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Originally published In Press as doi:10.1074/jbc.M302979200 on September 3, 2003

J. Biol. Chem., Vol. 278, Issue 47, 46895-46905, November 21, 2003
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Multiple Endoplasmic Reticulum-associated Pathways Degrade Mutant Yeast Carboxypeptidase Y in Mammalian Cells*

Roberta Mancini{ddagger}§, Markus Aebi¶, and Ari Helenius{ddagger}||

From the {ddagger}Institute of Biochemistry, Swiss Federal Institute of Technology, Hoenggerberg, CH-8093 Zurich and the Institute of Microbiology, Swiss Federal Institute of Technology, CH-8092 Zurich, Switzerland

Received for publication, March 24, 2003 , and in revised form, September 3, 2003.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
The degradation of misfolded and unassembled proteins by the endoplasmic reticulum (ER)-associated degradation (ERAD) has been shown to occur mainly through the ubiquitin-proteasome pathway after transport of the protein to the cytosol. Recent work has revealed a role for N-linked glycans in targeting aberrant glycoproteins to ERAD. To further characterize the molecular basis of substrate recognition and sorting during ERAD in mammalian cells, we expressed a mutant yeast carboxypeptidase Y (CPY*) in CHO cells. CPY* was retained in the ER in un-aggregated form, and degraded after a 45-min lag period. Degradation was predominantly by a proteasome-independent, non-lysosomal pathway. The inhibitor of ER mannosidase I, kifunensine, blocked the degradation by the alternate pathway but did not affect the proteasomal fraction of degradation. Upon inhibition of glucose trimming, the initial lag period was eliminated and degradation thus accelerated. Our results indicated that, although the proteasome is a major player in ERAD, alternative routes are present in mammalian cells and can play an important role in the disposal of both glycoproteins and non-glycoproteins.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
When newly synthesized proteins fail to fold or oligomerize properly in the endoplasmic reticulum (ER),1 they are as a rule retained and eventually degraded (1, 2). The proteolytic degradation of such proteins has been termed ERAD (ER-associated protein degradation). Although substrate selection for ERAD occurs in the ER or in the ER membrane, the actual proteolytic degradation steps frequently occur in the cytosol through the ubiquitin-proteasome pathway (for reviews, see Refs. 3–5). To be degraded, misfolded and unassembled proteins are therefore selectively retrotranslocated from the ER lumen or membrane to the cytosol (6–11). Transport occurs through the sec61 complex, followed by ubiquitination and rapid degradation in 26 S proteasomes (3, 10–12).

How proteins are sorted for ERAD is still poorly understood. Selection is based on conformational criteria, and it is likely to involve a variety of molecular chaperones and folding factors (13–15). There is reason to believe that distinct systems are employed for glycoproteins and non-glycoproteins. In addition to promoting proper folding and serving as retention markers, the N-linked oligosaccharide moieties of glycoproteins are used as specific signal tags for ERAD (16). Genetic evidence indicates that the ER mannosidase I, the carbohydrate trimming enzyme that generates the Man8 isomer B structure, is necessary for efficient degradation of soluble and membrane-bound glycoproteins in yeast (17). Inhibition of this mannosidase also blocks degradation of several ERAD substrates in mammalian cells (18–20). A possible role for mannose trimming is also suggested by the effect on ERAD observed when a putative mannose lectin, named EDEM in mammalian cells and HtmI/MnlI in yeast, is eliminated or overexpressed (21–25). For some glycoproteins, there is also evidence that trimming of glucose residues plays a role in regulating ERAD (11, 26).

To further characterize the molecular basis of substrate recognition and targeting to ERAD in mammalian cells, we expressed a folding-defective yeast carboxypeptidase Y (CPY*) in Chinese hamster ovary (CHO) cells. CPY* is commonly used as an ERAD substrate in Saccharomyces cerevisiae. The wild-type enzyme is a monomeric soluble glycoprotein (61 kDa) targeted via the Golgi complex to the yeast vacuole. The protein has five intrachain disulfide bonds, four N-linked glycans, and the crystal structure shows a globular overall structure (27). CPY is synthesized as a prepro-protein. In the ER, the signal sequence is cleaved, the glycans are added, and disulfide bonds are formed; in the Golgi apparatus, oligosaccharides are modified and the pro-CPY is sorted to the vacuole, where the enzyme is activated by proteolytic cleavage of the pro-sequence (28). Glycosylation is required for efficient intracellular transport but is not for folding, sorting to the vacuole, or for activation (29, 30). CPY*, the mutant protein used in our studies, has a point mutation in position 255 (Gly to Arg) that causes ER retention, retrotranslocation, and degradation via the ubiquitin-proteasome pathway in yeast (9, 31). Glycosylation and ER mannosidase I-catalyzed mannose trimming are necessary for efficient CPY* degradation in yeast (17, 31).

We found that, when expressed in CHO cells, CPY* was rapidly and efficiently degraded by ERAD. However, although some of the protein was degraded by proteasomes, a large fraction was degraded in a proteasome-independent fashion. Although the proteasome-dependent degradation was unaffected by inhibitors of mannose trimming by ER mannosidase I, the proteasome-independent degradation pathway(s) was inhibited.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Cell Culture and Transfection—Chinese hamster ovary (CHO) cells were maintained in minimal essential medium {alpha} supplemented with 8% fetal calf serum, 1% penicillin, and 1% streptomycin. Cells were transiently transfected with SuperFectTM transfection reagent (Qiagen) according to the instructions from the manufacturer.

