Correspondence to Davis T.W. Ng: dtn1{at}psu.edu
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Abbreviations used in this paper: CPY, carboxypeptidase Y; ERAD, ER-associated degradation; GT, glucosyltransferase; UPR, unfolded protein response.
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Introduction |
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The upstream events of substrate sorting and targeting to the translocation pore are less clear. In higher eukaryotes, folding and quality control functions of glycoprotein synthesis are integrated within the calnexin/calreticulin lectin cycle. A third ER lectin, EDEM (Htm1/Mnl1p in yeast), is used to direct misfolded proteins off-cycle and into the ERAD pathway (Molinari et al., 2003; Oda et al., 2003). The mechanism is not universal because many proteins (e.g., nonglycosylated) use other pathways that are less defined. In budding yeast, the process is murkier with the absence of a typical calnexin/calreticulin cycle. However, the requirement of Htm1/Mnlp for ERAD reflects its functional conservation and emphasizes that general strategies of protein quality control are shared among all eukaryotes (Jakob et al., 2001; Nakatsukasa et al., 2001).
The determinants used for sorting and targeting substrates have not been fully characterized. At first glance, tackling the problem seems simple. However, the range of possible substrates illustrates the enormity of the task. Trafficking through the ER includes soluble proteins, single and multi-spanning integral membrane proteins, and lipid anchored proteins. For their maturation, additional steps may include glycosylation, prolyl hydroxylation, disulfide bond formation, and assembly into complex oligomers. Within this backdrop is the vast number of conformations and configurations that the cell must determine are unfolded (in the process of folding), folded, and misfolded. Although the prevailing evidence indicates that chaperones recognize and bind unfolded and misfolded proteins, how the cell arrives at the decision to degrade individual molecules remains unknown. Defining the nature of substrate determinants is key to this understanding.
Previously, we reported that cytosolic and lumenal surveillance mechanisms coexist to monitor the range of proteins trafficking through the ER (Vashist and Ng, 2004). Because the pathways converge at the ubiquitylation/degradation step of ERAD, they were designated ERAD-C (cytosolic) and ERAD-L (lumenal). In this study, we focused our attention on glycoprotein substrates of ERAD-L. By systematically analyzing a series of substrate variants, we discovered that determinants used for sorting/retention (from folded proteins) could be distinguished from those used to target substrates to ERAD. Initially, the unfolded polypeptide alone suffices for efficient substrate recognition and retention. The decision to terminate the molecule, however, requires an additional structural determinant embedded in the substrate.
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Results |
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Unexpectedly, metabolic pulse-chase experiments showed that CPY1 turned over poorly by comparison to CPY* (Fig. 1, compare E with F). This indicated that CPY
1, though retained by ER quality control, is a poor substrate for ERAD. This view was confirmed by its failure to be further stabilized in the
cue1 ERAD mutant (Fig. 1 F). Cue1p is a critical component for CPY* degradation as it anchors the ubiquitin conjugating enzyme Ubc7p to the ER membrane (Biederer et al., 1997). The residual turnover was likely through alternative pathways that degrade misfolded proteins when ERAD is disrupted or saturated (Haynes et al., 2002; Spear and Ng, 2003). Together, the data show that ER quality control and ERAD are mechanisms that can be uncoupled at the substrate level. This raised the intriguing possibility that ER retention and entry into ERAD use distinct substrate determinants.
Nonetheless, we envisioned other equally plausible explanations that could account for CPY1's unusual behavior. Because the deletion is large, the severity of the lesion might increase the tendency of the remaining polypeptide to aggregate. Substrate solubility is an important prerequisite for ERAD. CPY* aggregates caused by faulty chaperone function were shown to degrade inefficiently (Nishikawa et al., 2001). To determine whether aggregate formation contributes to CPY
1 stability, microsomal membranes containing CPY* or CPY
1 were prepared from logarithmically growing cells. The membranes were solubilized in nonionic detergent under physiological conditions and subjected to centrifugation. Under these conditions, large protein aggregates sediment rapidly and separate from soluble proteins remaining in the supernatant. Detergent-insoluble (pellet) and detergent-soluble (supernatant) fractions were collected, proteins resolved by SDS-PAGE, and analyzed by immunoblotting. To control for the procedure, we specifically generated CPY* aggregates from cells severely limiting for ER chaperones (
ire1; Spear and Ng, 2003). This species was analyzed in parallel. As shown in Fig. 2, CPY* and CPY
1 were both recovered entirely from the supernatant fraction (B and C), whereas CPY* aggregates from
ire1 cells were found predominantly in the pellet fraction (A). In every case, the ER integral membrane protein Sec61p was recovered from the soluble fraction showing that membranes were completely solubilized. This experiment showed that the formation of detergent-insoluble aggregates was not a root cause of the CPY
1 ERAD defect. Furthermore, CPY
1 puncta, which would be characteristic of intracellular aggregates, were never observed in immunolocalization experiments (DePace et al., 1998). Instead, CPY
1 was always found to be evenly distributed throughout the ER in patterns indistinguishable from CPY* and Kar2p (Fig. 1 C).
