G2BR^AUP1 is required for ERAD

To assess the requirement of AUP1 in HRD1-dependent ERAD, we examined the degradation of a known ERAD-L HRD1 substrate, the NHK variant of α1ΑΤ, in HT1080 fibrosarcoma cells. The NHK variant of α1ΑΤ (hereafter referred to as NHK) contains a frameshift mutation that generates a nonsense codon, resulting in the translation of a truncated protein that is retained in the ER lumen and targeted for ERAD [35,51–53]. Consistent with the HRD1 pathway regulating NHK degradation [51], CRISPR/Cas-9 knockout (KO) of HRD1 in HT1080 cells (HRD1 KO) inhibited the degradation of NHK as assessed by cycloheximide (CHX) chase (Fig 1A).

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Fig 1. The G2BR of AUP1 is required for ERAD of NHK.

(A) HT1080 or cells in which HRD1 was knocked out by CRISPR/Cas-9 (HRD1 KO) were transfected with plasmid encoding carboxyl-terminally HA-tagged NHK. Cells were treated with CHX for the indicated times to inhibit protein synthesis and NHK-HA degradation monitored by western blot. GFP and HSP60 serve as transfection efficiency and gel loading controls, respectively. (B) HT1080 or cells in which either AUP1 or gp78 was knocked out (AUP1 KO or gp78 KO) were transfected with NHK-HA and their degradation assessed. (C) AUP1 KO cells were transfected with NHK-HA and empty vector or plasmid encoding carboxyl-terminally FLAG-tagged AUP1 and NHK degradation assessed. (D) Indicated cells were transfected with plasmid encoding carboxyl-terminally MYC-tagged INSIG-1 and assessed for INSIG-1 degradation. (E) Schematic representation of AUP1 in the ER membrane and deletion mutants used in subsequent experiments. Amino acid residues are numbered spanning each domain. (F) AUP1 KO cells were transfected with NHK-HA and plasmids encoding the indicated FLAG-tagged AUP1 WT or deletion mutants and assessed for NHK degradation. (G) HEK293 (Parental) or HEK293 AUP1 KO cells were assessed for degradation of NHK-HA. (H) HEK293 AUP1 KO cells were assessed for NHK-HA degradation as in (F). Markers represent apparent molecular weight in kDa. The data underlying this figure can be found in S1 Data. aa, amino acid; AUP1, ancient ubiquitous protein 1; CHX, cycloheximide; ER, endoplasmic reticulum; ERAD, endoplasmic reticulum–associated degradation; G2BR, UBE2G2 Binding Region; KO, knockout; NHK, Null Hong Kong; WT, wild type.

https://doi.org/10.1371/journal.pbio.3001474.g001

AUP1 was found to be associated with the HRD1 complex in mass spectrometry experiments [54]. However, published RNAi experiments had provided conflicting findings regarding a requirement for AUP1 in ERAD [35,39]. To establish whether, in our experimental system, AUP1 functions with HRD1 in the degradation of NHK, we utilized CRISPR/Cas-9 to generate an AUP1 KO in HT1080 cells (AUP1 KO). KO of AUP1 inhibited NHK degradation (Fig 1B), whereas a CRISPR/Cas-9 KO of the ERAD E3 gp78 had no effect on NHK turnover. Importantly, reexpression of full-length epitope-tagged AUP1 (AUP1-FLAG or wild type [WT]) restored NHK degradation in AUP1 KO cells (Fig 1C), ruling out possible clonal or off-target effects and definitively establishing a requirement for AUP1 in NHK turnover. These data confirm and extend previous findings on the effect of AUP1 loss in ERAD [35,40,41]. By contrast, degradation of INSIG-1, a known gp78 substrate [47,55], is dependent on gp78 and an intact G2BR^gp78 but was largely unaffected by loss of AUP1 (Fig 1D, S1A Fig). These findings establish the specificity of HRD1 and AUP1 in the targeting of NHK for degradation and also establish a role for the G2BR^gp78 in the degradation of an ERAD substrate other than gp78 itself.

To determine the requirements for the multiple cytosolic domains of AUP1 in NHK degradation, we generated deletion mutants of FLAG-tagged AUP1 for expression in cells (see schematic Fig 1E). These included deletions of either the CUE domain (aa 293 to 337; ΔCUE), the acyltransferase plus the CUE domains (aa 90 to 337; ΔATΔCUE), or 12 of 26 aa within a region that is homologous to the G2BR of gp78 (aa 386 to 397; ΔG2BR). Each variant was expressed in the AUP1 KO cells and assessed for its ability to restore NHK degradation, with reexpression of full-length AUP1-FLAG (WT) serving as a positive control (Fig 1F). The ΔCUE mutant of AUP1-FLAG fully restored NHK degradation in the AUP1 KO cells. Similarly, and despite its rapid degradation and substantially lower steady state level (see S1B Fig for long exposure), the ΔATΔCUE mutant also restored NHK degradation. Quantitative PCR (qPCR) confirmed that the reduced expression of this mutant is due to posttranscriptional effects (S1C Fig). In contrast to the other deletions, the ΔG2BR mutant failed to restore NHK degradation, demonstrating the singular importance of the G2BR in the degradation of this substrate. This requirement for NHK degradation was recapitulated in HEK293 AUP1 KO cells (HEK293 AUP1 KO; Fig 1G and 1H).

We next assessed whether the specific requirement for the G2BR^AUP1 applies to another soluble ERAD-L substrate. For this, we examined the turnover of a well-characterized truncated mutant of Ribophorin I (RI₃₃₂) [35, 56]. Based on knockdown experiments both SEL-1L, which is a component of the HRD1 ligase complex, and AUP1 have been implicated in degradation of RI₃₃₂ [35,57]. Consistent with the SEL-1L observations, HRD1 KO cells exhibited a dramatic stabilization of RI₃₃₂ (Fig 2A), which was reversed by reexpression of HRD1. As with NHK, RI₃₃₂ degradation required the G2BR^AUP1 but was unaffected by loss of the acyltransferase and CUE domains (Fig 2B). Similar results were found in HEK293 AUP1 KO cells (Fig 2C and 2D).

