Molecular Membrane Biology, 2010; Early Online, 1–16

Sorting things out through endoplasmic reticulum quality control

TAKU TAMURA1, JOHAN C. SUNRYD1,2 & DANIEL N. HEBERT1,2

1Department of Biochemistry and Molecular Biology, and 2Program in Molecular and Cellular Biology, University ofMassachusetts, Amherst, MA, USA

(Received 29 April 2010; and in revised form 13 May 2010)

AbstractThe endoplasmic reticulum (ER) is a highly organized and specialized organelle optimized for the production of proteins. It iscomprised of a highly interconnected network of tubules that contain a large set of resident proteins dedicated to thematuration and processing of proteins that traverse the eukaryotic secretory pathway. As protein maturation is an imperfectprocess, frequently resulting in misfolding and/or the formation of aggregates, proteins are subjected to a series of evaluationprocesses within the ER. Proteins deemed native are sorted for anterograde trafficking, while immature or non-native proteinsare initially retained in the ER in an attempt to rescue the aberrant products. Terminally misfolded substrates are eventuallytargeted for turnover through the ER-associated degradation or ERAD pathway to protect the cell from the release of adefective product. A clearer picture of the identity of the machinery involved in these quality control evaluation processes andtheir mechanisms of actions has emerged over the past decade.

Keywords: Protein folding, carbohydrates, molecular chaperones, ERAD

Introduction

Ribosome-attached secretory pathway cargo are co-translationally targeted to the endoplasmic reticulum(ER) translocon Sec61 by the signal recognitionparticle or SRP. Nascent chains co-translationallyemerge into the ER lumen through this ERmembranepore in a vectorial manner (N- to C-terminus). Asprotein maturation begins co-translational and trans-locationally, the vectorial nature of these processeshelps to taper the ensemble of possible intermediatesby starting maturation with shorter nascent chains.These early events, which are assisted by a groupof factors localized in close proximity to the Sec61entry site, include protein folding, processing andmodification. The majority of the secretory path-way customers are modified by N-linked glycans[Apweiler et al. 1999], and these modifications areused extensively as maturation and quality controltags that assist with the sorting process. This reviewwill focus on the machinery and mechanics for howthese sorting decisions are mediated with the help ofN-linked glycans and their related factors.

Getting started: Translocation and earlymaturation events

Proteins containing the consensus N-linked glyco-sylation site Asn-X-Ser/Thr are co-translationallymodified with a pre-assembled 14-member carbo-hydrate comprised of three glucoses, nine mannosesand two N-acetylglucosamines (Glc3Man9GlcNAc2;Figure 1A) through the action of the oligosaccharyltransferase (OST). The addition of these large bulkyhydrophilicmodificationsgreatlyalters the fundamentalproperties of the protein. The enzymatic transfer of theglycan requires that theSerorThr, situated tworesiduesC-terminal to the Asn attachment site, position theirhydroxyl side chain to render theAsnamide groupmorenucleophilic to support the progression of the transferreaction. Therefore, this reactionmechanism favors thetransfer of glycans that are situated on flexible regions.These regions are frequently located on the aqueousexposed surface of the protein. This optimally positionsglycans for their role as maturation and quality controltags to support the recruitment of protein folding anddegradation factors.

Correspondence: Prof. Daniel N. Hebert, Department of Biochemistry and Molecular Biology, University of Massachusetts, 710 N. Pleasant St., Amherst, MA01003, USA. Tel: +1 (413) 545 0079. Fax: +1 (413) 545 3291. E-mail: dhebert@biochem.umass.edu

ISSN 0968-7688 print/ISSN 1464-5203 online ! 2010 Informa UK LtdDOI: 10.3109/09687688.2010.495354

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The composition of the carbohydrate tag is con-trolled by a series of glycosidases and glycosyltrans-ferases that line the secretory pathway (Table I). In theER, glucosidase I initially removes terminal glucoseresidues creating diglucosylated (Glc2Man9GlcNAc2)side chains, which have recently been shown to sup-port the recruitment of the ER-resident protein mal-ectin [Schallus et al. 2008]. Malectin was identified asa protein immunoisolated in aquaporin-2 containingvesicles, suggestive of it possibly playing a role in itsmaturation [Barile et al. 2005]. As glucosidase I cleav-age occurs rapidly co-translationally [Chen et al.1995], this implies that malectin acts early in thematuration process perhaps during translocation(Figure 2). However, further studies are required tounderstand the spectrum of malectin substrates, thetiming and role in its assistance in cargo maturation.The second glucose is trimmed by glucosidase II

to generate monoglucosylated glycans. These carbo-hydrates serve as attachment sites for the lectinchaperones calnexin, a type I membrane protein,

and its soluble paralogue calreticulin [Ou et al.1993, Peterson et al. 1995]. The action of glucosi-dase II also occurs co-translationally supporting earlyco-translational binding to the lectin chaperones[Chen et al. 1995, Hebert et al. 1997, Deprez et al.2005] (Figure 2). Calnexin and calreticulin bindingserves a number of roles. First, as both chaperones areER retained through ER retention signals, binding toimmature proteins keeps client proteins in the ER[Rajagopalan et al. 1994]. Second, both chaperonesare associated with an oxidoreductase ERp57, sobinding recruits an oxidoreductase to the maturingchain that can assist in disulfide formation, reductionor rearrangement [Oliver et al. 1997, Zapun et al.1998, Solda et al. 2006]. Third, chaperone bindingconstrains or delays the folding of a protein in a region-specific manner; therefore, chaperone binding candirect the folding of a polypeptide by controlling thetemporal accessibility of regions to fold [Daniels et al.2003]. Finally, the lectin chaperones protect vulnera-ble nascent chains from aggregation and degradation[Hebert et al. 1996, Vassilakos et al. 1996]. Inhibitionof glucose trimming using competitive inhibitors ofglucosidases I and II such as castanospemine ordeoxynojirimycin abolished calnexin and calreticu-lin binding and accelerated the turnover for a numberof glycoproteins through the endoplasmic reticulum-associated degradation (ERAD) pathway [Kearse et al.1994, Keller et al. 1998, Ayalon-Soffer et al. 1999,Liu et al. 1999, Chung et al. 2000, Molinari et al.2002, Svedine et al. 2004]. While stable cell lineslacking calnexin, calreticulin, or ERp57 have beenestablished, the lectin chaperones and their associa-ted oxidoreductase are essential for the survival ofmulticellular metazoans underscoring their impor-tance in organismal homeostasis [Mesaeli et al.1999, Denzel et al. 2002, Coe et al. 2010].The cleavage of the final glucose by glucosidase II

disrupts binding to the lectin chaperones [Hebert et al.1995] (Figure 2). Glucosidase II cleavage of thesecond and third glucoses is not processive as thesubstrate appears to require repositioning betweencleavage events [Mackeen et al. 2009]. Glucosidase IIis comprised of a- and b-subunits. The a-subunit contains the catalytic activity, while theb-subunit possesses the ER retention signal as wellas a Mannose-6 phosphate Receptor Homology(MRH) domain that influences substrate specificity[Deprez et al. 2005, Stigliano et al. 2009]. The MRHdomain helps in substrate recruitment and perhapsin its activity as the b-subunit appears to be criticalfor the accelerated cleavage of the final glucose togenerate unglucosylated glycans [Deprez et al. 2005,Watanabe et al. 2009]. As small proteins may be ableto fold without the assistance of chaperones, it would

