Abstract
Ubiquitin ligases (E3s) embedded in the endoplasmic reticulum (ER) membrane regulate essential cellular activities including protein quality control, calcium flux, and sterol homeostasis. At least 25 different, transmembrane domain (TMD)-containing E3s are predicted to be ER-localised, but for most their organisation and cellular roles remain poorly defined. Using a comparative proteomic workflow, we mapped over 450 protein-protein interactions for 21 different stably expressed, full-length E3s. Bioinformatic analysis linked ER-E3s and their interactors to multiple homeostatic, regulatory, and metabolic pathways. Among these were four membrane-embedded interactors of RNF26, a polytopic E3 whose abundance is auto-regulated by ubiquitin-proteasome dependent degradation. RNF26 co-assembles with TMEM43, ENDOD1, TMEM33 and TMED1 to form a complex capable of modulating innate immune signalling through the cGAS-STING pathway. This RNF26 complex represents a new modulatory axis of STING and innate immune signalling at the ER membrane. Collectively, these data reveal the broad scope of regulation and differential functionalities mediated by ER-E3s for both membrane-tethered and cytoplasmic processes.
Introduction
The endoplasmic reticulum (ER) is the largest membrane-bound organelle in eukaryotic cells, comprised of a complex network of sheets, tubules, junctions and contact sites that can occupy more than 35% of the entire cell volume1 and a significant fraction of the total membrane surface area. The continuous lattice it forms with the nuclear envelope (NE) makes extensive and dynamic contacts through distal projections with mitochondria2, peroxisomes3, endosomes4,5, plasma membrane6 and lipid droplets (reviewed in7). Within this extensive network, the ER accommodates biogenic, metabolic and regulatory multi-subunit transmembrane domain (TMD)-containing protein complexes that span the lipid bilayer and simultaneously carry out processes essential for cellular homeostasis.
Post-translational modification by ubiquitin (Ub) targets proteins for degradation, promotes interactions, directs subcellular localisation, or drives signalling8. The enzymatic cascade conjugating and extending Ub chains on proteins throughout the mammalian cell uses one (or more) of the > 600 Ub ligases (E3s) to provide reaction specificity by bringing substrates and Ub conjugating enzymes (E2s) in proximity9. E3s distinguish substrates either directly through dedicated binding domains/surfaces, or indirectly by assembling co-factors into specialised multi-subunit complexes with recognition and recruitment capabilities (reviewed in10). Within aqueous environments of the cytoplasm and nucleus, freely diffusing E3s access substrates with reduced spatial impediment. In contrast, E3s embedded within lipid bilayers by virtue of one or more TMDs or lipid anchor, have pre-determined orientation and lateral motion restricted to the planar membrane where they reside. Eukaryotic membrane-embedded E3s are found in the inner nuclear membrane (INM)11, ER12, mitochondria9, Golgi13, endosomes and plasma membrane14. The RING and HECT domains of E3s coordinating Ub transfer are exclusively exposed to the cytoplasm (and nucleus), enabling the access of cytosolic, nuclear, and proximal membrane proteins. Moreover, lumenal proteins from within the ER also reach the RING domain of at least one membrane-bound E3 (Hrd1), by way of an aqueous channel it forms within the lipid bilayer15. Thus, membrane bound E3s arguably serve as broad platforms for ubiquitination within the cell. Understanding how E3 complexes recognise substrates in and around membranes, how they can coordinate efficient Ub conjugation, how they regulate access to them, and which cellular processes they regulate, are important biological questions that remain outstanding.
ER-associated degradation (ERAD) has been the principal modality for understanding E3 function in the ER. Secretory cargo transiently or terminally misfolded during biogenesis in the ER is prone to aggregation that can cause proteotoxic stress, necessitating removal from the organelle that ultimately ends in its degradation12,16,17. ERAD serves in tandem with chaperone-mediated folding and assembly processes, acting as an integral facet of the organelle’s quality control machinery18. How we envisage E3 function at the ER has been shaped by extensive studies on the evolutionarily conserved Hrd119-26. With as many as eight TMDs15, a cytoplasmic RING domain and an extended C-terminus with low complexity27, Hrd1 scaffolds specialised lumenal, integral membrane, and cytosolic co-factors such as the AAA-ATPase VCP/p9728,29 to form multi-component complexes that coordinate recognition, retrotranslocation, and ubiquitination of misfolded secretory cargo20,24.
E3 complexes not only control the quality of secretory cargo but also adjust the abundance (and hence activity) of ER-resident membrane proteins through “regulatory ERAD”30. As the primary site for phospholipid and sterol biosynthesis31, degradation of ER-resident ratelimiting enzymes such as HMGCoA reductase (HMGCR) by gp78/AMFR32, RNF139/Trc833 and RNF14534,35, and squalene monooxygenase (SM) by MARCH6/Doa1036,37, help tune the output of this pathway to maintain homeostasis. Both gp78 and RNF145 use Insig1/2 as adaptors to recruit and degrade HMGCR32,34. Other processes at the ER membrane regulated by E3s and their interactors/adaptors include calcium flux38, innate immune signalling39, antigen presentation40, endosome diffusion41, ER morphology42 and apoptotic signalling43. E3 complexes represent important post-translational regulatory modules tending to the protein landscape of the mammalian ER, but most have not been extensively characterised. We developed a comparative proteomic workflow to define interactions of ER-resident E3s that sought to identify networks involved in maintaining ER and/or cellular homeostasis. Among the E3 interactors confidently identified were proteins previously reported in connection with lipid regulation, calcium flux, quality control, as well as new interactors involved in innate immune signalling.
Results
Isolation and discovery of ER-resident E3 interactors
Of the > 600 E3s present in the human proteome, about 10% contain TMDs that tether affiliated processes to lipid bilayers. Starting from previous reports44-46 and topology predictions, we shortlisted 25 E3s demonstrated (or predicted) to reside in the ER membrane. Selected ER-resident E3s (ER-E3s) are topologically and structurally disparate, arise from different phylogenetic lineages, but commonly possess one or more TMDs and a cytosolic RING-domain (Fig. 1a). With a design to determine cognate ER-E3 co-factors and substrates, we developed a generalised expression protocol coupled to a comparative immunoprecipitation liquid chromatography tandem mass spectrometry (IP-LC-MS/MS) workflow that enriched for proteins interacting at the ER. We generated 25 stable, individual, HEK293 cell lines, each expressing a FLAG-HA (FH)-tagged E3 inducible by doxycycline (DOX) using Flp-In™ recombination (see Methods, Fig. 1b). Positioning of the FH-tag at either E3 terminus was determined by taking into account prior experimental observations44, predicted topology, and proximity to RING domains or other prominent structural features (Fig. 1a, Extended Data Table 1). Variable induction parameters sought to produce comparable E3 expression levels from each cell line (Extended Data Fig. 1a). More than 80% of ER-E3s tested (21/25) could be detected from whole cell lysates (WCL), accumulating in a DOX-dependent manner that reflects relative stability (Extended Data Fig. 1a). Including MG132 along with DOX enhanced detection of some E3s (e.g. RNF26, BFAR, TMEM129), indicative of intrinsic instability and constitutive turnover by proteasome-mediated degradation. E3s colocalised exclusively (or partly) with the ER-markers calnexin or KDEL (Extended Data Fig. 1b), consistent with their expected residency in the ER.
To assess whether tagged ER-E3s faithfully reproduce endogenous complexes, we compared the interaction profile of Hrd124,26,27,47-49 with that of DOX-induced Hrd1-FH. Cofactors including SEL1L, FAM8A1, OS-9, Herp, and UBE2J1 were comparably coimmunoprecipitated (co-IPed) by both Hrd1 and Hrd1-FH (Fig. 1c). Moreover, velocity sedimentation revealed that Hrd1-FH complexes migrate in fractions that overlap with those formed by endogenous Hrd1 (Extended Data Fig. 1c). From this, we anticipate that bona fide protein-protein interactions of candidate E3s will be recapitulated by the exogenously expressed E3s.
