Ida Signe Bohse Larsen,Lorenzo Povolo,Luping Zhou,Weihua Tian,Kasper Johansen Mygind,John Hintze,Chen Jiang,Verity Hartill,Katrina Prescott,Colin A. Johnson,Sureni V. Mullegama,Allyn McConkie-Rosell,Marie McDonald,Lars Hansen,Sergey Y. Vakhrushev,Katrine T. Schjoldager,Henrik Clausen,Thomas Worzfeld,Hiren J. Joshi,Adnan Halim
The SHDRA syndrome-associated gene TMEM260 encodes a protein-specific O-mannosyltransferase
glycosylation,congenital disorders of glycosylation,O-mannosylation,glycoproteomics,plexin
Significance We demonstrate that the TMEM260 gene encodes a novel protein-specific O-mannosyltransferase that selectively glycosylates a common protein domain shared among cMET, RON, and plexin receptors. Biallelic mutations in TMEM260 underlie structural heart defects and renal anomalies syndrome (SHDRA), a severe developmental disorder associated with congenital cardiac malformations and early childhood mortality. We show that disease-causing mutations impair the TMEM260 O-mannosyltransferase function, which affects proprotein maturation and intracellular trafficking of receptor substrates and epithelial morphogenesis. Our study uncovers a third biosynthetic pathway for protein O-mannosylation in higher eukaryotes and identifies SHDRA as a new congenital disorder of glycosylation.
The SHDRA syndrome-associated gene TMEM260 encodes a protein-specific O-mannosyltransferase

We demonstrate that the TMEM260 gene encodes a novel protein-specific O-mannosyltransferase that selectively glycosylates a common protein domain shared among cMET, RON, and plexin receptors. Biallelic mutations in TMEM260 underlie structural heart defects and renal anomalies syndrome (SHDRA), a severe developmental disorder associated with congenital cardiac malformations and early childhood mortality. We show that disease-causing mutations impair the TMEM260 O-mannosyltransferase function, which affects proprotein maturation and intracellular trafficking of receptor substrates and epithelial morphogenesis. Our study uncovers a third biosynthetic pathway for protein O-mannosylation in higher eukaryotes and identifies SHDRA as a new congenital disorder of glycosylation.

Congenital disorders of glycosylation (CDGs) are a continuously expanding group of metabolic disorders caused by deficiencies in glycosylation of proteins or lipids (1). More than 50 distinct CDGs are caused by loss of function of glycosyltransferase genes involved in different protein N- and as O-glycosylation pathways (1, 2), and in particular glycosyltransferase genes dedicated to protein O-mannosylation pathways have been identified as causing CDGs (3, 4). The clinical presentation of CDGs often involve multisystemic syndrome features without clear evidence pointing to glycosylation deficiencies, and discovery of novel CDGs often rely on identification of involved genes by whole-exome sequencing (1). It remains challenging to identify genes as glycosylation enzymes. However, new protein O- and C-mannosylation enzymes and novel CDGs have been identified within the last decade (5). Recently, biallelic mutations in the gene TMEM260 were identified in affected individuals of several consanguineous families characterized by structural heart defects, kidney abnormalities, neurological disorders, and perinatal death in a syndrome designated structural heart defects and renal anomalies syndrome (SHDRA) (OMIM: 617478) (6, 7). The causal role of the TMEM260 gene was validated in a zebrafish model by gene knockout (KO) and rescue (7), but the function of the encoded protein and molecular basis for effects remained obscure. We separately identified TMEM260 in a bioinformatic screen for putative multi-pass transmembrane (TM) glycosyltransferases using the Pfam database in course of searching for a novel type of protein O-mannosylation (4). The predicted domain structure of TMEM260 shows similarity to protein O-mannosyltransferases, POMT1 and POMT2 (8), for which deficiencies cause different forms of muscular dystrophy-dystroglycanopathies (9, 10), and the more recently discovered TM and tetratricopeptide repeat-containing proteins 1-4 (TMTC1-4) (11), for which deficiencies in TMTC3 cause brain malformations, e.g. cobblestone lissencephaly (12), and periventricular nodular heterotopia (13), while other TMTCs are associated with hearing loss (14, 15). The protein O-mannosyltransferase 1 and 2 (POMT1/2) and TMTC1-4 are distinct families of protein mannosyltransferase isoenzymes serving different classes of protein clients (4). POMT1/2 transfer O-linked mannose (O-Man) to alpha-dystroglycan (αDG) of the dystrophin-glycoprotein complex (DGC) where elongated complex O-Man glycans on αDG mediate interactions between extracellular matrix proteins, the DGC, and the cellular cytoskeleton (16). The TMTC1-4 family transfer O-Man to the extracellular cadherin (EC) domains of the cadherin superfamily, including E- and N-cadherin and clustered/non-clustered protocadherins (11, 17, 18). However, we previously identified a third class of O-mannosylated proteins characterized by extracellular immunoglobulin, plexin, transcription factor (IPT) domains, which were not dependent on the functions of the POMT1/2 and TMTC1-4 protein O-mannosyltransferase isoenzymes (4, 11). Thus, identification of TMEM260 as a protein with structural similarity to these mannosyltransferases prompted us here to investigate a putative role of TMEM260 in O-mannosylation of extracellular IPT domains. Using CRISPR/Cas9 genetic engineering and O-Man glycoproteomics, we demonstrate that TMEM260 encodes an ER-located protein that indeed is selectively required for O-mannosylation of extracellular IPT domains of cMET, RON, and multiple plexin receptors. We describe two novel patient families with TMEM260 mutations linked to the SHDRA syndrome (6), and demonstrate that these mutations selectively impair glycosylation of IPT domains.

