Endoplasmic reticulum (ER)-associated degradation (ERAD) is a proteolytic pathway for handling misfolded or improperly assembled proteins that are synthesized in the ER. Cytoplasmic peptide:N-glycanase (PNGase) is a deglycosylating enzyme that cleaves N-glycans that are attached to ERAD substrates. While the critical roles of N-glycans in monitoring the folding status of carrier proteins in the ER lumen are relatively well understood, the physiological role of PNGase-mediated deglycosylation in the cytosol remained poorly understood. We report herein the identification of endogenous substrates for the cytoplasmic PNGase in Saccharomyces cerevisiae. Using an isotope-coded glycosylation site-specific tagging (IGOT) method-based LC/MS analysis, 11 glycoproteins were specifically detected in the cytosol of PNGase-deletion cells (png1Δ). Among these molecules, at least five glycoproteins were clearly identified as ERAD substrates in vivo. Moreover, four out of the five proteins were found to be either deglycosylated by PNGase in vivo or the overall degradation was delayed in a png1Δ mutant. Our results clearly indicate that the IGOT method promises to be a powerful tool for the identification of endogenous substrates for the cytoplasmic PNGase.

Introduction

Proteins that go through the secretory pathway in eukaryotes are translocated into the lumen of the endoplasmic reticulum (ER) upon their synthesis. In the ER, newly synthesized proteins undergo correct folding with the aid of various ER-resident chaperones, lectins and enzymes [1]. When proteins fold or assemble properly, they exit from the ER via vesicular transport to be delivered to their respective destinations. On the other hand, misfolded or unassembled proteins are specifically recognized and are subjected to retrotranslocation from the ER to the cytosol, where proteasomes play a key role in their degradation. This protein surveillance system in the ER is referred to as ER quality control (ERQC) [1], whereas the degradation process, as a part of ERQC, is referred to as ER-associated degradation (ERAD) [2,3].

The cytoplasmic peptide:N-glycanase (PNGase; Png1 in yeast) is a deglycosylation enzyme, which cleaves N-glycans that are attached to misfolded proteins in the cytosol [47]. Under conditions where proteasome activity is compromised, several substrates were found to be deglycosylated by the cytoplasmic PNGase in vivo [810], indicating that this enzyme is involved in the ERAD process. Initially, it was thought that PNGase-mediated deglycosylation is critical for the efficient degradation of ERAD substrates, because N-glycans could serve as a bulky obstacle to proteolysis by proteasomes. In many of the model ERAD substrates tested, however, no severe defect in degradation efficiency was found, even in the presence of a PNGase inhibitor or upon the suppression of gene expression for PNGase [11,12]. Moreover, the deglycosylation of several glycosylated model ERAD substrates was not detected even under conditions in which proteasome activity was compromised [13,14]. Those observations made the clarification of functional importance for this enzyme very difficult. However, the recent discovery of human subjects bearing mutations in NGLY1, a human gene ortholog of PNG1, strongly indicates that this enzyme plays a critical role in the normal development of mammals [1518]. The subjects exhibited multiple severe symptoms, including global developmental delay, multifocal epilepsy, involuntary movements, abnormal liver/brain functions and the absence of tears. Mechanistic insights into the functional importance of PNGase, however, remain unclear.

One of the key issues remaining to be clarified regarding cytoplasmic PNGase is its endogenous substrates. It has been reported that the degradation of a model protein derived from the ricin toxin A-chain serves as a PNGase-dependent ERAD substrate, that is, PNGase-mediated deglycosylation is critical for its efficient degradation, both in Saccharomyces cerevisiae and in mice embryonic fibroblasts [1922]. On the other hand, endogenous, natural substrates for cytoplasmic PNGase have not been unequivocally demonstrated in any organism. It is therefore difficult to provide deeper insights into the functional importance of PNGase in the ERAD process, in the absence of such information.

The IGOT (isotope-coded glycosylation site-specific tagging) method for the comprehensive identification of glycoproteins was previously developed [23]. The method involves the affinity capture of glycoproteins using lectins, tryptic digests of protein mixtures and the PNGase-mediated incorporation of a stable isotope tag, 18O, into the N-glycosylation site, followed by an LC/MS (mass spectrometry)-based peptidomics analysis. Since IGOT is an excellent method for the comprehensive detection of N-glycosylated peptides, we aimed to apply this method to identify endogenous substrates for cytoplasmic PNGase. The rationale behind this approach is that the endogenous substrates for PNGase should accumulate in the cytosol of png1Δ cells, but not as much in isogenic wild-type cells. Indeed, we successfully identified 11 proteins that specifically accumulated in the cytosol of the png1Δ mutant through this approach. Of the 11 proteins, 8 were confirmed to be N-glycosylated and 5 were shown to be intrinsic ERAD substrates. Finally, a biochemical analysis revealed that Png1 was indeed involved in the deglycosylation/degradation of several novel ERAD substrate glycoproteins.

