development of hepatocellular carcinoma (HCC) globally. The pathogenic mechanisms of hepatocellular carcinoma with HCV infection are generally linked with inflammation, cytokines, fibrosis, cellular signaling pathways, and liver cell proliferation modulating pathways. HCV encoded proteins (Core, NS3, NS4, NS5A) interact with a broad range of hepatocytes derived factors to modulate an array of activities such as cell signaling, DNA repair, transcription and translational regulation, cell propagation, apoptosis, membrane topology. These four viral proteins are also implicated to show a strong conversion potential in tissue culture. Furthermore, Core and NS5A also trigger the accretion of the β-catenin pathway as a common target to contribute viral induced transformation. There is a strong association between HCV variants within Core, NS4, and NS5A and host single nucleotide polymorphisms (SNPs) with the HCC pathogenesis. Identification of such viral mutants and host SNPs is very critical to determine the risk of HCC and response to antiviral therapy. In this review, we highlight the association of key variants, mutated proteins, and host SNPs in development of HCV induced HCC. How such viral mutants may modulate the interaction with cellular host machinery is also discussed.

Characteristic structural features

The RHD consists of two hydrophobic regions, each 28-36 amino acids long, which are thought to be membrane-embedded regions, separated by a hydrophilic loop of 60-70 amino acids, and followed by a carboxy-terminal tail about 50 amino acids long (Figure 2a). Although much amino-acid identity has been lost over the course of evolution, the overall structure of the RHD has been preserved from plants to yeasts to humans. This suggests that three-dimensional protein structure is of greater importance than individual residues for RHD function. The RHD hydrophobic regions are unusually long for transmembrane domains: each spans approximately 30-35 amino acids, whereas most transmembrane domains are about 20 amino acids in length. This raises the interesting question of whether this longer length has significance for reticulon function. The topology of these hydrophobic regions within membranes is so far only partially defined.

The structure and membrane topology of reticulons. (a) Structure of reticulon proteins. Numbers refer to the exons that encode the protein regions. Black ovals represent hydrophobic regions. GenBank accession numbers are as in Figure 1. (b) Possible topologies of reticulon proteins in membranes. Although eight or more conformations are possible, only those for which evidence exists are depicted. Different topologies in different cell types and different membranes may enable reticulons to carry out diverse roles in the cell.

Reticulon topology

The RHD loop region has been detected both on the surface of cells and intracellularly, and it has been suggested that the RHD hydrophobic regions might either span the ER membrane or plasma membrane completely or might double back on themselves to form a hairpin (Figure 2b). Antibodies against the amino-terminal domain of RTN4 bind to the surface of chick oligodendrocytes in live spinal cord explants [8] and cultured oligodendrocytes interact specifically with both amino-terminal domain-specific antibodies and antibodies directed against the RTN4 66-amino-acid loop (66-loop) [16]. These findings suggest that the amino terminus and the 66-loop project into extracellular space, and therefore that the first RHD hydrophobic region must double back on itself in the membrane. However, other data suggest that the amino-terminal domain is intracellular. Antibodies against the 66-loop region of RTN4 detect small amounts of this epitope on the surface of live COS-7 cells, but antibodies against c-Myc tags fused to either the amino or the carboxy terminus do not bind to live cells [8].

More recent data from non-neuronal cells in which RTN4 is overexpressed strongly support a third model, in which most of both the amino-terminal domain and the 66-loop are cytoplasmic. In COS cells treated with maleimide polyethylene glycol, cysteines in the amino-terminal domain and the loop regions of ER-associated RTN4 were found to be modified by the reagent after detergent disruption of the plasma membrane but not the ER membrane [6]. Cysteines in the carboxy-terminal region were only partially modified. All these results suggest that mammalian reticulons might have different topologies in the ER and plasma membranes such multiple conformations may enable them to carry out multiple roles in the cell. Another protein with multiple membrane topologies is the mammalian prion protein (PrP) overexpression of a certain transmembrane form of the prion protein, Ctm PrP, causes neurodegenerative disease distinct from that caused by the natural pathogenic prion form PrP Sc [19, 20]. Another possibility is that reticulons assume different topologies in different cell types: the reticulon amino-terminal region has been detected only in the cytoplasm in COS-7 cells, but has been found on both the surface and in the cytoplasm of oligodendrocytes. Again, this may reflect the diverse roles of reticulon proteins in different cell types.