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Fig 2. The G2BR of AUP1 is required for ERAD of mutant ribophorin and GFP^u.

(A) HT1080 or HRD1 KO cells were transfected with plasmid encoding RI₃₃₂-MYC and with plasmid encoding HRD1 as indicated. Cells were subject to CHX chase and RI₃₃₂-MYC degradation monitored by western blot. GFP serves as a transfection efficiency control. (B) HT1080 or AUP1 KO cells were transfected with plasmid encoding RI₃₃₂-MYC and AUP1-FLAG WT or deletion mutants assessed for RI₃₃₂ degradation. Lower band (*) in MYC blot is nonspecific. (C, D) HEK293 or AUP1 KO cells were assessed for RI₃₃₂ turnover as in (B). (E) HT1080 or AUP1 KO cells were transfected with GFP^u plasmid, and its degradation assessed by emetine chase, which has previously been used to assess GFP^u turnover [40]. Luciferase and β-actin serve as transfection efficiency and gel loading controls, respectively. (F) HT1080 AUP1 KO cells were transfected with GFP^u and the indicated AUP1-FLAG WT or deletion mutants, and GFP^u degradation assessed as in (E). The data underlying this figure can be found in S1 Data. AUP1, ancient ubiquitous protein 1; CHX, cycloheximide; ERAD, endoplasmic reticulum–associated degradation; G2BR, UBE2G2 Binding Region; KO, knockout; WT, wild type.

https://doi.org/10.1371/journal.pbio.3001474.g002

To assess whether these requirements extend to substrates that engage the ERAD-C machinery, we followed the degradation of GFP^u, which is GFP rendered unstable in cells by fusion to the CL1 degron [45]. In human cells, degradation of CL1 fusion proteins has been reported to be dependent on AUP1 as well as on TRC8 and MARCH6 [40,41]. Loss of AUP1 prevented the rapid degradation of GFP^u (Fig 2E), which again did not require either the CUE or the acyltransferase domains (Fig 2F). GFP^u degradation was, however, completely abrogated by disruption of the G2BR^AUP1.

HRD1 is implicated in the basal degradation of 3-hydroxy-3-methylglutaryl CoA (HMG-CoA) reductase (HMGCR) [31]. In yeast, the targeting for degradation of the HMGCR ortholog, Hmg2p, by Hrd1p is an exemplar for the yeast ERAD-M pathway [8,58,59]. AUP1 has similarly been implicated in HMGCR degradation, although this has primarily been assessed in the context of its rapid degradation in response to high sterols [50], a process where at least four ERAD E3s have been reported to potentially play roles [22,55,60–62]. We have examined the basal degradation of endogenous HMGCR and find that it is also AUP1 dependent. As with the other substrates presented, HMGCR degradation uniquely required the G2BR^AUP1 (S1D and S1E Fig). Thus, of AUP1’s three identified cytosolic domains, only the G2BR^AUP1 is required for ERAD-L substrates, a presumed ERAD-M substrate, as well as a substrate that engages the ERAD-C machinery.

AUP1 binds specifically to UBE2G2

The G2BR^AUP1 sequence shares 41% identity to the G2BR of gp78 (G2BR^gp78), which binds to UBE2G2 but shows no discernable interaction with the most homologous mammalian E2, UBE2G1 [63]. Consistent with this, a GST fusion of AUP1 that includes the G2BR bound UBE2G2 but not UBE2G1 (Fig 3A). Importantly, UBE2J1/UBC6e, which is an ER-localized transmembrane E2 that has been implicated in NHK degradation and suggested to function with AUP1 and other components of the HRD1 ERAD-L machinery [39,54,64], also failed to show detectable binding in vitro. UBE2D2/UBCH5B and UBE2D3/UBCH5C [65], which are considered promiscuous E2s and are involved in a number of different cellular processes including ERAD [40], similarly showed no discernable binding to G2BR^AUP1.

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Fig 3. The AUP1–UBE2G2 interaction is required for ERAD.

(A) Equimolar amounts of GST-AUP1 (aa 292 to 410) or GST were immobilized on glutathione beads and binding to the indicated in vitro translated ³⁵S-labeled E2s was assessed. Left panel is 5% of input used for binding. E2s are referred to by their unique alpha-numeric designation (e.g., UBE2G1 and UBE2J1). (B) HT1080 or UBE2G2 KO cells were transfected with INSIG-1-MYC and degradation assessed by CHX chase. (C) The indicated cells were transfected with NHK-HA and empty vector, WT or catalytically inactive (C89S) N-terminally MYC-tagged UBE2G2, and NHK-HA degradation assessed. (D) Cells were transfected with NHK-MYC and HA-ubiquitin and ubiquitination of NHK assessed following proteasome inhibition and IP (right panel). Left panel is 10% of total cell lysate used for IP, indicating no discernable change in total ubiquitination. (E) Levels of endogenous HRD1 were monitored by CHX chase in the indicated cell types. (F) Ubiquitination of endogenous HRD1 was assessed following IP with HRD1 antiserum. Arrow in upper panel indicates migration of unmodified HRD1. Control immunoprecipitates using rabbit serum are shown for comparison. The data underlying this figure can be found in S1 Data. aa, amino acid; AUP1, ancient ubiquitous protein 1; CHX, cycloheximide; ERAD, endoplasmic reticulum–associated degradation; IP, immunoprecipitation; KO, knockout; WT, wild type.

https://doi.org/10.1371/journal.pbio.3001474.g003

Having established the requirement for G2BR^AUP1 and the specificity of the interaction between G2BR^AUP1 and UBE2G2, NHK degradation was assessed in HT1080-derived cells in which UBE2G2 expression was abrogated using CRISPR/Cas-9 (UBE2G2 KO). As expected, INSIG-1, which is a substrate of gp78 and UBE2G2, was stable in these cells (Fig 3B). NHK was also stable in UBE2G2 KO cells (Fig 3C). Reexpression of UBE2G2 restored NHK degradation, while a catalytically inactive form of UBE2G2 (C89S) did not. These results establish a requirement for active UBE2G2 in the degradation of this HRD1 substrate. A similar requirement for UBE2G2 was established for the HRD1 substrate RI₃₃₂ (S2A and S2B Fig).

