* Division of Cell and Molecular Pathology, Department of Pathology, University of Zürich, CH-8091 Zürich, Switzerland; and Mikrobiologisches Institut, ETH Zürich, CH-8092 Zürich, Switzerland
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Abstract |
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In Saccharomyces cerevisiae, transfer of
N-linked oligosaccharides is immediately followed by
trimming of ER-localized glycosidases. We analyzed
the influence of specific oligosaccharide structures for
degradation of misfolded carboxypeptidase Y (CPY).
By studying the trimming reactions in vivo, we found
that removal of the terminal 1,2 glucose and the first
1,3 glucose by glucosidase I and glucosidase II respectively, occurred rapidly, whereas mannose cleavage by mannosidase I was slow. Transport and maturation of
correctly folded CPY was not dependent on oligosaccharide structure. However, degradation of misfolded
CPY was dependent on specific trimming steps. Degradation of misfolded CPY with N-linked oligosaccharides containing glucose residues was less efficient compared with misfolded CPY bearing the correctly
trimmed Man8GlcNAc2 oligosaccharide. Reduced
rate of degradation was mainly observed for mis-
folded CPY bearing Man6GlcNAc2, Man7GlcNAc2
and Man9GlcNAc2 oligosaccharides, whereas
Man8GlcNAc2 and, to a lesser extent, Man5GlcNAc2
oligosaccharides supported degradation. These results
suggest a role for the Man8GlcNAc2 oligosaccharide in
the degradation process. They may indicate the presence of a Man8GlcNAc2-binding lectin involved in targeting of misfolded glycoproteins to degradation in S.
cerevisiae.
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Introduction |
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IN Saccharomyces cerevisiae, as in other eukaryotes, the
synthesis of asparagine-linked glycoproteins takes
place in the ER. After transfer to protein, the N-linked
oligosaccharide (NLO),1 while present in the ER, is subject to trimming reactions (see Fig. 1) involving glucosidase I, glucosidase II, and mannosidase I (Herscovics and
Orlean, 1993; Moremen et al., 1994
; Roth, 1995
). In higher
eukaryotes, a specific role of the trimming intermediate Glc1Man9GlcNAc2 oligosaccharide in the ER quality control process has been proposed (Helenius et al., 1997
). An
incorrectly folded glycoprotein bearing such an oligosaccharide structure is bound by specific ER resident proteins
and retained in a folding competent environment. Correctly folded glycoproteins can exit the ER, enter the
Golgi apparatus, and are delivered to their final destination. However, improperly folded glycoproteins are retained in the ER and are eventually degraded. In many
cases, degradation occurs via the ubiquitin-proteasome
pathway that requires their exit from the ER lumen to the
cytosol as shown both in higher eukaryotic cells and in
yeast (Jentsch and Schlenker, 1995
; Bonifacino, 1996
; Kopito, 1997
; Sommer and Wolf, 1997
; Varshavsky, 1997
).
For the export to the cytosol, constituents of the ER translocon play an important role (Pilon et al., 1997
; Plemper
et al., 1997
). Additionally, ER proteins such as the chaperone Kar2p (Plemper et al., 1997
), as well as the ubiquitin-conjugating proteins Ubc6p and Ubc7p, are thought to be
involved in this process (Biederer et al., 1996
; Hiller et al.,
1996
). In Saccharomyces cerevisiae, the proteolysis of nonglycosylated
-factor is ATP and cytosol-dependent (McCracken and Brodsky, 1996
) and also mutated and therefore misfolded carboxypeptidase Y (prc1-1, CPY*; Wolf
and Fink, 1975
; Finger et al., 1993
) has been shown to enter the ubiquitin-proteasome pathway (Hiller et al., 1996
).
The degradation of the misfolded protein appears to be
glycosylation dependent, since nonglycosylated CPY* remains stable in the ER (Knop et al., 1996
). Moreover, the degradation also appears to be mannosidase I-dependent
(Knop et al., 1996
). Despite this, the molecular signals required for the initiation of ER glycoprotein degradation
are not known.
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We investigated the possible role of specific oligosaccharide structures in degradation of CPY* by genetic tailoring of the protein-bound oligosaccharide structure. We
found that the Man8GlcNAc2 structure as the final product
of the trimming reaction in the ER in yeast (Byrd et al.,
1982) was mandatory for efficient degradation. Our results
suggest that the ER
1,2-mannosidase represents a key
enzyme for timing the onset of degradation. The period required for complete oligosaccharide trimming appears to
be the time frame for glycoproteins to fold correctly.