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To distinguish the models, we generated the remaining CPY* single-site glycosylation mutants to test if ERAD efficiency is related to carbohydrate density. By contrast to the D glycan, eliminating any of the other three carbohydrates had no effect on CPY* degradation (Fig. 4 C). These data show that only glycan D is essential for efficient degradation of misfolded CPY and rule out the notion of a carbohydrate threshold.
The findings were intriguing because Htm1/Mnl1p was proposed to be an ERAD-specific lectin (Jakob et al., 2001; Nakatsukasa et al., 2001). However, little was known regarding how it recognizes substrate. If Htm1/Mnl1p acts specifically through glycan D, the stability of ABCd-CPY*, which bears the other three glycans, should be unaffected in cells lacking HTM1/MNL1. To test the assertion, we measured CPY* and ABCd-CPY* turnover in wild-type and htm1/mnl1 cells. In metabolic pulse-chase experiments, CPY* degradation was dependent on HTM1/MNL1 as previously reported (Fig. 4 D; Jakob et al., 2001; Nakatsukasa et al., 2001). By contrast, ABCd-CPY* was degraded poorly in wild-type cells with no further stabilization in the
htm1/mnl1 cells (Fig. 4 E). Interestingly, the residual degradation of ABCd-CPY* in the presence or absence of Htm1/mnl1p required Cue1p (Fig. 4 E). This likely reflects a lectin-independent mode of ERAD revealed only upon simultaneous disruption of the lectin and its substrate determinant. We next assessed whether glycan D is sufficient to direct CPY* to ERAD in an Htm1/Mnl1p-dependent manner. For this, a mutant variant glycosylated only at site D was constructed (abcD-CPY*). When expressed in wild-type cells, abcD-CPY* was degraded efficiently (Fig. 4 F). In
htm1/mnl1 cells, however, abcD-CPY* was stabilized to the same extent as CPY* (Fig. 4 F). Together, these data show that a single, specific carbohydrate is necessary and sufficient in directing substrate into ERAD by way of Htm1/Mnl1p.
We wondered if the specificity of the glycan was unique to CPY* or a general feature of lectin-dependent ERAD. For this, we analyzed the ERAD substrate PrA* (Finger et al., 1993). PrA* is a mutant version of the endogenous vacuolar enzyme, proteinase A. PrA* contains two N-linked glycans, one near its NH2 terminus and the other near the COOH terminus at a position similar to CPY*'s glycan D. Each site was disrupted singly by replacing asparagine codons with glutamine. The mutant variants, Ab-PrA* and aB-PrA* (Fig. 5 A, follows the nomenclature of CPY* glycan mutants), were expressed in wild-type cells (deleted of endogenous PEP4 gene for detection of PrA*) and their turnover measured. As shown in Fig. 6, the Ab-PrA* was degraded indistinguishably from PrA*. Because PrA* is glycosylated at only two sites, its degradation depends on a single, specific glycan signal like CPY* or is carbohydrate independent. The question was answer by the results of two experiments. First, Ab-PrA* degradation is dependent on Htm1/Mnl1p to a similar extent as PrA* (Fig. 6, A and B). Second, turnover of reciprocal mutant, aB-PrA*, was strongly defective and little affected by the loss of Htm1/Mnl1p (Fig. 6 C). These data support the idea that single ERAD glycan determinants are preembedded in glycoproteins. However, a wider range of substrates must be tested to determine whether other configurations are used. Serendipitously, analysis of the PrA* model also ruled out COOH-terminal positioning being a requirement because its sole determinant is closer to the NH2 terminus.
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Discussion |
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Misfolded proteins recognized by ERQC are not always inevitably degraded by ERAD. Among a group of misfolded Ste6p mutants retained in the ER, some variants are degraded by ERAD, whereas others are stable (Loayza et al., 1998). Although the reasons for the difference are unknown, these studies demonstrated that retention and degradation are separable mechanisms. In the ERAD-L system, the theme is similar. Although both CPY* and CPY1 are retained by ERQC, only CPY* is degraded efficiently by ERAD (Fig. 1). Through systematic analysis of these substrates, we discovered that the deleted glycan of CPY
1 is a critical determinant for targeting CPY to ERAD when misfolded. Subsequent analysis using PrA* revealed an analogous signal demonstrating the generality of the mechanism. In either case, the determinant is used exclusively as other glycans naturally positioned along the molecules cannot substitute. By contrast, ERAD-C substrates are degraded independent of glycosylation state (Vashist and Ng, 2004).