CB

A

Man8B Man8C

Man7A

Asn-X-Ser/Thr

Man5

A. B.

α1,2

α1,2

α1,2

α1,3

α1,3

α1,3

α1,2

α1,3

α1,2

α1,6

α1,6

Figure 1. Schematic representation of N-linked glycan. (A) Struc-ture of N-linked glycan including glucoses (green), mannoses(blue) and N-acetylglucosamines (gray) are indicated by triangles,spheres and rectangles, respectively. The branches on the glycan(A, B and C) are designated. Carbohydrate linkages are indicated.(B) Proposed structures of the glycans serving as degradationsignals in ERAD. The glucoses have been trimmed after thecalnexin cycle. The outer most B-branch mannose is trimmed byER mannosidase I/Mns1p. The a1,2 mannose on the C-branch hasbeen proposed to be removed by EDEM1/Htm1p. Trimming ofC-branch mannose residues generates exposed a1,6 mannose,which has been proposed to be a degradation signal for the receptorssuch as Yos9p and OS-9/XTP3-B. This figure is reproduced incolour in the online version of Molecular Membrane Biology.

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be of interest to determine if glucosidase II trimmingcan be controlled to by-pass lectin chaperone bindingso that these chaperones can focus their attention onmore complex proteins that require their assistance for

proper maturation. The number and location of gly-cans appears to dictate to some extent the molecularchoreography or the chaperone binding profile for theindividual chaperones [Hebert et al. 1997, Molinari

Table I. ER maturation and quality control factors. The mammalian components are listed with Saccharomyces cerevisiae homologues denotedin parenthesis.

Category Component Description

Hsp70/90 BiP/GRP78 (Kar2) Hsp70 molecular chaperoneGRP94 Hsp90 molecular chaperone

Glycan processing and binding Glucosidase I (Gls1/CWH41) Trimming of terminal glucose residueGlucosidase II (Gls2/Gtb1) Trimming of second and third glucose residuesUGT1 Quality control sensor and glucose transferaseER mannosidase I (Mns1) Trimming of mannose residuesMalectin Association with Glc2 N-glycanCalnexin (Cne1) Membrane integrated chaperone that binds

monoglucosylated glycansCalreticulin Soluble chaperone that binds monoglucosylated glycansEDEM1-3 (Htm1/Mnl1) Mannosidase or mannose binding lectinOS-9 (Yos9) ERAD receptorXTP3-B (Yos9) ERAD receptor

Oxidoreductase PDI (Pdi1) Disulfide bond formation, reduction and rearrangementERp57 Arranging cargo disulfide bonds with calnexin/calreticulinERdj5 Reduction of ERAD substrate disulfide bonds,

associated with EDEM1 and BiPERFAD Facilitator of ERAD substrate dislocation

ERAD adapters SEL1L (Hrd3) ERAD scaffold proteinHERP (Usa1) ERAD scaffold protein

Translocon Sec61 (Sec61/Ssh1) Translocon for newly synthesized polypeptidesentering the ER and possibly for ERAD substrateretrotranslocation

Derlin1-3 (Der1) Putative ERAD translocation channelRetrotranslocation p97 (cdc48) AAA-ATPase involved in ERAD substrate extractionUbiquitination Hrd1 (Hrd1) E3 ligase for ERAD substrates

GP78 E3 ligase for ERAD substratesRNF5/RMA1 E3 ligaseTEB4/MARCHIV (Doa10) E3 ligase

CRT

UGT 1

Glucosidase IIGlucosidase II

Malectin

Glucosidase IAnterograde

transport

ERAD

CNX

Figure 2. Glucose-dependent early protein maturation. After the entry of polypeptides into the ER, glucosidase I trims the first glucose.Diglucosylated glycans mediate interactions with malectin and glucosidase II trims the second glucose. This monoglucosylated glycan is thetarget of lectin-chaperones, calnexin (CNX) and calreticulin (CRT). Glucosidase II removes the final glucose to generate an unglucosylatedglycan. Mature polypeptides are sorted for anterograde trafficking, while still immature or non-native proteins are reglucosylated by UGT1 tosupport calnexin/calreticulin rebinding. Eventually, terminally misfolded proteins are targeted to ERAD pathway for destruction. This figure isreproduced in colour in the online version of Molecular Membrane Biology.

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and Helenius 2000, Wang et al. 2005, 2008]. Molinariand Helenius (2000) proposed the cotranslationalmodel whereby prior interaction with BiP before thelectin chaperones only occurs if there is no glycanlocated within 50 amino acids of the N-terminus of theprotein. While non-glycosylated proteins or regionsmay be aided by the Hsp70 chaperone BiP and proteindisulfide isomerase family members such as PDI,glycans appear to provide the dominant chaperonesignal. Together, the region-specific recruitment ofthe carbohydrate binding chaperones and the bulkyhydrophilic nature of the glycan, likely inhibitHsp70 binding as this family of chaperones recognizeshydrophobic patches in extended conformations[Blond-Elguindi et al. 1993]. BiP can also serve as aback-up chaperone system in cases where lectin chap-erone binding has been inhibited [Zhang et al. 1997].The positioning of BiP at the translocon by the

J-domain of Sec63 and its ability to directly bindpolypeptides supports the early action of BiP in theprotein maturation process. However, N-linked gly-cans route glycoproteins to the lectin chaperone assis-tance program. The lectin chaperone binding profileinvolves the initial binding to calnexin, the membraneintegrated chaperone [Wang et al. 2005]. In manycases, this is followed by association with calreticulin,if the substrate is soluble or if its glycans eventuallyextend deep into the lumen away from the membrane.The level of calnexin associated with the transloconcomplex can be regulated by the phosphorylation of itsC-terminal cytoplasmic tail, which supports ribosomebinding and translocon association [Chevet et al.1999]. The evolution of larger more complex proteinsin multicellular organisms appears to have resulted inthe development of an additional sophisticated chap-erone system based on carbohydrate binding proper-ties. A better understanding of the organization of thetranslocon-associated factors will aid in our knowledgeof thematuration assembly line that appears to assist inthe efficient maturation of secretory cargo.