Materials—Protein A-Sepharose CL-4B beads, aprotinin, pepstatin, leupeptin, chymostatin, tunicamycin, phenylmethylsulfonyl fluoride, N-ethylmaleimide (NEM), dithiothreitol (DTT), cycloheximide, castanospermine, 1,4-dideoxy-1,4-imino-D-mannitol, and deoxymannojirimycin were purchased from Sigma; ALLN, MG132, lactacystin, okadaic acid, and swainsonine were supplied by Calbiochem; 4-acetamido-4'-maleimidylstilbene-2,2'-disulfonic acid (AMS) was from Molecular Probes; orthovanadate was from Axon Laboratory; kifunensine was obtained from Toronto Research Chemicals Inc.; SuperFectTM transfection reagent and cell culture reagents were from Qiagen and Invitrogen, respectively. CHAPS was obtained from Pierce. Promix [35S]methionine/cysteine was purchased from Amersham Biosciences. Glucosidase II was purified from bovine liver (provided by M. Kowarik, Swiss Federal Institute of Technology, Zurich, Switzerland). Anti-Golgi mannosidase II and anti-p58 were supplied by Dr. K. W. Moremen (University of Georgia, Athens, GA) and Dr. J. Saraste (University of Bergen, Bergen, Norway), respectively. Rabbit anti-calnexin, anti-CPY, and anti-BACE457{Delta} sera have been described elsewhere (15, 32–34).

Plasmid Constructs—The genes for CPY and CPY* were cloned into pSVSPORT1 (Invitrogen). CPY* was amplified by PCR using a forward primer (TTAATGAATTCATGGCCATCATTTATCTCATTCTCCTGTTCACAGCAGTGAGAGGG ATCTCATTGCAAAGACCGTTGG), which contains the influenza hemagglutinin (Japan strain) signal sequence (italic) substituting for the CPY* signal sequence (35). The PCR fragment was cloned into pSVSPORT1 to generate pSVHACPY*. To obtain pSVHACPY, the gene for the wild-type enzyme, the BglII-BsuI fragment containing the point mutation (G255R) was cut out and replaced by the BglII-BsuI fragment obtained from PCR on wild-type yeast genomic DNA. Plasmids for expression of BACE457{Delta} and HA-tagged EDEM are described elsewhere (23, 34).

Metabolic Labeling—Cells grown to 60–80% confluence were transfected, and after 20–24 h pulse-chase experiments were performed. Cell were washed twice with PBS containing Ca2+ and Mg2+ (PBS+), and starved for 30 min in serum-free medium lacking cysteine and methionine; labeling was performed in the same medium supplemented with [35S]methionine/cysteine for 10 min. Subsequent chase was performed in minimal essential medium {alpha} supplemented with methionine (5 mM) and cysteine (2.5 mM). At various chase times, labeling was stopped by washing cells in ice-cold PBS+ supplemented with 20 mM N-ethylmaleimide (NEM) on ice. Cells were lysed in 2% CHAPS, HBS (50 mM HEPES, pH 7.5, 200 mM NaCl), 20 mM NEM, 2 mM phenylmethylsulfonyl fluoride, and 10 µg/ml each of chymostatin, leupeptin, antipain, and pepstatin. When used, 10 mM AMS was added to washing and lysis solutions instead of NEM. Various inhibitors were added during the starvation time through all chase incubations unless indicated in the following concentrations: MG132 (50 µM), lactacystin (50 µM), ALLN (260 µM), leupeptin (10 µM), NH4Cl (50 mM), kifunensine (100 µM), swainsonine (100 µM), deoxymannojirimicyn (1 mM), castanospermine (1 mM), tunicamycin (5 µg/ml), okadaic acid (1 µM), and pervanadate (500 µM). Pervanadate was produced freshly before use as described elsewhere (36).

Immunoprecipitation—Cell lysates were cleared by centrifugation at 10,000 rpm for 10 min at 4 °C after solubilization with CHAPS. The postnuclear supernatant was pre-incubated for 1 h on protein A coupled to Sepharose CL-4B-beads at 4 °C. The nuclear pellets were washed once in HBS, resuspended in 1% SDS, and heated to 95 °C for 5 min. When anti-CPY immunoprecipitation was performed, the supernatant was denatured in 1% SDS for 5 min at 95 °C, and then the supernatant or resuspended nuclear pellets were diluted 10-fold in 1% Triton X-100 in HBS buffer (50 mM Hepes, pH 7.5, 200 mM NaCl). Anti-CPY was added together with protein A beads for overnight incubation at 4 °C. Immunoprecipitates were washed three times in 0.1% SDS, 0.05% Triton X-100, 0.3 M NaCl, 10 mM Tris-HCl, pH 8.6, and once in 10 mM Tris-HCl, pH 8.6. The immunocomplexes were eluted in sample buffer containing 100 mM DTT unless indicated otherwise. Anti-calnexin immunoprecipitation was performed as described (37). The immunocomplexes were then eluted in 1% SDS at 95 °C for 5 min and processed for anti-CPY immunoprecipitation as described above. Immunoprecipitates were subjected to 8% SDS-PAGE and phosphorimager analysis. Quantification was performed with ImageQuant software (Amersham Biosciences). Anti-BACE457{Delta} immunoprecipitations were carried out by adding anti-BACE457{Delta}-specific antibody and protein A beads to the postnuclear supernatants. The immunoprecipitates were washed three times in HBS and 0.5% CHAPS and then resuspended in sample buffer before 10% SDS-PAGE.

Sucrose Gradient Centrifugation—Lysates were prepared as described above. The postnuclear supernatant was loaded on top of 10–40% w/w linear sucrose gradient containing 2% CHAPS in HBS. The gradients were centrifuged for 7 h in SW50.1 rotor (Beckman Instruments, Inc., Fullerton, CA) at 40,000 rpm at 4 °C. Fractions were collected manually from the top of the gradient, and subjected to a 1-h pre-incubation on protein A-Sepharose, before CPY was immunoprecipitated under non-denaturing conditions. Pellets were washed twice in HBS, then dissolved in 1% SDS, diluted 10-fold in 1% Triton X-100 in HBS, and subjected to immunoprecipitation with anti-CPY.