Preserving native interactions between TMD-containing E3s and co-factors (or substrates) during sample processing was essential to ensure robust detection by liquid chromatography tandem mass spectrometry (LC-MS/MS). Cells were solubilised using LMNG (Lauryl Mannose Neopentyl Glycol)-containing buffer, a detergent shown previously to preserve labile ER-E3 complex interactions27. Immunoprecipitated E3-interactor complexes were washed and subsequently eluted from beads non-selectively by SDS to obtain the sample complexity necessary for subsequent comparative analyses (see below). All samples were prepared and processed for LC-MS/MS in parallel to facilitate comparative analysis (Fig. 1d, see details in Materials and Methods). ~1,600 individual protein groups were detected within each sample. In total, > 2,000 unique proteins were identified (Extended Data Table 2). To distinguish each E3’s most relevant interactors, we adapted the Bait-Specific Control Group (BSCG) method described previously for multiple sample processing50 (Fig. 1e, Extended Data Methods). This method defines the set of commonly detected interactors common as the “background proteome” and normalises each sample to it to facilitate multi-sample comparison. By permitting relative fold-change and p-values to be determined for each identified protein, normalisation enables an enrichment to be assessed. To enrich for factors with relevant change, each putative E3 interactor was individually evaluated for: (1) p-value <0.05; (2) positive fold-change; and (3) >1 unique peptide (Fig. 1d). Commonly identified contaminants (Extended Data Table 3) were excluded and removed from the dataset. Proteins meeting all criteria were designated as “high-confidence candidate interacting proteins” (HCIPs)51.
An inherent limitation of BSCG analysis is that candidate interactors detected in only one E3 sample evade classification as HCIPs because a relative fold-change cannot be calculated. Since exclusive ER-E3 interactors represent high-value candidates, we also analysed raw data by calculating the semi-quantitative spectral index quantitation (SINQ) score for each sample (as described in52). Filtering for sample-exclusive, high SINQ score proteins in each sample identified an additional 28 interactors co-precipitating uniquely by specific ER-E3 (Extended Data Table 4). Merging the modified BSCG and SINQ analyses revealed over 428 interactions with 21 E3s (Extended Data Fig. 1d), the majority of which are unreported. This composite dataset represents a systematic attempt to define E3 complexes at the ER membrane.
Interaction landscape of ER-resident E3s
From the 21 E3 ligases we comparatively examined, 218 different HCIPs that formed 400 interactions were identified (Extended Data Table 5). Hierarchical clustering of individual E3 interactomes (Fig. 2a) revealed that HCIPs were both exclusive to and shared between ER-E3s. Visualising HCIP networks for E3s individually (Extended Data Fig. 2) revealed interactors brought down with varying degrees of confidence and linked to a diverse range of activities, as exemplified by the networks for both Hrd1 (Fig. 2b) and RNF185 (Fig. 2c). While ER-E3 raw abundance differed markedly, this did not correlate with the number of HCIPs identified (Fig. 2d). Therefore, the number of interactions detected was a function of intrinsic E3 properties and not simply expression level. Importantly, over 50% of HCIPs (112/218) were significantly enriched by just one ER-E3 while those remaining associated with 2 (67/218), 3 (19/218) or > 4 (20/218) different ligases (Fig. 2e). HCIPs enriched by only one ER-E3 might represent specific cofactors or cognate substrates whereas interaction with multiple E3 could reflect proteins with generalised or adaptable functionality. We excluded HCIPs identified for RNF145, RNF150, Trim59 and RNF13 because bait peptide counts were insufficient (<2) or interactors failed to be enriched.
To concentrate subsequent analyses on interactions made at the ER, we searched the HCIP dataset for predicted protein features/domains associated with organelle targeting or residency, such as TMDs, signal sequences, N-linked glycosylation or disulphide bonds (UniProt). Approximately two-thirds of HCIPs (139/218) contained features consistent with localisation to the endomembrane system, validating our enrichment strategy for ER-associated proteins (Fig. 2f). Originality of E3-HCIP interactions is reflected by their underrepresentation in current protein-protein interaction resources (e.g. BioGRID 3.0), where only ~7% of E3-HCIP interactions have been reported previously53 (Fig. 2g). Unsurprisingly, most of these reported interactions (18/31) were with either of the two extensively characterised ER-E3s, Hrd1 and gp78/AMFR. This consistency provides additional assurance that the workflow could identify bona fide ER-E3 interactors.
ER-E3s, ERAD, and ER stress
E3 functionality at the ER is most commonly associated with the quality control (QC) process of ERAD (reviewed in12,17). We investigated whether any ER-E3s enriched for HCIPs implicated previously in ERAD, such as those that comprise the Hrd1 complex (e.g. SEL1L, UBE2J1, OS-9)24,26. While all were associated with Hrd1 (Fig. 2b), they were not HCIPs prominently enriched by other ER-E3s (Fig. 3a). Established gp78/AMFR interactors including UBE2G2, Derlin1/DERL1, and UBAC224,54, also did not feature among HCIPs of other E3s (Extended Data Fig. 2a, Fig. 3a). Thus, factors previously linked with ERAD do not appear to have generalised functionality that is readily adopted by other ER-E3s, except for VCP/p97 (discussed below).
One way the unfolded protein response (UPR) resolves ER stress is by coordinated upregulation of Hrd1 (and its cofactors) to increase ERAD capacity55. We investigated whether other ER-E3s respond similarly to ER stress by quantitatively monitoring transcriptional changes in ER-E3s and factors linked to ERQC in HEK293 cells treated with Tunicamycin (Tm) or the AB5 family bacterial toxin Subtilase cytotoxin (SubAB5) toxin56 (Extended Data Table 6). CGRRF1, RNF13, RNF170 and RNFT1 transcript levels increased with acute ER stress (~2 fold) as reported previously57, along with RNF5 (Tm only) and RNF139 (SubAB5 only) (Fig. 3b). When compared to the ~6-fold change (for Tm) observed for Hrd1, however, any responsive contribution made by other E3s to ER stress resolution may be nominal. Of the 226 HCIPs identified, 24 are among the 278 targets of the UPR transcription factors XBP1, ATF6, and ATF4 collated previously58, with a quarter (6/24) represented by the Hrd1 complex alone (Extended Data Table 7). Moreover, ER homeostatic maintenance did not require any individual ER-E3 since siRNA-mediated knockdowns of endogenous isoforms were not sufficient to induce the splicing of XBP1 (Extended Data Fig. 3a), consistent with findings from CRISPRi screens for ER stress induction59. Taken together, these findings are consistent with unique positioning of Hrd1 among E3s to resolve proteotoxic ER stress60.
Recruitment of VCP/p97 to ER E3 complexes
Binding and hydrolysis of ATP enables VCP/p97 to generate the force necessary to extract polypeptides from the ER membrane during ERAD29,61. VCP/p97 enrichment could therefore reflect a need for substrate extraction and a role in ERAD, so we searched for reported ER-E3s and HCIPs interactions with the AAA ATPase. VCP/p97 is recruited to protein complexes throughout the cell by factors that contain a UBX (Ub-regulatory X) domain or by a linear sequences such as the SHP motif (also known as binding site 1, BS1), the VCP-interacting motif (VIM), or the VCP-binding motif (VBM)62. These domains/motifs can be found in ER-E3s63,64, their cofactors (e.g. Derlins, VIMP)65,66 or ERAD-related enzymes (e.g RHBDL4)67. Surprisingly, only two ER-related HCIPs containing confirmed VCP-binding domains were found. FAF2/UBXD8 containing a UBX domain48 and Derlin1 containing a SHP domain65, were both associated with gp78/AMFR (Extended Data Fig. 2a, Fig. 3a) as previously reported24.
Differentiating bona fide recruitment of VCP/p97 from non-specific binding can be problematic in an IP-LC-MS/MS workflow because it is highly abundant, involved in a broad range of cellular processes, and can interact non-specifically. To address this, we compared VCP/p97 spectral counts determined for each E3 IP to assess whether any enriched the AAA ATPase. Among the highest were those for gp78/AMFR and Hrd1, in line with their established recruitment for ERAD, but RNF185, MARCH4 and RNF5 were also among the upper quartile (Fig. 3c). The soluble VCP/p97 cofactors UFD1 and NPLOC4 were also HCIPs of RNF185 (discussed below), which lends additional support for this E3 in an ERAD-related role68. RNF185 was recently highlighted among the set of prominent candidates identified in the VCP/p97 interactome69. Despite this, neither RNF185 nor its HCIPs contained canonical VCP/p97 binding domains/motifs that might justify recruitment. An unidentified factor or the presence of a non-canonical (or cryptic) VCP/p97 binding motif70 could be responsible, but this remains undetermined.