We hypothesized that the candidate protein would show structural similarity (Glycosyltransferase superfamily C (GT-C) folds) (19), but not necessarily significant sequence similarity (11) to the POMTs and TMTCs, or the DPY-19 protein C-mannosyltransferases (20), classified in the GT39, GT105, and GT98 families in CAZy, respectively (21). Candidate screening relied on the PMT yeast orthologs of POMTs. Yeast PMTs match the PMT_2 superfamily domain hmm model (cl21590), and specifically the PMT_2 PFAM model (pfam13231). Using the Conserved Domain Architecture Retrieval Tool (22), we identified sequences in early metazoans that matched the PMT_2 model and found a group of proteins that frequently matched with a sequence model for DUF2723. The sequence model was traced in higher organisms and matched with the TMEM260 gene predicted to encode a 79 kDa multipass TM protein. AlphaFold (23) predicts TMEM260 to share the GT-C fold characteristic for mannosyltransferases, including a conserved N-terminal GT-C module and a variable C-terminal module (Fig. 1) (24, 25). The N-terminal region of TMEM260 adopts the GT-C module (Fig. 1C) commonly found among enzymes that utilize lipid-linked donor sugars, including PMT1/PMT2 (26), ALG6 (25), PglB (27), STT3 (28, 29), ArnT (30) and DPY19 (24) glycosyltransferases. The N-terminal (residues 1 to 410) of TMEM260 is predicted to comprise 11 TM helixes of which TM1-TM7 are superimposable (Fig. 1E) with the conserved GT-C module of PMT1 (26), while the C-terminal region (residues 411 to 707) partially includes tetratricopeptide (TPR) repeats (31) as found in the TMTC1-4 (GT105) and OGT (GT41) glycosyltransferases (32, 33). The first predicted ER-luminal loop of TMEM260 aligns well with the corresponding loop of PMT1, with Asp52 of TMEM260 closely positioned to and oriented toward the same space as the catalytically important Asp77 of PMT1 (34), which faces the co-purified dolichol-phosphate lipid in the PMT1 structure (Fig. 1F), suggesting that Asp52 of TMEM260 may be important for catalytic functions. Like the TMTC1-4 isoenzyme family, the C-terminal region of TMEM260 protrudes into the ER-lumen (SI Appendix, Fig. S2), where the predicted TPR domain is likely involved in substrate recognition and selective recruitment of IPT-domains for O-Man glycosylation.