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Materials and Methods |
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Materials
Strains used are detailed in Table I. Wild-type denotes a strain with both normal biosynthesis of lipid-linked oligosaccharides and trimming of protein-bound oligosaccharides but harboring the prc1-1 mutation. Oligonucleotides (Microsynth, Balgach, Switzerland) used for gene deletion and screenings are listed in Table II. The integrative plasmid pRS306-prc1-1 containing the mutated CPY gene was provided by Dr. D.H. Wolf (University of Stuttgart, Germany). The antiserum against yeast hexokinase was provided by Dr. S. Schröder (Biozentrum, University of Basel, Switzerland).
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Yeast Manipulations
Standard protocols were followed for growth of yeast, mating, sporulation, and ascus dissection (Guthrie and Fink, 1991). If not otherwise
stated, the cells were grown at 30°C in either YPD medium (2% Bacto-Peptone, 1% Yeast extract [both from Difco Laboratories, Detroit, MI],
2% glucose) or for metabolic labeling experiments in MV medium (0.67%
Yeast nitrogen base [Difco Laboratories], 2% glucose and the appropriate
supplements).
Construction of Strains
Disruption of the MNS1 Locus.
The MNS1 locus ORF YJR131w (these
data are available from GenBank/EMBL/DDBJ under accession number
Z49631; Grondin and Herscovics, 1992) was inactivated by replacing a
major part of the locus with the KanMX cassette (Wach et al., 1994
). The
sequence of the kanamycin resistance gene was amplified by PCR by using the template pFA6a-KanMX4 plasmid (Wach et al., 1994
) and the primers MNS1forKan and MNS1revKan (Table II). The resulting DNA was transformed into strain SS328 and the cells were selected on G418
plates (200 µg/ml). Transformants were analyzed for correct integration
by whole cell PCR (Sathe et al., 1991
) using KanMXu and the MNS1-specific MNS1-68u and MNS1 + 431L primers.
Disruption of the ALG12 Locus.
The ALG12 locus ORF YNR030w
(these data are available from GenBank/EMBL/DDBJ under accession
number Z71645; Lussier et al., 1997) was inactivated by the same procedure using the primers ALG12forKan and ALG12revKan for amplifying
the KanMX cassette and KanMXu and ALG12for primers for verifying
the correct gene deletion (Table II).
Replacement of the PRC1 Locus with prc1-1.
The BglII-linearized plasmid pRS306-prc1-1 (Knop et al., 1996) containing the mutated form
(G255R) of CPY (Wolf and Fink, 1975
) was integrated into the PRC1 locus of various yeast strains, resulting in a duplication of the PRC1 locus.
Strains in which an excision of the duplication by homologous recombination had occurred were selected on 5-FOA plates and the resulting colonies screened by PCR for the prc1-1 locus. A fragment of the PRC1 locus was amplified by PCR using the primers CPY462u and CPY855L (Table II) giving raise to a product of 423 bp. Due to the prc1-1 mutation, a BstXI
restriction site is destroyed. Therefore, strains containing solely the prc1-1
locus were identified by the resistance of the PCR fragment towards
BstXI digestion. Western blot analysis confirmed that they only expressed
mutant CPY*.
Metabolic Labeling and Immunoprecipitations
Stationary grown cells from a YPD overnight culture were inoculated in
minimal medium and cultivated to an OD546nm of 1.0. The cells were harvested by centrifugation, washed in minimal medium containing 0.1% glucose and then incubated in the same medium at 30°C for at least 3 h. For
pulse-chase experiments, 2 × 107 cells per time point were labeled by the
addition of 50 µCi [35S]methionine (Tran35S-label, 10 mCi/ml; ICN Pharmaceuticals) for 10 min and then chased with a 100-fold excess of nonradioactive methionine. The chase was terminated by the addition of NaN3
(50 mM final concentration) and immediate freezing in liquid nitrogen.
Protein extractions, immunoprecipitation, and SDS-PAGE were performed as described (Franzusoff et al., 1991; te Heesen et al., 1992
). The
dried gels were exposed and analyzed using a PhosphoImager. The kinetics of CPY* degradation were calculated by setting the counts of time point zero as 100%. For the studies of the transport kinetics of CPY in the
strains with and without Mns1p, the cells were labeled for only 5 min at
26°C. The chase, protein extraction, and immunoprecipitation were performed as described above.