UGT1 as an early quality control gatekeeper

Glucose trimming can support a single round of lectinchaperone binding, however rebinding to these cha-perones or the so-called ‘calnexin cycle’ is controlledby the UDP-glucose:glycoprotein glucosyltransferase1 (UGT1) (Figure 2). UGT1 is a ~ 170 kDa solubleER resident protein comprised of two functionaldomains. The C-terminal domain provides the glu-cosyltransferase activity of the enzyme that uses theER UDP-glucose pool as a source to transfer glucosesonto the A-branch of glycans attached to immatureor non-native substrates (Figure 1A and Figure 2).

The majority of the enzyme is comprised of theN-terminal folding sensor domain for which the struc-ture is unknown. Together, these activities are criticalfor its role as an important quality control gatekeeperthat directs proteins for ER retention through rebind-ing to the lectin chaperones.Biochemical studies using purified UGT1 have

demonstrated that UGT1 recognizes surface-exposedhydrophobic regions on maturing proteins that areassociated with non-native or unassembled proteins[Sousa and Parodi 1995, Trombetta and Helenius2000, Keith et al. 2005]. The analysis of glycopeptidesreglucosylated by purified UGT1 discovered thatregions immediately C-terminal to efficiently reglu-cosylated sites possessed oscillating extended regionsof hydrophobicity [Taylor et al. 2003]. Positioning ofthe hydrophobic region closer to synthetic glycanshas also been shown to increase reglucosylationlevels [Totani et al. 2009]. These results suggestthat UGT1 optimally acts on glycans proximal tohydrophobic structural perturbations.In addition to retaining proteins in the ER, chap-

erone binding has also been proposed to perturb ordelay the folding of proteins in a region-specific man-ner [Daniels et al. 2003,Wang et al. 2005, Pearse et al.2008, Pearse and Hebert 2009]. For multidomainproteins, it can be advantageous to control the orderof domain folding. The non-sequential domain pro-tein influenza hemagglutinin (HA) has an N-terminalsegment that interacts with a C-terminal region, linkedthrough a large loop disulfide bond. Efficient calnexinbinding to the three glycans located on the N-terminalsegment protects this domain from immature oxida-tion to form non-native disulfide linkages, andtethers it to the membrane until a medial domain foldsand the C-terminal interacting region is translated[Daniels et al. 2003]. The efficient reglucosylationof the N-terminal region of HA supports persistentchaperone binding, which helps to direct its efficientfolding [Pearse et al. 2008]. These results are sugges-tive of a larger role for UGT1 beyond targeting pro-teins for ER retention, which includes actively playinga role in directing the folding of large multidomainproteins by controlling region-specific chaperonebinding.UGT1 reglucosylation appears to be positioned

within the ER to act at a late stage during ER matu-ration and quality control. Quantitative immunogoldelectron microscopy found UGT1 to be predomi-nantly located in transitional ER/exit sites or pre-Golgiintermediates [Zuber et al. 2001]. These resultswere consistent with more recent proteomics resultsthat observed UGT1 levels to be the highest in smoothER membrane preparations [Gilchrist et al. 2006].Finally, reglucosylation of HA was observed to solely

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occur post-translationally after release from theribosome-translocon complex [Pearse et al. 2008].Together, these results are indicative of UGT1 playinga late role in the folding process. This supports theinitial chaperone binding being directed by glucosi-dase II trimming prior to engaging the rebindingcycle or the reglucosylation activity that resides deeperinto the ER away from the entry sites found in therough ER.There is growing amount of evidence indicating

that reglucosylation by UGT1 and lectin chaperonerebinding can increase the maturation efficiency of asubset of glycoproteins as the maturation of a numberof glycoproteins is significantly diminished in ugt1-/-

cells [Molinari et al. 2005, Pearse et al. 2010]. Fur-thermore, augmenting UGT1 reglucosylation activityby increasing UDP-glucose levels has been shown toenhance the maturation of transferrin [Wada et al.1997]. A link between UGT1 and the sorting ofdefective secretory cargo to the ERAD pathway isless certain. In vitro studies with purified proteinshave demonstrated that UGT1 recognizes near nativeproteins [Caramelo et al. 2003, 2004], suggestive of itassisting proteins that are on pathway to reach theirnative state but perhaps just need a little more help ortime to mature properly. In contrast, the overexpres-sion of downstream ERAD factors such as EDEM1can accelerate defective cargo extraction from thecalnexin binding cycle and their subsequent degrada-tion, suggesting that ERAD cargo may be caught in afutile chaperone binding cycle awaiting recognitionfor ERAD [Molinari et al. 2003, Oda et al. 2003].Additional studies will be required to determine ifUGT1 is able to focus the efforts of the lectin chap-erone binding cycle to proteins that are near nativeand can eventually be rescued to fold properly, oralternatively, if it can also help to retain defectivecargo in the ER until recognition for ERAD sorting.

Mannose trimming for ERAD

Pharmacological and genetic analyses have revealedthat the clearance of misfolded glycoproteins from theER is linked to the processing of their N-linkedglycans or more specifically mannose trimming.The involvement of mannose trimming for misfoldedprotein turnover was first reported for the heterolo-gous expression of yeast prepro-a factor in rat pitu-itary cells as the inhibition of mannose trimming withdeoxymannojirimycin, a competitive inhibitor of ERa-mannosidases, significantly stabilized prepro-afactor [Su et al. 1993]. These results were furtherverified in both yeast and mammalian cells using bothinhibitors and the genetic deletion/knockdown of

mannosidase activity [Jakob et al. 1998, Liu et al.1999, Svedine et al. 2004, Bhamidipati et al. 2005,Avezov et al. 2008, Termine et al. 2009]. An importantcaveat to note is that these approaches supported theglobal retention of mannose residues on all glycopro-teins and was not limited to ERAD substrates. Thesefindings led to the hypothesis that mannose trimminggenerated an ERAD signal that was recognized bydownstream carbohydrate-binding proteins that aidedin sorting aberrant proteins for degradation. Manno-sidase activity was proposed to act slow; therefore, onlyglycoproteins that possessed non-native structures andhad been retained in the ER would be trimmed ofthese particular mannose residues [Helenius 1994,Cabral et al. 2001]. Initially, trimming was thoughtto only involve the removal of a single mannose fromthe B-arm to generate the so-calledMan8B glycoforms(Man8GlcNAc2; Figure 1B).More recent results suggest that the putative ERAD