Ubiquitin-related HCIPs and ER-E3s
ER-E3 HCIPs included factors implicated in Ub conjugation and binding, as well as those containing a Ub-like (UBL) domain. However, other than the two well-characterised E2-E3 pairs UBE2G2-gp78/AMFR and UBE2J1-Hrd1, our isolation protocol does not appear to have robustly preserved these interactions. As almost all E2s are soluble and bind with moderate or weak affinity to E3 RING domains during Ub transfer, their scarcity was not unexpected. The Ub precursor, ribosomal fusion protein RSP27A, was enriched by both RNF185 and gp78/AMFR, reflecting either its binding to or direct modification of the E3s. Hrd1 and RNF185 enriched UBL domain-containing proteins (and homologues) Herp/Herp2 (HERPUD1/HERPUD2) and TMUB1/TMUB2, respectively (figs. 2c, 3a). Both have been linked to ERAD; TMUB1 links Erlin1 to gp7871 while Herp binds to FAM8A127 and activates Hrd1 through a cytoplasmic domain72. Deubiquitinating enzymes (DUBs) were not among E3 HCIPs, which may not be surprising as the DUB interactome did not report interaction with any ER-E3s51. Some ER-E3s did co-precipitate (e.g. RNF185-RNF170, RNF185-RNF5), and may reflect organisation consistent with coordinated or sequential ubiquitination as part of ERAD73,74 or alternatively, an E3-substrate relationship.
ER-E3s and calcium-related HCIPs
Signalling from G-protein-coupled receptors (GPCRs) activates inositol 1,4,5-triphosphate (IP3)-receptors (IP3Rs), causing this calcium-gated Ca2+ channel to be ubiquitinated and turned over from the ER membrane75 by RNF170 and its cofactors Erlin1/Erlin238,76. We identified enrichment of the ERLIN1/2 heterodimer by RNF170 and RNF185, and Erlin1 alone by RNF5 and gp78 (Figs. 2c, 3a, Extended Data Fig. 2). RNF170 is an HCIP of both RNF185 and gp78, which suggests Erlin1/2 interactions could serve as a bridge for larger heterooligomeric E3 complexes. IP3R was only enriched by RNF170, in line with its previous identification as a cognate substrate and demonstrating that this methodology can also identify bona fide E3 substrates. Consistent with a larger hetero-oligomer, RNF170 and RNF185 share other HCIPs including the putative secreted factor c6orf120 and TMBIM6/BI-1/Bax-inhibitor 1 (Extended Data Fig. 3b). These form a Ca2+ leak channel in the ER that protects cells from ER stress77 by regulating Ca2+ release and interacting with TMBIM3/GRINA (see78). RNF185 enriched for TMUB1/TMUB2 and TMEM259/Membralin (Fig. 2c), a polytopic ER protein linked to motor neuron survival79 that appears to have been erroneously assigned as part of the Hrd1-gp78 ERAD network80. RNF185 interactions were validated by co-expression and pulldown with S-tagged HCIPs (Extended Data Fig. 3c). Interestingly, the Ca2+-load-activated Ca2+ channel TMCO1, which prevents overfilling in the ER81, was also enriched by RNF170. Collectively, RNF170 and RNF185 appear to associate with proteins linked to homeostatic maintenance of ER Ca2+ levels related to ER stress and apoptosis.
E3s and lipid-related HCIPs
Sterols and fatty acids are produced through coordinated biosynthetic reactions at the ER membrane. 3-hydroxy-3-methylglutaryl-CoA-reductase (HMGCR) and squalene epoxidase (SQLE), are among the best examples of rate-limiting enzymes degraded by ER-E3s through negative feedback to regulate biosynthetic activity. 25 different HCIPs were involved in the biosynthesis and regulation of cholesterol, fatty acids, or phospholipids, with 80% annotated as TMD-containing proteins residing in the ER (Fig. 3d). Sterol and fatty acid-related HCIPs were enriched by 16 different E3s with nearly a quarter of all interactions (11/48) made with gp78/AMFR. Among the HCIPs of gp78/AMFR was HMGCR, previously reported as its substrate33. Conventionally, lipid-related HCIPs associating with E3s would do so as substrates for regulatory ERAD processes and may include DHCR7 (Hrd1), ACSL4 (RNF185) or SOAT1 (RNF5). Additionally, ZDHCC5 (RNF128) is a palmitoyl-acyltransferase which itself is palmitoylated82,83 and involved in endosome-Golgi trafficking84. Notably, it is one of the few palmitoyl transferases localised to the endosomal system, consistent with RNF128 (GRAIL) also reportedly present beyond the ER in endocytic compartments85.
RNF26 is unstable and degraded by the UPS
ER-E3s co-precipitated uncharacterised proteins with high specificity and abundance, which may reflect essential functionality at the ER. One example is the HCIPs and complexes formed by RNF26, an ER-E3 implicated previously in innate immune signalling86 and organisation of perinuclear endosomes41. RNF26 is integrated into the ER membrane through six predicted TMDs (Fig. 4a). Its canonical C3-H-C4-type RING domain lying near its C-terminus is evolutionarily conserved (Extended Data Fig. 4a) and shares sequence and positional similarity with the nuclear SUMO-targeted Ub ligase (StUbL) RNF487,88, Inhibitor of Apoptosis Proteins (IAPs,89), and MDM290 (Fig. 4b), all of which form homo-/heterodimers through their RING domains. Despite efforts and consistent with published data41, endogenous RNF26 protein could not be detected from whole cell lysates, even though relative mRNA levels in HEK293 cells were comparable to other E3s such as Hrd1 (Extended Data Fig. 4b). Consequently, we used DOX-induced expression of FH-RNF26WT to facilitate detection by immunoblotting, which was enhanced by treatment with MG132 (Fig. 4c, top panel, Extended Data Fig. 1a). Stabilisation by MG132 reflected intrinsic instability of RNF26 and suggested constitutive disposal by ERAD or an ERAD-like process. RNF4 conjugates Ub using a penultimate tyrosine (Y189) to engage the E2-Ub thioester91. This aromatic residue is conserved in RNF26 (Y432, Fig. 4a,b, Extended Data Fig. 4a), where a neutralising mutation (Y432A) stabilises FH-RNF26 protein levels and renders it insensitive to MG132 (Fig. 4c, middle panel). Radiolabel pulse-chase assays confirmed the increased stability of newly synthesised FH-RNF26Y432A compared to FH-RNF26WT (t1/2 ~ 2 hrs. vs. < 0.5 hrs, Fig. 4d). Expression of FH-RNF26Y432A also enabled endogenous RNF26 to be co-precipitated and detected (Fig. 4e), consistent with the formation of stable and inactive heterodimers.
Like other E3 dimers, the increased stability afforded by MG132 and the E2-Ub activating mutant (Y432A) is consistent with RNF26 auto-ubiquitination. Accordingly, immunoprecipitated FH-RNF26WT produced FLAG-immunoreactive ladders and high-molecular weight smears, enhanced by MG132 and collapsed by the non-selective DUB Usp21 (Fig. 4f). In vitro ubiquitination assays using immunopurified FH-RNF26 (WT or Y432A) and recombinant UbcH5a, faithfully recapitulated these observations (Extended Data Fig. 4c). Curiously however, immunoprecipitated FH-RNF26WT subjected to a panel of linkage specific DUBs indicated RNF26 modification by Ub chains not conventionally linked with degradation, including K33-, K63- and K29-linkages as well as multiple mono-ubiquitination (Extended Data Fig. 4d). Linkage diversity in RNF26 ubiquitination may indicate effects not only on turnover, but also on scaffolding and interaction with other proteins.
Discovery of RNF26 interactors that enrich with stable mutant
The intrinsic instability of RNF26WT may have led low abundance interactors to be below detection thresholds, and so IP-LC-MS/MS was also performed using FH-RNF26Y432A, which was then introduced into the BSCG and SINQ analyses (Fig. 5a). The inclusion of FH-RNF26Y432A increased the total number of high-confidence protein-protein interactions from 428 to 457. A comparison of RNF26WT and RNF26Y432A HCIPs revealed many of the same interactors, albeit now with more specific enrichment (i.e. higher p-values) (Fig. 5b, Extended Data Table 8). We prioritized TMD-containing and ER-related HCIPs for validation, selecting first among those enriched by both forms. Prominent HCIPs of both RNF26 forms included TMEM43/LUMA (Fig. 5b, Extended Data Fig. 5a), an evolutionarily ancient multi-spanning membrane protein present in both the ER and INM92,93, and ENDOD1 (endonuclease domain containing-1), an uncharacterised polytopic protein containing a putative DNA/RNA nonspecific nuclease domain (Extended Data Fig. 5a). RNF26Y432A also enriched for TMED1, an atypical member of the p24 cargo receptor family94, an ER membrane-shaping factor TMEM3395, as well as the ERAD and lipid droplet-related proteins UBXD8 and AUP1 (Fig. 5a,b, Extended Data Fig. 5a).