Transient expression of tagged TMEM260 (TMEM260-3×FLAG) in BG1 human ovarian cancer cell line (35) indicated ER-localization (Fig. 1G), which is in agreement with the presence of di-arginine ER-retention motif at residues 20 to 22 (36) and consistent with the established subcellular localization of POMTs and TMTCs (37–39). We used anti-FLAG purification followed by bottom-up mass spectrometry analyses to demonstrate N-glycosylation at Asn569, which indicated that the most C-terminal region (residues 411 to 707) of TMEM260 faces the ER lumen (SI Appendix, Fig. S2). We used CRISPR/Cas9 mediated KO of TMEM260 in HEK293 SimpleCells (HEK293SC/KO:TMEM260), and employed human HEK293SC with KO of COSMC and POMGNT1 that do not elongate O-glycans to enable sensitive enrichment of O-Man glycopeptides by Concanavalin A (ConA) lectin chromatography (11, 18). Mass spectrometry analysis of total cell extracts from HEK293SC and HEK293SC/KO:TMEM260 differentially labeled with stable isotopes revealed selective loss of O-Man glycopeptides derived from IPT domains (Fig. 2 and Dataset S1), including plexins (plexin A1-4, B1-2 and D1), hepatocyte growth factor receptor (cMET), and the macrophage-stimulating protein receptor (RON). Notably, KO of TMEM260 did not influence O-Man glycopeptides derived from αDG and cadherins that are dependent on POMTs and TMTCs, respectively, as previously shown (Fig. 2) (11, 18). This analysis was replicated with cell lysates of HEK293WT and HEK293KO:TMEM260 cells (Dataset S1), which corroborated selective loss of O-Man glycopeptides derived from IPT domains in HEK293KO:TMEM260. We further reintroduced TMEM260 by site-directed knock-in (KI) in the HEK293KO:TMEM260 cells, which resulted in full rescue of O-Man glycopeptides from IPT domains (Fig. 2D and Dataset S1). Finally, we speculated that Asp52 of TMEM260 serves important functions based on the structural alignment of D52 with the catalytically important D77 of yeast PMT1 (Fig. 1F). To test this hypothesis, we generated a D52A mutant and knocked-in the TMEM260 D52A variant in HEK293KO:TMEM260 cells. Western blot (WB) analyses revealed that expression of TMEM260 D52A was indistinguishable from TMEM260 WT upon KI in HEK293KO:TMEM260 cells (Fig. 2C). In contrast, differential glycoproteomic analyses revealed that TMEM260 D52A was unable to restore O-Man glycosylation on IPT domains (Fig. 2C). We also expressed a full-length cMET-3×FLAG fusion protein in HEK293WT and HEK293SC/KO:TMEM260 cells and used label-free quantification to demonstrate high O-Man occupancy at Thr582 (68%), Thr676 (100%), and Thr761 (44%) on β-strands in three IPT domains of the mature cMET β-chain (SI Appendix, Fig. S3). The O-Man glycans were sensitive to jack bean alpha-mannosidase digestion, confirming α-anomeric configuration of the linkage (Manα1-O-Ser/Thr) (SI Appendix, Fig. S3). We utilized the open search Mass spectrometry (MS) Fragger tool (40) for unbiased mining of the MS/MS data and found no evidence of elongated O-Man glycans (SI Appendix, Fig. S4), which indicates that the TMEM260-directed O-mannosylation pathway may not undergo elongation. This is similar to the TMTC directed O-mannosylation, while the POMT directed O-mannosylation of α-DG undergo complex modifications (41). Taken together, these results demonstrate that the TMEM260 gene encodes a protein O-mannosyltransferase that serves a highly select group of protein clients with IPT domains that is distinct from the clients served by the POMTs and TMTCs. The TMEM260 enzyme will therefore be classified as a new distinct GT117 family in the CAZy database (21).

The SHDRA syndrome caused by mutations in the TMEM260 gene (6, 7) was predicted to have relative high carrier frequency in select populations with ranges from 0.0007 to 0.007 across ancestries. To confirm that predicted deleterious mutations affected the O-Man glycosylation function, we analyzed two reported homozygous TMEM260 mutations, c.1393C>T; p.Q465* and c.1698_1701del; p.Y567Tfs*27 (7), and two novel homozygous variants c.293 G>A; p.C98Y and c.1357T>C; p. C453R identified in two affected families (Fig. 3A and SI Appendix, Text and Fig. S1). The affected children carrying C98Y and C453R variants share overlapping phenotypes with previously reported SHDRA cases, including truncus arteriosus (TA), kidney defects and global developmental delays (Fig. 3B and SI Appendix, Table S1). We utilized HEK293KO:TMEM260 cells and introduced disease variants by site-directed KI (Fig. 3C). Western blot analyses indicated that all TMEM260 variants expressed but were unstable in comparison to the wild-type protein (Fig. 3D). Furthermore, KI of these variants revealed markedly reduced ability to rescue and induce O-Man glycosylation on extracellular IPT domains in HEK293KO:TMEM260 cells (Fig. 3E), verifying that the glycosylation function of TMEM260 is impaired. Interestingly, the TMEM260 variants from the two newly identified families presenting milder phenotypes (SI Appendix, Table S1) and carrying homozygous C98Y or C453R substitutions (SI Appendix, Fig. S1 and Table S1), appeared to exhibit some degree of rescue of enzyme function, as evidenced by the differential O-Man glycoproteomic analysis (Fig. 3E).

We next investigated the endogenous cMET, RON, and Plexin-B2 receptors in human ovarian cancer BG1KO:TMEM260 cells by Western blot analysis and observed that maturation of the cMET receptor was not influenced by TMEM260 KO (Fig. 4). However, proteolytic maturation of both RON and Plexin-B2 was partly affected, with accumulation of the pro-forms in total cell lysates (Fig. 4A). Immunofluorescence cytology revealed that Plexin-B2 accumulated in ER with marked reduction in cell surface expression in BG1KO:TMEM260 cells (Fig. 4B). Flow cytometry analysis further confirmed that cell surface expression of cMET was unaffected by loss of TMEM260 O-mannosylation, while cell surface expression of RON and Plexin-B2 receptors were markedly reduced in BG1KO:TMEM260 cells (Fig. 4C). Plexin receptors are important for controlled cell division and normal growth in epithelial cells (42, 43). Considering the kidney anomalies found in patients with TMEM260 deficiencies (7), we turned to a three-dimensional (3D) spheroid model using mouse inner medullary duct epithelial cells (mIMCD-3) (44) and introduced KO of Tmem260 (SI Appendix, Tables S2 and S3). Differential glycoproteomic analyses of mIMCD-3KO:Tmem260 cells confirmed selective loss of O-Man glycosylation in IPT domains of cMET and plexin A1, B1, B2, and D1 receptors (Dataset S1). Using two independent clones each of mIMCD-3WT and mIMCD-3KO:Tmem260 cells, we found that mIMCD-3WT produced normal spheroids with properly formed lumina and cellular junctions based on apical ZO-1 expression and adherence junctions visualization by E-cadherin immunofluorescence staining, while mIMCD-3KO:Tmem260 cells produced spheroids with profound growth and architectural defects (Fig. 5). The mIMCD-3WT cell clones formed an average of 76% and 70% normal cysts, whereas only 20% and 11% of the mIMCD-3KO:Tmem260 clones formed normal cysts (Fig. 5B). The mIMCD-3KO:Tmem260 spheroids (average area of 2.7 mm2 and 2.9 mm2) were notably larger in size compared to mIMCD-3WT spheroids, which had an average area of 1.0 mm2 (Fig. 5C), and we further noted that mIMCD-3KO:Tmem260 spheroids showed increased number of nuclei per cyst, with an average 46 and 38 nuclei vs. 10 and 13 nuclei for mIMCD-3WT (Fig. 5D). These findings indicate that TMEM260-directed O-mannosylation of IPT domains is critical for receptor functions and normal epithelial morphogenesis.