carbohydrate signal involves more extensive trimmingof mannose residues to glycoforms from Man7 toMan5 in which B and C branch mannoses aretrimmed (Figure 1B). The involvement of truncatedmannose residues in ERAD is supported by theanalysis of the carbohydrate binding specificities fordownstream quality control receptors, which will bediscussed more thoroughly below. Importantly, whenproteasome degradation is inhibited, ERAD sub-strates were retained in the ER with mannose residuestrimmed to Man5-7 glycoforms [Ermonval et al.2001, Frenkel et al. 2003, Hosokawa et al. 2003,Kitzmuller et al. 2003, Avezov et al. 2008]. As inhi-biting degradation resulted in the prolonged accumu-lation of the ERAD substrates in the ER, it cannot beruled out that the block supported more extensivemannosidase access and trimming. Further compli-cating the identification of the ERAD signal, cytosolicfree oligosaccharides apparently created by cyto-plasmic endoglycosidase cleavage of retrotranslocatedERAD substrates accumulated as Man8B glycans inyeast [Hirayama et al. 2010]. And in cells that transferMan5GlcNAc2 glycans, mannosidase inhibitors stillsupport the stabilization of ERAD substrates eventhough the glycans should not require further trim-ming for ERAD recognition [Ermonval et al. 2001].As mannose deprived side chains appear to mark

substrates for ERAD, the next question becomes whatis the identity of the mannosidase(s) responsible forthe remodeling of mannose arms of ERAD substrates?And how does this mannosidase distinguish non-native from native proteins? The ER possesses a smallgroup of mannosidase-like proteins, which includesER mannosidase I and the EDEM family of proteins.In budding yeast, these families are comprised oftwo proteins termed Mns1p (ER a-Mannosidase I

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orthologue) and Htm1p (the EDEM1 orthologue,also called Mnl1p) [Quan et al. 2008, Clerc et al.2009]. These two proteins appear to cooperate tosupport the extensive de-mannosylation of ERADsubstrates. The initial trimming to Man8B byMns1p created a substrate, which then could befurther trimmed by Htm1p to generate the ERADsignal that involved the exposure of a1,6-bondedmannose residues of the Man7 side chains(Figure 1B) [Clerc et al. 2009]. Deletion of Mns1supported ERAD substrate stabilization as the proteinwas no longer a substrate for the downstream man-nosidase activity of Htm1p.The trimming of mammalian ERAD substrates

appears to be more complex as the ER glycosyl hydro-lase family is expanded, and the characterization of themannosidase activity using purified proteins has beenenigmatic. The overexpression of ER a-Mannosidase Iin cells facilitated the turnover of misfolded proteins,and resulted in the accumulation of Man8B, as well asmore extensively trimmed glycans (Man7 or Man6)[Hosokawa et al. 2003]. In vitro studies with ERa-Mannosidase I have found that at biological levelsa single mannose is trimmed to create Man8B; how-ever, at higher levels of the enzyme and 24 h ofincubation additional a1,2-mannoses can be trimmedto produce Man5-7 glycans [Herscovics et al. 2002].As exogenously expressed ER a-Mannosidase I havebeen observed to accumulate in a perinuclear regionupon proteasome inhibition, this has led to an alter-native hypothesis that aberrant proteins are sorted toan ER quality control compartment where ER a-Mannosidase I is concentrated sufficiently to supportmore extensive trimming of the mannose residues onERAD substrates [Avezov et al. 2008, Lederkremer2009]. However, obtaining elevated levels of ERa-Mannosidase I would appear to be problematic asit is not induced by the unfolded protein response(UPR) in yeast or mammals, and its turnover hasbeen reported to be rapid [Travers et al. 2000,Hosokawa et al. 2001, Wu et al. 2007]. Intriguingly,ER a-Mannosidase I has been found to be stabilized byEDEM1 suggesting that ER a-Mannosidase I levelsmay be tightly regulated [Termine et al. 2009].Recent evidence suggests that the mammalian

ERAD system might work in a similar manner tothat proposed for the yeast system as EDEM familymembers may act as mannosidases situated down-stream of the activity of ER a-Mannosidase I.EDEM1 was first discovered as a mammalian ortho-logue to the yeast Htm1p protein that contained amannosidase-like domain but was believed to lackmannosidase activity [Hosokawa et al. 2001]. Theoverexpression of EDEM1 accelerated the turnoverof glycosylated ERAD substrates, whereas its

knock down stabilized them [Molinari et al. 2003,Gnann et al. 2004, Gong et al. 2005]. Elucidating therole of EDEM1 in the ERAD process has beenproblematic. While the overexpression of EDEM1in cells has been found to be associated with theaccelerated trimming of mannose residues, attemptsto identify any mannosidase activity using purifiedproteins have failed [Olivari et al. 2006, Cormier et al.2009, Mikami et al. 2010, Hosokawa et al. 2010b].This has led to a couple of contrasting models for thefunction of EDEM1 in ERAD.In the first model, EDEM1 possesses specialized

mannosidase activity that acts on aberrant proteins toaid in the generation of an ERAD signal. The over-expression of EDEM1 in cells was associated withincreased mannose trimming as determined by anincrease in mobility of glycosylated ERAD substratesby SDS-PAGE or the composition of glycans releasedfrom ERAD substrates as analyzed by HPLC[Olivari et al. 2006, Cormier et al. 2009, Hosokawaet al. 2010b]. EDEM1 overexpression also enhancedA-branch mannose trimming in B3F7 cells, as this celllines transfers Man5 glycans lacking B- and C-branch mannose residues [Olivari et al. 2006]. Incontrast, the overexpression of wild type but notmannosidase domain mutant forms of EDEM1supported the trimming of C-branch mannose residueof alpha-1-antitrypsin null Hong Kong to accumulateMan8C glycans (Figure 1B) [Hosokawa et al. 2010b].While the primary structure of EDEM1 and the anal-ysis of glycoproteins after EDEM1 overexpression areconsistent with EDEM1 possessing mannosidaseactivity, the demonstration of such an activity is lack-ing for studies using purified protein and conflictingresults are available as to the nature of the trimmedspecies (see above) [Olivari et al. 2006, Mikami et al.2010, Hosokawa et al. 2010b].An alternative model proposes that EDEM1 uses it

mannosidase-like domain to bind glycans as a qualitycontrol receptor that recognizes misfolded substratesand sorts them for degradation [Molinari et al. 2003,Oda et al. 2003, Wang and Hebert 2003]. EDEM1was identified as an ER stress-induced gene withhomology to conserved residues of ER MannosidaseI active site, however, it lacked a couple of cysteineresidues thought to be required for mannosidase activ-ity [Hosokawa et al. 2001]. Therefore, EDEM1 wasproposed to function as a quality control lectin thatbindsMan8B forms of N-glycan onmisfolded proteinsto promote ERAD [Hosokawa et al. 2001, 2003,Wang and Hebert 2003]. Co-immunoprecipitationresults have found that EDEM1 can differentiatebetween non-native and native proteins specificallybinding to aberrant substrates [Hosokawa et al.2006, Olivari et al. 2006, Cormier et al. 2009,