To validate interactions and gain insight into quaternary structures, S-tagged HCIPs were coexpressed with FH-RNF26WT and the resulting co-precipitation profiles compared. FH-RNF26 was reproducibly brought down by S-tagged TMEM43, TMED1, AUP1 and UBXD8, with the latter exhibiting a high molecular weight smear of RNF26 reminiscent of polyubiquitinated forms (Fig. 5c). For reasons that are not clear, exogenous expression of S-tagged ENDOD1 and TMEM33 proved difficult to detect, however S-tagged TMEM43 and TMED1 were able to bring down endogenous ENDOD1 and TMEM33 (TMED1 only), indicating that these HCIPs comprised larger macromolecular complexes. Neither ENDOD1 nor TMEM33 were co-precipitated by S-tagged AUP1 or UBXD8, suggesting the formation of heterogeneous complexes by RNF26. Velocity sedimentation of lysates from FH-RNF26 expressing cells revealed a profile of a complex migrating between ~200-300kDa (fractions 4-6) with endogenous TMEM43, TMED1, ENDOD1, TMEM33, UBXD8 and AUP1 co-sedimenting with FH-RNF26, albeit with varying degrees of overlap within fractions (Fig. 5d). The robustness of RNF26-HCIP interactions was assessed by performing IPs in Triton X-100 (TX-100) as well as the milder LMNG; detergents which differ in their ability to stabilise ER membrane protein complexes (27, Extended Data Fig. 5b,c). TMEM43 co-precipitated by FH-RNF26Y432A was independent of detergent conditions while other HCIP interactions were compromised to varying degrees in TX-100. These data support the formation of one or more heteromeric complexes containing RNF26 and HCIPs with a key interaction likely made through TMEM43.
To investigate organisation of RNF26 complexes in greater detail, validated siRNAs targeting HCIPs (Extended Data Figs. 5d-g) were introduced into FH-RNF26WT expressing cells and co-precipitation profiles monitored. Silencing TMEM43 disrupted the ability of FH-RNF26WT to coprecipitate other HCIPs, without substantially altering total cellular levels (Fig. 5e). A reduction in immunoprecipitated RNF26 (both unmodified and ubiquitinated) resulted from ENDOD1 knockdown, which might underlie the lower levels of other HCIP levels in RNF26 pulldowns, in particular TMED1. Although the pattern of HCIPs co-precipitated by RNF26 was unaffected by depletion of either TMEM33 or TMED1, detection of RNF26 and RNF26-Ubn forms was markedly enhanced by the loss of TMED1. These profile changes, together with the fact that the RNF26-TMEM43 interaction is resistant to solubilisation in TX100, indicate TMEM43 plays a key role in RNF26 complex formation, while ENDOD1 and TMED1 exert influence over RNF26 abundance.
We next asked whether HCIPs affected RNF26 stability (Fig. 4e). In cycloheximide (CHX) chase assays, siRNA-mediated depletion of TMEM43, TMEM33 or ENDOD1 did not markedly stabilise RNF26 nor did they alter turnover of immature CD147 (Fig. 5f), a well-characterised Hrd1-dependent ERAD substrate96. TMED1 knockdown slowed degradation of both RNF26 and CD147, suggesting it may impact an ER-resident process, such as trafficking, more generally. In contrast to RNF26, its HCIPs appeared stable and were unaffected by loss of their counterparts (Fig. 5f), even though in some cases they were no longer in a complex (Fig. 5e). Of note, VCP/p97 inhibition by NMS-873 only partially restored RNF26 while stabilising CD147 equivalently to MG132 (Fig. 5f, 4a). RNF26 was not among those E3s that enriched VCP/p97 (Fig. 3c), potentially distinguishing its mechanism of turnover from that of a canonical, misfolded ERAD substrate. Neither MG132 nor NMS-873 increased basal levels of RNF26 HCIPs, consistent with relative stability in the ER membrane. These data indicate RNF26 instability is intrinsic, unaffected by its interactions with more stable HCIPs.
RNF26 interactors regulate STING-dependent innate immune signalling
The immune signalling adaptor STING (STimulator of INterferon Genes, also known as MITA/TMEM173) is a polytopic ER-resident protein activated by the cyclic dinucleotide cGAMP, a product of cyclic GMP-AMP-synthase (cGAS) generated in response to the presence of cytosolic double-stranded DNA97 (Fig. 6a). Activation of dimerised STING by cGAMP triggers higher order oligomerisation in the ER98, subsequent efflux into ERGIC-derived vesicles99,100, and type I interferon (IFN) signalling through recruitment of TBK1 and IRF3, leading to its eventual turnover by p62/SQSTM-dependent autophagy101. In the ER, STING is a target for ubiquitination by multiple E3s including RNF2686,102,103, which are reported to modulate its IFN signalling capability through UPS-dependent degradation. STING and RNF26 reportedly interact through their TMDs and the absence of RNF26 (or its RING domain) counterintuitively increases STING turnover and attenuates IFN signalling86. As anticipated, endogenous STING co-precipitated with FH-RNF26 in an interaction reduced following activation by directly adding cGAMP (Fig. 6a), consistent with departure of STING from the ER that reduces its proximity to RNF26. MG132 stabilised RNF26 and consequently restored co-precipitation of STING even in the presence of cGAMP, although concurrent impairment to UPS-dependent degradation of the ER-resident population of STING might also be a contributing factor.
We next considered whether RNF26 HCIPs influenced STING and observed that like RNF26 deficiency, TMED1 siRNAs reduced STING protein levels by ~30% whereas those targeting TMEM43, ENDOD1 or TMEM33 had little to no impact (Fig. 6b,c). Directly introducing cGAMP activates STING and promotes its trafficking and subsequent degradation, reflected by reduced detection of STING protein (Fig. 6b,c). Reductions in STING abundance were proportional in all knockdowns (~50%), indicating that the influence of RNF26 and TMED1 did not depend on whether or not STING could be activated.
To ascertain whether RNF26 HCIPs also influence STING-mediated signalling in response to cGAMP, we monitored transcription of the interferon-stimulated gene (ISG) IFIT1 (interferon-induced protein with tetratricopeptide repeats 1) after silencing HCIPs alone and together with RNF26. STING activation by cGAMP increased IFIT1 transcription ~3-4 fold in WT cells (Fig. 6d,e) and did so in a dose-dependent manner (Extended Data Fig. 6a). RNF26 knockdown (Fig. 6d, Extended Data Fig. 6b) attenuated the increase in IFIT1, consistent with previous findings86. Expression of FH-RNF26Y432A also inhibited IFIT1 upregulation (Extended Data Fig. 6c), demonstrating that the IFIT1 response requires ubiquitination by RNF26 and the Y432A mutant functions as a dominant negative. Knockdown of either TMED1 or ENDOD1 also dampened the cGAMP-induced IFIT1 response (Fig. 6d), but lower STING levels only coincided with TMED1 loss and not ENDOD1 (Fig. 6b,c), suggesting these two HCIPs impact STING and signalling through it, by different mechanisms. In marked contrast, depleting cells of either TMEM43 or TMEM33 enhanced IFIT1 responses to cGAMP, resulting in 5-7-fold increases that were nearly twice the magnitude elicited from CTRL cells (Fig. 6d). These results were recapitulated with an independent set of siRNAs (Extended data Fig.6d). Codepleting RNF26 along with TMED1 or ENDOD1 did not attenuate IFIT1 responses further, and co-depleting with either TMEM43 or TMEM33 reduced the enhanced IFIT1 responses by ~50% (Fig. 6e), indicating that the impact of these HCIPs on STING activation was at least in part through RNF26. These data describe four novel modulators of STING-dependent immune signalling and define the RNF26 complex as an immunoregulatory unit.