Materials and methods

Yeast strains and media

We used the following yeast strains: BY4741 (MATahis3Δ1 leu2Δ0 met15Δ0 ura3Δ0), AHY201 (hrd1Δ::hphNT1 BY4741) [24], AHY202 (png1Δ::hphNT1 BY4741) [25], AHY273 (doa10Δ::hphNT1 BY4741; the present study), AHY274 (hrd1Δ::hphNT1 doa10Δ::natNT2 BY4741; the present study), AHY262 (pdr5Δ::natNT2 BY4741) [24] and AHY263 (pdr5Δ::natNT2 png1Δ::hphNT1 BY4741) [24]. Standard yeast media and genetic techniques were used [26,27]. Where indicated, cells bearing a pdr5Δ mutation were incubated with 50 µM MG-132 (3175-V; Peptide Institute, Japan) to inhibit proteasomal degradation.

Plasmid construction

All genes (ASI3, EPS1, KRE2, MMM1, MNN1, NPR1, PMT6, SCS2, SNG1 and SUC2) were inserted into pAG425GPD-ccdB-HA (Addgene, MA, USA) by the Gateway cloning system. Detailed information on primer sequences is available upon request. The DNA sequences of the constructs were confirmed using BigDye version 3.1 and an ABI DNA sequencer (3730xl).

Purification of glycopeptides for LC–MS/MS analysis by IGOT

Wild-type and png1Δ cells were grown in YPD liquid medium. Cultured cells (10 g) were washed twice with TBS buffer [50 mM Tris–HCl (pH 7.5) and 150 mM NaCl], resuspended with 10 ml of lysis buffer [TBS buffer containing 1 M sorbitol and a protease inhibitor cocktail (complete, EDTA-free) (Roche)] and were lysed with glass beads. The lysate was centrifuged for 5 min at 1000 × g to remove cell debris. The supernatant fraction thus obtained was subjected to ultracentrifugation at 100 000 × g for 60 min at 4°C to obtain the cytosolic fraction. Trichloroacetic acid was added (final concentration 10%, w/v) to the supernatant on ice for 30 min. The samples were centrifuged at 15 000 × g for 15 min at 4°C and pellets were washed with acetone. The pellets were solubilized in 50 ml of guanidine—HCl buffer [7 M guanidine–HCl, 50 mM Tris–HCl (pH 8.6) and 20 mM EDTA]. Dithiothreitol was added (final concentration 50 mM) to the solubilized fraction, and the resulting suspension was incubated at room temperature for 30 min. For alkylation, 2-iodoacetamide was added (final concentration 400 mM) and the resulting suspension was incubated in the dark at room temperature for 60 min. The samples were dialyzed against 10 mM ammonium bicarbonate three times. The samples were concentrated to 5 mg/ml by evaporation. TPCK Trypsin (Worthington) was added 1:100 (w/w) and the suspension was incubated overnight at 37°C. TPCK Trypsin was re-added 1:100 (w/w) and incubation was continued for 5 h at 37°C. AEBSF was then added (final concentration 1 mM) to stop the protease reaction. Glycopeptides were collected using ConA agarose (J-OIL MILLS) as follows. A 5 ml sample of a tryptic digest, prepared as described above, was applied to a column that contained 1 ml of ConA agarose resin equilibrated with equilibration buffer (50 mM Tris–HCl, pH 7.5, 150 mM NaCl2, 1 mM MgCl2, 1 mM CaCl2). A flow-through sample was applied again to the same ConA agarose column. The ConA agarose resin was then washed three times with 4 ml of equilibration buffer and the glycopeptides were then eluted by washing the column with 2 ml of equilibration buffer containing 0.5 M methyl α-d-mannoside. The samples were desalted using Sep-Pak C18 (Waters) according to the manufacturer's instruction and were concentrated by evaporation.