Our study elucidated the function of the TMEM260 gene underlying the SHDRA syndrome and uncovered the genetic and biosynthetic basis for a third distinct type of protein O-mannosylation in higher eukaryotes. We identified TMEM260 as a single O-mannosyltransferase specifically serving IPT domains in cMET, RON and plexin receptors without obvious potential redundancy from paralogs. TMEM260 shares the GT-C fold and overall structure with the POMT and TMTC enzyme families without significant sequence similarities, and the three classes of O-mannosyltransferases serve highly distinct classes of protein clients. Identification of TMEM260 as a glycosyltransferase also establishes SHDRA as a new member of the CDGs (1). The TMEM260 gene is conserved in metazoans with distant homologs in bacteria, and the TMEM260-directed O-mannosylation pathway expands the known types of distinct protein glycosylation pathways in mammals to a total of 14 (2, 4). Protein O-mannosylation was originally considered restricted to yeast with a family of seven PMTs that collectively serve a wide set of protein clients (8). A detailed view of the yeast O-glycoproteome revealed that O-Man glycans are widely distributed on many different classes of proteins and positions (45), which much resemble how higher eukaryotes decorate proteins with GalNAc-type O-glycans (2). Interestingly, the two POMT1/2 orthologs of the yeast PMTs appear to serve a quite restricted set of clients, including α-DG and KIAA1549, where O-Man glycosites are positioned in dense mucin-like regions (4, 11, 17, 18). Since these substrate regions are largely unstructured, it is yet unclear how the POMTs drive such high protein client selectivity, but studies have suggested that upstream protein regions may serve as recognition of substrates (46). In contrast, the TMEM260 as well as TMTC1-4 have evolved clear protein domain fold-directed acceptor substrate specificities to serve highly distinct classes of proteins and biological functions (4, 11). Many other types of protein glycosylation, including O-glucose, O-fucose, O-GlcNAc, and C-Man, are directed by isoenzymes having similar selectivity for specific protein domains (2, 47). The same is clearly not the case for N-glycosylation and the GalNAc-type O-glycosylation that are widely found on proteins and different in different positions and protein folds (2). Our analysis of disease-causing TMEM260 mutations clearly confirmed that loss of O-mannosylation underlies the complex phenotypic traits observed in the SHDRA syndrome (6). Moreover, previous studies in zebrafish shows that tmem260 is required for development in early metazoans (7). Biallelic deleterious nonsense/frameshift mutations in TMEM260 often lead to early childhood deaths, and patients that survive the first few years of life display neurological phenotypes with global developmental delays (6). We presented two new patients carrying homozygous missense mutations (C98Y and C453R) that are not predicted to affect the catalytic function, but rather appear to influence the stability of TMEM260 (Fig. 3D). These mutant enzymes induced partial rescue of O-mannosylation of IPT-domains in our KO cell model (Fig. 3E), which may explain the milder phenotype observed in the two affected individuals. Both were diagnosed with TA at birth and have now survived past the age of 7. They display varying degrees of cognitive impairment with one (C98Y) being nonverbal with autism and the other (C453R) capable of reading/writing and managing mainstream schooling. Interestingly, the rare truncus arteriosus heart defect consistently observed in individuals with bi-allelic TMEM260 mutations, is also a prominent phenotype in plexinD1−/− mice (48) suggesting that O-mannosylation is critical for Plexin-D1 function and related to the heart malformations observed in SHDRA (6). Glycosylation serves wide roles in folding, transport, and fine-tuning of protein functions (2), and most types of protein glycosylation initiated in the ER affect protein folding and/or stability and transport from ER (49–52). The O-mannosylation initiated in ER by POMTs and TMTCs have so far not been demonstrated to affect folding and transport of client proteins (11, 18). O-mannosylation of α-DG is clearly critical for subsequent building of the matriglycan chain that mediates laminin binding essential for the dystrophin glycoprotein complex (53), but our knowledge of molecular functions of the TMTC and TMEM260 directed types of O-mannosylation is still highly limited and so far these only involve single mannose glycans that are not elongated or modified. Here, we did find that loss of TMEM260-directed O-mannosylation impaired maturation and ER exit of select receptors (Plexin-B2 and RON) (Fig. 4). These observations are reminiscent of the secretion defects and reduced cell-surface expression of receptors that undergo O-Glc and O-Fuc glycosylation by POGLUT1/2 and POFUT1/2, respectively (50, 54). Our use of a 3D spheroid model further support functional roles for O-mannosylation of plexins (and cMET) in epithelial morphogenesis (Fig. 5) but further studies are clearly needed to understand how loss of these single O-Man glycans on specific protein domains have so wide biological effects. The EC domains (55) served by TMTCs and the IPT domains are characterized by varying numbers of antiparallel β-strands assembled into a β-sandwich frame (56, 57). While EC-domains are O-mannosylated by TMTCs on B- and G-strands (11), only the B-strands of IPT domains are O-mannosylated by TMEM260, which may indicate a common role for O-Man in guidance of cadherin and plexin cis-interactions at the cell surface. In conclusion, we provide evidence that TMEM260 encodes a novel protein O-mannosyltransferase dedicated to IPT domains in select receptor proteins, which serves critical functions during development.

