N–linked glycosylation is a highly conserved and essential protein modification process in eukaryotic cells. Oligosaccharyltransferase (OST) complexes in the endoplasmic reticulum (ER) initiate N–glycosylation by catalyzing the attachment of high-mannose oligosaccharides to specific asparagine residues at acceptor sites, which are further modified within the ER and Golgi complex. The resulting N–linked glycans play essential roles in protein folding, assembly, trafficking, stability, and various other functions (1, 2).

In mammalian cells, STT3A−OST and STT3B−OST complexes perform co- and post-translational glycosylation, respectively. Each OST complex consists of an isoform-specific catalytic subunit (STT3A or STT3B) along with multiple accessory proteins, which include six shared and one isoform-specific subunit. Ribophorin I (RPN1) is a non-catalytic subunit in both OST complexes, and is a type I transmembrane protein with a short C–terminal cytoplasmic tail. In the STT3A−OST complex, its cytoplasmic tail adopts a four-helix bundle structure, whereas it remains unstructured in STT3B−OST (3). This bundle in STT3A−OST is directly associated with a membrane-bound translating ribosome (4), which may explain the co-translational glycosylation by STT3A−OST, but not by STT3B−OST. RPN1 depletion decreases the expression of STT3A and STT3B, and severely reduces OST activity (5). RPN1 binds to newly synthesized OST substrates to facilitate N–glycosylation (6-8). Additionally, RPN1 functions as an N–glycosylation-dependent chaperone that facilitates protein folding. Thus, RPN1 promotes the maturation of opioid receptors (the rhodopsin G protein-coupled receptor [GPCR] family members) and their transport from the ER to the cell surface in an N–glycosylation-dependent manner (9).

Ring finger protein 128 (RNF128), also known as gene related to anergy in lymphocytes (GRAIL), is a type I transmembrane E3 ligase containing a luminal protease-associated domain and a cytosolic RING finger domain (10). It is primarily localized in the ER, and is known for regulating T cell anergy, cytokine production (10-12), and contributing to cancer progression. However, its role in N–glycosylation remains unknown. In this study, we used a proximity-dependent biotin labeling proteomic technique (13-15) to investigate the function of RNF128 in N–glycosylation by identifying RPN1 as its target, and evaluating its involvement in the glycosylation process.

RNF128 promotes RPN1 ubiquitination and degradation

To discover substrates of RNF128, we employed a proximity-dependent biotinylation method that we previously developed (14) to capture proteins ubiquitinated by a nearby E3 ligase. The ubiquitinated proteins were then identified by proteomic analysis using LC−MS/MS, as described in the Methods section. We detected RPN1 as a potential RNF128 substrate (Fig. 1A, B). To validate RPN1 as a target of RNF128, HeLa cells were co-transfected with RNF128−BirA−HA, RPN1−FLAG, and AP−Ub. RPN1−FLAG exhibited strong biotinylation by RNF128−BirA−HA, but not by BirA−HA or the ligase-dead mutant RNF128 H297/300N−BirA−HA, indicating that RPN1 is a specific substrate of RNF128 (Fig. 1C).

To further confirm that RPN1 is a substrate of RNF128, we investigated whether RNF128 influences RPN1 expression. Co-transfection of HeLa cells with RPN1−FLAG and either RNF128−HA or the ligase-dead mutant RNF128 H297/300N−HA revealed that RNF128−HA significantly reduced RPN1−FLAG levels, while RNF128 H297/300N−HA had no effect (Fig. 1D). Transfection with RNF128−HA, but not with the RNF128 H297/300N−HA mutant, also reduced the level of endogenous RPN1 (Fig. 1E). The RNF128-mediated reduction in RPN1 was inhibited by treatment with the proteasome inhibitor MG132 or the lysosome inhibitor NH₄Cl (Fig. 1F). Co-immunoprecipitation (Co–IP) assays further demonstrated that RNF128 directly binds to RPN1 (Fig. 1G).