IGOT-based LC–MS/MS analysis

N-Glycosylated peptides were treated with PNGase F (TakaraBio) in a buffer prepared with 18O-labeled water (H218O) to remove N-glycans and to concomitantly label the glycosylated Asn side chain as described previously [28]. The resulting stable isotope-labeled peptides were analyzed by LC–MS and identified by a Mascot search as described previously [29] with minor modifications. Briefly, the peptide mixture was injected into a C18 trap column (0.5 mm × 1 mm, packed in-house). After washing, the column was connected to a nanoflow LC system (flow rate: 100 nl/min), and the peptides were separated on a reverse-phase (C18) tip column (150 μm × 70 mm, packed in-house) by a linear gradient of MeCN (0−35% in 0.1% formic acid) for 70 min. The eluted peptides were sprayed directly into a quadrupole time-of-flight hybrid mass spectrometer (Q-TOF Ultima; Waters, Milford, MA). The spectra were obtained in the data-dependent MS/MS mode and were processed using the MassLynx software (version 4.0; Waters) to create peak list files after smoothing by the Savitzky−Golay method (window channels, ±3). The files were processed by the MASCOT algorithm (version 2.4.1; Matrix Science, Boston, MA) to assign peptides using a protein sequence database (orf_trans.fasta obtained from Saccharomyces Genome Database, 5884 entries, downloaded on 6 March 2008, http://downloads.yeastgenome.org). The database search was performed with previously described parameters [28]. Briefly, the following parameters were used: enzyme, trypsin; maximum missed cleavage, 2; fixed modification, carbamidomethylation (Cys); variable modifications, deamination (pyroGlu, peptide N-terminal Gln), oxidation (Met) and IGOT (deamidation incorporating 18O, +3 Da, Asn); peptide mass tolerance, 200 ppm and fragment mass tolerance, 0.5 Da. All results of the peptide search were exported in CSV files and processed by Microsoft Excel. We initially selected the peptides with rank 1 with an expectation value within 0.05 as ‘identified peptides.’ If a prospective ‘identified peptide’ contained one or more 18O-labeled aspartic acid residues in the consensus tripeptide sequence for N-glycosylation (Asn-Xaa-[Ser/Thr/Cys], where Xaa is any amino acid except Pro), the peptide was accepted as an ‘N-glycopeptide’.

In order to clarify the PNGase F-dependency on the incorporation of 18O into peptides, control experiments without PNGase F treatment were carried out. To this end, half of the ConA-captured tryptic glycopeptides were further purified by hydrophilic interaction chromatography on a TSKgel Amide-80 column (2.0 mm id × 10 mm, TOSOH, Japan), as described recently [30]. After evaporation, one-fourth of the obtained N-glycosylated peptides were dissolved in PNGase buffer, 50 mM Tris–HCl, pH 8.6, prepared with H218O (98% atoms 18O, Taiyo Nippon Sanso), and the resulting solution was incubated at 37°C overnight (with or without 0.4 mU PNGase F).

The samples thus prepared were analyzed using a nanoflow LC–MS system, essentially as described in a recent report [31]. The incubated samples were separated by the LC system (5–35% MeCN/45 min gradient, flow rate: ∼300 nl/min) using a reversed-phase ODS tip column (0.15 mm i.d. × 75 mm, Nikkyo Technos, Japan), and the effluent was electrosprayed directly into the mass spectrometer (LTQ-Orbitrap Velos, Thermo Scientific), which was operated in the positive ion and data-dependent acquisition mode (spray voltage: 2.0 kV, ion transfer tube temperature: 250°C). Full MS scans were acquired by means of the Orbitrap analyzer (resolution: 30 000 at m/z 400, mass range: 440–1 500, maximum ion accumulation time: 500 ms, micro scan: 1), the 10 most intense ions were fragmented by collision-induced dissociation (selection threshold for MS–MS: 10 000 counts, normalized collision energy: 35, activation q: 0.25, dynamic exclusion duration time: 60 s), and the MS–MS spectra were then analyzed by ion trap MS.

Proteins were identified by a Mascot database search as described recently [31] with minor modifications. Raw data files were converted into Mascot generic format files using the Protein Discoverer software (version 1.4; Thermo Fisher Scientific) and then processed using the Mascot server (version 2.4; Matrix Science) to assign peptides by searching the NCBI Refseq for S. cerevisiae S288c (downloaded on 3 February 2016, 5915 sequences). The database search was performed using the parameters described above, with a peptide molecular weight tolerance of 7 ppm; peptide charges of +2–+4; MS/MS tolerances of 0.8 Da; significance threshold, 0.05; false discovery rate, <0.02. The peptide search results were exported in the form of a CSV file and processed by Microsoft Excel for further analysis, as described above. In the case of the PNGase F treatment, 95% (74 out of 78 proteins) of the N-glycoproteins from wild-type (WT) cells were identified and 90% (78 out of 87 proteins) of those from png1Δ cells were identified, validating the consistency of the new analyses. The control sample without PNGase F treatment was also subjected to the same analysis to identify the ‘N-glycopeptide’.

Preparation of yeast cell extracts and western blotting

Preparation of yeast cell extracts and western blotting were carried out as described previously [22]. A different procedure was used to extract multiple membrane-spanning proteins, such as Pmt6 and Sng1. After adding the sample buffer, the samples were allowed to stand at 37°C for 1 h. Antibodies were used at the following dilutions: 1:10 000 for anti-Pgk1 (22C5; Invitrogen, CA, USA) and 1:20 000 for anti-HA (H9658; Sigma, MO, USA).

Cycloheximide decay assay

Cells were grown and cycloheximide (CHX) (01810; Sigma) was added to the cultures (final concentration 4 µg/ml). The cultures were collected at the indicated times, and the cells were subjected to western blotting analysis.