N-linked carbohydrates perform many functions and their participation in ERQC is well established. In higher eukaryotes, a subset of glycoproteins relies on the ER lectins calnexin and calreticulin for folding (for reviews see Ellgaard and Helenius, 2003; Sitia and Braakman, 2003). They work by binding trimmed, monoglucosylated N-linked glycans of newly synthesized proteins and provide a platform for the participation of accessory folding enzymes. Removal of the remaining glucose residue by glucosidase II frees the substrate from the lectin. Should the substrate remain unfolded, UDP-glucose/glycoprotein glucosyltransferase (GT) reglucosylates the glycan for another round of lectin binding. In this mode, GT is the folding sensor and substrates remaining in the cycle are retained in the ER as a consequence. Proteins that cannot fold properly go off-cycle and enter an ERAD pathway. Although used by many organisms, this mechanism is absent in yeast due to the lack of GT.
Even if the calnexin cycle is not universally conserved, current evidence indicate that all eukaryotes have adapted N-linked glycans for use in ERAD. For example, eliminating the N-linked glycosylation sites of CPY* disrupted its degradation in yeast cells (Knop et al., 1996). Although their role in ERAD was unclear, it was suggested that the carbohydrates were needed to maintain substrate conformations favorable for ERAD. A different view emerged from genetic and pharmacological studies that assessed the effects of compromised carbohydrate processing. The trimming of protein-linked Glc3Man9GlcNAc2 core carbohydrate(s) to the Man8GlcNAc2 form (Glc, glucose; Man, mannose; GlcNAc, N-acetylglucosamine) was shown to be required for efficient substrate degradation (Jakob et al., 1998; Tokunaga et al., 2000). This led to the proposal that N-linked glycans can function as signals for targeting ERAD substrates. The failure of Man9GlcNAc2, Man7GlcNAc2, and Man6GlcNAc2 glycoforms to substitute for Man8GlcNAc2 implied a degree of specificity expected of a ligandreceptor interaction (Jakob et al., 1998). This model is particularly appealing because the crucial mannose trimming step was found to be much slower than the preceding processing steps. This led to the idea of a mannosidase Idependent mechanism that provides newly synthesized proteins a fixed window of time for folding. Should the protein remain unfolded by the time the enzyme acts, a lectin receptor was hypothesized that targets Man8GlcNAc2-containing substrates to ERAD (Jakob et al., 1998).
The discovery of an ER lectin-like protein required for ERAD provided support for the proposed mechanism. Known variously as Htm1p1, Mnl1p, or EDEM, the results of four independent groups indicate that glycoprotein substrates are directed into ERAD by way of this membrane bound factor (Hosokawa et al., 2001; Jakob et al., 2001; Nakatsukasa et al., 2001; Molinari et al., 2003; Oda et al., 2003). The genetic experiments that showed the importance of Man8GlcNAc2 would also suggest that Htm1/Mnl1p plays a role in recognizing the glycan (Jakob et al., 1998). Even as the Htm1p/Mnl1p/EDEM lectin filled a gap in the model, other important questions remained. Most importantly, how can substrates be distinguished by this mechanism when properly folded glycoproteins also bear Man8GlcNAc2 (Gemmill and Trimble, 1999)? The discovery that ERAD glycan determinants are constrained to single, specific sites helps address the conundrum.
The preference for specific carbohydrates reveals new insight into how substrates are recognized by the glycan-dependent ERAD system. The simple explanation of a general positional constraint along any polypeptide chain was ruled out because respective signals were found in different positions within PrA* and CPY*. This finding, together with the inability of other glycans to substitute, suggest that a yet uncharacterized component of the signal is located somewhere along the polypeptide chain. Either of two scenarios could account for how the determinant would act. In one, it directs modifications to specific glycans much like the enzyme UDP-GlcNAc/lysosomal enzyme N-acetlyglucosamine-1-phosphotransferase that participates in the addition of mannose 6-phosphate to prelysosomal hydrolases (Baranski et al., 1990). The modified glycan would then serve as the ligand for the lectin receptor. Recent experiments from the Jakob group have ruled out this possibility. In agreement with substrate requirements from their previous genetic studies, direct analysis of CPY* glycans purified from wild-type cells showed that they are primarily Man8GlcNAc2 (C. Jakob, ETH Switzerland, personal communication).