Endoglycosyidase H (Endo H) Digestion—Anti-CPY immunoprecipitates were washed as described. After the last wash, 50 mM sodium citrate, pH 5.5, was added plus 1 unit of endoglycosidase Hf (New England Biolabs). The samples were incubated at 37 °C for 1 h, followed by one wash in 0.5% CHAPS in HBS and one in HBS only. Samples were boiled in reducing sample buffer before SDS-PAGE analysis.

Glucosidase II and Jack Bean {alpha}-Mannosidase Assay—Anti-CPY immunoprecipitates were washed as described, followed by incubation with glucosidase II in HBS buffer for 1 h at 37 °C or jack bean {alpha}-mannosidase (Sigma) in 50 mM sodium citrate, pH 4.5, overnight at 37 °C. After glucosidase II treatment, immunocomplexes were washed once in 0.5% CHAPS in HBS and once in HBS alone before boiling in reducing sample buffer and SDS-PAGE.

Immunofluorescence—CHO cells grown on 12-mm coverslips were transfected with pSVHACPY* plasmid DNA as described above. Twenty-four hours after transfection, the cells were fixed with 2.5% formaldehyde in serum-free medium supplemented with 10 mM HEPES, pH 7.4, for 20 min at room temperature. The cells were washed twice with serum-free medium, incubated 10 min at room temperature in PBS+, and permeabilized in PBS+ containing 10% goat serum, 15 mM glycine, and 0.05% saponin for 15 min at room temperature. Washing steps and antibody decoration were carried out with the same buffer. The cells were incubated with a monoclonal anti-CPY (Molecular Probes) at 40 µg/µl and an antibody against marker proteins of various cellular organelles at dilution of 1:100 at room temperature for 1 h. After extensive washing, the cells were incubated with Alexa label-conjugated secondary antibodies (Molecular Probes) diluted 1:200 at room temperature in the dark for 1 h. Further washings were performed in permeabilization solution and once in double-distilled H2O, after which coverslips were mounted on glass slides with Mowiol. Fluorescence microscopy was performed using a Zeiss Axiovert microscope, and image processing with a Hamamatsu CCD camera and the OpenLab software (InVision).


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
CPY* Is Retained in the ER and Degraded—Yeast CPY and CPY* were expressed in CHO cells using a transient transfection system. To ensure proper translocation of these heterologous proteins into the ER of the mammalian cells, we replaced the signal peptide sequence in the yeast enzyme with the signal sequence peptide from influenza virus hemagglutinin, which has been shown to support efficient translocation of CPY into the mammalian ER (35). Twenty-one hours after transfection with the wild-type or mutant constructs (pSVHACPY or pSVHACPY*, respectively), the cells were subjected to pulse labeling with [35S]methionine/cysteine for either 2 or 10 min, and chased for time periods up to 4 h. The cells were then solubilized with CHAPS, and postnuclear lysates were subjected to immunoprecipitation using anti-CPY antibodies. The medium was also collected and immunoprecipitated.

SDS-PAGE of the immunoprecipitates followed by phosphorimager analysis showed that most of the wild-type CPY was rapidly secreted into the medium. The protein disappeared from the cell lysate (Fig. 1A, lanes 1–5) and appeared in the medium fraction (lanes 7–10) with a half-time of less than 30 min. Judging by the difference in gel mobility of the secreted and intracellular CPY (compare lanes 1 and 7–10), and by the acquisition of Endo H resistance (data not shown), the N-linked glycans were processed in the Golgi complex. Thus, although expressed as a heterologous protein, CPY was properly targeted to the ER, it passed the quality control, but instead of being transported to the lysosomes (vacuole) as in yeast it was rapidly secreted (35).



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  		FIG. 1.
Pulse-chase analysis of wild-type (CPY) and mutated (CPY*) carboxypeptidase Y. A, wild-type CPY was transiently expressed in CHO cells. After a 2-min pulse, cells were chased for the indicated times. Aliquots of cell lysates and media were immunoprecipitated with anti-CPY antibody and analyzed by SDS-PAGE under reducing conditions. sCPY indicates secreted CPY. B, mutant CPY (CPY*) was similarly expressed and examined after 10-min pulse and indicated chase periods. Lysates and media were treated as in A. C, aliquots of immunoprecipitated CPY* were subjected to endoglycosidase H digestion (H) or Peptide:N-glycosidase F (PNGase F) digestion (F). The latter cleaves off the entire N-linked glycan also after Golgi processing. Sample in lane 2 was incubated in Endo H buffer without enzyme. deCPY* refers to deglycosylated CPY*. D, kinetics of degradation of CPY* in CHO cells. The graph shows the percentage of CPY* bands relative to time 0. Each point represents the average of at least four independent experiments. E, the amount of immunoprecipitated CPY* detected in the postnuclear supernatant (PNS) and the pellets was quantified and plotted. Aliquots of lysate and pellet correspond to the same amount of cells. Each bar represents the average of five independent experiments. F, wild-type (wt) and mutant (*) CPY were transiently expressed in CHO cells. After a 5-min pulse, cells were chased for the indicated times. Cell lysates were subjected to immunoprecipitation, and immunocomplexes were analyzed by SDS-PAGE under reducing (Red) or non-reducing (NonRed) conditions as indicated. The arrow points to the Golgi-processed wild-type CPY. G, CHO cells expressing CPY* were labeled for 10 min in absence or presence of 5 mM DTT, washed, and lysed in presence or absence of AMS. After immunoprecipitation, samples were analyzed by non-reducing SDS-PAGE. Red CPY* indicates the fully reduced, conjugated CPY*. CPY*-AMS refers to different oxidized forms, which are modified by AMS to various extent. H, CHO cells expressing wild-type and mutant CPY were labeled for 1 h. Postnuclear supernatants were loaded on 10–40% linear sucrose gradients as described under "Experimental Procedures." CPY was recovered by immunoprecipitation from each fraction and analyzed by reducing SDS-PAGE. The relative amount of wild-type and mutant CPY is quantified as a percentage of the sum of the amount in the individual fractions. Fractions were loaded from top (1) to bottom (13). P refers to pellet fraction.