Results

Identification of candidate of novel ERAD substrate glycoproteins

In order to identify endogenous ERAD substrates in yeast, we compared the cytosolic glycoproteins/glycopeptides in wild-type cells with those of png1Δ cells. To this end, glycoproteins/glycopeptides were purified from the yeast cytosolic fraction using ConA lectin and tryptic peptides obtained were subjected to IGOT-based LC–MS analysis [23]. In the IGOT method, N-glycosylated asparagine residues are converted to aspartic acids after PNGase digestion. As the reactions were carried out in the presence of H218O, N-glycosylated asparagine was converted to 18O-labeled aspartic acid during the PNGase F digestion. Through the analysis of samples prepared from both growth and early stationary phases, numerous 18O-labeled peptides were obtained from the cytosol of WT and png1Δ cells; a total of 89 proteins were identified as N-glycoproteins (Table 1 and Supplementary Tables S1 and S2). Among these 89 proteins, 11 proteins (Asi3, Eps1, Kre2, Mmm1, Mnn1, Npr1, Pmt6, Scs2, Sng1, Suc2 and YBR016W) were specifically detected in png1Δ cells (Supplementary Table S1 and Figure S1). The known properties of these proteins were summarized in Table 2. In order to unequivocally confirm that the incorporation of 18O into the peptides was due to the PNGase F treatment, a control incubation without PNGase F was carried out. In the latter case, no N-glycopeptides were identified from WT cells, while only one glycopeptide from Ecm33 was identified from png1Δ cells (data not shown). This result clearly suggests that the non-enzymatic incorporation of 18O into the peptide is negligible, further validating the IGOT strategy used to identify the N-glycopeptides.

Table 1
Glycoproteins detected in wild-type and png1Δ cells

The column of ‘Both strains’ shows glycoproteins detected from both strains. The column of ‘Wild type’ shows glycoproteins detected from wild-type cells. The column of ‘png1Δ’ shows glycoproteins detected only from png1Δ cells. The row entitled ‘Total’ shows combined data from log- and stationary phases.

 Both strains Wild type png1Δ 
Log phase 55 20 
Stationary phase 53 30 
Total 76 11 
 Both strains Wild type png1Δ 
Log phase 55 20 
Stationary phase 53 30 
Total 76 11 
Table 2
Properties of ERAD substrate candidates

‘No. of NxS/T (IGOT)’ indicates number of putative N-glycosylation sites and bold number in parentheses indicate asparagine residues detected as N-glycosylation site by IGOT. TM refers to the transmembrane region. The number of TM was indicated by several references [45,47,48] and predicted by TMHMM. INM, ER and PM refer to the inner nuclear membrane, endoplasmic reticulum and plasma membrane, respectively. The column entitled ‘Deglycosylation by Png1’ and ‘Degradation delay’ are based on the results shown in Figure 3 

Name No. of NxS/T (IGOT) No. of TM Localization Function Deglycosylation by Png1 Degradation delay 
Asi3 6 (17INM Ubiquitin ligase Yes – 
Eps1 5 (299ER PDI Yes Yes 
Kre2 1 (197Golgi Mannosyltransferase – – 
Mmm1 4 (50,55ER ERMES complex subunit Yes – 
Mnn1 3 (225Golgi Mannosyltransferase – – 
Npr1 16 (54Cytosol Protein kinase – – 
Pmt6 4 (404ER Mannosyltransferase – – 
Scs2 3 (202ER Regulate transcription factor – – 
Sng1 9 (190PM Unknown – Yes 
Suc2 14 (369Extracellular Invertase – – 
YBR016W 1 (31PM Unknown – – 
Name No. of NxS/T (IGOT) No. of TM Localization Function Deglycosylation by Png1 Degradation delay 
Asi3 6 (17INM Ubiquitin ligase Yes – 
Eps1 5 (299ER PDI Yes Yes 
Kre2 1 (197Golgi Mannosyltransferase – – 
Mmm1 4 (50,55ER ERMES complex subunit Yes – 
Mnn1 3 (225Golgi Mannosyltransferase – – 
Npr1 16 (54Cytosol Protein kinase – – 
Pmt6 4 (404ER Mannosyltransferase – – 
Scs2 3 (202ER Regulate transcription factor – – 
Sng1 9 (190PM Unknown – Yes 
Suc2 14 (369Extracellular Invertase – – 
YBR016W 1 (31PM Unknown – – 

While we attempted to clone all genes into the pENTR D-TOPO plasmid, we were not able to clone the YBR016W gene possibly due to its repetitive sequence. We therefore used the other 10 genes for further analyses to determine whether some of them are, in fact, deglycosylated by Png1 in vivo. Accordingly, those genes were cloned into pAG425GPD-ccdB-HA by means of the Gateway cloning system and were expressed in WT yeast cells. The N-glycosylation of these candidate proteins was confirmed by western blotting with or without an Endo H treatment. It was found that, out of these 10 proteins, 8 (Asi3, Eps1, Kre2, Mmm1, Mnn1, Pmt6, Sng1 and Suc2) were indeed N-glycosylated, as judged by their Endo H sensitivity (Figure 1). On the other hand, no evidence was found for N-glycosylation on Npr1 and Scs2 (Figure 1). This is not surprising for Npr1, since it is predicted to be a cytosolic protein. We have no explanation for why the molecular weight of Npr1 increases upon Endo H treatment (Figure 1). The result, however, was highly reproducible and therefore cannot be assumed to be a technical error.