A ubiquitination assay in HeLa cells co-expressing RNF128−HA, RPN1−FLAG, and T7−Ub, followed by immunoprecipitation with RPN1−FLAG, showed that RNF128, but not the RNF128 H297/300N mutant, produced a ubiquitination smear on RPN1 (Fig. 1H), indicating specific ubiquitination by RNF128. Immunofluorescence analysis confirmed that RNF128 reduced RPN1 expression in cells transfected with the wild-type, but not with ligase-dead RNF128 (Fig. 1I). Collectively, these results demonstrate that RNF128 ubiquitinates RPN1, leading to its degradation via proteasome- and lysosome-dependent pathways, identifying RPN1 as a novel RNF128 target.

RNF128 regulates N–glycosylation through RPN1 modulation

Since RPN1 depletion reduces the levels of STT3A and STT3B (5, 6), we used two well-characterized model glycoproteins: sex hormone–binding globulin (SHBG), and asialoglycoprotein receptor 1 (ASGR1), to investigate whether RNF128 affects N–glycosylation. SHBG is an STT3B-dependent substrate with two glycosylation sites located at the extreme C–terminus (16). ASGR1 is a STT3A- and STT3B-dependent substrate with two glycosylation sites (6).

We employed immunoblotting and pulse–chase labeling to analyze the N–glycosylation patterns of these proteins. Cells expressing wild-type RNF128 exhibited significant alterations in the glycosylation patterns of SHBG and ASGR1, characterized by a reduction in the fully glycosylated forms, and an increase in the noN–glycosylated and monoglycosylated forms of both proteins (Fig. 2A, C). In contrast, no changes were observed in cells expressing the ligase-dead RNF128 mutant. Pulse–chase labeling of newly synthesized glycoproteins revealed a decrease in the fully glycosylated form, and an increase in the monoglycosylated form of newly synthesized SHBG (Fig. 2B). For ASGR1, there was a slight reduction in the fully glycosylated form, accompanied by a marked increase in the non-glycosylated form (Fig. 2D). Subsequently, we utilized a stable RPN1-knockdown HeLa cell line to assess whether RPN1 is required for N–glycosylation of these proteins. RPN1 knockdown led to reduced glycosylation of SHBG and ASGR1, which was restored by the expression of an shRNA-resistant form of RPN1 (Fig. 2E, F). These findings suggest that RNF128 regulates N–linked glycosylation by promoting the degradation of RPN1.

RNF128 affects the transport of opioid receptor mu 1 (OPRM1) to the cell surface by regulating RPN1 expression

N–glycosylation is crucial for the cell surface expression of proteins, including opioid receptors. The absence of N–glycosylation at any one of the five glycosylation sites prevents the export of OPRM1 to the plasma membrane (9). RPN1 is required for the cell surface trafficking of OPRM1 by functioning as its N–glycosylation-dependent chaperone. Given that RNF128 modulates the protein levels of RPN1, we sought to investigate whether RNF128 affects the export of OPRM1 to the plasma membrane, using confocal microscopy to monitor OPRM1−GFP expression in cells expressing RNF128 wild type or RNF128 H297/300N. OPRM1−GFP remained localized intracellularly in cells overexpressing RNF128, whereas it was observed both intracellularly and on the surface of mock-transfected cells and RNF128 H297/300N-overexpressing cells (Fig. 3A). Consistent with these observations, cells treated with RPN1-specific siRNA were also unable to translocate OPRM1 to the plasma membrane (Fig. 3B). These results suggest that RNF128 inhibits the export of OPRM1 to the cell surface, similar to the effect observed with RPN1 deficiency.