Discussion
The ER accommodates a range of functional modalities; many of which fall under the regulatory remit of ubiquitination by resident E3-containing complexes. Comparative proteomic strategies have generated extensive and informative interaction networks of ubiquitination machinery51,104. We have applied those principals to expand the functional ERAD modules described for Hrd1 and gp78/AMFR24 by mapping over 450 interactions to form the landscapes for 21 ER-E3s. The ER membrane is increasingly appreciated as a diverse site of regulation for metabolic and homeostatic processes, which is reflected in the diversity of interactions made by resident E3s.
HCIPs represent cofactors and substrates of ER-E3s
ER-E3s assemble multi-subunit complexes from protein-protein interactions made through lipid bilayer, lumenal, and cytosolic contacts, making these units highly adaptable at conjugating Ub proximal to the membrane. Because ER-E3s appear structurally and topologically diverse, the minimal redundancy their HCIP networks exhibit (Fig. 2a) is consistent with assembling complexes from dedicated cofactors that would favour selective cellular responsibilities. More than 70% of HCIPs identified contain predicted TMDs or signatures associated with ER residency, among which will be both E3 cofactors and substrates. E3 cofactors are exemplified by Hrd1 interactors such as SEL1L and FAM8A1, but also by those validated for RNF26 (Fig. 5) and proposed for RNF185 (Extended Data Fig. 3b). Substrates are typified by the Wnt receptor Evi/WLS/GPR177, which after initially being identified early in this study was established as novel target for regulated ERAD by CGGRF1105. Other promising candidate substrates might include DHCR7, the terminal enzyme of cholesterol biosynthesis found with Hrd1 and a validated ERAD substrate106,107 and ACSL4, an RNF185 HCIP catalysing long chain polyunsaturated CoA synthesis108 which global turnover studies indicate has a short half-life109. SLC39A14/ZIP14, a stress-regulated Zn2+ and Mn2+ transporter at the plasma membrane, was identified as a Hrd1 HCIP. As it was detected under ER-stress free conditions, Hrd1 may be responsible for constitutive degradation of ZIP14 to maintain metal homeostasis. Like these, additional E3-substrate relationships are likely to be present within the dataset but will require detailed kinetic analyses to confirm them. Identifying and differentiating cognate cofactors from substrates of E3s remains a significant challenge in ubiquitin biology. Proteomics-based studies such as this, cannot readily distinguish E3 substrates from cofactors participating in substrate recruitment, complex assembly, localisation or ubiquitin conjugating activity. As the example of RNF26 HCIPs illustrates, biochemical, genetic and functional validation remains essential to accurately define complexes within each individual network. Although not providing comprehensive insight into all ER-E3 complexes that form, our analysis establishes a starting point from which to systematically elucidate their molecular organisation and functional responsibilities.
Ubiquitin linkages and machinery associated with ER-E3s
Ubiquitination at the ER is often envisaged in terms of conventional ERAD, where associated ubiquitin linkages (e.g. K48, K11) added to substrates by one (or more) ER-E3 facilitates recognition by ubiquitin-binding proteins associated with VCP/p97110 and proteasomes111. But our findings highlight the potential of some resident E3s to function differently. For example, RNF26 reportedly modifies STING with K11 chains86 but our data demonstrate its own turnover from the ER is rapid and is associated with K33-, K63- and/or K29-linked ubiquitin chains not conventionally associated with degradation or targeting to proteasomes. K29-linkages can be part of heterotypic branched and mixed Ub chains112, which can hallmark some ERAD substrates113. What role linkage heterogeneity plays in RNF26 turnover is not yet clear but might indicate linkages other than K48- and K11- also possess the ability to target proteins from the ER to proteasomes111.
E2s are determinants of poly-ubiquitin linkages while associated DUBs trim and prune them with linkage-specificity, yet most pairwise relationships with E3s remain undefined. As there are ~40 E2s and nearly ~100 DUBs in the human proteome, we would expect one (or more)to partner with each E3 to expand its substrate range. Envisioning ER-E3 prey profiles that enriched UPS components, we instead found that E2s and DUBs were not prominent among HCIPs. While key interactions for ERAD were confirmed (UBE2J1-Hrd124,48 and UBE2G2-gp78114), the often transient, low affinity E3-E2 and E3-DUB interactions appear not to have been widely preserved. Yeast-2-hybrid screens used to define human E2-E3 pairs have reported ER-E3s (e.g. RNF26, BFAR, RNF5) as interactors of multiple E2s from prey libraries115,116, but these have yet to be validated in vivo and were not identified as HCIPs. Similarly, functional roles in ERAD were identified for USP13, Atx3, VCPIP and USP19, but ER-E3s were not identified among the DUB interactome51. Detailed mapping of the ubiquitin linkage landscape attributable to individual ER-E3s and the UPS machinery required, remains an outstanding question for future studies.
The specialised role of Hrd1 among ER-E3s
ERAD is responsible for misfolded protein disposal during ER stress to restore organelle homeostasis. Induction of any ER-E3s by pro-survival UPR branches might have signalled complementary contributions to ERAD and ER stress resolution provided by the Hrd1 complex. Although ER stressors upregulated some ER-E3s (e.g. CGRRF1, RNF170, RNF5; Fig. 3b,57), their upregulation was modest relative to that observed for Hrd1. This is consistent with the observation that Hrd1 was the only E3 upregulated by direct activation of either the XBP1 or ATF6 transcription factors117. Moreover, there were only 24 HCIPs among known UPR target genes58 with at least a quarter belonging to the Hrd1 complex (Extended Data Table 7,24). Thus, evolutionary expansion of E3s in the mammalian ER has not extensively supplemented adaptive stress resolving capabilities functionally redundant with Hrd1, consistent with ubiquitination by the Hrd1 complex being essential for survival under proteotoxic ER stress conditions60,118. With misfolding of secreted rather than membrane proteins being the principal instigators of ER stress, an exclusivity of access to lumenal substrates via SEL1L offers an explanation why Hrd1 appears indispensable. Instead, other ER-E3s already known to oversee specific metabolic or regulatory functions consistent with higher order organismal functions, appear more likely to have evolved more selective client ranges of integral membrane proteins. ER-E3s may only hold strategic importance in selected cells or tissues, where a particular substrate/s are physiologically relevant. The enrichment of VCP/p97 only by only some ER-E3s (e.g. Hrd1, gp78, RNF185) supports a model where ERAD-like degradation is only one of the processes E3s oversee at this membrane interface.
Protein-conducting channels within ER-E3 complexes
An essential facet of ERAD is delivery of misfolded ER proteins to cytosolic proteasomes. Recent Cryo-EM structures containing recombinant yeast Hrd1p compellingly show the presence of an aqueous cavity formed by its TMDs that could provide a retrotranslocation conduit for misfolded polypeptide transport across the ER membrane15,119. With functionally demonstrated roles in ERAD of membrane proteins, other ER-E3s might be expected to form structurally analogous protein-conducting channels to dislodge substrate TMDs. Some ER-E3s (e.g. MARCH6, Trc8/RNF139) encode a sufficient number of TMDs from which such a channel could reasonably be formed from monomers, while others (e.g. RNF185) would require higher-order oligomers or large multi-subunit, TMD-containing HCIP/cofactor assemblies. If analogous cavities are formed, the absence of HCIPs common to ERAD-related E3s might indicate the evolution of specialised strategies to selectively remove TMD-containing substrates. So far only Hrd1 has been implicated in “retrotranslocation” of misfolded lumenal proteins, whereas multiple ER-E3s (e.g. MARCH6, Trc8/RNF139, TMEM129, gp78/AMFR) as well as Hrd1, appear sufficient to “dislocate” TMDs of integral membrane substrates33,37,46,47,120. Perhaps only the Hrd1 protein-conducting channel is optimised to accept lumenal substrates, thus providing a key function for ER stress resolution. The ER’s lipid bilayer presents a formidable barrier for downstream degradation machinery. With lateral access, membrane-bound substrates seem poised to access an ER-E3 conduit differently or engage proteases to cleave TMDs and release fragments. Perhaps this is why coordinated involvement of intramembrane proteases such as SPP or the rhomboid RHBDL4 in ERAD has always been conceptually appealing67,121. In all kingdoms of life, membrane-spanning protein complexes are capable of translocating polypeptides represent specialised, macromolecular apparatuses122. If and how ER-E3s mediate this process is a rich area for future exploration.