We next assessed ubiquitination of NHK in cells lacking either AUP1 or UBE2G2. Loss of either protein resulted in a marked decrease in substrate ubiquitination (Fig 3D), consistent with reduced NHK turnover in both AUP1 and UBE2G2 KO cells. This diminution of ubiquitination is not due to a reduction in the levels of HRD1 (Fig 3E). If anything, the steady state level of this E3 is increased in UBE2G2 KO cells. This increase in HRD1 is consistent with the induction of an ER stress response in these cells, which lack a critical component of the ERAD machinery (S2C and S2D Fig). In yeast, there is evidence that ubiquitination of Hrd1p itself is critical to its role in substrate retrotranslocation and degradation [66–68]. We found that in HT1080 cells, ubiquitination of HRD1 is readily detectable and that there is no apparent alteration in this ubiquitination with loss of either AUP1 or UBE2G2 (Fig 3F, S2E and S2F Fig). Despite its robust ubiquitination, there is no evidence that HRD1 is degraded over a period of 6 hours (Fig 3E). This suggests that this AUP1 and UBE2G2-independent ubiquitination does not target HRD1 for degradation, whereas AUP1- and UBE2G2-mediated ubiquitination is required for substrate degradation.

Structural and biophysical characterization of UBE2G2:G2BR^AUP1

We have determined the crystal structure of UBE2G2 in complex with G2BR^AUP1 at 1.74-Å resolution (PDB entry 7LEW, Table 1). The UBE2G2:G2BR^AUP1 structure reveals UBE2G2 residues 2 to 96 and 107 to 165, G2BR^AUP1 residues 377 to 404, and 158 oxygen atoms of water molecules. Residue 1 and residues 97 to 106 of the extended loop characteristic of UBE2G1 and UBE2G2, as well as yeast Ubc7p, are disordered without observable electron density. The overall structure exhibits a high degree of similarity to the previously described structure of UBE2G2:G2BR^gp78 [17, 30] with a root–mean–square deviation (RMSD) of 0.5 Å for 167 out of 182 pairs of Cα atoms. This RMSD remains for 145 out of 154 pairs of UBE2G2 Cα atoms. Fig 4A depicts virtually identical ribbon depictions of the two G2BR helices resulting from the superposition of the two UBE2G2 Cα traces.

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Fig 4. Crystal structure of the UBE2G2:G2BR^AUP1 complex.

(A) Ribbon representation of the superimposed UBE2G2:G2BR^AUP1 (in cyan and magenta, this work) and UBE2G2:G2BR^gp78 (in orange, PDB entry 3H8K) complexes. The UBE2G2 in complex with G2BR^gp78 is not shown for high similarities between the two structures. The left and right views are related by a 180° rotation around the vertical axis. (B) Intermolecular hydrogen bonds and salt bridges between UBE2G2 and G2BR^AUP1 are shown as dotted lines. Involved aa residues are shown as stick models in atomic color scheme (N in blue, C in cyan or magenta, O in red, and S in orange). The left and right views are related by a 180° rotation around the vertical axis. (C) Contacts between UBE2G2 (in cyan) and G2BR^AUP1 (in magenta) are conserved in the UBE2G2:G2BR^gp78 (in orange) complex. The UBE2G2 residues involved in hydrogen bonds and/or salt bridges are indicated with a red star. Black lines link each G2BR residue to its reciprocal contact(s) in UBE2G2, and “x” denotes G2BR^AUP1 residues that do not contact UBE2G2 directly. (D) Calculated dissociation constants (K_d, in nM) of G2BR peptides for UBE2G2 from ITC, FP, or MST, experiments. Mean and standard deviations or 95% CI for at least two titrations are reported. aa, amino acid; CI, confidence interval; FP, fluorescence polarization; G2BR, UBE2G2 Binding Region; ITC, isothermal titration calorimetry; MST, microscale thermophoresis.

https://doi.org/10.1371/journal.pbio.3001474.g004

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Table 1. Data collection and refinement statistics^a.

https://doi.org/10.1371/journal.pbio.3001474.t001

The UBE2G2:G2BR complex is stabilized with two types of interactions across the interface. One type includes hydrogen bonds and salt bridges, electrostatic in nature (Fig 4B), and the other type includes van der Waals contacts, i.e., hydrophobic interactions. The high degree of similarity of the UBE2G2:G2BR^AUP1 and UBE2G2:G2BR^gp78 structures is the result of a largely conserved set of G2BR residues involved in direct contacts with UBE2G2 (Fig 4C). It is not surprising that this set of residues in the two G2BR sequences forms an almost identical pattern of interactions with the E2 molecule in the UBE2G2:G2BR^AUP1 and UBE2G2:G2BR^gp78 complexes.

To quantify the affinity between UBE2G2 and G2BR^AUP1, we performed isothermal titration calorimetry (ITC) measurements with purified UBE2G2 and either the G2BR^gp78 or the G2BR^AUP1 peptide. This resulted in measured K_d values of 2.7 ± 1.0 nM and 3.1 ± 2.0 nM, respectively (Fig 4D, S3A Fig), which is in accordance with the previously reported high-affinity interaction between UBE2G2 and G2BR^gp78 [17,30]. We also utilized both microscale thermophoresis (MST) and fluorescence polarization (FP) assays with purified UBE2G2 and fluorescently labeled G2BR^gp78 or G2BR^AUP1 peptides. Both methods confirmed the low nanomolar K_d values (Fig 4D, S3B and S3C Fig). Based on these results, we conclude that the affinity of G2BR^AUP1 for UBE2G2 is similar to that of G2BR^gp78.