To confirm that the RNF128-mediated inhibition of OPRM1 export is due to RPN1 degradation, rather than other RNF128 substrates, we used three RPN1 mutants with C–terminal deletions in the cytosolic region containing 14 lysine residues, potential ubiquitination sites for RNF128 (Fig. 3C, left panel). Two deletion mutants (amino acids 1−577 and 1−523), retaining ten and six lysine residues, respectively, were degraded by RNF128, whereas the mutant with all 14 lysines removed (RPN1 1−464) was not affected (Fig. 3C, right panel). We also generated a K0 mutant of RPN1, substituting all 14 lysine residues with arginine. Like RPN1 1−464, RPN1 K0 was resistant to RNF128-mediated degradation (Fig. 3D). Further, we assessed whether these ubiquitination-resistant RPN1 mutants could prevent RNF128 from inhibiting OPRM1 export to the plasma membrane. Co-transfection of RNF128 with RPN1 K0 permitted OPRM1−GFP surface expression, suggesting that the absence of ubiquitination by RNF128, and consequently the prevention of RPN1 degradation, is crucial for OPRM1 export to the cell surface (Fig. 3E). Unexpectedly, co-expression of RNF128 with RPN1 1−464, which lacks the C–terminal domain, predominantly retained OPRM1 intracellularly, indicating that this truncation impedes OPRM1 export. These findings demonstrate that RNF128 reduces RPN1 levels, thereby inhibiting OPRM1 transport to the plasma membrane, and highlight the essential role of the RPN1 cytoplasmic tail in facilitating OPRM1 export to the cell surface.

RNF128 inhibits colorectal cancer (CRC) cell migration by regulating RPN1

Given that RNF128 affects the N–glycosylation and cell surface expression of glycoproteins, we sought to investigate its biological relevance. Notably, RNF128 is significantly upregulated in colon cancer tissues (17). To evaluate the impact of RNF128 and RPN1 on colon cancer cell migration, stable cell lines were here employed. HT29 cells, which exhibit high endogenous RNF128 expression, were used to generate stable knockdowns of RNF128 and RPN1, while DLD1 cells, with low RNF128 expression, were engineered to overexpress RNF128 (Fig. 4A). RPN1-depleted HT29 cells displayed a significantly increased migration rate, whereas RNF128 knockdown in HT29 cells, accompanied by elevated RPN1 expression, resulted in a marked decrease in migration (Fig. 4B, C). Similarly, RNF128 overexpression in DLD1 cells led to reduced RPN1 levels and the enhanced migration of cells (Fig. 4D, E). These findings suggest that RNF128 promotes cancer cell migration by downregulating RPN1.

N–glycosylation is a prerequisite for exit from the ER, and subsequent transport to the cell surface of selective N–glycosylated plasma membrane proteins, including a subset of GPCRs (18, 19). Depletion of RPN1 affects N–glycosylation of a subset of OST substrates (5, 6, 20), implying that N–glycosylation of a group of plasma membrane proteins may be highly sensitive to RPN1 expression. Given that RNF128 affects RPN1 expression, conditions of upregulated RNF128 (such as the induction of T cell anergy) or those of RPN1 downregulation likely inhibit the cell surface expression of selective glycoproteins to impact specific cellular functions.

Our data shows that RNF128 decreased the RPN1-dependent N–glycosylation of SHBG and ASGR1, which indicates that RNF128 affects the degradation of RPN1 integrated into the OST complex to reduce N–glycosylation activity. Recently, ankyrin and suppressor of cytokine signaling box 11 (SOCS11) were reported to ubiquitinate RPN1 to increase its turnover by functioning as substrate recognition subunits in elongin–cullin–SOCS E3 ligase (21). Although ASB11-mediated regulation of N–glycosylation has not been explored, our results strongly suggest that the cellular levels of RPN1 are diversely regulated by ubiquitination. This is consistent with the notion that N–glycosylation is not a constitutive process, but is instead tightly regulated (22). On-demand regulation of N–glycosylation and cell surface expression of selective glycoproteins may contribute to physiological processes, including T cell anergy and tumor progression.

RPN1 preferentially binds to misfolded proteins (23) and associates with newly synthesized membrane proteins (7), indicating its potential role as a molecular chaperone. RPN1 overexpression rescues an OPRM1 mutant that is intrinsically deficient in export, and increases its cell surface expression as a chemical chaperone (9). However, it is unclear whether RPN1 works together with other subunits of the OST complex to mediate this chaperone function. Our data showed that RNF128 affected the cell surface expression of OPRM1 by downregulating RPN1.