N-Glycosylation of ERAD substrate candidates.

Figure 1.
N-Glycosylation of ERAD substrate candidates.

Wild-type cells harboring ERAD substrate candidate proteins expression plasmid were extracted and digested with (+) or without (−) Endo H, and the products resolved by SDS–PAGE. Proteins were visualized by immunoblotting using anti-HA antibody.

Figure 1.
N-Glycosylation of ERAD substrate candidates.

Wild-type cells harboring ERAD substrate candidate proteins expression plasmid were extracted and digested with (+) or without (−) Endo H, and the products resolved by SDS–PAGE. Proteins were visualized by immunoblotting using anti-HA antibody.

We used the eight proteins that were verified as being N-glycosylated for further analyses.

Asi3, Eps1, Mmm1, Pmt6 and Sng1 are ERAD substrates

Next, in order to confirm whether the eight proteins are degraded by proteasomes, a CHX decay assay was carried out. Eight HA-tagged candidate glycoproteins were expressed in pdr5Δ cells and the cells were treated with 50 µM MG-132. MG-132 is a proteasome inhibitor and the pdr5Δ cells were used as a drug-permeable strain so that a proteasome inhibitor treatment can be applied to yeast cells [32]. As shown in Figure 2, five proteins (Asi3, Eps1, Mmm1, Pmt6 and Sng1) were stabilized in the presence of MG-132 (Figure 2A,B,D,F,G). These results indicate that those proteins are degraded in a proteasome-dependent fashion in vivo. On the other hand, in the case of the other three glycoproteins (Kre2, Mnn1 and Suc2), delayed degradation was not obvious under the experimental conditions employed (Figure 2C,E,H).

Proteasomal degradation of ERAD substrate candidates.

Figure 2.
Proteasomal degradation of ERAD substrate candidates.

Eight glycoproteins were expressed in pdr5Δ cells. These cells were treated with MG-132 for 1 h before the CHX decay assay. CHX was added at t = 0 min. Samples were collected at the indicated times. Samples were subjected to SDS–PAGE, followed by immunoblotting using anti-HA antibody. The immunoblot was also probed with anti-Pgk1 antibody as a loading control. Three independent experiments were performed. Quantification data of the rate of protein degradation were represented as a graph. (A) CHX decay assay of Asi3 (n = 3). (B) CHX decay assay of Eps1 (n = 3). (C) CHX decay assay of Kre2 (n = 3). (D) CHX decay assay of Mmm1 (n = 3). (E) CHX decay assay of Mnn1 (n = 3). (F) CHX decay assay of Pmt6 (n = 3). (G) CHX decay assay of Sng1 (n = 3). (H) CHX decay assay of Suc2 (n = 3). Student's paired t-test was applied.

Figure 2.
Proteasomal degradation of ERAD substrate candidates.

Eight glycoproteins were expressed in pdr5Δ cells. These cells were treated with MG-132 for 1 h before the CHX decay assay. CHX was added at t = 0 min. Samples were collected at the indicated times. Samples were subjected to SDS–PAGE, followed by immunoblotting using anti-HA antibody. The immunoblot was also probed with anti-Pgk1 antibody as a loading control. Three independent experiments were performed. Quantification data of the rate of protein degradation were represented as a graph. (A) CHX decay assay of Asi3 (n = 3). (B) CHX decay assay of Eps1 (n = 3). (C) CHX decay assay of Kre2 (n = 3). (D) CHX decay assay of Mmm1 (n = 3). (E) CHX decay assay of Mnn1 (n = 3). (F) CHX decay assay of Pmt6 (n = 3). (G) CHX decay assay of Sng1 (n = 3). (H) CHX decay assay of Suc2 (n = 3). Student's paired t-test was applied.