Assay for Degradation of CPY* by Western Analysis
Yeast strains were grown at 30°C in YPD or minimal medium containing
the appropriate supplements into stationary phase. 3 × 108 cells were harvested and broken with glass beads in 50 mM Tris-HCl, pH 7.5, 1% SDS,
2 mM PMSF (Franzusoff et al., 1991; te Heesen et al., 1992
). Protein extract equivalent to 7 × 106 cells was subjected to reducing SDS-PAGE,
transferred to nitrocellulose membranes, and probed with specific antibodies. Binding was visualized by chemiluminescence (SuperSignal ULTRA; Pierce Chemical Co., Rockford, IL). The x-ray films were scanned
and the intensity of the protein bands was determined. The antibody conjugates on the nitrocellulose membranes were stripped by treatment in
62.5 mM Tris-HCl, pH 6.7, 2% SDS, 100 mM
-mercaptoethanol at 65°C for 45 min, and the membranes were reprobed with another antibody. As
an additional control for equal protein concentrations, protein contents
were determined using the method of Sailer and Weissmann (1991)
.
Analysis of Lipid-linked and Protein-linked Oligosaccharides
The analysis of lipid- and protein-linked oligosaccharides has been described (Cacan et al., 1993; Zufferey et al., 1995
; Jakob et al., 1998
). For
pulse-chase labeling of the oligosaccharides, typically 3 × 109 cells of a
logarithmically growing culture were pelleted, washed with YP0.1D (2%
Yeast extract, 1% Bactopeptone, 0.1% glucose), and resuspended in 450 µl
YP0.1D containing 400 µCi 2-[3H]mannose (30 Ci/mmol; ICN Pharmaceuticals). The oligosaccharides were labeled for 1 min at 26°C and the radioactivity was chased by adding nonradioactive (±)D-mannose (111 mM
final concentration). At the given time points, 5 × 108 cells were removed,
placed in 1 ml of CM 3:2 (chloroform/methanol 3:2 vol/vol) and mixed by
vortexing. Extraction, work-up and analysis of lipid-linked oligosaccharides (LLO) and NLO was as described above. For detailed verification of
oligosaccharide structure, endo H-released NLO were further digested
with
1,2-specific mannosidase from Aspergillus saitoi (15 µU; Oxford
Glycosystems, Abingdon, UK) in the supplied buffer. After the digest the
NLO were separated by HPLC (see above).
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Results |
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Trimming of Protein-bound Oligosaccharides In Vivo
To understand in more detail the role of NLO in glycoprotein degradation, we determined the kinetics of protein-bound oligosaccharide trimming in the ER in vivo. For
this, two yeast strains that carry the sec18-50 mutation
were used. The sec18-50 mutation results in a temperature-sensitive phenotype and prevents the fusion of ER-derived vesicles with the Golgi apparatus at nonpermissive temperature. No processing of protein-bound oligosaccharides by Golgi glycosyltransferases was observed in sec18
mutant strains at nonpermissive conditions (Novick et al.,
1980; Eakle et al., 1988
). We performed the experiments at
the permissive temperature for the sec18-50 mutation,
nevertheless, the export rate of secretory proteins to the
Golgi apparatus was slower in sec18-50 cells as compared
with wild-type cells and we were able to analyze the trimming of the NLO in the ER. One of the strains carried in
addition a deletion in the GLS2 locus inactivating glucosidase II. No growth phenotype was associated with the
gls2 mutation. Cells were labeled with 3H-mannose for 1 min at 26°C and the radioactivity was chased by adding an
excess of nonradioactive (±)D-mannose. At the given time points, the chase was terminated, NLO were released
from protein by endoglycosidase H (endo H), and then analyzed by HPLC (see Materials and Methods section).
In the gls2 strain, only two trimming events occur in
the ER: the removal of the terminal
1,2-linked glucose
residue by glucosidase I and the cleavage of an
1,2-linked
mannose residue by mannosidase I. As expected, a protein-bound oligosaccharide with the putative structure
Glc2Man9GlcNAc2 (G2M9)2 was found, which was slowly
converted to Glc2Man8GlcNAc2 (G2M8) with a half-life of
~10 min (Fig. 2, A and B, left). We were unable to detect
protein-bound Glc3Man9GlcNAc2.
|
We analyzed the structure of the different oligosaccharides
in more detail. The endo H-released Glc2Man9GlcNAc2
NLO comigrated with the Glc2Man9GlcNAc2 oligosaccharide obtained from endo H-treated LLO of a alg10 strain
(Fig. 2 C). This strain accumulates lipid-linked Glc2Man9
GlcNAc2 due to the inactivation of the
1,2 glucosyltransferase (Burda and Aebi, 1998
). Digestion of the two major
protein-derived oligosaccharides from a
gls2 sec18-50
strain by exo-
1,2-mannosidase (from Aspergillus saitoi)
converted both of them to a single species (Fig. 2 C).