BG1 cells were cultured in RPMI 1640 medium (Sigma) supplemented with 10% Fetal Bovine Serum (FBS) (Gibco) and 1% GlutaMAX (Gibco). HEK293 cells were cultured in DMEM medium (Gibco) supplemented with 10% FBS (Gibco) and 1% GlutaMAX (Gibco). mIMCD-3 cells (25) were cultured in DMEM/F12 medium (Gibco) supplemented with 10% FBS (Gibco) and 1% GlutaMAX (Gibco).

CRISPR/Cas9 targeted gene KO TMEM260 cell lines were generated as previously reported (11). Briefly, cells were seeded in 6-well plates, transfected the following day with 1.5 µg of PBKS-Cas9-2A-eGFP plasmid (Addgene #68371) and 1 µg gRNA plasmid using Lipofectamine 3000. eGFP positive cells were fluorescence-activated cell sorting (FACS)-enriched 24 h after transfection and single-cell sorted after 1 wk. KO clones were identified by Indel Detection by Amplicon Analysis (11) and validated by sanger sequencing (SI Appendix, Tables S2–S4). Site-directed KI of human TMEM260 full coding cDNA into the human AAVS1 safe harbor site was performed using a modified ObLiGaRe gene KI strategy as previously published (58). The donor plasmid pAAVS1 Insulator ObLiGaRe/EPB71 was designed and synthesized by Genewiz, USA (Addgene #90018), and contains CMV-gene of interest-polyA. The full-length TMEM260-3×FLAG WT sequence of donor plasmid was commercially synthesized and cloned in the donor plasmid (Genewiz, USA), and mutations were introduced by site-directed plasmid mutagenesis. Mutations Q465* and Y567Tfs*27 were generated using InFusion DNA polymerase (TaKaRa) and C98Y and C453R using KOD DNA polymerase (Sigma-Aldrich), followed by DpnI (NEB) digestion of donor plasmid and transformation in Stellar competent cells (TaKaRa). Plasmids encoding TMEM260(Q465*)-3×FLAG and TMEM260(Y567Tfs*27)-3×FLAG were generated from the donor constructs using primers in SI Appendix, Table S4. All plasmids were confirmed by Sanger Sequencing and a total of seven TMEM260 variant constructs were used for stable cell lines generation. HEK293 TMEM260 KO cells were used as parental strain to generate TMEM260 variant KI cell lines. In brief, 3 µg of donor plasmid and 1.5 µg of each AAVS1 CompoZr Zinc-finger nuclease (ZFN) plasmids (Sigma) encoding ZFN1/2-2A-GFP/E2-Crimson (58) were co-transfected in TMEM260 KO cells using Lipofectamine 3000 (Thermo) according to the manufacturer’s protocol. One day after transfection, cells expressing both GFP and E2-Crimson were enriched by FACS (Sony Cell Sorter SH800S), and after 1 wk the bulk populations were single-cell sorted in 96-well plates. Correct stable genomic integration was confirmed using junction PCR of A) the region between the 5′ of AAVS1 genomic locus upstream of the ZFNs cutting site and the 3′ end of the donor construct B) the region between the 5′ end of the donor construct and the 3′ of AAVS1 genomic locus downstream of the ZFNs cutting site.