RNF26 is an intrinsically unstable ER-E3
We found that RNF26 auto-ubiquitination coincided with its rapid turnover from cells (Fig. 4c). Sequence similarity between the RNF26 and RNF4 C-termini, notably in a conserved penultimate Tyr residue required for Ub transfer (Fig. 4b), explains its recalcitrance to detection at the protein level. RNF4 homo-dimerisation allows activated E2 to bind one monomer while engaging the other to activate Ub-transfer to substrate. The Ub Ile44 hydrophobic patch binds Tyr189 at the RNF4 dimer interface, permitting the E2-Ub oxyester to be efficiently hydrolysed88. Evidence of dimerisation along with the observation that the Y432A mutant phenocopies ubiquitination defects like those observed when directly disrupting the RING domain or E2 binding (e.g. C401S, I382R41,86), suggests that RNF26 embraces a Ub conjugating mechanism like that of RNF4 and related E3s (e.g. IAPs).
STING regulation by RNF26
Chronic activation of STING is linked to autoimmune and auto-inflammatory disorders123, denoting an imperative for tight regulation and fine control over this signalling cascade. Ubiquitination offers complex spatiotemporal, post-translational control of STING abundance, activation and consequently, signalling. Multiple E3s, including RNF26, reportedly ubiquitinate STING while in the ER and after it is trafficked into endolysosomal vesicles following cGAMP-induced oligomerisation98. RNF26 knockdown lowered STING levels and dampened IFIT1 upregulation (Fig. 6b,c,d). Expression of FH-RNF26Y432A exerted the same effect on IFIT1 levels (Extended Data Fig. 6c), placing RNF26-mediated ubiquitination as a potent modulator of IFN signalling86. One model posits that RNF26 competes with RNF5 to extend K11-rather than K48-Ub chains on Lys150 of STING86, where dynamic balancing of ER-E3 ubiquitination reactions governs STING proclivity for degradation, based on affinities of Ub-binding proteasome subunits for different linkages. Lys150 lies in the connector helix loop just after the final TMD (TM4,98) and so would be accessible by different ER-E3s. Domain mapping demonstrated RNF26 and STING interact through TMDs86, but further investigation is required to clarify how RNF26 might ubiquitinate STING and not itself, perhaps by recruiting a substrate-selective cofactor (e.g. an HCIP) or by engaging different E2s. We found RNF26 capable of modifying itself with K63-, K33-, and/or K29-Ub chains and not K48- and K11- as shown previously for STING86, which distinguishes the ubiquitination reactions occurring in cis- and trans- by RNF26 and raises the potential involvement of different E2s.
An RNF26 complex scales the IFN response through STING
We identified four RNF26 interactors capable of scaling the IFN response through STING. These modulators were not found previously by RNF26 proteomics, which only used the RING-containing, cytoplasmic C-terminus as bait41. Together with RNF26, TMEM43, TMEM33, ENDOD1, and TMED1 form a membrane-bound complex that appears capable of influencing STING activation, most likely through its membrane-spanning TMDs. Although these RNF26 HCIPs do not appear structurally similar or functionally orthologous, they share the ability to scale IFN signalling through cGAS-STING pathway in an RNF26-dependent manner (Fig. 6e), suggesting they act collectively through one or more complexes. How each RNF26 HCIP accomplishes this is not yet clear, as their individual functions are not yet fully appreciated.
Loss of TMEM43 or TMEM33 increases the IFIT1 response (Fig. 6), consistent with roles as negative regulators of STING activation. As TMEM43 was required for RNF26 interaction with TMEM33 (Fig. 4e), enhanced signalling through STING could be explained by the loss of TMEM33 from the RNF26 complex. TMEM33 is the human ortholog of S. pombe Tts1, an ER-shaping protein that helps sustain high-curvature ER membranes124 and is involved in nuclear envelope remodelling during mitosis125. Functional conservation in metazoans could indicate that local ER membrane curvature is an important determinant of STING activation. Alternatively, its role in organising the peripheral ER and as a binding protein of reticulons could mean that TMEM33 influences localisation of RNF26 and STING throughout the ER/INM network, which then determines signalling capability and regulation. TMEM33 has been linked recently to regulation of intracellular calcium homeostasis through an interaction with polycystin 2 (PC2), suggesting its influence extends beyond its contribution to a complex that modulates STING or RNF26. TMEM43/LUMA localises in both the INM and the ER and interacts with proteins such as Lamin A/B and Emerin92. Mutations in TMEM43 are genetically linked to the heritable cardiomyopathy autosomal dominant arrhythmogenic right ventricular cardiomyopathy/dysplasia (ARVC/D, S358L)126 and the autosomal recessive myopathy Emery-Dreifuss Muscular Dystrophy (EDMD, Q85K, I91V,127). Whether these rare conditions are in some way attributable to the suppression of STING activation by TMEM43 mutants is not known. TMEM43 has been linked previously to immune signalling and NF-kB through an interaction with CARMA3/CARD10 and EGFR128, but not previously to STING.
Silencing TMED1 or ENDOD1 phenocopied the reduction of IFIT1 levels upon cGAMP treatment that is observed with loss of RNF26, consistent with both HCIPs functioning to either permit or enhance STING activation. TMED1/tp24 contains a GOLD domain and is part of the p24 family of trafficking proteins (TMED1-10). It forms monomers and dimers instead of hetero-oligomers with other p24 family members for function94. TMED1 may influence vesicle trafficking from the ER as a cargo receptor or it may be linked to vesicle coat formation, potentially placing it as a gatekeeper for budding ER vesicles containing proteins such as STING. Notably Kelch-like 12 (KLHL12), a CUL3 adaptor and protein linked to ER-Golgi trafficking through regulation of COPII vesicle size129, was an HCIP enriched by RNF26Y432A. ENDOD1 is predicted to contain a non-specific endonuclease domain shown to have nuclease activity in vitro using an orthologous domain from Paralichthys olivaceus (Japanese flounder)130. Importantly, this study identified ENDOD1 among the genes that are important for innate immunity in fish, suggesting a role that is evolutionarily conserved. It is not yet clear from our study whether ENDOD1 and STING interact directly or what may be the implications of having endonuclease activity near a hub for immune signalling.
Our data reveal that RNF26 nucleates an immuno-regulatory complex, which was discovered through constructing the interaction landscape of the ER-resident E3s. Information within this landscape provides a resource to uncover E3 functions within various cellular processes. Understanding the mechanisms modulating abundance and activity of its membrane-embedded proteins will help to determine if they represent tractable targets that may be leveraged for potential therapeutic benefit.
Methods & Materials
Plasmids and Transfections
All cDNAs encoding individual ER-E3s (Extended Data Table 9) were amplified and appended with restriction site-containing linkers by PCR, and subsequently subcloned into a pcDNA™/5/FRT/TO vector (Invitrogen) containing a FLAG-HA (FH) tag in frame (N- or C-terminal) by restriction digest and ligation. Sequences for HCIPs (Extended Data Table 10) were obtained and processed similarly but subcloned instead into pcDNA3.1(-) vectors containing either an N- or C-terminal S-tag in frame as described previously27. All plasmids were transfected into recipient cell lines using Lipofectamine™2000 (Thermo Fischer Scientific) according to manufacturer’s guidelines.
Cell culture and generation of stable cell lines
Flp-In™ T-REx™ human embryonic kidney 293 cells (Thermo Fischer Scientific, referred to as Flp-In™293) were used to generate stable cell lines individually expressing selected E3s. Briefly, each FH-E3-pcDNA5/FRT/TO construct was co-transfected with the Flp recombinase vector pOG44 (3:1 ratio) as described above. Cell pools stably recombining and expressing E3s were selected by resistance to Hygromycin B (100 μg/ml, InvivoGen). All Flp-In™293 cell lines were cultured in DMEM (Lonza, BE12-604F) supplemented with 10% (v/v) fetal bovine serum (FBS) and glutamine (2 mM). All cells were grown at 37°C and 5% CO2.