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Kosmaoglou et al. 2009]. However, while EDEM1efficiently bound to ERAD substrates lacking gly-cans, it did not accelerate the turnover of thesenon-glycosylated substrates [Cormier et al. 2009,Kosmaoglou et al. 2009]. Furthermore, mutation ofthe mannosidase domain of EDEM1 still supportsthe accelerated degradation of ERAD substrates[Cormier et al. 2009, Termine et al. 2009, Hosokawaet al. 2010a]. These results are suggestive of a bi-partite binding function for EDEM1 that involvesboth protein-protein interactions involving proteindefects (perhaps exposed hydrophobic domains) andmannose trimmed glycans. While one form ofbinding might support interactions as probed by co-immunoprecipitation of lysed cells, apparently bothbinding forms must be engaged for EDEM1 to accel-erate the turnover of an ERAD substrate.There are a number of additional mysteries in

regard to EDEM1 characterization and function. First,EDEM1 was original identified as a type II membraneprotein that contained an N-terminal hydrophobicsignal anchor sequence [Hosokawa et al. 2001].More recently a soluble form of EDEM1 hasbeen observed generated by the cleavage of its N-terminal sequence [Olivari et al. 2005, Zuber et al.2007]. Second, EDEM1 has been shown to associatewith calnexin, directly positioning it to extract ERADcargo from the calnexin binding cycle [Oda et al.2003]. In contrast, morphological studies thatexplored the localization of endogenous EDEM1found that it did not co-localize with calnexin and itcould be found in the vesicle-like structures that exitthe ER [Zuber et al. 2007]. Third, the half-life ofEDEM1 is short and autophagy or an autophagy-like pathway has been implicated in its rapid turnover[Cali et al. 2008, Le Fourn et al. 2009]. The signif-icance of this turnover route and accelerated rate isunknown but it may help explain why EDEM1 can befound in vesicles exiting the ER. Finally, themannosidase-like domain of EDEM1 that has beenproposed to mediate substrate recognition or trim-ming, has also been implicated in supportingEDEM1 binding to a downstream ER membraneadapter protein, SEL1L [Cormier et al. 2009].EDEM1 binding to endogenous SEL1L was dimin-ished by mutating conserved residues in themannosidase-like domain of EDEM1, or treatmentwith the mannosidase inhibitor kifunensine. Theseresults suggest that the mannosidase-like domainmay be deployed to target EDEM1-substrate com-plexes to ER membrane complexes involved in dislo-cation and ubiquitination. Future studies usingpurified proteins to recapitulate the substrate selectionand targeting events would help to further understandthese processes.

Mammalian EDEM1 has two paralogues termedEDEM2 and EDEM3 that are also upregulated byUPR and possess ER a-Mannosidase I homologydomains [Mast et al. 2005, Olivari et al. 2005, Hiraoet al. 2006]. EDEM2 and EDEM3 have N-terminalcleavable signal sequences that generate soluble ERresident proteins. Only EDEM1 and EDEM3 havebeen found to be associated with increased mannosetrimming when overexpressed in cells [Mast et al.2005, Olivari et al. 2006, Hirao et al. 2006, Cormieret al. 2009, Hosokawa et al. 2010b]. The overexpres-sion of EDEM3 enhanced the mannose trimming ofN-glycan of ERAD substrates to Man6 and Man7[Hirao et al. 2006, Hosokawa et al. 2009]. However, asthe demonstration of mannosidase activity for isolatedEDEM family members has not been demonstrated, itcannot be ruled out that EDEM interacting or stabi-lized proteins are responsible for the observed increasein cellular mannose trimming. In support of thispossibility, ER a-Mannosidase I has been found tobe stabilized by EDEM1 [Termine et al. 2009].

Quality control sorting receptors

Mannose trimming appears to generate an ERADsignal that must then be recognized by quality controlreceptors to target the defective cargo for ERAD. Thefamily of possible mannose-dependent quality controlreceptors includes EDEM1 (as discussed above) anda group of ER resident MRH domain containingproteins that include Yos9p/OS-9 and XTP3-B.These MRH domain proteins bind to mannosesand interact with downstream ER membrane com-plexes involved in ERAD, implicating them in playingcritical roles in the selection and delivery of defectivesecretory cargo to the ERAD pathway.Yos9 was initially identified as an ERAD-related

gene by genome screening of Saccharomyces cerevisiaeas a Yos9 deletion strain stabilized CPY* [Buschhornet al. 2004]. It appeared to act specifically onglycosylated proteins since it did not stabilize non-glycosylated CPY* [Buschhorn et al. 2004, Kaneharaet al. 2010]. Its MRH domain is important for ERADas mutating this domain suppressed CPY* turnover[Bhamidipati et al. 2005]. Yos9p recognized ERADsubstrates bearing Man8B and Man5 N-glycan[Szathmary et al. 2005]. In contrast, frontal affinitychromatography analysis, a quantitative methoddeveloped to analyze carbohydrate-protein interac-tions, demonstrated that the MRH domain ofYos9p has high affinity for Man5 glycoforms butnot Man8B [Quan et al. 2008]. This later resultsuggested that Yos9p binds proteins that have beenextensively trimmed by Mns1p and Htm1p to expose

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a1,6-linked mannose residues [Quan et al. 2008,Clerc et al. 2009]. Since Yos9p also associated withnon-glycosylated protein, it appears to possess bothcarbohydrate and protein-protein selection processes[Bhamidipati et al. 2005, Denic et al. 2006]. Further-more, Yos9p has also been found to associate withHrd3p that nucleates the formation of an ERADcomplex in the ER membrane [Carvalho et al.2006, Denic et al. 2006, Gauss et al. 2006]. Alto-gether, these results suggest that Yos9p is equippedwith at least two functional domains: An MRHdomain that supports aberrant glycoprotein recogni-tion; and a protein-binding domain that binds toHrd3p or misfolded proteins. These binding proper-ties position Yos9p as an important link betweensubstrate recognition and delivery to a downstreamER membrane ERAD complex.There are two mammalian homologues of Yos9p

called OS-9 and XTP3-B, which are both upregulatedby UPR through Ire1/Xbp-1 activation [Bernasconiet al. 2008, Alcock and Swanton 2009]. OS-9 andXTP3-B contain one and twoMRH domains, respec-tively [Munro 2001]. Like Yos9p, these proteins alsoappear to serve as quality control receptors that areable to selectively bind non-native substrates and sortthem to the ERAD pathway through their associationwith an ER membrane adapter protein.OS-9 has three spliced variants termed OS-9.1