Therefore, the two oligosaccharides found on protein in a
gls2 sec18 strain differed by one
1,2-linked mannose.
This observation, the mobility of the exo-
1,2-mannosidase digestion product and the comigration of one oligosaccharide with the Glc2Man9GlcNAc2 marker showed
that protein bound Glc2Man9GlcNAc2 and the corresponding mannosidase product Glc2Man8GlcNAc2 were
present in the
gls2 sec18-50 strain. The minor peak observed after exo-
1,2-mannosidase digestion (Fig. 2 C,
left) was possibly due to incomplete digestion, since the
presence of glucose residues on oligosaccharides reduced the efficiency of the enzyme (Burda, P., unpublished
data). Taken together, the results confirmed that release
of the
1,2-linked glucose from protein-bound oligosaccharide by glucosidase I was a rapid process in the
gls2
strain. However, trimming of the oligosaccharide by endogenous
1,2-mannosidase (Mns1p) occurred much slower.
When we analyzed the NLO processing in a strain fully
competent for trimming (sec18-50; Fig. 2, right), we were
again unable to detect the complete protein-bound Glc3
Man9GlcNAc2 oligosaccharide, even in preparations obtained shortly after the pulse (Fig. 2, A and B, right). The
largest oligosaccharide, detected after 2 min of chase (1-min pulse), comigrated with the Glc2Man9GlcNAc2 oligosaccharide but represented a minor fraction (<10%) of
the total NLO. In contrast, significant amounts of Glc1
Man9GlcNAc2 were detected at this time point (the structural analysis of this oligosaccharide is described below).
However, from this oligosaccharide one or two hexose
units were rapidly trimmed. To determine whether this
trimming was due to glucosidase II or mannosidase I activity, we analyzed the NLO preparation from the 5-min
chase point by digestion with the exo-1,2-mannosidase
(Fig. 2 C). This preparation contained small amounts of
oligosaccharides comigrating with the Glc1Man9GlcNAc2
standard and significant levels of oligosaccharides that migrated as expected for Man9GlcNAc2 and Man8GlcNAc2
oligosaccharide. Indeed, digestion by exo-
1,2-mannosidase revealed that only the Glc1Man9GlcNAc2 oligosaccharides contained a protective glucose residue and was
converted to Glc1Man7GlcNAc2, whereas the majority of
the oligosaccharides was trimmed to Man5GlcNAc2. These
results showed that the Glc1Man9GlcNAc2 protein-bound
oligosaccharide was converted primarily to Man9GlcNAc2
and that the peak representing this oligosaccharide contained no significant amounts of mannosidase I-trimmed,
monoglucosylated oligosaccharide.
The analysis of the structure of protein-bound oligosaccharide species as well as their temporal appearance
showed that the removal of the terminal 1,2-glucose on
protein-bound oligosaccharides by glucosidase I was a
rapid process in vivo. Similarly, since we observed only
small amounts of diglucosylated oligosaccharides (Fig. 2,
A and B, 2-min chase), the hydrolysis of the first
1,3-linked glucose by glucosidase II was a rapid process. We
concluded that under our experimental conditions, the
monoglucosylated oligosaccharide Glc1Man9GlcNAc2 was
converted to the Man9GlcNAc2 oligosaccharide with a
half-life of ~2 min and that this occurred before processing by mannosidase I, which was a relatively slow process
(half-life 10 min). Evidently, removal of glucose-linked
residues was not a prerequisite for mannosidase I action
because mannose hydrolysis occurred with approximately
the same kinetics in both glucosidase II-proficient or -deficient strains (Fig. 2 B).
Role of N-linked Oligosaccharides in Glycoprotein Processing
Removal of a mannose residue by 1,2-mannosidase concludes the trimming of NLO in the ER of S. cerevisiae
(Byrd et al., 1982
). Since this cleavage occurred at a slow
rate, we speculated that it represents a rate-limiting step
and thus is important for efficient glycoprotein transport
and maturation. Therefore, we analyzed this aspect in detail by studying the processing of vacuolar proteinase CPY. In the ER, CPY receives four N-linked oligosaccharides (p1CPY, glycosylated proCPY, 67 kD) that are modified in the Golgi apparatus (p2CPY, 69 kD). Upon reaching the vacuole, CPY maturates by proteolytic cleavage of
the propeptide (mCPY, 63 kD). In a pulse-chase experiment, we compared the transport rates of CPY, from ER
to Golgi and to vacuole in wild-type and
mns1 strains
lacking
1,2-mannosidase activity. We observed that CPY
was transported at the same rates (Fig. 3). This demonstrated that trimming of NLO by mannosidase I was not
required for export of glycosylated CPY to the Golgi apparatus and the transport to the vacuole.