Antibodies and compounds
The following primary antibodies were used for detection by Western blot: anti-Hrd1 (Bethyl, #A302-946A), anti-SEL1L (Santa Cruz Biotechnology, #sc-48081), anti-UBE2J1 (Abcam, #ab39104), anti-OS-9 (kind gift from R. Kopito, Stanford), anti-Herp (Abcam, #ab150424,) anti-Derlin1 (kind gift from Y.Ye, NIH), anti-tubulin (Sigma, #T6074), anti-FLAG (Sigma, #F3165 and #F7425), anti-HA (Sigma, #H9658; Cell Signaling Technologies, #3724) antiubiquitin (Cell Signaling Technologies, #3933 and #P4D1), anti-S-tag (Thermo Scientific, #MA1-981), anti-TMEM43 (Abcam, #ab184164), anti-TMED1 (Abcam, #ab224411), anti-ENDOD1 (Abcam, ab121293), anti-AUP1 (Atlas Antibodies, #HPA007674), anti-UBXD8 (Proteintech, #16251-1-AP), anti-STING (Cell Signaling Technologies, #D2P2F), anti-CD147 (Santa Cruz Biotechnology, #sc-25273). Anti-FAM8A1 has been reported previously27. The secondary antibodies used for western blot and IF include: goat anti-rabbit HRP (1:10,000, BioRad), goat anti-mouse HRP (1:10,000, Santa Cruz Biotechnology), donkey anti-goat HRP (1:10,000, Santa Cruz Biotechnology), goat anti-rabbit-Alexa 488 (1:400, Life Technologies), goat anti-mouse-Alexa 568 (1:400, Life Technologies). The following compounds were used in this study; MG132 (10 μM, Merck Millipore), Tunicamycin (500 ng/ml, Sigma), NMS-873 (10 μM, Sigma), cycloheximide (100 μg/ml, Abcam), N-ethylmaleamide (NEM, Acros Organics), DAPI (Sigma), doxycycline (DOX, Sigma), dithiolthreitol (DTT, Sigma), iodoacetamide (IAA, Roche) and SubAB56.
siRNA transfections
Flp-In™293 cell lines seeded in 12-well plates were transfected with individual siRNAs (50 nM, Sigma, Extended Data Table 11) using Lipofectamine RNAiMax (Thermo Fischer Scientific) at a ratio of 1:4 according to the manufacturer’s instructions. Cells were expanded 24 h post-transfection and harvested following another incubation of 24 h.
Mass Spectrometry and Proteomic analysis
Each FH-E3-expressing Flp-In™293 cell line was seeded in 15 cm plates and treated with DOX (1-1000ng/ml, 18 h). Cells were harvested at ~80% confluence, washed and subsequently resuspended in solubilisation lysis buffer (SLB: 150 mM NaCl, 50 mM Tris-HCl pH7.4, 5 mM EDTA) containing 1% Lauryl Maltose Neopentyl Glycol (LMNG, Anatrace) and supplemented with cOmplete™ protease inhibitor cocktail (Roche). Lysates were clarified by centrifugation (20,000 x g, 30 min.) and pre-cleared using CL-4B Sepharose beads (50μL of 50:50 slurry, Pharmacia/GE). The resulting clarified whole cell lysate (WCL, 10 mg) was used as source material for immunoprecipitations with anti-FLAG agarose (M2, Sigma, A2220) for 2 h. Immunoprecipitated complexes were washed twice with SLB (without detergent), twice with TBS, and eluted by 2 x SDS + 10% β-Mercaptoethanol. Eluates were reduced by DTT, alkylated by IAA, and subject to double chloroform-methanol precipitation. Precipitated proteins were subject to tryptic digest prior to purification using C18 Sep-Pak cartridges (Waters). Purified peptides were analysed by LC-MS/MS with a tandem mass spectrometer (Q Exactive™ HF, Thermo Fischer Scientific) with an EASY-Spray™ C18 LC Column (2 μm, 100Å, 75 μm x 50 cm, Thermo Fischer Scientific) over a 63 min 2-35% acetonitrile gradient in 5% DMSO (v/v)/0.1% formic acid (v/v). The data were acquired with a resolution of 70,000 full-width half maximum at mass/charge 400 with lock mass enabled (445.120025 m/z), Top 15 precursor ion selection, Dynamic Exclusion of 27 s, and fragmentation performed in Higher-energy C-trap dissociation (HCD) mode with Normalized Collision Energy of 28. Samples were analysed twice to generate technical duplicates. Chromatogram alignment and peptide intensity were determined by Progenesis-QI (nonlinear Dynamics). Peptides were identified and matched using the SwissProt database. Each bait sample was assigned with more than 2,000 protein IDs. Assignment of p-values to identified proteins was accomplished by adapting the comparative BSCG method described previously50. High-confidence interacting proteins (HCIPs) were defined by an ability to meet four criteria: 1) p-value <0.05, 2) positive foldchange, 3) identified by >1 peptide, and 4) not classified as ‘common’ contaminants (Extended Data Table 3). SINQ analysis52 was also carried out to identify interactors unique to one E3 only, which would not be assigned a p-value with the comparative analytical method. Raw data have been deposited in the PRIDE database (submission ongoing).
Immunoprecipitation, SDS-PAGE and western blotting
Cells rinsed in phosphate-buffered saline (PBS) were mechanically lifted, harvested, and lysed in SLB + 1% LMNG or Triton X-100 (TX-100, Fischer Scientific), as described above. Lysates were clarified by centrifugation (17,000 x g, 30 min.) and pre-cleared using CL-4B Sepharose beads (50μL of 50:50 slurry, Pharmacia/GE), with subsequent affinity and immunopurifications carried out using the resulting lysates. Beads were washed thrice in SLB and subsequently resuspended in 2 x Laemmli buffer + 20 mM DTT after the final wash, separated by SDS-PAGE and transferred to PVDF membrane for western blotting. Western blots were performed by incubating membranes in PBST blocking buffer (PBS + 1% Tween-20 supplemented with 5% non-fat dry milk), with subsequent primary and secondary antibody incubations in PBST + 5% non-fat dry milk. Secondary antibodies conjugated with horseradish peroxidase (HRP) were used to detect proteins bound to primary antibodies for enhanced chemiluminescence (ECL) with images captured either on X-ray film (FujiFilm, SuperRX) or by CCD camera (Chemidoc, BioRad) for quantification.
Quantitative transcript analysis by NanoString
RNA from Flp-In™293 cells was extracted using the RNeasy kit (Qiagen) in accordance with the manufacturer’s instructions that included the genomic DNA digestion with DNaseI (Qiagen). Isolated RNA from each sample (150 ng) was hybridized to a Reporter CodeSet and Capture ProbeSet (10μL each) for a selected set of genes (Extended Data Table 6) by incubating in hybridization buffer (65°C, 18 h) and loaded in the nCounter® PrepStation according to manufacturer’s instructions. Hybridized probe/target complexes were immobilized on the nCounter® Cartridge and imaged in the nCounter® MAX Digital Analyzer (high-resolution setting). Data were processed and analysed according to the manufacturer’s guidelines (NanoString Technologies Inc.). All experiments were performed in biological triplicate (n=3).
Radiolabelling and pulse-chase
Radiolabel pulse-chase assays of Flp-In™293 cells stably expressing either FH-RNF26WT or FH-RNF26Y432A were carried out as previously described22,27. Briefly, following DOX treatment (18 h) cells were starved in DMEM (Lonza) lacking methionine (Met) and cysteine (Cys) + 10% dialysed FBS for 10 min, metabolically labelled by supplementing starvation medium with35S-Met/Cys (EXPRE35S35S Protein Labelling Mix (PerkinElmer), 80 μCi/6 cm plate) for 10 min, rinsed thrice in PBS, and chased for indicated time points in DMEM supplemented with Met and Cys (50 mM each). Cells were lysed in SLB containing 1% TX-100, and the detergentsoluble, post-nuclear lysates pre-cleared using CL-4B Sepharose beads followed by immunoprecipitation with anti-HA-antibody (12CA5) and Protein G agarose (Roche). Beadbound radiolabelled substrates were resuspended in 2 x Laemmli buffer (+20 mM DTT), separated by SDS-PAGE and imaged using a phosphoimager (BioRad).
cGAMP transfection for IFIT1 qRT-PCR
Flp-In™293 cells seeded in 24-wells were stimulated by transfecting 5 μg/ml cGAMP using Lipofectamine 2000 at a ratio of 1.25:1. Cells were harvested 6 h post-transfection and the extracted RNA (RNeasy, Qiagen) reverse transcribed to produce cDNA (QuantiTect, Qiagen) according to manufacturer’s instructions. Taqman probes targeting human GAPDH (Hs02758991_g1) and IFIT1 (Hs03027069_s1) were purchased from Life Technologies. qRT-PCR data were collected on a StepOnePlus™ Thermal Cycler (Thermo Fischer Scientific) and analysed by the ΔΔCt method, normalising IFIT1 levels to GAPDH. Averages and S.E.M. were determined from triplicate assays from at least three independent experiments (n=3).