Reticulon tertiary structure

The solution structure of the RHD loop of RTN4, known as Nogo66, has recently been probed by circular dichroism (CD) and nuclear magnetic resonance (NMR). Nogo66 is soluble in pure water and consists of three alpha helices, two short flanking one long, spanning residues 6-15, 21-40 and 45-53, followed by the unstructured residues 55-60 [21, 22]. The Nogo66 loop is involved in several RTN4-specific signaling cascades, including interaction with the Nogo receptor (NogoR) to inhibit neurite outgrowth [23], and with the cell adhesion molecule contactin-associated protein (Caspr) [24] to mediate the localization of potassium channels at axonal paranodes. The human RTN1 and RTN3 66-loops share 71% and 63% identity with the RTN4 loop mouse RTN1 and RTN3 identity with human RTN4 is 67% and 59%. Despite this high degree of identity, the RTN1 66-loop does not bind to NogoR, and the function of the 66-loops in RTN1 and RTN3 is unknown in both mammals and lower organisms.

As mentioned above, the amino-terminal regions of different reticulons are highly divergent in sequence. The amino-terminal domains of the human RTN4 isoforms appear to be highly unstructured, even under physiological conditions. In silico analysis and measurements by CD and NMR of the human isoforms RTN4A and RTN4B reveal a high degree of disorganization, with only short alpha helices and beta sheets that exist transiently [25]. Recent studies have shown that intrinsically unstructured proteins (IUPs) are more likely to form multiprotein complexes than are proteins with stable tertiary structure [26], are better able to 'moonlight' - carry out alternative functions [25] - and may fold upon binding to their partners [27]. It has been shown that up to 33% of eukaryotic proteins contain long disordered regions, compared with 2% of archeal proteins [25]. The characterization of RTN4 as an intrinsically unstructured/disordered protein may explain its involvement in many physiological processes, as explained below.

VII. Peroxisomal solute transporters

A number of metabolic pathways span peroxisomes and other subcellular compartments metabolites and cofactors must therefore move across the peroxisomal membrane, and at least some of these movements should rely on membrane transporters. To date, several peroxisomal membrane proteins have been identified to specifically transport β-oxidation substrates (i.e. fatty acids, OPDA, CA and IBA) and large cofactors (i.e. ATP, NAD + and CoA (Charton et al., 2019 )).

The Arabidopsis full-size ABC transporter PXA1/CTS/PED3 can cleave acyl-CoA and import free FAs into the peroxisomal matrix. PXA1 also interacts with long-chain acyl-CoA synthetases LACS6 and LACS7, which reactivate free FA to acyl-CoA to feed into β-oxidation (De Marcos Lousa et al., 2013 ). In addition, PXA1 transports precursors for the biosynthesis of IAA, JA, BA and the benzenoid moiety of ubiquinone (Figs 5 & 6, 5 & 6) (Block et al., 2014 Li et al., 2016 ). PXA1 is also important in acetate metabolism, as a loss-of-function PXA1 mutant named acn2 (acetate non-utilizing 2) is compromised in metabolizing acetate in seedlings (Hooks et al., 2007 ). PXA1 interacts with CGI-58, a regulator of lipid metabolism and signaling (Park et al., 2013 ). Consistent with its broad function, PXA1 knockout mutants have defects in lateral root development, seed dormancy and germination, fertilization and leaf necrosis (Li et al., 2016 ).

Besides transporters, Acyl-CoA-binding proteins (ACBPs) can also mediate lipid transfer across membranes (Xiao & Chye, 2011 ). Arabidopsis ACBPs have not been found in the peroxisome, but rice ACBP6 is peroxisomal and its overexpression partially recovers the β-oxidation defects of pxa1, suggesting that rice ACBP6 can contribute to the import of FA into peroxisomes for degradation (Meng et al., 2014 ). Whether this divergence in ACBP function between Arabidopsis and rice is due to different metabolic features of monocot and dicot species remains to be determined.

Arabidopsis PNC (PNC1 and PNC2) and PXN proteins are peroxisomal ATP and NAD + carriers, respectively. PNC1 and PNC2 can catalyze the counter-exchange of ATP with ADP or AMP and complement yeast mutants deficient in peroxisomal ATP import. The pnc mutants are impaired in β–oxidation, suggesting that the proteins are essential for supplying peroxisomes with ATP (Linka et al., 2008 ). PXN can transport many substrates in vitro, including NAD + , NADH, AMP, ADP, CoA and acetyl-CoA (Agrimi et al., 2012 Bernhardt et al., 2012 ), and contributes to optimal fatty acid degradation during seedling establishment, and photorespiration under fluctuating and high light conditions (Bernhardt et al., 2012 J. Li et al., 2019a ). However, exogenously expressed AtPXN in yeast strains can import NAD + into peroxisomes in exchange for AMP but cannot transport CoA or mediate NAD + /NADH exchange (van Roermund et al., 2016 ).