Sample preparation was performed as previously described (11) with minor modifications. Approximately 0.5 mL packed cell pellets were lysed in 1 mL 0.1% RapiGest SF Surfactant (Waters) in 50 mM ammonium bicarbonate by incubation on ice and probe sonication. Extracts were cleared by centrifugation (1,000 g, 10 min), reduced (10 mM dithiothreitol, 37 °C, 30 min) and alkylated (25 mM iodoacetamide, RT, 30 min) before digestion with 25 µg trypsin at 37 °C, 16 h (Roche). Digests were labeled with stable dimethyl isotopes using light or medium dimethyl labels on C18 Sep-Pak columns (Waters), and mixed (1:1 ratio), digested by PNGase F (Sigma, 16U in 150 µL 100 mM Tris, pH 7.4, 37 °C, 16 h), and subjected to ConA lectin weak affinity chromatography (LWAC).

HEK293 total cell extracts were prepared using S-Trap midi columns (Protifi) with minor modifications to protocol from vendor. Briefly, 0.2 mL packed cells were lysed by probe sonication in 500 µL 5% Sodium dodecyl sulfate (SDS), 100 mM ammonium bicarbonate, cleared by centrifugation (13,000 g, 10 min), reduced (20 mM Dithiothreitol (DTT), 80 °C, 10 min), and alkylated (100 mM iodoacetamide, 30 min, room temperature (RT)). Phosphoric acid was added [final 1.2% (v/v)] and mixed with 3.3 mL 100 mM ammonium bicarbonate in 90% methanol before applied to a S-Trap column. Trypsin (25 µg in 350 µL 100 mM ammonium bicarbonate) was gently applied (37 °C, 18 h), and tryptic peptides eluted by centrifugation [4,000 g, 1 min including a second 50% acetonitrile (ACN), 0.2% formic acid (FA) elution]. Eluates were evaporated, desalted (Sep-Pak C18 100 mg), Speedvac dried, and labeled with stable diethyl isotopes in light (acetaldehyde) or heavy channels (13CH313CHO, Cambridge Isotope Laboratories), as previously described (59). Samples were mixed (1:1 ratio), digested with PNGase F, and subjected to ConA LWAC.

Proteins were extracted and digested with trypsin in 0.1% RapiGest, 50 mM ammonium bicarbonate as described above. Digest were desalted (Sep-Pak C18), quantified (Nanodrop 205 nm), and 500 µg peptides (each channel) reconstituted (50 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), pH 8.0), and labeled with 800 µg tandem mass tags (6-plex TMT kit, Thermo). miMCD-3 WT clone #1 and clone #2 were labeled with TMT-126 and TMT-127, respectively. miMCD-3 Tmem260 KO clone #1 was labeled with TMT-128 and TMT-129. miMCD-3 Tmem260 KO clone #2 was labeled with TMT-130 and TMT-131. Labeling efficiency and mixing was evaluated by mass spectrometry. The final 6-plex–labeled tryptic peptide mixture (3 mg) was digested with PNGase F as described above and subjected to ConA LWAC.

Digests diluted to 1.5 to 2 mL (25 mM Tris, pH 7.4, 300 mM NaCl, 0.5 mM MgCl2, 0.5 mM MnCl2) were loaded onto a ConA-agarose column (3 m, 100 µL/min) (Vector Laboratories), washed until UV 210 nm absorbance was <5 mAU in same buffer, and eluted with 0.5 M methyl-α-D-mannopyranoside. Fractions were desalted by in-house packed Stage tips (Empore disk-C18, 3M).

Mass spectrometric analysis of O-Man glycopeptides was carried out as previously described (11). Briefly, desalted samples were individually injected using a EASY-nLC 1000 system (Thermo Fisher Scientific) interfaced via a nanoSpray Flex ion source to an Fusion Tribrid or Fusion Tribrid Lumos mass spectrometer (Thermo Fisher Scientific). The EASY-nLC 1000 was operated using a single analytical column setup (PicoFrit Emitters, 75 μm inner diameter; New Objectives, Woburn, MA) packed in-house with Reprosil-Pure-AQ C18 phase (1.9-μm particle size; Dr. Maisch) at 200 nL/min with gradient step-wise elution using solvent A (0.1% FA) and solvent B (ACN, 0.1% FA) and 2 to 20% B (95 min), 20 to 80% B (10 min), and 80% B (15 min). Precursor MS1 scan (m/z 355 to 1,700) was acquired in the Orbitrap at a resolution setting of 120,000, followed by Orbitrap Higher-energy dissociation (HCD)-MS/MS and electron-transfer/collision-induced dissociation (ETciD)-MS/MS of multiply charged precursors (z = 2 to 6) in the MS1 spectrum; a minimum MS1 signal threshold of 10,000 to 50,000 ions was used for triggering data-dependent fragmentation events; MS2 spectra were acquired at a resolution of 60,000 (HCD and EtciD). For bottom-up analyses of in-gel digests, gradient elution was achieved using 2 to 25% (65 min), 25 to 80% B (10 min), and 80% B (15 min). Data analyses was carried out using Proteome Discoverer 1.4 software (Thermo Fisher Scientific). Data files (.raw) were processed using the Sequest HT or MS Amanda nodes and searched against the canonical human proteome downloaded (January 2013) from the UniProtKB database (http://www.uniprot.org/). Precursor mass tolerance was set to 10 ppm (Sequest HT) or 5 ppm (MS Amanda) and fragment ion mass tolerance to 0.02 Da. Up to 2 missed trypsin (full- and semi-specific) cleavages were allowed. Carbamidomethylation (cysteine: 57.02146 Da), dimethyl (light: 28.0313 Da, medium: 32.0564 Da), diethyl (light: 56.0626 Da, heavy: 60.07602 Da) and TMT (229.1629 Da) modifications of peptide N-termini and lysines set as fixed modifications. Oxidation (methionine: 15.9949 Da) was set as variable modification. In addition, Hex (162.0528 Da) was set as variable modification for serine and threonine residues. Peptide confidence levels were calculated using the Target Decoy PSM Validator node and results were filtered for high-confidence (P < 0.01) identifications only. All spectral matches were inspected manually for validation. Open search was performed via the Fragpipe (v17.1) user interface with MS fragger (v3.4) installed (40). The “glyco-O-open-hybrid” method was used; 162.0528 Da and 324.1056 Da masses were included in the Mass Offsets and Y ion masses lists (MS Fragger). The mass spectrometry proteomics data have been deposited to the ProteomeXchange Consortium via the PRIDE (60) partner repository with the dataset identifier PXD032328. The data may be accessed using the following; username: reviewer_pxd032328@ebi.ac.uk and password: 7pPu1FvF.