Png1 is involved in the deglycosylation of Asi3, Eps1, Mmm1 and Sng1

Having confirmed that five glycoproteins were degraded by the ERAD process in vivo, we next attempted to confirm that these proteins underwent deglycosylation. To this end, a CHX decay assay was carried out using WT and png1Δ cells. As shown in Figure 3A, the degradation of Eps1 was delayed slightly in png1Δ cells and this result was statistically significant (P = 0.019). To confirm the deglycosylation of Eps1, we performed a CHX decay assay using pdr5Δ and pdr5Δ png1Δ cells. Our recent results indicated that a small amount of deglycosylated CPY* can be detectable in the presence of a proteasome inhibitor [24]. The deglycosylated Eps1 clearly accumulated in pdr5Δ but not in pdr5Δ png1Δ cells in the presence of MG-132 (Figure 3B). These data indicate that Png1 is required for both the efficient degradation and deglycosylation of Eps1. As shown in Figure 3C, the degradation of Sng1 was delayed slightly in png1Δ cells and this result was statistically significant (P = 0.016). To confirm the deglycosylation of Sng1, we performed a CHX decay assay using pdr5Δ and pdr5Δ png1Δ cells. However, the deglycosylated Sng1 could not be detected (data not shown). As shown in Figure 3D, a delayed degradation of Asi3 was not detected in png1Δ cells. However, a small amount of deglycosylated Asi3 was observed in pdr5Δ but not pdr5Δ png1Δ cells in the case of an MG-132 treatment (Figure 3E), indicating that Png1 indeed cleaves N-glycans from Asi3 in vivo. Delayed degradation of Mmm1 was not detected in png1Δ cells (Figure 3F). However, the time-dependent reduction in deglycosylated/non-glycosylated Mmm1 was observed in png1Δ cells when compared with WT cells (Figure 3F, arrow). Therefore, the ratio of non- or deglycosylated Mmm1 to total Mmm1 was compared between WT and png1Δ cells. The finding showed that non- or deglycosylated Mmm1 was significantly reduced in png1Δ cells at the 60 min time point (P = 0.019). This result suggests that at least some deglycosylated Mmm1 was produced by Png1 in WT cells. As shown in Figure 3G, neither a deglycosylation defect nor a delayed degradation of Pmt6 was detected in png1Δ cells.

Degradation and deglycosylation of novel ERAD substrates.

Figure 3.
Degradation and deglycosylation of novel ERAD substrates.

(A) Eps1-HA was expressed in WT and png1Δ cells. CHX was added at t = 0 min. Samples were collected at the indicated times. Samples were subjected to SDS–PAGE, following immunoblotting and quantification, as described in Figure 2 (n = 3). (B) Eps1-HA was expressed in pdr5Δ and pdr5Δ png1Δ cells. The MG-132 treatment, the CHX decay experiment, SDS–PAGE and immunoblotting were carried out as described in Figure 2. Arrows indicate deglycosylated Eps1-HA (n = 1). Bar graph indicates the ratio of non-/deglycosylated Eps1-HA relative to total Eps1-HA. (C) Sng1-HA was expressed in WT and png1Δ cells. CHX decay experiment, SDS–PAGE, immunoblotting and quantification were carried out as described in Figure 3A (n = 3). (D) Asi3-HA was expressed in WT and png1Δ cells. CHX decay experiment, SDS–PAGE, immunoblotting and quantification were carried out as described in Figure 3A (n = 3). (E) Asi3-HA was expressed in pdr5Δ and pdr5Δ png1Δ cells. The MG-132 treatment, the CHX decay experiment, SDS–PAGE and immunoblotting were carried out as described in Figure 2. Arrows indicate deglycosylated Asi3-HA (n = 1). Bar graph indicates the ratio of non-/deglycosylated Asi3-HA relative to total Asi3-HA. (F) Mmm1 was expressed in WT and png1Δ cells. CHX decay experiment, SDS–PAGE, immunoblotting and quantification were carried out as described in Figure 3A. Additional bar graph indicates the rate of deglycosylated Mmm1-HA to total Mmm1-HA at 60 min (P = 0.019) (n = 3). (G) Pmt6-HA was expressed in WT and png1Δ cells. CHX decay experiment, SDS–PAGE, immunoblotting and quantification were carried out as described in Figure 3A (n = 3). Student's paired t-test was applied.

Figure 3.
Degradation and deglycosylation of novel ERAD substrates.

(A) Eps1-HA was expressed in WT and png1Δ cells. CHX was added at t = 0 min. Samples were collected at the indicated times. Samples were subjected to SDS–PAGE, following immunoblotting and quantification, as described in Figure 2 (n = 3). (B) Eps1-HA was expressed in pdr5Δ and pdr5Δ png1Δ cells. The MG-132 treatment, the CHX decay experiment, SDS–PAGE and immunoblotting were carried out as described in Figure 2. Arrows indicate deglycosylated Eps1-HA (n = 1). Bar graph indicates the ratio of non-/deglycosylated Eps1-HA relative to total Eps1-HA. (C) Sng1-HA was expressed in WT and png1Δ cells. CHX decay experiment, SDS–PAGE, immunoblotting and quantification were carried out as described in Figure 3A (n = 3). (D) Asi3-HA was expressed in WT and png1Δ cells. CHX decay experiment, SDS–PAGE, immunoblotting and quantification were carried out as described in Figure 3A (n = 3). (E) Asi3-HA was expressed in pdr5Δ and pdr5Δ png1Δ cells. The MG-132 treatment, the CHX decay experiment, SDS–PAGE and immunoblotting were carried out as described in Figure 2. Arrows indicate deglycosylated Asi3-HA (n = 1). Bar graph indicates the ratio of non-/deglycosylated Asi3-HA relative to total Asi3-HA. (F) Mmm1 was expressed in WT and png1Δ cells. CHX decay experiment, SDS–PAGE, immunoblotting and quantification were carried out as described in Figure 3A. Additional bar graph indicates the rate of deglycosylated Mmm1-HA to total Mmm1-HA at 60 min (P = 0.019) (n = 3). (G) Pmt6-HA was expressed in WT and png1Δ cells. CHX decay experiment, SDS–PAGE, immunoblotting and quantification were carried out as described in Figure 3A (n = 3). Student's paired t-test was applied.