 
In contrast, no secretion could be observed when CPY* was expressed (Fig. 1B, lane 1); labeled CPY* was only found in the cell lysate fraction after pulse and chase (lanes 2–7). The mobility difference in the CPY* band, observed before and after treatment with Peptide:N-glycosidase F (PNGase F) or Endo H (Fig. 1C, compare lanes 1, 3–5, and 2), indicated that the CPY* was glycosylated and therefore correctly targeted to the ER. Judging by the size of the shift after glycan removal, all four consensus glycosylation sites were probably used. That CPY* was undergoing some glycan trimming was suggested by a small, gradual drift in the position of the band with time (Fig. 1B, lanes 3–7). The small band appearing at lower molecular weight probably corresponded to a minor fraction of unglycosylated, untranslocated CPY*.

Endo H digestion showed that the CPY* remained sensitive throughout the 4-h chase period (Fig. 1C). This indicated that the protein was retained in a pre-Golgi compartment. Indirect immunofluorescence using anti-CPY showed that the CPY* staining pattern overlapped with that of the ER visualized by co-staining with an anti-calnexin antibody (Fig. 2, A–C). CPY* localization was distinct from that of Golgi marker, Golgi mannosidase II, in agreement with the fact that CPY* remains Endo H-sensitive, and thus never reaches the medial Golgi (Fig. 2, G–I). Co-staining with the ERGIC marker p58, showed that the majority of CPY* did not overlap with p58, although we could not exclude some co-localization (D-F).



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  		FIG. 2.
Intracellular localization of CPY*. CPY*-expressing cells were double-stained with anti-CPY antibody (A, D, and G) (red) together with anti-calnexin (B) (green) or anti-p58 (E) (green) or anti-Golgi mannosidase II (H) (green) antibodies.

 
The cause for CPY* retention in the ER was most likely its inability to fold properly. To analyze the folding status of CPY*, we performed SDS-PAGE under non-reducing conditions comparing wild-type CPY and CPY*. Both showed slightly heterogeneous migration patterns. That they had mobilities somewhat faster than the fully reduced protein suggested that they had acquired intrachain disulfide bonds (Fig. 1F) (38).

To test whether all intramolecular disulfides had formed in CPY*, we treated it with the thiol-conjugating reagent 4-acetamido-4'-maleimidylstilbene-2,2'-disulfonic acid (AMS), which reacts with free sulfhydryl groups increasing the molecular mass of the target protein by ~0.5 kDa for each AMS molecule added (39). After radiolabeling, CPY*-expressing cells were lysed in presence of AMS and immunocomplexes were visualized in SDS-PAGE under non-reducing conditions. The protein displayed an even more heterogeneous migration with several discrete bands. The mobility of these bands (Fig. 1G, lane 1) compared with the band obtained when the fully reduced CPY* was conjugated (lane 3) confirmed that CPY* occurred in the ER as a mixture of differently oxidized forms lacking the full complement of disulfide bonds.

In the non-reduced SDS-PAGE, there was no indication of covalent disulfide cross-linked aggregates like those often seen for misfolded proteins in the ER (Fig. 1F). To test whether CPY* was present in non-covalently associated aggregates, aliquots of CHAPS lysates from pulse-labeled cells were subjected to sucrose gradient velocity centrifugation (Fig. 1H). CPY* and CPY were found in a slowly sedimenting peak (S = 3.6) corresponding to the monomeric enzyme. Taken together, the results indicated that CPY* was retained in the ER as a mixture of incompletely folded, heterogeneously oxidized monomers.

Degradation Occurs by Multiple Pathways—Although CPY* was not secreted, the amount of labeled enzyme in the cell lysates decreased rapidly with time (Fig. 1, B, lanes 3–7, and D). Because only insignificant amounts of CPY* were found in the nuclear pellet (Fig. 1E), the loss must have been caused by degradation. A detailed time course indicated that degradation started after a 45-min lag, and proceeded rapidly to virtual completion within the next 3 h (Fig. 1D). Half of the protein was degraded after 88 min.

In yeast, CPY* is degraded after retrotranslocation from the ER lumen by proteasomes in the cytosol (9). To investigate whether this was also the case in CHO cells, we performed the pulse-chase experiments described above in cells treated with the reversible proteasome inhibitor MG132. We consistently found, as shown in Fig. 3A (lane 6), some inhibition; 20–30% of the protein was protected by the inhibitor over a 4-h period. Additionally, in the presence of the more specific irreversible proteasome inhibitor lactacystin (40), degradation was only partially blocked (compare lanes 4 and 8). ALLN, another non-specific inhibitor of the proteasome, had no effect at all (data not shown). To test whether the inhibitors needed longer time to be functional, we included a 90-min pre-incubation step in the presence of the proteasome inhibitors. Even in this case the inhibition was weak.



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  		FIG. 3.
Effect of various protease inhibitors on CPY* degradation. A, CPY*-expressing cells were pre-incubated for 30 min, pulsed for 10 min, and chased up to 4 h in the absence or presence of different protease inhibitors (MG, MG132; LC, lactacystin; L, leupeptin; N, NH4Cl). Cell lysates were processed as described in Fig. 1A. B, quantification of CPY* bands was performed on three different experiments. Percentage of the amount recovered under different treatments was expressed as referred to the amount at time 0 for each experimental condition. C, CHO cells co-transfected with CPY* and BACE457{Delta} were labeled for 10 min and chased for 4 h in the absence or in the presence of MG132. Equal amounts of post-nuclear supernatant were immunoprecipitated with anti-BACE457{Delta} and anti-CPY, respectively. Quantification of the bands of BACE457{Delta} and CPY* (data not shown) was expressed as percentage of time 0 (100%).