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Role of N-linked Oligosaccharides in Degradation of Misfolded CPY
Previous studies have shown that a specifically mutated
form of vacuolar proteinase CPY (CPY*) is retained in
the ER and degraded by the proteasome (Hiller et al.,
1996) in an oligosaccharide-dependent manner (Knop et al.,
1996
). Moreover, deletion of the MNS1 locus affects the
degradation of CPY*. These observations suggested that
trimming of the oligosaccharides is required for processing of misfolded CPY in the ER. To precisely define the NLO
moieties important for proteasome-dependent degradation of CPY*, we generated yeast strains containing CPY*
carrying defined NLO structures on glycoproteins (for details, see Fig. 1 and Table III).
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In a first step, we investigated the influence of glucose
residues of oligosaccharides on CPY* degradation (Fig. 4).
For that purpose, we constructed mutant strains that produced the following NLO structures: Glc2Man8GlcNAc2
(G2; alg10
gls2), Glc1Man8GlcNAc2 (G1;
alg8
gls2),
Man8GlcNAc2 (G0; wild-type and
alg6
gls2). It is important to note that
1,2-mannosidase can act on glucosylated oligosaccharides in vivo (Fig. 2). When we analyzed
the processing of CPY* by pulse-chase experiments, a differential degradation of CPY* was observed depending on
the number of glucose residues present on the NLO (Fig.
4). The G2 CPY* (
alg10
gls2) was degraded at the slowest rate, whereas the G0 CPY* was degraded at the same rate as CPY* in a strain with normal oligosaccharide biosynthesis and trimming (Fig. 4, wild-type). By Western
blot analysis, another G2 CPY*,
gls2, behaved similarly
as G2 CPY* (
alg10
gls2, see below and Fig. 5 C). For
the G1 CPY*, we observed an intermediate degradation
rate (Fig. 4). From the initial degradation rates (time
points 30 and 60 min), we calculated the half-life of CPY* in the various strains (Table IV). In the wild-type and the
G0 cells it was 21 min, similar to published data (Hiller et
al., 1996
). In comparison, the half-life of CPY* in the G1
cells was 43 min and in the G2 cells 82 min under our experimental conditions. Thus, the larger the number of glucose residues the NLO of CPY* contained, the slower its
degradation rate.
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Next, we determined whether mannose residues of the
NLO influenced degradation of CPY*. The core mannose
structure of LLO is synthesized by a set of sequentially
acting ER mannosyltransferases (Alg3p, Alg9p, Alg12p;
see Fig. 1, Orlean, 1997). Due to the fact that incompletely
assembled oligosaccharides can be transferred to protein,
albeit with a reduced efficiency, we were able to obtain
yeast strains with incomplete, but defined NLO structures, namely Man5GlcNAc2 (M5,
alg3; Aebi et al., 1996
),
Man6GlcNAc2 (M6,
alg9; Burda et al., 1996
), Man7
GlcNAc2 (M7,
alg12, Burda and Aebi, manuscript in
preparation), Man8GlcNAc2 (M8, wild-type with respect
to NLO; M8,
alg6; Reiss et al., 1996
), and Man9GlcNAc2 (M9,
mns1; Puccia et al., 1993
; see also Table III).
When the half-lives of mutant CPY* bearing various oligosaccharide structures were analyzed, we found that
CPY* containing Man8GlcNAc2 NLO was rapidly degraded (Fig. 5 A, lanes 1 and 6 and B). The Man5GlcNAc2
CPY* (Fig. 5, A lane 2 and B) was degraded at a reduced
rate compared with the Man8GlcNAc2 CPY*. However, a
greatly reduced rate of CPY* degradation was observed
when the NLO were of Man6GlcNAc2 (Fig. 5 A, lane 3 and
B), Man7GlcNAc2 (Fig. 5 A, lane 4 and B), and Man9
GlcNAc2 (Fig. 5 A, lane 5 and B) structures. By Western
blot analysis, CPY* with diglucosylated oligosaccharides
(gls2) was also degraded at a slower rate than Man8
GlcNAc2 CPY* (Fig. 5 C, compare with Fig. 4). However,
the stabilization was not as prominent as for Man9
GlcNAc2 CPY* (
mns1; Fig. 5 C). Taken together, our results showed that efficient degradation of CPY* was dependent on the NLO structure. Man8GlcNAc2 CPY* was rapidly degraded, whereas the trimming intermediate
Man9GlcNAc2 CPY* and the incompletely assembled
Man6-7GlcNAc2 CPY* were inefficiently degraded.