Mouse M2 antibody (1 mg, F1804-5MG, Sigma-Aldrich) was buffer exchanged (0.1 M sodium phosphate, pH 7.4) using Zebaspin columns (2 mL, Thermo Fischer Scientific), diluted with 500 µL 3M ammonium sulfate in 0.1 M sodium phosphate (pH 7.4), mixed with 50 mg of pre-washed Dynabeads M-270 epoxy (#14302D, Thermo Fisher Scientific), and incubated with rotation at 30 °C (20 to 24 h). Beads were washed (4 × 2 mL Phosphate buffered saline (PBS), 2 × 2 mL 0.5% Triton X-100 in PBS, 2 × 2 mL PBS), and stored in 10% glycerol, 0.02% NaN3, in PBS up to 3 mo at 4 °C.

Cell pellets (300 µL) were lysed in 600 µL (50 mM HEPES, pH 7.4, 50 mM K-Acetate, 200 mM NaCl, 2 mM MgCl2, 1 mM DTT, 0.1% Tween-20, 0.5% Triton X-100), cleared (21,000 g, 10 min, 4 °C), mixed with 60 µL M2-Dynabeads slurry, and incubated with rotation at 4 °C, 45 min. Beads were washed with 3 × 12 mL (50 mM HEPES, pH 7.4, 50 mM K-Acetate, 200 mM NaCl, 2 mM MgCl2, 1 mM DTT, 0.1% Tween-20), and eluted (2×) by incubation in 25 µL with 1 µg/µL FLAG peptide (#F4799, Sigma-Aldrich) (10 min shaking, RT). The eluate was reduced (10 mM DTT, 37 °C), treated with 1 µL PNGase F (on, 37 °C), diluted in NuPAGE lithium dodecyl-sulfate (LDS) sample buffer, heated (72 °C, 10 min), and separated on a 4 to 12% BIS-Tris sodium dodecyl-sulfate polyacrylamide gel electrophoresis (SDS-PAGE) gel in 3-(N-morpholino) propanesulfonic acid (MOPS) running buffer, and stained with Imperial protein stain (#24615, Thermo Fisher Scientific). Bands migrating at as predicted for pro-MET and mature cMETβ were excised, destained by 3 × 200 µL (50% ACN in 100 mM ammonium bicarbonate, 15 min, 37 °C), followed by addition of 100% ACN, 10 min, RT, to dehydrate. Gel pieces were soaked in 50 µL 5% ACN, 100 mM ammonium bicarbonate containing 0.5 µg trypsin (Roche) at 37 °C ON before addition of 25 µL 100% ACN, 0.1% FA. Extraction was repeated with 2 × 50 µL 50% ACN, and peptides dried (speedvac) and labeled by diethyl stable isotope. Samples immunoprecipitated from HEK293WT were labeled with heavy diethyl (13CH313CHO), while samples from HEK293SC/KO:TMEM260 were labeled with light diethyl (CH3CHO) reagents. Samples were stage-tipped, mixed in equal amounts and subjected to liquid chromatography (LC)-MS/MS. Treatment with α1-2,3,6-mannosidase (Jack bean, New England Biolab) was performed on immunoprecipitated cMET from HEK293WT cells. In-gel trypsin digest of bands corresponding to cMETβ were split in two and diethyl labeled with heavy diethyl isotopes (mock treatment) and light diethyl isotopes (mannosidase treated). The samples were combined, stage-tipped, and subjected to LC-MS/MS.