 
The effect of proteasome inhibitors was thus limited to a partial block in CPY* degradation only. That the MG132 preparations used were functional was confirmed by following the degradation of BACE457{Delta} (15, 24) co-transfected together with CPY* in CHO cells. The proteasome inhibitor could block BACE457{Delta} disposal 2.8-fold more than CPY* degradation (Fig. 3C). We concluded that, although some of the CPY* was degraded by proteasomes, the majority was degraded by a pathway that did not depend on the proteasome.

To characterize the proteasome-independent, alternative pathway, we tested inhibitors of lysosomal proteolysis, leupeptin and NH4Cl (Fig. 3A, lanes 9–12). Neither had a significant effect on degradation of CPY* (Fig. 3B). Recently it has been described that the PI Z variant of human {alpha}1-antitrypsin is degraded via a proteasome-independent pathway that involves tyrosine phosphatases (41). To test this possibility, we added the tyrosine-phosphatase inhibitor sodium pervanadate during 4-h chase. The inhibitor showed no effect on CPY* degradation (data not shown). We obtained the same result incubating cells with okadaic acid, a general serine/threonine phosphatase inhibitor (data not shown).

It has been reported that protein synthesis inhibitors like cycloheximide prevent degradation of some proteins (42–44). We tested the effect of cycloheximide, adding it to the chase medium, but did not observe any protective effect (data not shown). Thus, we could not find any obvious similarity in respect to inhibitor sensitivity between the pathway by which the majority CPY* was degraded and those previously described.

CPY* Oligosaccharides Are Processed in CHO Cells—During the chase period, the increase in electrophoretic mobility indicated that CPY* was undergoing gradual modification (Figs. 1B, lanes 3–7, and 3A3, lanes 1–4). It clearly involved changes in the oligosaccharides because removal of the N-linked glycans with Endo H produced molecules with identical mobility (Fig. 1C).

To test whether glucose trimming was involved, we added castanospermine to block glucosidases I and II (45). The lower mobility observed (Fig. 4A, compare lanes 1 and 3) confirmed that glucose trimming was at least partly responsible for the shift in CPY* mobility. Addition of an ER mannosidase I inhibitor, kifunensine, which blocks the removal of a middle branch (B-branch) mannose (45), also decreased the shift in CPY* mobility (Fig. 4A, compare lanes 4 and 6), indicating that CPY* was a substrate for ER mannosidase I. The same result was obtained with the inhibitor of both ER mannosidase I and II activities, deoxymannojirimycin (DMJ), whereas the specific inhibitors of ER mannosidase II, swainsonine and 1,4-dideoxy-1,4-imino-D-mannitol, did not block the shift in CPY* mobility (data not shown). However, when castanospermine and kifunensine were applied together, a slight shift was still observed at 4 h of chase time. This may suggest removal of mannoses from the C-branch by ER mannosidase II (Fig. 4A, compare lanes 3 and 4 with lanes 7 and 8). Taken together, the results indicated that glucoses in the A-branch and a mannose residue in B-branch, and possibly a mannose in the C-branch, were trimmed from CPY*.



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  		FIG. 4.
Kifunensine blocks proteasome-independent CPY* degradation. A, CHO cells expressing CPY* were starved for 30 min, pulsed for 10 min, and chased for 4 h with the indicated inhibitors eventually in combination (C, castanospermine; K, kifunensine; MG, MG132; L, leupeptin). Cells were then lysed, and then CPY* was immunoprecipitated with anti-CPY antibody before SDS-PAGE analysis. B, the radioactivity was detected and quantified using a phosphorimager device. The figure represents the result of the average of three identical experiments. C, CHO cells expressing CPY* alone or with HA-tagged EDEM were starved for 30 min, pulsed for 10 min, and chased for the indicated times. Post-nuclear supernatants were subjected to anti-CPY immunoprecipitation. D, kinetics of CPY* degradation has been compared in control cells in the absence (–) or presence of MG132 (MG132) and in cells co-expressing HA-tagged EDEM in the absence (EDEM) or presence of MG132 (EDEM+MG132). E, CHO cells were treated as in A, except that in the MG132 treatment, before the starvation period, cells were pre-incubated for 30 min in presence of the same compound. TM, tunicamycin.

 
Furthermore, we noticed that, immediately after the pulse period, CPY* had the same electrophoretic mobility in the presence and absence of kifunensine, and it was faster than after castanospermine pretreatment (Fig. 4A, compare lanes 1, 3, and 5). This suggested that the removal of the two outermost glucoses had already occurred during the pulse, and that it took place before the mannose trimming.

Inhibitor of ER Mannosidase I Inhibits Non-proteasomal Degradation—Kifunensine not only changed the electrophoretic mobility of CPY* but it also inhibited degradation. This is evident when comparing lanes 4 and 6 in Fig. 4A, or when viewing the quantitative data in Fig. 4B. After 4 h, half of the CPY* remained undegraded. An effect by this ER mannosidase I inhibitor was not unexpected, because it has been previously observed to inhibit ER degradation of a variety of glycoproteins, and there is growing evidence that mannose trimming plays a role in the recognition of glycoprotein substrates for ER degradation by the proteasome pathway (46, 47).

To determine whether it was the proteasome-mediated or the proteasome-independent degradation pathway that was affected by kifunensine, we added the drug together with the proteasome inhibitor MG132 (Fig. 4, A, lanes 9 and 10 and quantification in B) or lactacystin (data not shown). In both cases, degradation of CPY* was inhibited almost entirely. When kifunensine was combined with leupeptin, there was no such synergic effect (Fig. 4A, lanes 11 and 12).