These results indicate that Png1 is at least involved in the proteasomal degradation of Asi3, Eps1 and Mmm1. The degradation of Sng1 was also delayed in png1Δ cells. Taken the above results into account, we safely conclude that we successfully identified many natural substrates for Png1 by using the IGOT-based glycoproteomics approach.

Discussion

While it is widely accepted that cytoplasmic PNGase is involved in the ERAD process, its functional importance remains largely unknown. Part of the reason for this is the lack of information on the endogenous substrates for this enzyme. Even for many available model ERAD substrates in S. cerevisiae, RTA/RTL is the only one that has been clearly shown to be degraded in a PNGase-dependent manner, that is, both a lack of PNGase-mediated deglycosylation and a delay in degradation were clearly observed [21,22]. It is therefore imperative to identify the bona fide endogenous substrates for this enzyme to provide deeper insights into the functional importance of the cytoplasmic PNGase.

IGOT was developed as a method for comprehensive glycoproteome analysis [23]. This method can be applied to any organisms, provided the N-glycan structures produced are not resistant to PNGase digestion [23,2831,3335]. In S. cerevisiae, N-glycan structures remain as the high mannose type throughout the secretory pathway, and no modification of the proximal GlcNAc residue, which often renders the glycans resistant to PNGase F digestion [36], has been observed. These characteristics make this organism the perfect choice for an IGOT analysis. In the present study, we applied the IGOT method for the detection of cytosolic N-glycoproteins, 11 of which specifically accumulated in png1Δ cells. In the present study, we characterized 10 of these 11 proteins.

No conclusive evidence was found for the occurrence of N-glycans in 2 (Npr1 and Scs2) out of the 10 proteins tested (Figure 1). It is still possible, however, that those proteins are N-glycosylated with a very low efficiency. It should also be noted that 18O incorporation into Asn may also occur non-enzymatically. Indeed, 18O incorporation into Asn not in the sequon (N-X-S/T, X ≠ P) can also be observed (we found three positions in three proteins), and it should occur through natural deamidation during the PNGase F digestion. Our control experiment without PNGase F treatment, however, clearly suggests that the non-enzymatic deamidation on Asn in the sequon rarely happens, further validating our identification strategy. Npr1 is believed to be a cytosolic protein; therefore, it may be unlikely to assume that this protein undergoes N-glycosylation in vivo.

Among the glycoproteins identified, five (Asi3, Eps1, Mmm1, Pmt6 and Sng1) were found to be degraded intracellularly in a proteasome-dependent fashion (Figure 2). The lack of evidence for proteasomal degradation for other proteins, however, does not exclude the possibility that those proteins could still be endogenous substrates for Png1, as the ERAD process may involve the processing of only a minor portion of those proteins. In any case, our results imply that at least the five proteins are naturally degraded by proteasomes and therefore can be regarded as endogenous ERAD substrates. Moreover, in three of them (Asi3, Eps1 and Mmm1), the formation of deglycosylated forms was reduced in png1Δ cells, clearly indicating that these proteins were bona fide substrates. Moreover, the degradation of Eps1 and Sng1 was found to be delayed in the png1Δ strain. These results clearly indicate that at least four of those proteins are very likely to be deglycosylated by Png1 in vivo (Table 2) and accordingly would be specifically detected in the cytosolic fractions of png1Δ cells. To the best of our knowledge, these are the first examples of the identification of natural substrates for the yeast Png1 enzyme.

It is a bit puzzling that a delay in degradation, however, was not observed for two of the Png1 substrates, namely, Asi3 and Mmm1. This situation resembles the ones for various mammalian glycoprotein substrates, for which deglycosylation by PNGase can be observed upon the inhibition of proteasome activity, but a delay in degradation was not obvious, even when PNGase activity was inhibited [11,12]. Consistent with these observations, the N-glycans on glycoproteins were shown to not impair the proteasomal degradation of several model glycoprotein substrates [37]. As glycopeptides are detected only in png1Δ cells, the accumulation of those proteins in png1Δ was expected. It may be possible that specific proteolytic intermediates that were not detected by western blotting may have accumulated in png1Δ cells. In this connection, it should be noted that a delay in the degradation of CPY* or its derivative, CTL, was observed [4,22], while their deglycosylation in vivo was not initially evident. Very recently, however, it was found that Png1 is indeed involved in the deglycosylation of CPY*, but limited proteolysis by an endoprotease activity hindered the detection of the intact deglycosylated CPY* [24].