The G2BR^AUP1 allosterically increases the affinity of UBE2G2 for the gp78 RING domain

Our previous studies of UBE2G2:G2BR^gp78 revealed a significant change in nuclear magnetic resonance (NMR) chemical shifts for the backbone ¹⁵N-¹H^N resonances of UBE2G2 upon binding of G2BR^gp78 [17,24]. The recognition that the backbone structure of UBE2G2 changes very little between the apo and G2BR^gp78 bound states suggested that the chemical shifts were due to variations in the hydrogen bond strengths that stabilize the core structure. Nevertheless, subtle changes in populations of dynamic conformers at the gp78 RING:UBE2G2 interface take place to promote the significant allosteric effect observed for binding of gp78 RING to UBE2G2 when G2BR^gp78 is bound [69]. We explored whether the same effects are present in the interaction with G2BR^AUP1. The ¹⁵N-¹H^N chemical shifts of UBE2G2 exhibit the same dramatic shifts upon binding G2BR^AUP1 as upon binding G2BR^gp78, suggesting similar allosteric effects (Fig 5A–5C, Biological Magnetic Resonance Data Bank deposition https://doi.org/10.13018/bmrbig33). Furthermore, consistent with the very similar crystal structures of UBE2G2:G2BR^gp78 and UBE2G2:G2BR^AUP1 that we report here, the UBE2G2 chemical shifts in the ¹⁵N-¹H^N HSQC spectrum showed a very similar shift from the apo UBE2G2 (Fig 5C).

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Fig 5. Titration of UBE2G2:G2BR with gp78 RING.

Superposition of ¹⁵N-¹H^N HSQC NMR spectra for apo-UBE2G2 and UBE2G2 bound to G2BR^AUP1 (A) or G2BR^gp78 (B). Spectra for UBE2G2 (150 μM) bound to G2BR were acquired at a 1:1.1 ratio. (C) Superposition of the spectra for apo-UBE2G2, UBE2G2:G2BR^AUP1, and UBE2G2:G2BR^gp78, illustrating the equivalence of perturbations between G2BR^AUP1 and G2BR^gp78. (D) Expansion of overlaid resonances corresponding to UBE2G2 residues showing fast exchange binding kinetics for gp78 RING binding to both UBE2G2:G2BR^AUP1 (left panels) and UBE2G2:G2BR^gp78 (right panels). Spectra were acquired at ratios of RING to UBE2G2:G2BR of 0, 0.33, 0.67, 1.0, 2.0, and 3.0. (E) Binding curves as a function of [RING]/[UBE2G2:G2BR] ratio and analyses for representative residues E12, Q15, L66, and V113 of UBE2G2:G2BR^gp78 (orange) and UBE2G2:G2BR^AUP1 (magenta) UBE2G2:G2BR binding to gp78 RING. Errors are reported from the regression analysis of the data in S1 Table (B, D). G2BR, UBE2G2 Binding Region; NMR, nuclear magnetic resonance.

https://doi.org/10.1371/journal.pbio.3001474.g005

We have previously found that the addition of G2BR^gp78 or U7BR to their cognate E2s resulted in a significant increase in E2:RING affinity as assessed by NMR, which correlated with enhanced ubiquitination in vitro [17,19]. Taking advantage of the well-characterized gp78 RING domain, we assessed the relative effects of G2BR^gp78 and G2BR^AUP1 on the affinity of UBE2G2 for this domain. Titration of UBE2G2 and UBE2G2:G2BR^AUP1 with the gp78 RING confirmed that the gp78 RING exhibited mostly fast exchange kinetics and that the increased affinity of gp78 RING was enhanced to a similar degree by G2BR^AUP1 as by G2BR^gp78. Through chemical shift perturbation (CSP) mapping, backbone amide resonances of G2BR-bound UBE2G2 were examined upon titration of the RING domain (Fig 5D and 5E, Biological Magnetic Resonance Data Bank deposition https://doi.org/10.13018/bmrbig33). Titration of UBE2G2:G2BR^gp78 and of UBE2G2:G2BR^AUP1 resulted in measured affinities of 20.3 ± 1.5 μM and 22.5 ± 1.8 μM, respectively. These results indicate equivalent enhancements of approximately 10-fold for G2BR^gp78 and G2BR^AUP1 on the affinity of UBE2G2 for the gp78 RING, which was measured as 207.4 ± 6.4 μM. Consistent with this increase in affinity, the G2BR^AUP1, like the G2BR^gp78, increased ubiquitination in an in vitro autoubiquitination assay employing UBE2G2 with either the gp78 RING domain or those of HRD1 and TRC8 (S4A and S4B Fig). Thus, as assessed both biophysically and functionally, G2BR^AUP1 has similar effects on UBE2G2 activity to what is observed with G2BR^gp78.

The AUP1–UBE2G2 interaction is required for ERAD

Inspection of the UBE2G2:G2BR^AUP1 interface reveals several contact residues that are likely responsible for their high-affinity binding. Specifically, five positively charged or polar residues of the G2BR^AUP1—R382, Q383, K390, R398, and R400—form salt bridges or hydrogen bonds with negatively charged or polar groups on UBE2G2 (Fig 4B and 4C). We first generated alanine mutations on these G2BR^AUP1 residues to neutralize positively charged side chains and assessed their effects on UBE2G2 binding and activity in cells by GST pulldown and CHX chase, respectively. Beginning at the N-terminal end of the G2BR^AUP1 helix, the R382A/Q383A mutation (2A) was unable to disrupt UBE2G2 binding (Fig 6A). Charge reversal of these residues (R382E/R383E, or 2E) also failed to disrupt UBE2G2 binding. Mutation of all five of the positively charged or polar G2BR^AUP1 residues to glutamate (5E), however, led to a loss of detectable UBE2G2 binding. Consistent with the importance of binding to UBE2G2, expression of a full-length AUP1 bearing the 2E mutation restored degradation of NHK in the AUP1 KO cells, to a degree comparable to WT AUP1 (Fig 6B). By contrast, expression of the AUP1 5E mutant failed to restore NHK degradation in these cells. These results support a requirement for UBE2G2 binding in AUP1 function in cells.

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Fig 6. Disruption of UBE2G2:G2BR^AUP1 interaction abrogates AUP1 function in cells.