To demonstrate that the differences in degradation rate
of CPY* were due to the altered oligosaccharide structure,
we transformed the yeast cells with plasmids complementing the deleted gene loci. The alg3,
alg9, and
alg12
cells were transformed with plasmids containing the
ALG3 (pALG3), ALG9 (pALG9) and ALG12 (pALG12)
locus. Further, the CPY* wild-type strain was transformed
with the vector plasmid (YEp352). The NLO structure-dependent CPY* degradation phenotype could be reverted by complementing the deleted alg gene loci with
the appropriate plasmids (Fig. 5 D, lanes 3-8), but the degradation in wild-type cells was not influenced by the transformation with the empty plasmid (Fig. 5 D, lanes 1 and 2).
Incomplete assembly of the LLO leads to a reduced oligosaccharide transfer to protein by the oligosaccharyltransferase (Sharma et al., 1981; Silberstein and Gilmore,
1996
), which is then apparent by underglycosylation of glycoproteins (Huffaker and Robbins, 1981
, 1983
; Stagljar et al.,
1994
; te Heesen et al., 1994
; Burda et al., 1996
; Burda and
Aebi, 1998
). The equal degradation of Man8GlcNAc2
CPY* in glycosylation wild-type cells and in
alg6 cells
(Fig. 5 A, lanes 1 and 6) indicated that the degradation was
not due to impaired synthesis of LLO, but rather to the altered oligosaccharide structure.
Also, in contrast to wild-type CPY, CPY* was efficiently
glycosylated in both alg3 (Man5GlcNAc2) and
alg9
(Man6GlcNAc2) cells under the conditions used. We observed a single CPY* glycoform in
alg3 prc1-1 and
alg9
prc1-1 cells (Fig. 6, lanes 1 and 3), whereas correctly folded
CPY was incompletely glycosylated in
alg3 and
alg9
cells (Fig. 6, lanes 7 and 9). This hypoglycosylation of CPY
was visualized by the distinct bands upon Western blot analysis representing different glycoforms of this protein
(Stagljar et al., 1994
). In addition, this experiment showed
that such forms of CPY* did not reach the vacuole because endo H treatment resulted in the deglycosylated
pro-form of CPY* in
alg9 cells (Fig. 6, lanes 3 and 4,
compare with lanes 9 and 10). As expected, the oligosaccharides in
alg3 cells were resistant to endo H digestion, whereas in
alg9 cells they were endo H sensitive (Fig. 6,
lanes 8 and 10).
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In alg3 or
alg9 cells, a much higher steady state level
of CPY was observed compared with CPY* in various mutant cells (Fig. 6). This indicated that degradation of CPY*
was not completely blocked by the altered oligosaccharide
structure. However, our results demonstrated that oligosaccharide structures specifically affected the degradation of misfolded CPY accumulating in the ER, whereas
processing and secretion of wild-type CPY was not altered.
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Discussion |
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A Defined Oligosaccharide Structure Was Required for Efficient Degradation of Misfolded Glycoprotein
We observed a significant effect of the structure of N-linked
oligosaccharides on degradation of misfolded CPY*, a
model protein for degradation of glycoproteins retained in
the ER (Wolf and Fink, 1975; Finger et al., 1993
; Hiller et
al., 1996
; Knop et al., 1996
). When expressed in
gls2 or
mns1 cells, we found a reduced degradation of CPY*
(Fig. 5 C). The effect of the mannosidase I inactivation
was significantly stronger than that of the glucosidase II
deletion. A similar stabilization of CPY* as in
mns1 cells was observed in
alg9 and
alg12 cells, where incompletely mannosylated oligosaccharides (Man6GlcNAc2 and
Man7GlcNAc2, respectively) were transferred to protein.