through OS-9.3. Downregulation of OS-9 by RNAistabilized glycosylated ERAD substrates; however, asimilar fate was also observed upon overexpression[Bernasconi et al. 2008, Christianson et al. 2008,Hosokawa et al. 2009]. This may be caused by theoverexpressed protein binding to substrate but beingineffective in substrate presentation or downstreamdelivery due to the lack of a sufficient level of anassociated factor. Overexpression of OS-9 may createan orphan subunit of the quality control receptor thatproduces a dominant negative effect on ERAD. TheMRH domain of OS-9 has been shown to be criticalfor its role in ERAD [Hosokawa et al. 2009, Mikamiet al. 2010]. Recombinant human OS-9 MRHdomain was recently found to associate preferentiallywith glycans missing C-arm mannose residues.Bound glycoforms include glycans that ranged fromGlc1Man8 to the more extensively trimmed glycanssuch as Man4. These results were consistent withexposure of the a1,6-linked mannose on the C-armbeing critical for OS-9 substrate recognition. Further-more, OS-9 co-immunoprecipitated with mutant, butnot wild type alpha-1-antitrypsin, demonstrating itsability to selectively bind defective substrates.XTP3-B is a soluble ER protein with two MRH

domains and two spliced variants [Hosokawa et al.2008]. The shorter variant is missing a small segment

of the linker that connects the two MRH domains.The C-terminal MRH domain appears to be involvedin substrate recognition as it is required for glycan-dependent binding to null Hong Kong alpha-1-antitrypsin [Yamaguchi et al. 2010]. However,knockdown of XTP3-B does not affect the degrada-tion of null Hong Kong alpha-1-antitrypsin, sugges-tive of redundant cellular factors being involved in itsturnover [Christianson et al. 2008, Hosokawa et al.2008]. The C-terminal MRH domain of XTP3-Bpreferentially binds to Lec1 cells, which express pro-teins bearingMan5-6GlcNAc2 glycans on their surface[Yamaguchi et al. 2010]. Inhibitory studies usingvarious disaccharides revealed that Mana1,6Man effi-ciently inhibited substrate binding consistent with thesecond MRH domain of XTP3-B binding to termi-nally exposed a1,6-linked mannose residues. Theseresults suggest that OS-9 and XTP3-B possess similarglycan binding requirements but as the two proteinsare not associated with one another their roles appearto be distinct [Christianson et al. 2008]. Interestingly,the simultaneous knockdown of both OS-9 andXTP3-B by RNAi specifically stabilized solubleproteins possessing luminal lesions [Bernasconiet al. 2010]. Future studies will be required to deter-mine if the two proteins are involved in redundantpathways or possess different binding requirements.Similar to Yos9p, OS-9 and XTP3-B were found

to associate with the mammalian Hrd3p homologueSEL1L, an ER membrane ERAD adapter protein[Christianson et al. 2008, Hosokawa et al. 2008,Mueller et al. 2008]. Therefore in addition to select-ing glycosylated ERAD substrates for degradation,these luminal quality control receptors also appear tosupport delivery of the defective substrates to thedislocation site in the ER membrane. The nature ofthe quality control receptor-adapter protein interac-tion is uncertain. It was reported that OS-9 andXTP3-B are recruited to the SEL1L complex bytheir MRH domains as mutation of the MRHdomain of OS-9 and XTP3-B decreased their inter-action with SEL1L but did not affect its associationwith ERAD substrates [Christianson et al. 2008](Figure 3B). However, contradicting results arealso available that suggest that OS-9 and SEL1Lassociate through protein-protein interactions(Figure 3A) [Hosokawa et al. 2009, Yamaguchiet al. 2010]. Similar uncertainty has also beenobserved with EDEM1 binding to SEL1L. Themannosidase-like domain of EDEM1 supportsbinding to the glycosylated SEL1L and not nullHong Kong alpha-1-antitrypsin as mutations of themannosidase-like domain or kifunensine diminishedEDEM1 binding to SEL1L and not to the ERADsubstrates [Cormier et al. 2009].

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Preparation of ERAD substrates forretrotranslocation

Once marked for ERAD and selected by qualitycontrol receptors for degradation, the non-nativeand possibly aggregated substrates need to be pre-sented to an ERAD complex in a translocation com-petent form for their dislocation to the cytoplasm. AsERAD substrates can contain disulfide bonds, theseevents can require the protein-assisted reduction ofdisulfide bonds and unfolding.The ER contains a family of oxidoreductases of

which a number of these enzymes appear to playimportant roles in the reduction of intra- and inter-molecular disulfides to create ERAD substrates thatare competent for dislocation. The extensively oxi-dized ERAD substrate BACE457 was found in acomplex with PDI and BiP prior to dislocation tothe cytoplasm [Molinari et al. 2002]. Furthermore,PDI disassembled cholera toxin dimers to facilitatetheir retrotranslocation [Tsai et al. 2001]. PDI hasalso been observed in a complex with Htm1p in

yeast further implicating it in the ERAD process[Clerc et al. 2009, Sakoh-Nakatogawa et al. 2009].Interestingly, PDI stimulated the retrotranslocation ofglycan and cysteine-free prepro (a) factor using mam-malian and yeast microsome systems [Gillece et al.1999, Wahlman et al. 2007]. PDI appears to aid insubstrate recognition and preparation for dislocationby both reduction, as well as by using its peptidebinding properties to unfold misfolded and aggre-gated proteins.The ER also contains a newly identified

thioredoxin-domain containing flavoprotein calledERFAD [Riemer et al. 2009]. The knockdown ofERFAD expression by RNAi stabilized ERAD sub-strate turnover and decreased the cellular level ofpoly-ubiquitinated proteins. ERFAD also interactedwith a number of ERAD factors including OS-9,SEL1L and ERdj5 as shown by co-immunoprecipi-tation. ERdj5 possesses four thioredoxin domains,and a J-domain that supports the recruitment ofBiP. ERdj5 was also identified as an EDEM1 inter-acting protein using a modified yeast two-hybrid andco-immunoprecipitation procedures [Ushioda et al.2008]. Its overexpression accelerated turnover ofERAD substrates by suppressing aggregation of mis-folded proteins possessing non-native disulfide bonds.The activities of ERFAD, ERdj5, and their interactingpartners, support them playing roles in the reductionand possibly unfolding of ERAD substrates so that theycan be efficiently threaded through an ER translocon.Adenine nucleotide regulated ER chaperones also

appear to be involved in generating translocation com-petent ERAD substrates. BiP has been localized toERAD complexes containing EDEM1 and ERdj5, aswell as with XTP3-B [Christianson et al. 2008,Ushioda et al. 2008]. In addition, the ERHsp90 familymember GRP94 was observed in a complex withOS-9 using mass spectrometry to analyze interactingpartners of overexpressed tagged OS-9 [Christiansonet al. 2008]. The knockdown of GRP94 stabilized nullHong Kong alpha-1-antitrypsin. As both BiP andGRP94 are molecular chaperones, they may play arole in ERAD substrate recognition and/or their ATP-dependent binding cycle may aid in the unfolding ofthe misfolded substrates.