If CPY*'s four N-linked carbohydrates share the same structure and yet only the COOH-terminal glycan can function to signal ERAD, it can be deduced that the signal also contains a specific peptide component. By extension, we propose a model by which the lectin binds Man8GlcNAc2 carbohydrates only in conjunction with a specific protein determinant in the unfolded state. This would explain why only single, specific glycans in CPY* and PrA* are used to target the substrates to the ERAD pathway. This simple mechanism would provide the means to distinguish folding proteins (Man9GlcNAc2/unfolded determinant: no binding), folded proteins (Man8GlcNAc2/folded determinant: no binding), and "misfolded" proteins (Man8GlcNAc2/unfolded determinant: binding). Operationally, the cell need not define "misfolded" in structural terms. Any molecule exceeding its time limit to fold is simply degraded whether it could eventually fold or not. In line with the proposed mechanism, our preliminary studies have determined a short peptide segment from CPY* that fulfills the criteria for an additional determinant. It is required for lectin-mediated ERAD and is functionally transposable to other parts of the polypeptide (unpublished data).
This study shows that determinants for ER quality control (sorting and retention) and ERAD (targeting and degradation) can be modular in their function. In the Htm1/Mnl1p arm of ERAD, single, specific carbohydrates are required to target substrates for degradation but are dispensable for their recognition and retention as misfolded proteins (e.g., CPY1, ABCd-CPY*). As these substrates represent but a small fraction of all classes of misfolded proteins, our understanding of ERQC/ERAD determinants is far from complete. With the list of ERQC/ERAD factors rapidly expanding, it has never been more important to uncover the nature of their substrates to understand how aberrant proteins are sorted for degradation.
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Materials and methods |
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Site-directed mutagenesis of CPY* and CPY1
Site-directed mutagenesis was performed using a PCR-based approach as described previously (Ng et al., 1996). Primers used for site-directed mutagenesis are listed in Table III. All mutants were confirmed by DNA sequence analysis performed by the Penn State DNA sequence core facility.
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Indirect Immunofluorescence
Indirect immunofluorescence experiments were performed as described previously (Spear and Ng, 2003). Polyclonal rabbit anti-Kar2p and HA.11 mAb (Covance Inc.) were used as primary antibodies and Alexa Fluor 488 goat antirabbit and Alexa Fluor 546 goat antimouse (Molecular Probes, Inc.) were used as secondary antibodies. Formaldehyde-fixed cells were visualized using an Axioplan epifluorescence microscope (Carl Zeiss MicroImaging, Inc.) with a Plan-Neofluar 100x objective (1.3 NA; Carl Zeiss MicroImaging, Inc.) and immersion oil (Carl Zeiss MicroImaging, Inc.). Image acquisition was performed using a SPOT 2 cooled CCD camera using SPOT v. 3.5.5 software (Diagnostic Instruments, Inc.). Images were archived and converted to gray scale using Adobe Photoshop 4.0 (Adobe Systems).
Analysis of protein aggregates
Wild-type cells expressing CPY* (pDN436) or CPY1 (pES57) were grown in synthetic complete media supplemented with 2% glucose and lacking leucine and uracil, respectively, to logarithmic phase. Control
ire1 cells containing the GAL-CPY* gene (plasmid pES67; Spear and Ng, 2003) were grown to logarithmic phase in synthetic complete containing 3% raffinose and 50 µg/ml myo-inositol. CPY* overexpression was induced for 7 h by the addition of galactose to 2%. Cells (5.0 OD600 units) were collected by centrifugation and washed once with ice-cold water. The cell pellet was resuspended in 500 µl TNE (50 mM Tris, pH 7.4, 150 mM NaCl, 5 mM EDTA) containing protease inhibitor cocktail (Sigma-Aldrich) and 1 mM PMSF, and transferred to a 1.5-ml screw-cap tube on ice. Zirconium beads (0.4 ml of 0.5-mm diam) were added and cells were homogenized using a vortex mixer at full speed for 1 min, followed by 1 min on ice. This cycle was repeated 810 times in the cold. The lysate was cleared by centrifugation at 750 g for 5 min and repeated. Triton X-100 was added to 1% vol/vol and incubated for 5 min at RT. A 50 µl portion (T) was saved before ultracentrifugation at 100,000 g for 15 min at 4°C. The supernatant (S) was removed and the pellet was resuspended in 450 µl 3% SDS, 50 mM Tris, pH 7.5, and incubated at 100°C for 5 min. Total (T), supernatant (S) and pellet (P) fractions were resolved by SDS-PAGE, transferred to nitrocellulose, and probed using specific antisera (1:10,000 anti-HA; 1:5,000 anti-Sec61) and HRP-conjugated secondary antibodies. Proteins were visualized by ECL (Pierce Chemical Co.).
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Acknowledgments |
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This work was supported by a grant from the National Institutes of Health to D.T.W. Ng (GM059171).
Submitted: 23 November 2004
Accepted: 3 March 2005
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References |
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