The simplest interpretation was that the proteasome-independent pathway was blocked by kifunensine, and the proteasome-dependent pathway was not. If so, targeting to the proteasome-independent pathway required the removal of a mannose form the B-branch. Sorting to the proteasome pathway did not require mannose removal. With the proteasome-independent pathway inhibited by kifunensine, the proteasome-dependent pathway became, in fact, somewhat more efficient in degrading CPY*.

The role of the Man8GlcNAc2 isomer B glycan and of the putative mannose lectin EDEM in ERAD has been previously described for other glycoprotein substrates (23, 24, 48). However, when we co-expressed a HA-tagged version of EDEM with CPY*, we could not detect any effect on the kinetics of degradation (Fig. 4, C and D, compare Control (- in C) and EDEM). EDEM expression was verified by immunoprecipitation and Western blot with an antibody against the HA tag (data not shown). Because our data showed that it was the proteasome-independent pathway that required the formation of the Man8GlcNAc2 isomer B, we applied the proteasome inhibitor MG132 to cells expressing both CPY* and EDEM. In this condition, we observed the same weak inhibitory effect on CPY* degradation at late chase time that we detected in control cells treated with MG132 (D, compare MG132 and EDEM+MG132). Moreover, the non-proteasomal degradation occurring in presence of MG132 was not significantly affected by expression of HA-tagged EDEM (D, compare MG132 and EDEM+MG132).

DMJ, an inhibitor reported to block both ER mannosidases I and II, had a similar effect, but was less efficient. To reach an inhibitory effect comparable to kifunensine with DMJ, we either had to add fresh inhibitor every hour during the chase, or at much higher concentrations (2–5 mM). Further, we verified that, even in the presence of cycloheximide, CPY* degradation was prevented by the addition of kifunensine (data not shown). This indicated that the effect could not have been indirect via other newly synthesized proteins.

To exclude unspecific effects of kifunensine, and to confirm that its action was related to the presence of glycans, pulse-chase experiments were performed in the presence of tunicamycin, which inhibits the transfer of N-linked oligosaccharides to proteins. The non-glycosylated CPY* was degraded similarly to the control, but the degradation was not affected by kifunensine. Proteasome inhibitors blocked non-glycosylated CPY* degradation at the same level as the glycosylated form (Fig. 4C). This suggested the presence of proteasome-independent pathways as well for this non-glycosylated protein, an observation that we did not pursue further.

The Role of Glucose Residues and Calnexin—As with other incompletely folded glycoproteins, CPY* could be expected to enter the calnexin/calreticulin cycle when expressed in a mammalian cell. In S. cerevisiae, where the protein is normally expressed, this cycle does not exist. To test the association with calnexin in CHO cells, we used a double immunoprecipitation protocol in which precipitates obtained with anti-calnexin were solubilized and re-precipitated with anti-CPY. We found that some labeled CPY* was in fact co-immunoprecipitated with calnexin. The presence of CPY*-calnexin complexes persisted over the entire 4-h period of chase (Fig. 5A, lanes 1–3). Because of the difficulty in performing the sequential immunoprecipitation quantitatively, it was not possible to estimate exactly how large the fraction bound to calnexin was.



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  		FIG. 5.
CPY* associates with calnexin. A, CPY*-expressing cells were pre-incubated for 30 min, pulsed for 10 min, and chased up to 4 h in the absence or presence of different inhibitors (K, kifunensine; MG, MG132; LC, lactacystin). Cell lysates were subjected to anti-calnexin immunoprecipitation, followed by anti-CPY immunoprecipitation after washing under non-stringent conditions. Immunocomplexes were analyzed by reducing SDS-PAGE. B, cells were treated as in Fig. 4A in the absence or in the presence of kifunensine (K). Immunocomplexes were subjected to treatment with glucosidase II (GII), jack bean {alpha}-mannosidase (JBM), or both as described under "Experimental Procedures." C, kinetics of degradation of CPY* was determined in control cells and in cells treated with castanospermine from the starvation through the whole chase period (C) or only during the chase (T). D, quantification of gel in C. The graph is representative of three identical experiments. E, CPY*-expressing cells were treated with castanospermine from the starvation through all chase periods (C) or only during the chase (T) eventually in combination with MG132 (MG). Samples were analyzed in SDS-PAGE under reducing conditions after performing immunoprecipitation.

 
The presence of CPY*-calnexin complexes meant that some of the CPY* oligosaccharides must have been monoglucosylated, i.e. they must have contained only the innermost glucose essential for binding to this lectin chaperone (49). The presence of glucose was confirmed by digestion of the immunocomplexes with jack bean {alpha}-mannosidase (50), and by digestion with purified ER glucosidase II (Fig. 5B). Here, we analyzed CPY* mobility on SDS-PAGE at different chase times after a 10-min pulse. Soon after the pulse, glucosidase II treatment caused a small but reproducible shift in the CPY* mobility, suggesting the presence of glucose containing glycans (Fig. 5B, compare lanes 1 and 2). When the glucosidase II-treated samples were incubated with jack bean {alpha}-mannosidase, the CPY* band migrated faster compared with CPY* from samples treated with jack bean {alpha}-mannosidase alone (Fig. 5B, compare lanes 3 and 4). This demonstrated that the CPY* glycans still contained glucose residues. The combined glucosidase II and jack bean {alpha}-mannosidase treatment showed that the innermost glucose was present in most of the CPY*. After 2 h of chase the situation was similar (lanes 5–8). Thus, the majority of CPY* in the ER occurs in glucosylated form until it is degraded, and at least some of it is bound to calnexin. In presence of kifunensine, we showed that CPY* remained monoglucosylated at least up to 2 h of chase (Fig. 5B, lanes 9–16).