Notably, the export of OPRM1 requires the cytosolic domain of RPN1. The cytosolic domain of RPN1 within the STT3A−OST complex (but not within the STT3B−OST complex) directly interacts with the ribosome (3). This observation provides a potential explanation for the role of STT3A−OST, but not STT3B−OST, in facilitating co-translational processes. Consequently, it is likely that at least one of the five N–glycosylation sites of OPRM1 undergoes modification in a co-translational manner. Furthermore, considering that the nascent polypeptide chain exiting the Sec61 translocon may interact with RPN1 in the ER lumen, it is anticipated that the large luminal domain of RPN1 possesses chaperone activity.

Lastly, RNF128 has been reported to be involved in several cancers (24-26). However, information is still lacking on the role of RPN1 in cancer cell metastasis. The data of this study shows that the overexpression of RNF128 and depletion of RPN1 can inhibit the migration of CRC cells. Further study is required to determine the mechanism underlying the function of RNF128 and RPN1 in cancer progression.

In summary, we provide evidence that targeting RPN1 for ubiquitination and degradation by RNF128 affects the N–glycosylation, maturation, and transport of specific glycoproteins. This is the first report of an E3 ubiquitin ligase that directly regulates the N–glycosylation process in the ER. Our findings greatly enhance understanding of the biological and functional roles of RNF128- and RPN1-dependent N–glycosylation.

Plasmids

The full-length cDNAs of RNF128, RPN1, OPRM1, and ASGR1 acquired from the DNASU Plasmid Repository (Tempe, AZ, USA), and that of SHBG from the Korea Human Gene Bank (Daejeon, Korea), were amplified by PCR. Amplified DNA fragments were inserted into pcDNA3.1 (Invitrogen, Waltham, Massachusetts, USA), pEGFP−N1 (CLONETECH, Mountain View, California, USA), or pYR vectors individually to generate the expression plasmid of each gene (13, 15). The cDNA of RNF128 was additionally inserted into another expression vector encoding C–terminally FLAG-tagged BirA, as previously described (13, 14). The deletion mutants of RPN1 (amino acids 1−464, 1−523, and 1−577) were generated by PCR amplification using specific primer sets. To generate the K0 mutant of RPN1, the cytosolic K0 fragment of RPN1 provided by M.Biotech Ltd. (Korea) and the N–terminal fragment of RPN1 were amplified, and fused with each other using PCR with a unique primer set. The mutant with point mutations, RNF128 H297/300N, was generated from the RNF128 plasmid using a QuickChange Site-directed mutagenesis kit, following the manufacturer’s protocol (Agilent, Santa Clara, California, USA). RNF128 FLAG-tagged PCR-amplified cDNA was subcloned into pCDH_CMV_GFP＋Puro (System Biosciences, Palo Alto, CA, USA). The RPN1 shRNA expression vector was generated by the subcloning of RPN1 specific shRNAs into pLKO1 hygro (Addgene, Watertown, MA, USA). Tables S1 and S2 of the Supplementary Information (SI) list the details of primers utilized in the study.

Cell culture and transfection

HeLa and HEK293T cells were cultured in DMEM (WELGENE, Gyeongsangbuk-do, Korea), while HT29 and DLD1 cells were cultivated in RPMI 1640 medium (WELGENE), supplemented with 10% FBS (Gibco Carlsbad, California, USA), 1% penicillin/streptomycin, and maintained at 37°C and 5% CO₂. Lipofectamine 2000, Lipofectamine RNAiMAX (Thermo Fisher Science, Waltham, Massachusetts, USA), and Polyethylenimine (Sigma−Aldrich, St. Louis, Missouri, USA) were used for transient transfection experiments. MG132 and tunicamycin (TM) were purchased from Cayman Chemical Co. (Ann Arbor, MI, USA) and Sigma−Aldrich, respectively.