Among the natural Png1 substrates identified, Asi3 is a member of the Asi complex located in the inner nuclear membrane (INM) [38]. It was previously reported that Asi3 is required for the proteasomal degradation of several proteins that are located in INM [39,40]. Our results indicate that Asi3 is not only an ERAD component for degradation of INM proteins, but is also an ERAD substrate. It was reported that Png1 was localized not only in the cytosol, but also in the nucleus [4]. In addition, Png1 contains a putative nuclear localizing signal on its C-terminal side (295-FITKRLRYS-303) by cNLS Mapper [41]. It is therefore possible that N-glycans on Asi3 localized in INM could somehow be removed by nucleus-localized Png1.

Eps1 is a member of protein disulfide isomerase family in the ER and is required for ERAD [42,43]. In the present study, Eps1 was found to be an ERAD substrate and to be deglycosylated by Png1 (Figures 2B and 3B). While the amount of deglycosylated Eps1 detected was very low, this may be due to this protein undergoing proteolysis during the ERAD, as was the case with CPY* [24]. Eps1 was degraded in an Hrd1-dependent manner (data not shown), implying that the ER luminal side or transmembrane region of Eps1 is recognized by ERAD components.

Sng1 is a multiple membrane-spanning protein that is localized in the plasma membrane and required for resistance to 6-azauracil [44]. Sng1 is the first Png1-dependent multiple membrane-spanning ERAD substrate identified in yeast. On the other hand, in png1Δ cells, a delay in its degradation was observed, while the deglycosylation in wild-type cells was not so obvious. These seemingly contradictory results may be partly due to the technical difficulty associated with identifying PNGase-mediated deglycosylation, as was the case of CPY*/CTL [4,22].

Mmm1 is localized in the ER membrane of the ER–mitochondria contact site and is required for the connection of these organelles as an ERMES (ER–mitochondria encounter structure) complex member [45]. In png1Δ cells, delayed degradation of Mmm1 was not observed (Figure 3F). However, a time-dependent reduction in the formation of deglycosylated Mmm1 was observed in png1Δ cells, suggesting that Png1 is required for the deglycosylation of Mmm1.

Recently, human patients bearing mutations in NGLY1 (mammalian PNGase gene ortholog) have been identified [1518]. Given the severe symptoms observed for such patients, clarification of the pathophysiology of an NGLY1-deficiency is urgently awaited. To this end, the identification of endogenous substrates for the cytoplasmic PNGase will be a key. In the present study, we successfully applied an IGOT-based glycoproteomic analysis to the identification of several Png1 substrates in yeast. Mammalian cells produce another deglycosylating enzyme that acts on N-glycans in the cytosol (endo-β-N-acetylglucosaminidase [46]); therefore, this situation is not as simple as the case with yeast. Nevertheless, we believe that the IGOT method will be a powerful tool for the identification of any glycoproteins that specifically accumulate in tissues or cells derived from patients. The identification of endogenous substrates for NGLY1 will be critical for developing an understanding of the pathophysiology of an NGLY1 deficiency.

Abbreviations

CHX, cycloheximide; ER, endoplasmic reticulum; ERAD, ER-associated degradation; ERMES, ER–mitochondria encounter structure; ERQC, ER quality control; H218O, 18O-labeled water; IGOT, isotope-coded glycosylation site-specific tagging; INM, inner nuclear membrane; MS, mass spectrometry; PDI, protein disulfide isomerase; PNGase, peptide:N-glycanase; WT, wild type.

Author Contribution

A.H. performed all the cell culture studies and wrote the paper. M.F. prepared the MS samples. A.T. performed the MS study. H.K. performed the MS study and also contributed to the discussion section of the data. T.S. designed and directed the project and also wrote the paper.

Funding

The work was supported in part by a Grant-in-Aid for Scientific Research (C) [25440058] (to A.H.) and a Grant-in-Aid for Scientific Research (B) [25291030] (to T.S.) from the Ministry of Education, Science, Sports and Culture of Japan.

Acknowledgments

We thank the members of the Glycometabolome Team (RIKEN) for fruitful discussions. We thank Ms. Yae Tsuchiya and Tsugiyo Matsuda (RIKEN) for technical support.

Competing Interests

The Authors declare that there are no competing interests associated with the manuscript.

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doi:

Author notes

*

Present address: Graduate School of Agriculture, Shinshu University, 8304 Minamiminowa, Kamiina, Nagano 399-4598, Japan.

Supplementary data