In
alg3 cells, which contain protein-bound Man5Glc
NAc2, degradation of CPY* was reduced, however, not to the same extent as in
alg9 or
alg12 cells. Interestingly,
in
alg6 cells, where nonglucosylated oligosaccharide
Man9GlcNAc2 was transferred to protein and trimmed
to Man8GlcNAc2, no effect on the stabilization of CPY*
was observed. We concluded, that the protein-bound
Man8GlcNAc2 structure was an important recognition element in the degradation pathway of CPY*. Our results
were best explained by a model (Fig. 7) where the
Man8GlcNAc2 oligosaccharide on a misfolded glycoprotein acted as a positive signal for degradation. This oligosaccharide structure might be recognized by a lectin, since any changes in its structure reduced the efficiency of
degradation. Such a carbohydrate-binding protein has also
been postulated in the degradation of glycoproteins in
mammalian cells (Yang et al., 1998
) and might represent
"the additional signal to direct them (soluble misfolded
proteins) to the dislocation and the ubiquitation machinery" (Kopito, 1997
). Our results showed that glucosylated oligosaccharides also reduced the degradation rate of
CPY*, albeit to a lesser extent than the Man9GlcNAc2,
Man7GlcNAc2 and Man6GlcNAc2 structures. We concluded that the
1,2-
1,2-dimannose branch of the Man8
GlcNAc2 oligosaccharide was a less important structural element for oligosaccharide recognition than both the
1,6- and
1,3-branch affected by the alg3, alg9 and alg12
mutations (see Fig. 1). The observation that the Man5
GlcNAc2-producing
alg3 mutation had a less severe effect on degradation was explained by the hypothesis that
this oligosaccharide structure represents an intermediate
in the degradation of glycoprotein as shown in higher eukaryotic cells (Villers et al., 1994
; Ermonval et al., 1997
).
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Whether the postulated lectin additionally recognizes
unfolded protein domains, as does UDP-glucose/glycoprotein glucosyltransferase, involved in the ER quality control pathway of higher eukaryotes (Sousa and Parodi,
1995), is not known. It is possible that the binding of both,
the chaperones and the postulated lectin constitute a signal which targets the glycoprotein to the degradation pathway. Indeed Kar2p, the yeast homologue of BiP, transiently binds to wild-type CPY (te Heesen and Aebi, 1994
;
Simons et al., 1995
). Furthermore, this Hsp70 protein was
shown to be involved in the degradation of misfolded protein by the proteasome pathway (Plemper et al., 1997
).
It has been proposed that the trimming of the protein-bound oligosaccharide in the endoplasmic reticulum represents a biological timer for the protein maturation in the
ER of higher eukaryotes (Helenius et al., 1997). This timer
function might be required to prevent permanent residence of misfolded glycoproteins in the ER due to the
binding to calnexin and calreticulin in higher eukaryotes. Our results are fully compatible with this timer model: as
in higher eukaryotic cells (Hubbard and Robbins, 1979
),
the protein bound oligosaccharide underwent a step-wise
trimming process. Removal of the terminal
1,2-glucose
by glucosidase I and the first
1,3-glucose by glucosidase
II was a very rapid process, whereas the second
1,3-glucose was removed more slowly. The same difference in
glucose hydrolysis was observed for glucosidase II of
higher eukaryotic cells (Hubbard and Robbins, 1979
) and it has been proposed that two different substrate binding
sites of glucosidase II are responsible for this difference:
the high affinity site would be responsible for the hydrolysis of the first glucose, the low affinity site for the hydrolysis of the second glucose (Alonso et al., 1993). Similar to
the findings in higher eukaryotic cells, the mannose trimming was a slow process as compared with hydrolysis of
the glucose residues of the protein-bound oligosaccharide. The removal of one specific
1,2-mannose residue by ER-
1,2-mannosidase might represent the time-point after
which a misfolded protein is routed to the degradation
pathway. We speculated that the processing by mannosidase I determined the time-scale in which a protein had to
be correctly folded. If this was not achieved, the glycoprotein was degraded. Importantly, maturation and transport
of correctly folded CPY is oligosaccharide-independent
(Schwaiger et al., 1982
; Winther et al., 1991
) and was also
not influenced by trimming or oligosaccharide structure
(Figs. 3 and 6). It was also postulated that there is a selective export of proteins out of the ER in yeast (Kuehn and
Schekman, 1997
). Selective export of only correctly folded
proteins in combination with degradation of misfolded
proteins, timed by oligosaccharide trimming, might therefore represent an effective quality control system for glycoprotein folding in the ER of S. cerevisiae, where the
calnexin/calreticulin cycle (reglucosylation of misfolded
proteins) has not been found (Fernandez et al., 1994
, Jakob et al., 1998
).
In support of our model, we found that the degradation
of a mutant form of the oligosaccharyltransferase component Stt3p, a glycoprotein with multiple transmembrane
domains (Zufferey et al., 1995) was also controlled by oligosaccharide trimming (Bodmer, D., U. Spirig, and M. Aebi, manuscript in preparation), suggesting that resident
ER glycoproteins are subject to the same degradation system as are glycoproteins that are exported from the ER.