(A) GST or indicated GST-AUP1 fusions (aa 292 to 410, expression shown in Coomassie-stained gel) were immobilized on glutathione beads and binding to ³⁵S-UBE2G2 was assessed as in Fig 3A. Mutations include 2A (R382A/Q383A), 2E (R382E/Q383E), and 5E (R382E/Q383E/K390E/R398E/R400E). (B) AUP1 KO cells were transfected with NHK-HA and forms of full-length AUP1-FLAG with the indicated point mutations from (A) and NHK degradation assessed by CHX chase. (C) Binding assay performed as in (A) with AUP1 mutants predicted to disrupt hydrophobic interactions with UBE2G2. (D) Cellular assay performed as in (B) with AUP1-FLAG mutants that disrupt interactions with UBE2G2 in vitro in (C). The data underlying this figure can be found in S1 Data. aa, amino acid; AUP1, ancient ubiquitous protein 1; CHX, cycloheximide; G2BR, UBE2G2 Binding Region; KO, knockout; WT, wild type.

https://doi.org/10.1371/journal.pbio.3001474.g006

As mentioned, the pattern of hydrophobic interactions in the UBE2G2:G2BR^AUP1 and UBE2G2:G2BR^gp78 complexes are highly similar (Fig 4C), suggesting that the van der Waals forces resulting from a perfect landscape match of the 2 molecules is also required for the stabilization of the complex. To test this hypothesis, we mutated the G2BR^AUP1 residues L386, Y394, and A397 to aspartate to disrupt this hydrophobic landscape and assessed effects on in vitro binding of UBE2G2. While the Y394D mutation alone reduced binding, mutation of L386 and Y394 together eliminated detectable binding to UBE2G2 (Fig 6C). Binding was similarly lost with a single point mutation, A397D, which makes multiple hydrophobic contacts with UBE2G2 (Fig 4C). Consistent with the in vitro binding results, reexpression of AUP1 harboring either the L386D/Y394D or A397D mutations failed to restore NHK degradation in cells lacking AUP1, confirming the functional importance of these residues (Fig 6D). Taken together, our mutational analysis demonstrates that the high-affinity binding between G2BR^AUP1 and UBE2G2, driven by both electrostatic and hydrophobic interactions, is critical for the function of AUP1 in ERAD.

The G2BR protects UBE2G2 from degradation

Previous studies have demonstrated that yeast Cue1p protein and its U7BR, which binds its E2 in an overall similar manner as the G2BR, protects Ubc7p from a degradative process that is dependent on the activity of the E2 [18,19,70]. The exact mechanism by which Cue1p protects Ubc7p remains unclear. Strikingly, despite the continued expression of gp78 and its G2BR, HT1080 cells lacking AUP1 showed substantially reduced levels and increased turnover of UBE2G2 (Fig 7A). On the other hand, cells lacking gp78 expression maintained normal levels of endogenous UBE2G2. Transcript levels of UBE2G2 were not affected by loss of AUP1 (Fig 7B). By transfecting either UBE2G2 or the catalytically inactive C89S mutant, we established that the observed turnover of UBE2G2 requires its E2 activity (Fig 7C). Increased distribution of AUP1 from the ER to lipid droplets has been reported in cells loaded with oleic acid [35]. This raises the interesting possibility that accumulation of lipid droplets in cells, such as observed in obesity, could affect ERAD by altering the availability or level of UBE2G2. However, in our system, we find that loading HT1080 cells with oleic acid, and thereby increasing lipid droplets, affected neither UBE2G2 levels nor the degradation of NHK (S5A and S5B Fig).

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Fig 7. G2BR^AUP1 protects UBE2G2 from degradation.

(A) Indicated cells were assessed for steady state levels and turnover of endogenous UBE2G2 by CHX chase. (B) Transcript levels of endogenous UBE2G2 in AUP1 KO cells compared to HT1080 as determined by qPCR. Mean and standard deviation from three independent experiments is shown. (C) Turnover of transfected MYC-UBE2G2 or a catalytically inactive mutant (C89S) was monitored in AUP1 KO cells. (D) AUP1 KO cells were transfected with plasmids encoding MYC-UBE2G2 and GFP fusions of either a scrambled G2BR (SCR), G2BR^gp78, or G2BR^AUP1, and the turnover of UBE2G2 was assessed. (E) Relative abundance of gp78 (left panel) and AUP1 (right panel) in HT1080 cells was determined by resolving indicated amounts (in pmoles) of purified GST-gp78 (aa 429 to 611) or GST-AUP1 (aa 61 to 410) fusions alongside approximately 1 × 10⁴ cellular equivalents of cell lysate and immunoblotting with gp78 or AUP1 antibodies. For each antibody, the amount of immunoreactivity in the lysate was compared to a defined quantity of a GST fusion protein of gp78 or AUP1 spanning the epitope recognized by the probing antibody. This analysis suggests there is close to 40-fold as much cellular AUP1 (150 pmole) as gp78 (4 pmol). The data underlying this figure can be found in S1 and S8 Data. aa, amino acid; AUP1, ancient ubiquitous protein 1; CHX, cycloheximide; G2BR, UBE2G2 Binding Region; KO, knockout; qPCR, quantitative polymerase chain reaction; WT, wild type.

https://doi.org/10.1371/journal.pbio.3001474.g007

To determine whether the G2BR^AUP1 is responsible for preventing UBE2G2 from undergoing constitutive degradation, we generated constructs encoding GFP fusions with either G2BR^AUP1 or G2BR^gp78 and transfected these into AUP1 KO cells. When compared to the control (a scrambled (SCR) G2BR sequence fused to GFP), either GFP-G2BR^AUP1 or GFP-G2BR^gp78 protected UBE2G2 from degradation and increased the steady state level of this E2 (Fig 7D; see S5C Fig for quantification of representative GFP-G2BR overexpression). A similar loss of UBE2G2 and rescue by GFP-G2BR overexpression was observed in HEK293 AUP1 KO cells (S5D and S5E Fig), and a dependency of E2 stability on AUP1 was also observed in M17, a neuroblastoma cell line (S5D Fig). The question then arises as to why endogenous gp78 is inadequate to maintain UBE2G2 levels in the absence of AUP1. A possible explanation could be the relative amounts of the two G2BR-containing proteins. By making use of existing AUP1 and gp78 antibodies and GST fusion proteins, we estimate that there is nearly 40-fold more AUP1 relative to gp78 in HT1080 cells (Fig 7E). This difference in prevalence between AUP1 and gp78 and, by extension, their G2BRs, provides at least one plausible explanation for the differential role of AUP1 in protecting UBE2G2. A substantially higher level of expression of AUP1 relative to gp78 is also observed in HEK293 and other human cell lines that we evaluated (S5F Fig). While these findings collectively suggest why AUP1 is important for maintaining UBE2G2 levels, the molecular basis for G2BR-mediated protection of UBE2G2 remains to be determined.