ER membrane ERAD adapter complexes

Once a protein has been deemed misfolded by anERAD receptor, the misfolded protein needs to betargeted to a dislocation complex in a translocationcompetent form for the extraction of the misfoldedprotein to the cytoplasm. This ERAD complexappears to consist of a number of proteins that aid

Proteasome

AAA-ATPase

TransloconAdapter

ERADreceptor

A. B.

E3

Figure 3. Two models for quality control receptor-substrate target-ing to an ER membrane ERAD complex. (A) ERAD receptorsrecognize aberrant cargo displaying carbohydrate based ERAD sig-nals using their sugar-binding domains (MRH or mannosidase-like domains). The receptor-substrate complex is then delivered toan ERAD complex in the ER membrane consisting of an adapterprotein, an E3 ligase and a translocation channel or translocon. (B)In an alternativemodel for selection and delivery, the ERAD receptorselects client proteins based on the folding status of the proteinthrough protein-protein interactions. Once a misfolded protein hasbeen recognized the receptor uses its sugar-binding domain to dockto a glycan on the adapter protein to support delivery of the substrateto the ERAD complex. This figure is reproduced in colour in theonline version of Molecular Membrane Biology.

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in the delivery, translocation and ubiquitinationprocesses [Carvalho et al. 2006, Denic et al. 2006,Gauss et al. 2006, Christianson et al. 2008]. Centralto the formation of this ER membrane complex areadapter proteins that are integrated into the ER mem-brane and serve as the link between substrate recog-nition/delivery machinery and factors involved indislocation and ubiquitination.The yeast and mammalian ER contain ER

membrane adapter proteins that possess a numberof luminal-exposed tetratricopeptide repeat (TPR)domains [D’Andrea and Regan 2003]. The yeastprotein termed Hrd3p has nine TPR domains whilethe mammalian protein, SEL1L, contains a total ofeleven. Structures of known TPR domains displayhelical folds comprised of two helices connected by ashort linker. Adjacent TPR domains come togetherto form concave and convex surfaces, with protein-protein interactions generally occurring on the con-cave surface [Das et al. 1998]. The well studiedcytoplasmic TPR-containing protein Hop providesa link between Hsp70 and Hsp90 chaperones byusing two separate clusters of TPR domains to recruitthe individual chaperones to form a large proteinfolding complex [Scheufler et al. 2000]. This cyto-plasmic complex facilitates the passage of a maturingsubstrate from the Hsp70 to the Hsp90 chaperonesystem. Similarly, these ER membrane TPR domainproteins appear to create a connection between qual-ity control receptors and dislocation/ubiquitinationmachinery to support the selective translocation ofERAD substrates to the cytoplasm for eventuallydestruction.Affinity purification of different ERAD compo-

nents followed by mass spectrometry was used todiscover that Hrd3p resides in a large molecularweight complex, interacting with luminal (Yos9p),ER membrane (Usa1p, Hrd1p, Ubx2p) as well ascytosolic ERAD components (Cdc48p). This impli-cates Hrd3p serving as a scaffold or adapter proteinthat nucleates an ERAD translocation complex[Carvalho et al. 2006, Denic et al. 2006]. Truncationstudies with Hrd3p suggest that it binds Hrd1p andYos9p through two distinct domains, implying thatthe TPR domains of Hrd3p direct distinctinteractions with ERAD machinery [Gauss et al.2006]. Furthermore, Hrd3p also has the ability todirectly bind misfolded proteins independent ofYos9p [Denic et al. 2006]. The interaction betweenHrd3p and Hrd1p may have multiple functions. First,Hrd3p appears to aid the delivery of misfolded pro-teins to the ER membrane E3 ligase, Hrd1p. Second,deletion of Hrd3p destabilized Hrd1p [Plemper et al.1999, Gardner et al. 2000, Gauss et al. 2006], imply-ing that Hrd3p might also be needed to stabilize the

translocation complex, insuring that uncomplexedligases do not accumulate.The 11 TPR domains of SEL1L are arranged into

three clusters comprised of four, five and two TPRdomains moving from the N- to the C-terminus.SEL1L is N-linked glycosylated and has an N-terminal fibronectin type II domain [Biunno et al.2006]. SEL1L is known to interact with a large numberof proteins including Hrd1, Herp, Derlin-1/2,EDEM1, OS-9 and XTP3-B (see below) [Lilleyet al. 2006, Christianson et al. 2008, Hosokawaet al. 2008, Mueller et al. 2008, Cormier et al.2009]. Kopito and co-workers showed that SEL1Lcan bind to both OS-9 and XTP3-B without its trans-membrane domain, but deletion of the two C terminalmembrane proximal TPR domains abolished binding[Christianson et al. 2008]. The role of SEL1L inERAD was initially discovered by studying thecytomegalovirus-induced degradation of major histo-compatibility complex class I heavy chains [Muelleret al. 2006]. This study also demonstrated that knock-down of SEL1L stabilized a broader range ofmisfoldedproteins.As previously mentioned, both EDEM1 and

XTP3-B have the ability to bind nonglycosylated mis-folded proteins [Christianson et al. 2008, Hosokawaet al. 2008, Cormier et al. 2009, Kosmaoglou et al.2009]. Mutations in their respective MRH ormannosidase-like domains abolished binding betweenOS-9/XTP3-B/EDEM1 and SEL1L [Christiansonet al. 2008, Cormier et al. 2009]. This suggests thatthe MRH or mannosidase-like domains of the ERADmachinery help support SEL1L binding, rather thansubstrate recognition (Figure 3B). The glycans ofSEL1L are all positioned either within or in closeproximity to the TPR domain clusters, consistentwith the possibility that the glycans and TPR domainswork in concert to support protein interactions. Unlikefor Hrd3p, SEL1L does not appear to directly bindERAD clients [Christianson et al. 2008].Moreover therepertoire of SEL1L interacting proteins was recentlyexpanded with the addition of UBC6e (an ER mem-brane anchor for an E2 ligase), UBXD8 (recruiterof the cytoplasmic AAA-ATPase p97) and AUP1(ubiquitin-binding protein) [Mueller et al. 2008].The Hrd3p complex in yeast also appears to

contain an additional adapter protein termed Usa1p[Carvalho et al. 2006]. Usa1p is needed for thedegradation of luminal ERAD clients and serves asa linker for membrane proteins including Hrd1p andDer1p [Carvalho et al. 2006, Horn et al. 2009]. Themammalian homolog of Usa1, Herp, is required fordegradation of kappa light chain, a non-glycosylatedsubunit that is recognized by BiP rather thanER quality control lectin receptors [Okuda-Shimizu

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and Hendershot 2007]. Like Usa1p, Herp is an ERmembrane protein with both termini residing in thecytoplasm [Kokame et al. 2000]. Interestingly, Herpwas also shown to bind the H8 and S1 subunits of theproteasome, implying that Herp connects the trans-location and ubiquitination processes to the protea-some [Okuda-Shimizu and Hendershot 2007].Together, these results indicated that adapter proteinsplay critical roles in the recruitment of ERADmachinery within the lumen, cytoplasm, as well asthe ER membrane.