Immuno- and affinity purification of ubiquitinated proteins
FH-RNF26WT and FH-RNF26Y432A -expressing Flp-In™293 cells were induced with DOX (18 h) and where indicated, samples were additionally treated with 10 μM MG132 for 2 h. Cell pellets were lysed in TUBE lysis buffer (20 mM sodium phosphate pH 7.5, 1% NP-40 (v/v), 2 mM EDTA, supplemented with cOmplete protease inhibitor cocktail (Roche), PhosSTOP (Roche), NEM (50 mM), and DTT (1 mM)). Lysates were centrifuged (as above) and the detergent-soluble fraction subsequently incubated overnight with 15 μL magnetic GST resin (Thermo Fischer Scientific) conjugated to 50 μg 1x UBA-His6 binder. Bound resin was washed thrice with TUBE lysis buffer and each sample split equally to accommodate control/untreated or USP21 deubiquitinase treatment.
Deubiquitination assays
Bead-bound material from UBA-His6 binder pulldowns (above) was resuspended in deubiquitinating-buffer (50 mM HEPES pH 7.5, 100 mM NaCl, 2 mM DTT, 1 mM MnCl2, 0.01 % Brij-35) without or with 0.5 μM USP21 (Ubiquigent). Samples were incubated (1 h, 30°C) in a thermoshaker (VWR, 750 rpm) and subsequently denatured by incubation (65°C, 20 min) with 2 x Laemmli buffer + DTT (20 mM) followed by separation on SDS-PAGE. For UbiCREST analysis131, FH-RNF26WT from stably expressing Flp-In™293 cells (4mg) was immunoprecipitated by anti-FLAG agarose, washed, divided and individually incubated with the panel of recombinant dUbs, according to the manufacturer’s instructions (Boston Biochem).
In vitro ubiquitination assay
FH-RNF26WT and FH-RNF26Y432A -expressing Flp-In™293 cells were treated and lysed in TUBE lysis buffer as described above. Resulting supernatants were incubated with anti-FLAG M2 magnetic beads (Sigma, 3 h at 4°C). Beads were washed thrice with TUBE lysis buffer, dividing samples in half prior to the last wash and then washed once with in vitro ubiquitination (IVU) buffer (50 mM Tris-HCl, pH 7.5, 2.5 mM MgCl2). Assays were carried out by resuspending beads in IVU buffer supplemented with; ubiquitin (10μM, Boston Biochem), E1 enzyme (150 nM, Enzo), UbcH5a (1 μM, Enzo), and DTT (0.5 mM), with or without ATP (4 mM, pH 8.0, Sigma) and incubated in a thermoshaker (15 min, 30°C, 750 rpm). Samples were denatured with 2 x Laemmli buffer + 20 mM DTT (65°C, 20 min).
Velocity sedimentation
Velocity sedimentation was carried out as previously described27. Briefly, DOX-induced Flp-In™293 cells (1μg, 18 hrs) were mechanically harvested and lysed in SLB containing 1% LMNG (as described above). Post-nuclear, pre-cleared WCLs (1 mg total) were layered onto either a continuous sucrose gradient (10-40% or 5-30%) prepared using a Gradient Master 108™ (BioComp). Sucrose was dissolved in a physiological salt solution (150 mM NaCl, 50 mM Tris-HCl pH 7.4, 5 mM EDTA, 1 mM PMSF) + 1% LMNG. Samples were centrifuged in an SW.41 rotor (OptimaTM L-100 XP, Beckman Coulter, Brea, CA) at 39,000 rpm for 16 h at 4°C. Thirteen fractions (940 μL each) were collected manually and proteins precipitated by addition of 190 μL 50% (v/v) trichloroacetic acid (TCA). Following acetone washes, precipitated proteins were resuspended in 2 x Laemmli buffer + 20 mM DTT and separated by SDS-PAGE. If necessary, samples were neutralised with 1 M Tris-HCl (pH 9). All samples were heated (10 min, 56°C) and separated by SDS-PAGE. Gel filtration standards (Gel Filtration Markers Kit, MWGF1000, Sigma Aldrich) were separated on similar gradients to estimate protein complex size and included: alcohol dehydrogenase (150 kDa), β-amylase (200 kDa), apoferritin (443 kDa) and thyroglobulin (663 kDa). Standards were processed as above and detected by Coomassie staining.
Immunofluorescence and microscopy
For detection of FH-E3s, Flp-In™293 cells were seeded onto 13 mm poly-L-lysine coated cover slips and induced with DOX (18 h). Cells were fixed with 4% paraformaldehyde (PFA, 20 min at room temperature (RT)), permeabilised with PBS containing 0.2% TX-100 for 5 min at RT and blocked with PBS containing 0.2% PBG (fish skin gelatin) for 30 min. Coverslips were incubated with 1° antibodies diluted in 0.2% PBG (1 h, RT), rinsed twice in PBS and incubated with fluorescent 2° antibodies (0.2% PBG in PBS, 1h, RT). Coverslips were incubated with DAPI (5μg/mL, 10 min, RT) and mounted using ProLong® Gold antifade reagent (Life Technologies). All images were captured on a Zeiss LSM710 confocal microscope and processed in Image J (NIH) and Photoshop (Adobe).
Statistical analysis
Statistical significance within NanoString data was determined using multiple t-tests (Holm-Sidak method, α = 0.05) that compared fold-change in E3 transcripts from untreated and Tunicamycin-treated (Tm, 500 ng/ml, 8 h) or SubAb5-treated (10ng/ml, 8h) samples. All other data (e.g. qPCR, protein quantification) were analysed using a two-tailed paired t-test comparing non-targeting control siRNA (siNTC) to each siRNA target. All statistical analyses were carried out and plotted using GraphPad Prism (Version 7.0). Detailed statistical information is available in Extended Data Table 12.
Bioinformatic analysis
Primary amino acid sequences for all E3s and HCIPs were obtained from UniProt (http://www.uniprot.org/), with common motifs annotated using Pfam (http://pfam.xfam.org/)132, TMDs predicted by TOPCONS (http://topcons.net/)133 and N-linked glycosylation sites predicted by NetNGlyc 1.0 (http://www.cbs.dtu.dk/services/NetNGlyc/). E3 interactions were compared against those previously reported in BioGRID (https://thebiogrid.org/).
Conflict of interest
The authors declare no conflict of interest.
Author contributions
E.F. and F.L. designed, performed and analysed experiments; R.F., M-L.T. and B.M.K. performed LC-MS/MS; P.D.C analysed LC-MS/MS raw data; M.G.H. provided reagents and designed cGAMP-STING experiments; A.W.P. and J.C.P provided reagents; K.B. performed in vitro ubiquitination experiments; J.C.C. designed, performed and analysed experiments and wrote the manuscript.
Extended Data Tables (in corresponding Excel File)
Extended Data Table 1: ER-resident E3 features and positioning of FLAG-HA epitopes
Extended Data Table 2: Unique protein signatures of ER-resident E3 interactors identified by LC-MS/MS and BSCG analysis
Extended Data Table 3: Common contaminants removed from BSCG analysis
Extended Data Table 4: Unique ER-resident E3 interactors identified by SINQ analysis of LC-MS/MS dataset
Extended Data Table 5: High-confidence candidate interacting proteins (HCIPs) of ER-resident E3s
Extended Data Table 6: NanoString quantitative transcriptomics of ER-resident E3s in response to ER stress
Extended Data Table 7: Targets of UPR transcription factors among ER-resident E3 HCIPs
Extended Data Table 8: BSCG analysis of RNF26Y432A compared to RNF26WT
Extended Data Table 9: MGC clones for ER-resident E3
Extended Data Table 10: MGC clones for HCIPs
Extended Data Table 11: siRNA sequences
Extended Data Table 12: Statistical analysis
Acknowledgements
We are grateful to Dr. Norbert Volkmar and Dr. Dönem Avci for critical discussions. We also thank Dr. Jan Rehwinkel for technical assistance. E.F. was supported by a fellowship from the Medical Research Council. P.D.C was supported by an EPSRC grant (nr EP/N034295/1) and by the Chinese Academy of Medical Sciences (CAMS) Innovation Fund for Medical Science (CIFMS), China (grant number: 2018-I2M-2-002), awarded to B.M.K. J.C.C. was supported by a grant from the Medical Research Council (MR/L001209/1) and by the Ludwig Institute for Cancer Research.
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