Affinity purification of biotinylated proteins and ubiquitinated-peptides, and Mass spectrometry

HeLa cells were transfected with the expression constructs of RNF128−BirA−FLAG and AP−HA-Ub. At 24 h post-transfection, cells were treated with biotin and MG132 for 4 h. Following the methods previously described (13, 14), the biotinylated proteins in the cell membrane fraction were processed, and the purified biotinylated-ubiquitinated peptides were analyzed by mass spectrometry using nanoelectrospray LC−MS/MS. The most intense peaks were selected for further MS/MS analysis, and Mascot software was used to search the resulting spectra against the National Center for Biotechonology Information (NCBI) databases.

Immunoprecipitation and Immunoblotting

After transient transfection with plasmids for the indicated time, cells were lysed with lysis buffer (50 mM Tris–HCl (pH 7.4), 150 mM NaCl, 1 mM EDTA, 1% Nonidet P−40, 1 mM dithiothreitol, 0.2 mM PMSF), and processed for immunoprecipitation or immunoblotting, as previously described (13). Table S3 provides all the antibodies used.

Ubiquitination assay

The plasmid constructs expressing T7−Ub and RPN1−FLAG were co-transfected with the constructs expressing pRNF128−HA, or pRNF128 H297/300N−HA, or the control vector, into HeLa cells for 24 h. Cells were treated with 25 μM of MG132 for 4 h, and harvested for the ubiquitination assay, as previously described (13).

Immunofluorescence

Cells on the coverslips were transfected with the indicated genes for 48 h. The processes for the immunofluorescence experiments were conducted as previously described (13), and Table S3 provides the antibody information.

Lentiviral packaging, and generation of the stable cell lines

HEK293T cells were transfected with pLKO.1 hygro; pLKO.1 hygro shRPN1; pCDH_CMV_GFP+Puro or pCDH_CMV_GFP＋Puro RNF128−FLAG with psPAX2 and pMD2.G (Addgene). The lentiviral supernatant produced was used for the transduction of HeLa, HT29, or DLD1 cells, following the protocol previously described (27). Stable clones constitutively expressing RNF128 were selected with 3 μg/ml Puromycin (Sigma−Aldrich), while clones expressing shRNF128 or shRPN1 were selected with 175 μg/ml Hygromycin (AG Scientific, San Diego, CA, USA).

Pulse–chase radiolabeling method

Twenty-four hours after transfection of the cells with the expression plasmids, the medium was changed to Methionine and Cysteine-free DMEM media (Gibco) supplemented with 10% dialyzed FBS. Twenty minutes later, cells were labeled with 100μCi ³⁵S Protein Labeling Mix/ml (PerkinElmer, Waltham, MA, USA) for 5 min. Medium containing unlabeled methionine and cysteine was added to stop the labeling, and cells were further incubated for 15 min. Cells were harvested and lysed with RIPA buffer for 30 min at 4°C. After centrifugation, the supernatant was incubated overnight with anti-FLAG M2 agarose (Sigma−Aldrich) at 4°C. Beads were washed three times with RIPA buffer. Proteins were eluted with gel loading buffer, and separated by SDS−PAGE. Dry gels were exposed to a film (Agfa, Mortsel, Belgium), to detect the labeled proteins.

Wound-healing assay

The migration assay was carried out as previously reported (27). Briefly, HT29 and DLD1 stable cell lines were seeded onto a 6-well plate. When the cell monolayers reached 90% confluency, mitomycin C 5 (μg/ml) was used to inhibit proliferation. After 4 h of treatment, a 200 μL sterile pipette tip was used to generate a linear wound, which was then imaged under a camera mounted on a microscope at appropriate times. The wound-healing rate was calculated based on the average area of cell migration determined using ImageJ software v1.46r (28). GraphPad Prism v.8.0.2 was used for statistical analysis. Statistical significance was estimated by using unpaired Student’s t-test for DLD1 cell lines test and one-way ANOVA with Dunnett's multiple comparisons test for HT29 cell line test.

The authors declare that this work was supported by grants from the National Research Foundation of Korea (NRF-2021R1A2C1011189) and the Ministry of Science and ICT (RS-2023-00273665) by the Korean government.

The authors have no conflicting interests.