Translocation and ubiquitination

The final ER events for ERAD include the translo-cation of the ERAD substrate across the ER mem-brane and its poly-ubiquitination at the cytoplasmicface of the ER membrane by membrane localizedE3 ligases. Machinery involved in these importantprocesses appears to be components of the largemembrane protein complex nucleated by the ERmembrane adapter proteins (Figure 4). The organi-zation of these complexes insures the efficient transferof ERAD substrates from quality control receptors, totranslocons and their ubiquitination upon arrival inthe cytoplasm.The polytopic ER membrane protein Derlin-1 and

its yeast homologue Der1p associate with the SEL1L/HRD1 and Hrd3p/Hrd1p adapter protein complexes,respectively [Lilley and Ploegh 2005, Schulze et al.2005, Ye et al. 2005, Carvalho et al. 2006]. Thehuman cytomegalovirus initiated turnover of majorhistocompatibility class I heavy chain involves the useof Derlin proteins, as Derlin-1 was discovered asan US11-interacting protein that accelerated thedegradation of heavy chain [Lilley and Ploegh

2004]. Two additional homologues of Derlin havebeen discovered termed Derlin-2 and Derlin-3. Theoverexpression of the Derlin family members accel-erated the degradation of ERAD substrates, whereasits knockdown stabilized them [Oda et al. 2006,Sun et al. 2006]. Additional studies will be requiredto determine if Derlins serve as translocation pores, asthis role is largely speculative and based on theirtopology that includes four putative transmembranedomains.The Sec61 translocon that is responsible for the

insertion of proteins into the ER has also been pro-posed to play a role in the translocation of ERADsubstrates in the reverse direction [Wiertz et al. 1996].This demonstration is complicated by the need todelineate the protein import and export processes.However, yeast strains possessing mutations inSec61p have been described that support efficienttranslocation into the ER but are deficient in theretrotranslocation of proteins out of the ER [Pilonet al. 1997, Plemper et al. 1997, 1998, Scott andSchekman 2008]. Furthermore, yeast and mamma-lian ERAD substrates have been observed to associatewith Sec61 during late stages of quality control[Wiertz et al. 1996, Pilon et al. 1997, Plemperet al. 1997]. Future studies will be required to deter-mine the range of proteins that can serve as retro-translocons, and if each of the translocons supportsthe transit of different types of substrates. Addition-ally, while the AAA-ATPase p97 (Cdc48p in yeast)has been implicated in driving the translocation of thesubstrate once the termini of the substrate reachesthe cytoplasm, it is unclear what the mechanism isfor the initial threading of the ERAD substrate inthrough the pore. Cotranslational translocation isaided by chain elongation driven by the ribosome,which is nestled on the translocon. However, a

TransloconsSec61Derlin 1-3

E3 LigasesHRD1GP78TEB4/MARCHIVRNF5/RMA1TRC8RFP2

MammalsYeast

Adapters

E3 LigasesHrd1pDoa10p

Sel1LHerp

AdaptersHrd3pUsa1p

TransloconsSec61pDer1p

Figure 4. ER membrane ERAD complexes. Proteins that reside in the yeast and mammalian ERAD translocation complex are designated.This figure is reproduced in colour in the online version of Molecular Membrane Biology.

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post-translocation event would appear to require aninitial push from the lumen for the chain to reach thecytoplasmic AAA-ATPase involved in pulling thechain into the cytoplasm.During the arrival in the cytoplasm, ERAD sub-

strates are rapidly ubiquitinated by a family of ERlocalized E3 enzymes. The topology of the ERADsubstrate and the location of the defect appear todetermine the identity of its modifier. In yeast, mem-brane proteins possessing cytoplasmic defects or sol-uble cytoplasmic proteins were modified by Doa10p[Vashist and Ng 2004]. In contrast, soluble or mem-brane proteins possessing luminal lesions used Hrd1p[Carvalho et al. 2006, Denic et al. 2006]. The trans-membrane region of Hrd1p also appears to supportthe direct recognition of proteins with defects that liewithin the membrane [Sato et al. 2009]. Altogether,Hrd1p appears to manage the formation of a numberof ERAD complexes that support the processing ofdifferent substrate classes [Kanehara et al. 2010].In mammalian cells, the family of E3 ligases

involved in ERAD has expanded beyond the smallset of yeast modifiers. In addition to the yeast Hrd1phomologue HRD1, mammalian E3 ligases currentlyinclude GP78, RNF5/RMA1, TEB4/MARCHIV,TRC8 and RFP2 [Fang et al. 2001, Hassink et al.2005, Lerner et al. 2007, Morito et al. 2008, Stagget al. 2009]. The selectivity of many of the ERE3 ligases is poorly defined. The selection processappears to differ from the yeast system whereHrd1p ubiquitinates substrates with defects withinthe membrane or luminal compartments [Vashistand Ng 2004]. Recent results suggest that solubleERAD substrates that are dependent on HRD1, canbe rendered HRD1-independent by simply tetheringthem to the membrane with a single hydrophobicdomain [Bernasconi et al. 2010]. The change intopology apparently alters the E3 ligase involved indirecting turnover. The larger spectrum of ERADsubstrates found in multicellular systems has resultedin the expansion of E3 ligases and diversification ofthe selection process. In some cases, the association ofligases with specific adapter complexes appears todictate the substrate selection process, whereas forother ligases substrates may be selected directly by theligase.

Conclusion

The quality control process evaluates mutant proteinsthat are defective and targets them for degradation. Inaddition, as one third of the genome products traversethe secretory pathway and maturation is an errorprone process, the quality control machinery must

also test all the secretory client proteins. While manyquality control components have been identifiedalong with the complexes in which they reside, howthis relatively small group of proteins efficientlyscreens the thousands of different cargo proteins isstill poorly understood. In the cytoplasm, hundreds ofE3 ligases are responsible for the recognition, mod-ification and turnover of cytosolic factors [Deshaiesand Joazeiro 2009]. Since recognition and modifica-tion of secretory pathway proteins occurs in twoseparate locations, this would appear to provide anadditional layer of complexity; however, the machin-ery involved appears to be simplified. Future studieswill be required to determine how a glycan-basedquality control process can efficiently manage sucha large group of diverse customers that includes wildtype and mutant proteins, misfolded or unassembledcargo, and factors that are degraded through regu-lated proteolysis.

Declaration of interest: This work was supportedby US Public Health grants CA79864 andGM086874 (DNH) and The Uehara MemorialFoundation (TT). In addition, JCS was partiallysupported by an NIH Chemistry-Biology Interfacetraining grant (T32GM00815). The authors reportno conflicts of interest. The authors alone are respon-sible for the content and writing of the paper.

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