Non-selective peroxisomal membrane channels may allow the passage of small solutes, such as organic acids. Yeast Pex11 has a pore-forming function, but whether plant PEX11s also possess this function has not been reported (Mindthoff et al., 2016 ). Several additional peroxisomal membrane proteins, such as PMP22 (peroxisomal membrane protein of 22 kDa), CDC (Ca 2+ -dependent carrier) and SMP2 (short membrane protein 2), also have potential pore-forming activities (summarized in Pan & Hu, 2018a ).

Function of ER in reviewing mutated proteins - Biology

Cells: The Basic Unit of Life

Cell Theory: All known living things are made up of cells. All cells come from preexisting cells by division. The cell is structural and functional unit of all living things.

Cell Structural Overview: The major parts of a cell are the nucleus, cytoplasm, and cell membrane.

  • The nucleus contains a nucleolus and is separated from the cytoplasm by the nuclear envelope.
  • The nucleus contains the cell’s DNA, a type of nucleic acid.
  • The nucleolus is like a “tiny nucleus” inside the actual nucleus. It contains RNA, a type of nucleic acid.
  • The nucleus communicates through holes in the envelope called nuclear pores.
  • The nucleus decides what the cell needs and uses DNA to print out instructions for the rest of the cell to produce that need.


  • Hold the cell’s DNA in the nucleus.
  • The nucleus contains genetic information in the form of DNA (the universal genetic code).
  • The DNA does not hang around loosely in the nucleus. The DNA is packaged with proteins and wound up.
  • Recall that the role of nucleic acids is to carry genetic information, which is inherited by an organism’s offspring.
  • These wound up DNAprotein structures are called chromosomes.

Cytoplasmic Organelles: Are compartmentalized structures that perform a specialized function within a cell.

Golgi apparatus: ships packages around the cell.

  • The golgi is made up of flattened, folded sacs.
  • Packages (e.g. containing proteins) are carried to the golgi in vesicles.
  • The golgi receives an incoming vesicle, tags the package, and sends the vesicle to its final destination.
  • Lysosomes contain an environment made to destroy waste.
  • Vesicles carry the waste (bacteria, old organelles, etc.) into the lysosome.
    Once inside, the waste is destroyed and its parts recycled.
  • Smooth ER is NOT attached to the nucleus and DOES NOT have attached ribosomes (thus smooth).
  • Smooth ER synthesizes carbohydrates (sugars) and lipids (fats).

Mitochondria: produce energy to power the cell.

  • The mitochondria convert carbohydrates (sugar) taken from food into ATP.
  • The mitochondria are unique in that it has two protective shells.
  • The ribosome reads the DNA strand instructions to make proteins for the cell to use in its normal activities.
  • The units clasp around a strand of nucleic acid instructions from the nucleus.
    Each ribosome is made of two protein subunits.

Rough endoplasmic reticulum: The two types of ER make different building blocks for the cell.

  • Rough ER is found attached to the outside of the nucleus. It appears rough because of the ribosomes on its surface.
  • Rough ER helps the attached ribosomes in finishing protein synthesis.
  • Plasma Membrane, the cell’s membrane is made of phospholipids, which have carbohydrate heads and lipid tails.
  • Embedded proteins are anchored to the cell membrane.
  • Exterior of the plasma membrane touches water polar heads touch water on the inside of the cell and water on the outside of the cell.
  • Interior Blocks Passage However, water and other molecules cannot pass through to either side because of the nonpolar tails.
  • Provides a stabilized environment, which protects and maintains the cell’s internal environment, separate from the environment outside.
  • Proteins embedded into the membrane send and receive signals to communicate with other cells.

Three types of passive transport are osmosis, diffusion, and facilitated diffusion. Osmosis is the natural movement of water from a high concentration of water to a lower concentration of water. Diffusion is the natural movement of molecules from a higher concentration to a lower concentration. Facilitated Diffusion is the natural movement of molecules from a higher concentration to a lower concentration with the help of a transporter protein embedded on the cell membrane.