UBE2G2 membrane recruitment and activation by G2BR are necessary for ERAD

One simple explanation for the requirement for AUP1 in degradation of HRD1 substrates is that the dramatic reduction in UBE2G2 in the absence of AUP1 prevents efficient ERAD. However, as already shown, the degradation of INSIG-1, an established gp78 substrate that requires UBE2G2 (Fig 3B), was largely unaffected by AUP1 loss (Fig 1D). Even so, perhaps UBE2G2 levels are too low to support NHK degradation in the absence of AUP1. If this were the case, then simply overexpressing UBE2G2 might be expected to correct this deficit and restore NHK degradation. However, overexpression of MYC-tagged UBE2G2 to levels exceeding that of the endogenous protein did not restore NHK degradation in AUP1 KO cells (Fig 8A). This outcome establishes a requirement for AUP1 for ERAD function, beyond maintaining UBE2G2 levels. Notably, as seen also in Fig 7D, the overexpressed E2 was also unstable in AUP1 KO cells, although it is clearly present at higher levels than the endogenous E2 (Fig 8A).

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Fig 8. Recruitment of UBE2G2 to the ER membrane and activation by G2BR^AUP1 is required for ERAD.

(A) HT1080 or AUP1 KO cells were transfected with NHK-HA and either empty vector or MYC-UBE2G2 and NHK turnover assessed by CHX chase. (B) AUP1 KO cells were transfected with NHK-HA and GFP fusions of either a scrambled G2BR sequence (SCR) or G2BR^AUP1 and NHK turnover and levels of endogenous UBE2G2 assessed. Luciferase and β-actin served as transfection efficiency and gel loading controls, respectively. (C) AUP1 KO cells were transfected with NHK-HA, MYC-UBE2G2, and either GFP-SCR or GFP-G2BR^AUP1 and NHK turnover assessed. (D) Membrane fractionation was performed on HT1080 or AUP1 KO cells transfected with either empty vector or AUP1-FLAG and localization of UBE2G2 assessed by western blotting. Localization of β-actin (cytosol) and Calnexin (ER membrane) were used as controls for cytosolic (C) and membrane (M) fractions, respectively. (E) Membrane fractionation of AUP1 KO cells transfected with either MYC-UBE2G2 or TM-UBE2G2. (F) HT1080 cells were transfected with TM-UBE2G2 and its localization assessed by confocal microscopy. Images shown are from an optical section after deconvolution. HA-tagged UBE2J2, an ER-localized E2, serves as an ER marker. (G) U-2 OS cells were transfected as in (F), and localization assessed by SIM. (H) AUP1 KO cells were transfected with NHK-HA and TM-UBE2G2 and either GFP-SCR or GFP-G2BR^AUP1 and NHK turnover assessed. TM-UBE2G2 is indicated by the arrow, an incomplete form is indicated by *. (I) AUP1 KO cells were transfected with RI₃₃₂-MYC, TM-UBE2G2, and either GFP-SCR or GFP-G2BR^AUP1 and RI₃₃₂ turnover assessed. Endogenous UBE2G2 and TM-UBE2G2 are indicated, as is the incomplete form of the latter (*). The data underlying this figure can be found in S1 Data. AUP1, ancient ubiquitous protein 1; CHX, cycloheximide; ER, endoplasmic reticulum; ERAD, endoplasmic reticulum–associated degradation; G2BR, UBE2G2 Binding Region; KO, knockout; SIM, structured illumination microscopy.

https://doi.org/10.1371/journal.pbio.3001474.g008

Based on the specific, high-affinity interaction we observe between UBE2G2 and G2BR^AUP1, we asked whether expression of G2BR^AUP1 as a GFP fusion is sufficient to both increase UBE2G2 levels and restore ERAD in AUP1 KO cells. Overexpression of GFP-G2BR^AUP1, despite protecting UBE2G2 from degradation and increasing the levels of endogenous UBE2G2, was insufficient to reconstitute NHK degradation (Fig 8B). Expression of an AUP1 construct lacking only the hairpin membrane anchor was similarly unable to restore NHK degradation (S6A Fig). We next overexpressed UBE2G2 and GFP-G2BR^AUP1 together. Again, this did not restore NHK degradation, despite supraphysiological levels of UBE2G2 (Fig 8C; for a direct comparison of the level of endogenous UBE2G2 in HT1080 cells to the levels of transfected UBE2G2 levels at the zero time points, see S6B Fig, lanes 1 to 3).