Is there a similar role of the Man8GlcNAc2 oligosaccharide in the quality control process of glycoproteins in the
ER of higher eukaryotic cells? In the trimming process of
protein-bound oligosaccharides, removal of the glucose
residues precedes the hydrolysis mannose trimming (Hubbard and Robbins, 1979). There is an
1,2-mannosidase activity that leads to the same Man8GlcNAc2 oligosaccharide as the yeast MNS1 enzyme in the ER of higher eukaryotic cells (Bischoff and Kornfeld, 1983
). However,
there are additional ER mannosidase activities in the ER
of higher eukaryotic cells and a different Man8GlcNAc2
oligosaccharide isomer, where the
1,2-mannose linked to
the
1,6-mannose is removed, can be produced (Weng and
Spiro, 1993
, 1996
; Moremen et al., 1994
). The trimming
process in higher eukaryotes is therefore more complex
than in yeast. Nevertheless, our results obtained in yeast
are compatible with reports on inhibition of
-mannosidase trimming by deoxymannojirimycin that stabilizes specific misfolded glycoproteins in the ER (Su et al., 1993
;
Daniel et al., 1994
; Liu et al., 1997
; Yang et al., 1998
). On
the other hand, degradation of some glycoproteins in the
ER was not affected by mannosidase inhibition (Yang et al.,
1998
).
Alternative Pathways for ER Degradation?
Previous work has shown that CPY* remains in the ER, is
ubiquitinated and then degraded in a proteasome-dependent pathway (Hiller et al., 1996). When we compared the
level of wild-type CPY and mutant CPY* in both the
alg9 and
mns1 cells (Figs. 5 A and 6), it was apparent
that more mature (vacuolar) CPY was present in wild-type
cells than CPY* in the prc1-1 cells. A major portion of
CPY* was apparently degraded in these cells. Our results showed that alterations of the oligosaccharide structure
did not completely block degradation of CPY*. Moreover,
mutations reported to affect CPY* degradation do not
completely block CPY* degradation either (Hiller et al.,
1996
; Knop et al., 1996
; Plemper et al., 1997
). Taken together, these results suggest an alternative, glycosylation-independent degradation pathway for misfolded glycoproteins in the ER of S. cerevisiae.
Evidence for Posttranslocational N-Glycosylation of CPY*
N-linked oligosaccharides are added co- and posttranslocationally in yeast. For CPY, competition between glycosylation and folding has been reported (Holst et al., 1996).
In cells containing alg mutations, incompletely assembled
oligosaccharide is transferred to nascent protein (Stagljar
et al., 1994
), albeit at a reduced rate (Sharma et al., 1981
).
This is reflected by incomplete use of potential N-glycosylation sites. However, we noticed that in both
alg3 and
alg9 cells, only CPY* with all four potential N-glycosylation sites occupied accumulated, whereas in the
alg3 and
alg9 cells, wild-type CPY lacking one or two oligosaccharides were found (Fig. 6). Therefore, we postulated that
the prolonged exposure of misfolded CPY* to the oligosaccharyltransferase compensated for the reduced affinity of the oligosaccharyltransferase towards incompletely
assembled oligosaccharide and resulted in fully glycosylated CPY* in both
alg3 and
alg9 cells.
The assembly of the lipid-linked oligosaccharide and its transfer to selected asparagine residues of polypeptides in the ER is a highly conserved process. The selective processing of the protein-bound oligosaccharide supports the idea that conservation of the transferred oligosaccharide structure is due to the function of specific trimming intermediates in glycoprotein maturation. Genetic tailoring of NLO structures will provide a useful tool to identify additional roles of the oligosaccharide in glycoprotein processing.
![]() |
Footnotes |
---|
Received for publication 19 May 1998 and in revised form 31 July 1998.
Address all correspondence to Markus Aebi, Mikrobiologisches Institut, ETH Zürich, Schmelzbergstrasse 7, CH-8092 Zürich, Switzerland. Tel.: 41 1 632 64 13. Fax: 41 1 632 11 48. E-mail: aebi{at}micro.biol.ethz.chWe thank Dr. D.H. Wolf (University of Stuttgart, Germany) for providing the prc1-1 integrative plasmid. The hexokinase antiserum was kindly provided by Dr. S. Schröder (Biozentrum, University of Basel, Switzerland). We would also like to thank Drs. A. Helenius and S. te Heesen for helpful discussions and comments on the manuscript and anonymous reviewers for invaluable remarks.
This work was supported by the Swiss National Science Foundation grants 3100-040350 (to M. Aebi) and 31-50835.97 (to J. Roth).
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Abbreviations used in this paper |
---|
CPY, carboxypeptidase Y; CPY*, mutated, misfolded CPY; endo H, endoglycosidase H; LLO, lipid-linked oligosaccharides; NLO, N-linked oligosaccharides.
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