When cells were incubated with kifunensine, CPY* binding to calnexin at 4 h increased, and the bound fraction increased further if proteasome inhibitors were also present (Fig. 5A, lanes 4–12). We interpreted these findings as indicating that CPY* accumulates in calnexin-complexes when degradation is blocked or slowed down.

Published data on the role of calnexin in ERAD have been conflicting. Although some have suggested calnexin-substrate complexes as intermediates toward ERAD (18, 25), other reports indicate that calnexin binding protects glycoproteins from degradation (26, 51). To test the situation in our system, we used castanospermine to block glucosidase I and II and to prevent CPY* binding to calnexin. The results shown in Fig. 5 (C and D) indicated that CPY* was still efficiently degraded but the initial lag phase during which the protein is almost stable no longer appeared. This means that the half-time of degradation was 30 min instead of 88 min. The simplest interpretation of this result was that, when CPY* is allowed to associate with the calnexin/calreticulin cycle, it is indeed protected. The effect is temporary, however, and only manifested during the lag phase. If calnexin and calreticulin binding as such is what protects CPY* during the lag, one would expect that prolonging the time of interaction with these lectins would extend the lag period. To prolong association with the lectins, we added castanospermine only after the pulse and not already before it. Delayed addition of a glucosidase inhibitor freezes glycoproteins in the monoglucosylated form, thus prolonging their interaction with calnexin and calreticulin (52). We verified the formation of calnexin-CPY* complexes by co-immunoprecipitation (data not shown) and determined the rate of degradation. We found that CPY* was degraded with the same kinetics as in controls in which no castanospermine was added (Fig. 5D). We also showed that the proteasome was still partially responsible for degradation of CPY* in cells treated with castanospermine (Fig. 5E, compare lanes 8 and 10 and lanes 15 and 17).

The result indicated that the mechanism that leads to the lag in degradation of CPY* is a complex one. Although clearly dependent on trimming of one or more of the three glucose residues, it was apparent that it was not merely caused by the trimming-induced association of CPY* with calnexin and calreticulin. It is probable that degradation is regulated by interplay of glucose and mannose trimming enzymes and multiple lectins in the ER that recognize, bind, and modify the different glycoforms. Further studies are needed to clarify the underlying mechanisms.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
Given the high volume of proteins synthesized in the ER and the large fraction of misfolded and unassembled proteins generated, the ERAD plays a central role during active secretion, cell growth, and normal turnover in eukaryotic cells. The only well established, central pathway for ERAD so far depends on retrotranslocation of misfolded protein substrates and degradation by cytosolic proteasomes. Our results with CPY* indicate that in mammalian cells there are other pathways, and that ERAD is more complex than initially anticipated.

When expressed as a heterologous protein in CHO cells, we established that, unlike wild-type CPY, the CPY* mutant failed to undergo complete oxidative folding and secretion. The protein was recognized as misfolded by the ER quality control system, it was retained, and after a lag period of ~45 min it was efficiently degraded by ERAD. Although misfolded and heterogeneous in its disulfide pattern, CPY* was a "well behaved" model protein in the sense that it remained monomeric in the ER. The four glycans, moreover, underwent synchronized changes during ER trimming. Glucoses were removed, and there was gradual loss of mannoses as a result of ER mannosidase I and possibly ER mannosidase II. A fraction of the CPY* was always associated with calnexin, indicating that the glycans in most of the proteins were monoglucosylated.

The effect of proteasome inhibitors showed that, in contrast to the situation in S. cerevisiae, only approximately one third of the CPY* was degraded by proteasomes. The remaining two thirds were degraded by a distinct pathway, characterized by its insensitivity to all proteasome inhibitors tested and by its sensitivity to the ER mannosidase I inhibitor kifunensine. Another ER mannosidase I inhibitor, deoxymannojirimycin, although less efficient, was also inhibitory. The pathway was not affected by reagents that inhibited lysosomal proteases, protein synthesis, or tyrosine phosphatases.

The existence of alternate ERAD pathways has been previously proposed particularly in studies on mammalian cells (41, 42, 44, 53–55). Sifers and co-workers found that misfolded human {alpha}1-antitrypsin can be disposed by distinct pathways depending on the allele; the null Hong Kong mutant is degraded by the proteasome (18), whereas the PI Z mutant is degraded by yet unidentified protease(s) and degradation is sensitive to tyrosine-phosphatase inhibitors (41). Human thyroperoxidase has also been reported to be degraded by two pathways, one sensitive to proteasomal inhibitors, and the other to inhibitors of protein synthesis and cysteine/serine proteases (44). Moreover, CPY* itself has been proposed to be degraded in part via a proteasome-independent pathway in yeast (17).

Aside from the lack of inhibition by proteasome inhibitors, the alternative pathway for CPY* degradation had no obvious resemblance with these reported alternate pathways. It is, however, reasonable to think that cells may be equipped with several possible ways to avoid problems with ER overloading, and that the use of these systems may depend on the properties of the substrate proteins. The presence of glycans, the aggregation state, membrane association, and the association with distinct chaperones may, for example, be important. Alternative degradation may take place in the cytosol, where, for example, the giant protease TPP II may take over proteasome functions (56), or it can occur inside the ER lumen. The nature and location of the proteases involved in CPY* degradation by the alternate pathway in our system remain open.

That mannose trimming can serve as a determinant for glycoprotein ERAD is already known from inhibitor studies and mutant analysis in yeast and mammalian cells (17, 18, 20, 57). Glycoprotein degradation in the proteasome has been found to be sensitive to kifunensine and blocked by mutations in ER mannosidase I. These findings have led to the notion that generation of Man8GlcNAc2 isomer B serves as a signal for degradation. It cannot be the only signal, however, because proteasome-mediated degradation of certain glycoproteins proceeds normally in the presence of ER mannosidase I inhibitors (19, 41).