We have shown that an internal deletion mutant of AUP1, which lacks both the acyltransferase and CUE domains but retains its membrane anchor and the G2BR, restores NHK degradation in AUP1 KO cells (Fig 1F). Therefore, it is possible that, in the context of the substrates being analyzed, the primary function of AUP1 is to recruit UBE2G2 to the ER membrane. In support of a role for AUP1 in ER membrane recruitment, UBE2G2 was overwhelmingly found in the membrane fraction in both WT HT1080 cells and in AUP1 KO cells when AUP1 was reexpressed (Fig 8D). By contrast, UBE2G2 was distributed between the membrane and cytosolic fractions in the AUP1 KO cells. To address the importance of G2BR^AUP1 in recruiting UBE2G2 to the ER membrane, we fused MYC-tagged UBE2G2 to the transmembrane domain of the yeast ERAD accessory protein Cue1p [14] to generate TM-UBE2G2. This transmembrane domain has been previously used to localize the cytoplasmic region of gp78 to the ER, which was found to be sufficient to restore degradation of gp78 substrates in cells where gp78 has been knocked down [28]. TM-UBE2G2 is inserted into membranes as assessed by fractionation (Fig 8E) and localizes to the ER as revealed by fluorescence microscopy (Fig 8F and 8G). Moreover, TM-UBE2G2 co-immunoprecipitated both HRD1 and SEL-1L, as previously shown for AUP1 (S6C Fig) [35,39]. Importantly, membrane-tethered UBE2G2 is functional in vivo as it reconstituted ERAD of NHK in UBE2G2 KO cells (S6D Fig). By contrast, when TM-UBE2G2 was expressed in AUP1 KO cells it did not, by itself, restore ERAD (Fig 8H, lanes 4 to 6). However, degradation of NHK was fully restored when TM-UBE2G2 was co-expressed with G2BR^AUP1 (GFP-G2BR^AUP1) (Fig 8H, lanes 7 to 9; for a direct comparison of the level of TM-UBE2G2 to endogenous UBE2G2, see S6B Fig, lanes 1, 4, and 5). Similarly, TM-UBE2G2 together with GFP-G2BR^AUP1 restored degradation of RI₃₃₂ (Fig 8I, lanes 7 to 9). This requirement for the G2BR builds upon the in vitro biophysical and biochemical findings in this and previous studies and demonstrates that the G2BR indeed has an activating role in cells. Taken together, our results suggest three distinct cellular roles for G2BR^AUP1: (1) stabilization of UBE2G2 by preventing its apparent self-targeting for degradation; (2) recruitment of UBE2G2 to the ER membrane; and (3) activation of UBE2G2 as an ERAD E2.

Cell lines

HT1080 (ATCC, CCL-121), HEK293, U-2 OS (ATCC, HTB-96), 293T (ATCC, CRL-3216), and HeLa (CCL-2) were maintained in DMEM supplemented with 10% (v/v) fetal bovine serum, 100 units/ml penicillin, 100 μg/ml streptomycin, and 2 mM glutamine in a humidified incubator at 37°C and 5% CO₂. BE(2)- M17 (ATCC, CRL2267) cells were additionally supplemented with MEM nonessential aa (HyClone), 100 μM sodium pyruvate, and 50 μM 2-mercaptoethanol (Gibco, Waltham, MA, USA). CRISPR/Cas-9 KO variants of HT1080, M17, and HEK293 were generated using Santa Cruz Double Nickase Plasmid kits: AUP1 (sc-410699-NIC), gp78 (sc-402346-NIC), and UBE2G2 (sc-404851-NIC), Ultra Cruz transfection reagent and plasmid medium (Santa Cruz, Dallas, TX, USA). Transfected cells were selected with 2 μg/mL puromycin (Millipore Sigma, Burlington, MA, USA) for approximately 3 weeks. Individual clones were screened, and loss of protein expression was confirmed by western blotting. sgRNA targeting HRD1 early in the protein sequence were identified using the sgRNA Scorer 2.0 algorithm [78] and 6 candidates were in vitro transcribed, complexed with Cas-9 protein and evaluated in 293T cells for editing efficiency as previous described [79]. The best candidate was then cloned into the LentiCRISPRv2GFP vector and transfected into parental cells using Lipofectamine 2000 (Thermo Fisher, Waltham, MA, USA). LentiCRISPRv2GFP was a gift from David Feldser (Addgene plasmid #82416; http://n2t.net/addgene:82416; RRID:Addgene_82416) [80]. Transfected cells were sorted by fluorescence activated cell sorting, and individual clones were expanded for sequencing to confirm HRD1 gene disruption. PCR primers (forward: 5′-TCCCTACACGACGCTCTTCCGATCTACACCAGTTCTACCCCACTG-3′ and reverse: 5′-GTTCAGACGTGTGCTCTTCCGATCTTGAAGAGTGCAACAAAGCGG-3′, where bold sequence corresponds to locus-specific sequence) were used to amplify around the region of interest and amplicons were subjected to Illumina sequencing. Loss of HRD1 protein expression was also confirmed by western blotting.