Cells were lysed on ice in buffer (50 mM HEPES pH 7.4, 150mM NaCl, 1% Triton X-100) with 1× complete™ Protease Inhibitor Cocktail (Merck) for 10 min. Lysates were cleared by centrifugation (10 min, 4 °C) and protein amounts were adjusted after BCA assay (#23225, Thermo Scientific), and equal extracts separated on a Bis-Tris 4 to 12% 10-well Mini Protein Gel (Invitrogen) using MOPS or 2-(N-morpholino)ethanesulfonic acid (MES) running buffers. Transfer was performed for 11 min at 20 V to a nitrocellulose membrane using iBlot™ 2 Gel Transfer Device (Thermo Fisher Scientific). Membranes were blocked in a 5% solution of Skim Milk in Tris Buffered Saline with Tween (TBS-T) buffer for 2 h or 30 min. Antibodies used for blotting were diluted according to SI Appendix, Table S5. For all blots, primary antibodies were incubated overnight at 4 °C while secondary antibodies were incubated for 1 h at RT with gentle agitation. Blots were developed using SuperSignal™ West Pico PLUS Chemiluminescent Substrate (Thermo Scientific) and imaged by ImageQuant™ LAS 4000 (GE Healthcare). 1× ReBlot Plus Strong Antibody Stripping Solution (#2504, Merck Millipore) was used for stripping before β-Actin reprobing of membranes. Densitometry analysis was performed using the Gel Analyser function in ImageJ and expressed by quantifying pro- and mature form intensities over the total intensity (pro-form + mature). Statistical analysis was performed by one-way ANOVA, Tukey test.

BG1 WT and TMEM260 KO cells were grown to 80% confluence, on 10 mm-diameter cover slides, in 24 well plates. The cells were washed once in PBS and fixed with 4% paraformaldehyde for 10 min at RT and quenched with 100 mM NH4Cl in PBS for 10 min. Cells were permeabilized with 0.05% Saponin in PBS with 2 mg/mL bovine serum albumin for 10 min. Cells were stained with primary antibody diluted in permeabilization buffer for 1 h at RT (SI Appendix, Table S5). Cells were washed and incubated for 1 h with fluorophore-conjugated secondary antibodies (SI Appendix, Table S5) and 4',6-diamidino-2-phenylindole (DAPI). Coverslips were mounted on Mowiol (Calbiochem). Image acquisition was performed using LSM710 (Carl Zeiss) confocal microscope. BG1 WT cells were transiently transfected with TMEM260-3×FLAG, using jetPRIME (PolyPlus) according to manufacturer’s protocol. Cells were cultured on 10 mm-diameter cover slides, in 24 well plates. 50 µL of jetOPTIMUS buffer along with 0.5 µg DNA and 1 µL of jetOPTIMUS reagent were mixed and incubated for 10 min before addition to the cells. Media were changed 6 h after transfection and cells were incubated for 24 h before fixation and staining as described above.

Family I: A trio whole-exome sequencing was performed on family I using samples from the affected child and both parents. Using genomic DNA from the proband and parents, the exonic regions and flanking splice junctions of the genome were captured using the IDT xGen Exome Research Panel v1.0 (Integrated DNA Technologies, Coralville, IA). Massively parallel (NextGen) sequencing was done on an Illumina system with 100 bp or greater paired-end reads. Reads were aligned to human genome build GRCh37/UCSC hg19, and analyzed for sequence variants using a custom-developed analysis tool. Additional sequencing technology and variant interpretation protocol has been previously described (61). The general assertion criteria for variant classification are publicly available on the GeneDx ClinVar submission page http://www.ncbi.nlm.nih.gov/clinvar/submitters/26957/). A homozygous variant in TMEM260 (c.293 G>A p C98Y) was identified. No other variants were reported. Additional genetic testing included Array CGH, testing for the FMR1 expanded CGG repeat, were normal. Family II: Whole-exome sequencing was performed using SureSelect Human All Exon V6 (Agilent Technologies) using DNA samples extracted from peripheral blood from the affected individual, an unaffected sibling and both parents. Libraries were sequenced on an Illumina HiSeq 3000 instrument with 6 samples pooled per lane. Reads were aligned to GRCh37 reference genome using Novoalign (Novocraft Technologies, Selangor, Malaysia). Data analysis was performed assuming a homozygous mutation as the most likely cause of disease. Genome Analysis Toolkit HaplotypeCaller was used for variant calling and Ensembl Variant Effect Predictor for variant annotation. Copy number variation (CNV) analysis of WES data was performed using the Exome Depth program in the affected individual. A homozygous TMEM260 variant; 14:57088379T>C, p.C453R, was identified in the affected child and was heterozygous in both parents. In silico tools were supportive of pathogenicity and the CADD score was 25.7. The variant was rare in the Gnomad database (2 of 249,420 alleles, no homozygotes). Two other rare homozygous variants were recognized in the affected individual (in SULT4A1 and C2Orf88), neither of which could be linked to the phenotype. Segregation of the variant identified in TMEM260 was confirmed using conventional Sanger sequencing as homozygous in the affected individual, and heterozygous in both parents and an unaffected sibling.

Click here for additional data file. Click here for additional data file.