In contrast to previous reports, our observations indicated that inhibition of ER mannosidase I with kifunensine selectively blocked the proteasome-independent and not the proteasome-dependent pathway of CPY* degradation. Kifunensine had no effect on ER-degradation of non-glycosylated CPY* confirming that the effect was mediated by the changes in the oligosaccharide moieties. Thus, it appears that the sorting principle used in other cells and for other proteins to tag substrates for delivery to proteasomes is used for CPY* targeting to an alternative pathway.

It has been hypothesized the system that recognizes the Man8GlcNAc2 isomer B tag on glycoproteins is a mannose lectin (18). A candidate protein to fulfill this function, called EDEM (HtmI/MnlI in yeast), has recently been identified, and shown to be responsible for efficient degradation of some glycoproteins (21–23). Overexpression of EDEM in mammalian cells accelerates ERAD (23, 24, 48). However, it remains obscure how the lectin distinguishes between native and non-native substrates. So far the only ER factor known to serve as a glycoprotein-specific folding sensor is the UDP-glucose:glycoprotein glucosyltransferase (58), but there is no evidence that this enzyme takes part in the ERAD sorting directly.

We observed that expression of heterologous EDEM did not significantly accelerate CPY* degradation. This was also true when we monitored the proteasome-independent pathway after the addition of proteasome inhibitors. This observation suggests that the pathway may not involve EDEM as a rate-determining factor.

The glucose trimming status of glycoproteins has also been reported to influence degradation of certain glycoproteins, suggesting an involvement of the calnexin/calreticulin cycle in ERAD. However, no consistent picture has emerged; in some cases accelerated degradation is observed when a glycoprotein is prevented from entering the cycle (26, 51), whereas in other cases interaction with the cycle accelerated degradation (18, 25). There are also cases in which glucose trimming has been shown to have no effect on ERAD (20, 43).

Our results showed that glucose trimming influenced the degradation of CPY* by securing an initial lag period of 45 min during which the protein, although misfolded, was protected. Such a "timer effect" has often been seen for newly synthesized proteins in the ER, and is probably important because it gives proteins an initial period during which they can fold, mature, and assemble without risk of degradation (2, 59). How this lag is regulated has not been previously analyzed. Because inhibition of glucose trimming also prevented CPY* entry into the calnexin/calreticulin cycle, it is likely that inclusion into the cycle somehow protects CPY* from degradation. Exit of a glycoprotein from the cycle is regulated by the interplay between two opposing enzymes, glucosidase II and UDP-glucose:glycoprotein glucosyltransferase (60), which in turn are sensitive to the mannose trimming status of the glycans (61, 62). Consistent with this explanation, we observed that the addition of kifunensine to block the alternative degradation pathway resulted in an increase in the fraction of CPY* co-immunoprecipitated with calnexin. However, that post-pulse addition of castanospermine, known to prolong glycoprotein interaction with calnexin and calreticulin (63), did not extend the lag phase, indicating that there is more to the regulation than we can explain by simple models.

From our data we could, further, conclude that the proteasomal pathway of CPY* degradation is not dependent on the presence or the processing of N-linked glycans. Degradation of non-glycosylated CPY* chains was as sensitive to proteasome inhibitors as degradation of glycosylated CPY*. The importance of N-linked glycans appeared in the alternative route, which was blocked by kifunensine. Both glycosylated and non-glycosylated CPY* were degraded in a non-proteasomal manner, but only disposal of the glycosylated protein was sensitive to kifunensine. A possible explanation could be that both forms are degraded via the same protease, but the kifunensine-sensitive step works upstream in the process where the glycosylated CPY* are sorted for degradation. Alternatively, a different protease is responsible for the turnover of non-glycosylated CPY*.

In conclusion, our data strengthen the growing evidence that ERAD in mammalian cells involves distinct degradation pathways. Retrotranslocation and the cytosolic proteasomes are an important element in ERAD, but they do not constitute the only possible pathway. Moreover, our data confirm that the sorting process that targets glycoproteins for ERAD depends on specific mannose and glucose trimming events. However, in the case of CPY* in CHO cells these changes affect the alternate pathways, not the proteasomal one. We also found that trimming of the core glycans is used as part of a system that sets the timer for the onset of degradation of the newly synthesized glycoproteins. Further studies are required to elucidate the nature of the alternative route(s) and their regulation.


   FOOTNOTES
 
* This work was supported in part by grants from the Swiss National Foundation. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

§ Supported by the Swiss Federal Institute of Technology, Zurich. Back

|| To whom correspondence should be addressed. Tel.: 41-1-6326817; Fax: 41-1-6321269; E-mail: ari.helenius{at}bc.biol.ethz.ch.

1 The abbreviations used are: ER, endoplasmic reticulum; CPY, carboxypeptidase Y; CPY*, mutant carboxypeptidase Y; ERAD, endoplasmic reticulum-associated degradation; PBS, phosphate-buffered saline; CHO, Chinese hamster ovary; NEM, N-ethylmaleimide; DTT, dithiothreitol; AMS, 4-acetamido-4'-maleimidylstilbene-2,2'-disulfonic acid; MG132, carbobenzoxy-L-leucyl-L-leucyl-L-leucinal; ALLN, N-acetyl-leucyl-leucyl-norleucinal; DMJ, deoxymannojirimycin; HBS, Hepes-buffered saline; Endo H, endoglycosidase H; HA, hemagglutinin; CHAPS, 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonic acid. Back


   ACKNOWLEDGMENTS
 
We thank Dr. M. Kowarik for purified glucosidase II, Dr. J. Saraste for anti-p58, Dr. K. Nagata for the EDEM-expressing vector, and Dr. M. Molinari for BACE457{Delta} construct and antibody.



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