Active transport requires energy to occur. Active transport is “forced” movement of molecules from a lower concentration to a higher concentration. The most common type of active transport is a pump. Pumps are proteins embedded in the cell membrane, which use ATP energy to work.

Different Cell Types: Prokaryote and Eukaryote.

  • Prokaryotic: Bacteria and other microscopic organisms are made up of prokaryotic cells. Prokaryotic cells do not have any complex organelles (not even a nucleus). However, prokaryotes do have ribosomes.
  • Eukaryotic: Two types of eukaryotic cells are plant and animal cells.

The cell contains a nucleus, which contains the genetic material necessary for reproduction. Within the cytoplasm of the cell are the organelles the cell requires to reproduce, energy production, and removal of waste.

Key concepts about how cells obtain and import the necessary nutrients for survival along with the energy requirements of these processes will be presented.

Specific Tutorial Features:

  • Detailed description of the function of each organelle within cells is discussed.
  • The role of the nucleus as a command center will be covered along with the location of the cellular DNA within chromosomes.
  • Concept map showing inter-connections of new concepts in this tutorial and those previously introduced.
  • Definition slides introduce terms as they are needed.
  • Visual representation of concepts.
  • Examples given throughout to illustrate how the concepts apply.
  • A concise summary is given at the conclusion of the tutorial.

The definition of a cell: The smallest unit of an organism that can live independently.
The nucleus of the cell:

  • Nucleus
  • Nucleolus
  • Nuclear envelope
  • Chromsomes
  • Golgi apparatus
  • Lysosome
  • Smooth endoplasmic reticulum
  • Mitochondria
  • Nucleus
  • Ribosomes
  • Rough endoplasmic reticulum
  • Provides a stable internal cell
  • Transport across the cell
  • Prokaryotic vs. Eukaryotic
  • Cell Levels of Organization.

See all 24 lessons in Anatomy and Physiology, including concept tutorials, problem drills and cheat sheets: Teach Yourself Anatomy and Physiology Visually in 24 Hours


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Kinase, an enzyme that adds phosphate groups (PO4 3− ) to other molecules. A large number of kinases exist—the human genome contains at least 500 kinase-encoding genes. Included among these enzymes’ targets for phosphate group addition ( phosphorylation) are proteins, lipids, and nucleic acids.

For protein targets, kinases can phosphorylate the amino acids serine, threonine, and tyrosine. The reversible phosphorylation of proteins, by the antagonistic (opposing) action of kinases and phosphatases, is an important component of cell signaling because the phosphorylated and unphosphorylated states of the target protein can have different levels of activity. Edwin Gerhard Krebs and Edmond H. Fischer received the Nobel Prize for Physiology or Medicine in 1992 for their work in establishing this paradigm.

Phosphorylation of lipid molecules by kinases is important for controlling the molecular composition of membranes in cells, which helps to specify the physical and chemical properties of the different membranes. Inositol, a compound similar in structure to a carbohydrate, is phosphorylated by kinases to create a diverse array of phosphoinositol and phosphoinositide lipids. These molecules then function as second messengers to propagate signaling information throughout the cell.

Nucleotides, the fundamental units of RNA (ribonucleic acid) and DNA (deoxyribonucleic acid), contain a phosphate molecule attached to a nucleoside, a compound made up of a ribose moiety and a purine or pyrimidine base. In polymers of RNA and DNA, the backbone is composed of repeating phospho-ribose units. Kinases attach the phosphate to the nucleoside, creating a nucleotide monophosphate. For example, an enzyme called nucleoside phosphorylase serves this role when cells switch to synthesizing nucleotides from recycled purines instead of from new starting materials. Mutations in the gene encoding nucleoside phosphorylase can cause a severe form of immune deficiency.

Metabolism of dietary sugars ( glycolysis) involves several different steps of phosphorylation by distinct kinases. These phosphate groups are ultimately used to form the high-energy compound known as ATP (adenosine triphosphate).

Inhibitors of kinases can be important treatments for human diseases in which hyperactive processes need to be dampened. For example, one form of human leukemia, CML ( chronic myelogenous leukemia), is caused by excess activity of the Abelson tyrosine kinase. Imatinib (Gleevec) is a chemical that binds to the active site of this kinase, thereby blocking the enzyme’s ability to phosphorylate targets. Imatinib has been useful in the initial treatment of CML however, in many cases the kinase enzyme mutates, rendering the drug ineffective.


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