Plasmid constructs

Sequences encoding aa 61 to 410, 292 to 410, and 379 to 410 of AUP1 were amplified by polymerase chain reaction (PCR) from pEGFP-AUP1 (a generous gift from Hidde Ploegh) [54] and subcloned into pGEX-6P-1 (GE Healthcare, Chicago, IL, USA) from BamHI to XhoI. For co-expression of G2BR^AUP1 and UBE2G2 in Escherichia coli, sequence encoding aa 379 to 410 of AUP1 was subcloned from BamHI to HindIII sites within the first multiple cloning site (MCS) of pETDuet-GST [81]. UBE2G2 was subcloned from NdeI to XhoI within the second MCS of the vector. Sequence encoding full-length AUP1 was amplified by PCR from pEGFP-AUP1 and subcloned into pcDNA3-FLAG from BamHI to XhoI. All mutations in pGEX-6P-1 and pcDNA3-AUP1 were generated by site-directed mutagenesis using Quikchange XL (Agilent Technologies, Santa Clara, CA, USA), including 2A (R382A/Q383A), 2E (R382E/Q383E), 5E (R382E/Q383E/K390E/R398E/R400E), L386D, Y394D, L386D/Y394D, and A397D. Deletions of the transmembrane region (ΔTM, aa 25 to 45), G2BR (ΔG2BR, aa 386 to 397), CUE domain (ΔCUE, aa 293 to 337), and both the acyltransferase and CUE domains (ΔATΔCUE, 90 to 337) were generated in pcDNA3-AUP1-FLAG using QuikChange XL. Nucleotide sequences encoding AUP1 (aa 378 to 404), gp78 (aa 574 to 600) or a scrambled (SCR) gp78 sequence (SKANDSRELQFRKMRLRAQRQVQELDKL) were synthesized (Integrated DNA Technologies, Coralville, IO, USA) and subcloned into pEGFP-C1 vector (Clontech, San Jose, CA, USA) from BamHI to XhoI to generate GFP-tagged G2BRs. Sequence encoding the cytoplasmic domain of HRD1/SYVN (aa 236 to 617) was subcloned from pcDNA3.1(+)-HRD1-Myc/6His A(−) [31] into pGEX-4T-3 (GE Healthcare) from BamHI to EcoRI. Sequence encoding HRD1 (aa 236 to 444) was subsequently PCR amplified and subcloned into pGEX-6P-1 from BamHI to EcoRI. The gp78-RING construct (aa 313 to 393) was generated by inserting a stop codon in the pGEX-4T-2-gp78C plasmid [25]. gp78 (aa 429 to 611) was PCR amplified from pGEX-4T-2-gp78C and subcloned into the pGEX-6P-1 from BamHI to SalI. Site-directed mutagenesis of sequence corresponding to aa 595 to 600 within the G2BR (KRFLNK to AGAAGG) in pCINeo-gp78 [25] was used to generate a plasmid encoding gp78 with a disrupted G2BR. Disruption of this region in G2BR^gp78 results in a >5 × 10³ decrease in its affinity for UBE2G2 [17]. Sequence encoding the cytoplasmic domain of TRC8 (aa 513 to 664) was PCR amplified from HA-TRC8 [82] and subcloned into pGEX-4T-3 from BamHI to EcoRI. pcDNA3-NHK-HA was provided by Jeff Brodsky and was a gift from Ron Kopito [39]. NHK was PCR amplified and cloned into pcDNA4/MYC-6His A from HindIII to XhoI to generate NHK-MYC-6His. Ribophorin I (aa 1 to 332) was PCR amplified from pCMV-SPORT6-RPN1 (Dharmacon, Lafayette, CO, USA) and cloned into pcDNA4/MYC-6His A from EcoRI to XbaI. pcDNA3.1(+)-Cue1pTM-MYC-UBE2G2 (TM-UBE2G2) was generated by PCR of sequence encoding aa 1 to 44 of Cue1p from pRS316-Cue1p-Ubc7-HA [18] and subcloning into pcDNA3.1(+)-MYC-UBE2G2 from BamHI to EcoRI. This construct was modified by site-directed mutagenesis to add a carboxyl-terminal FLAG tag for efficient immunoprecipitation, generating pcDNA3.1(+)-Cue1pTM-MYC-UBE2G2-FLAG (TM-UBE2G2-FLAG). Sequence encoding the cytoplasmic domain of UBE2J1 (aa 1 to 283) was PCR amplified from an EST (IMAGE #4137664) and cloned into pET3a (Novagen, Burlington, MA, USA) from NdeI to BamHI. A Cys residue was inserted between the PreScission Protease cleavage sequence and HA-tag (LEVLFQGPLGSCYPYDVPDYA) on pGEX-6P-1-HA-ubiquitin [81] by site directed mutagenesis to allow labeling with Cy5. The deduced aa sequence of all coding regions was confirmed by sequencing.

The following plasmids have been previously described: pGEX-4T-2-UBE2G2, pcDNA-HA-UBE2J2, pcDNA-MYC-UBE2G2 WT and C89S [29]; pET3a-UBE2G2 [17]; pcDNA3.1(+)-HA-ubiquitin [83]; pET15b-UBE2D2 and pET15b-UBE2D3 [65]; pCIneo-gp78 [25]; pCMV14-INSIG-1-MYC was a gift from Joseph Goldstein [84]; pET11d-UBE2G1 was a gift from Simon Wing [85]; GFP^u was a gift from Ron Kopito [45]; and LentiCRISPRv2GFP (Addgene #82416) was a gift from David Feldser [80].

Antibodies

Antibodies were obtained for HRD1/SYVN1 (D3O2A #14773), AUP1 (D5M9Q #35055), UBE2G2 (D8Z4G #63182), CHOP (L63F7, #2895), BiP (C50B12, #3177), Phospho-eIF2α (S51, D9G0, #3398), eIF2α (#9722), and β-actin (8H10D10, #3700) from Cell Signaling (Danvers, MA, USA); FLAG (M2, F1084) and HA-Peroxidase (clone 3F10) from Millipore Sigma; GFP (B-2, sc-9996), Calnexin (AF18, sc-23954), HA (Y-11, sc-805), Sel-1L (F-3, sc377350) and ubiquitin (P4D1, sc-8017) from Santa Cruz; HSP60 (ab46798) from Abcam (Waltham, MA, USA); Luciferase (G7451) from Promega (Madison, WI, USA); HRP-linked anti-Mouse (NA931) and anti-Rabbit (NA934) IgG from GE Healthcare; Fluorescein-conjugated anti-Mouse IgG (F2761) and Texas Red-X-conjugated anti-Rabbit IgG (T6391) from Invitrogen (Waltham, MA, USA); and Clean-Blot (HRP) from Thermo Fisher. Rabbit antibodies recognizing the G2BR^gp78 (Ab2) was raised to aa 574 to 611 of gp78 and affinity purified using a peptide corresponding to aa 574 to 597 of gp78. In S1A Fig, where the G2BR^gp78 is disrupted, affinity-purified rabbit Ab1 antibodies, raised against aa 505 to 529 of gp78, were utilized [25]. Rabbit antiserum against HRD1 [23], antibodies against gp78 (Ab2) [28], and ubiquitin [86] were previously described. Anti-MYC (9E10, ATCC CRL-1729) was purified on a MYC column from hybridoma supernatant. Preparation and use of mouse monoclonal antibody directed against HMGCR (A9) has been described [55]. Anti-HA Affinity Gel (E6779), Anti-c-MYC Affinity Gel (A7470), and Anti-FLAG M2 Magnetic Beads (M8823) were from Millipore Sigma.

Peptides

G2BR peptides (Table 2) were synthesized as follows for in vitro and biophysical measurements.

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Table 2. Amino acid sequences of G2BR peptides.

https://doi.org/10.1371/journal.pbio.3001474.t002

G2BR^gp78 was synthesized with the addition of an N-terminal Trp residue for peptide quantification. Scrambled peptide (SCR) is a mutated sequence based on aa 378 to 404 of AUP1 and used as a negative control in experiments. Lyophilized peptides were resuspended and dialyzed extensively against 50 mM Tris-HCl, pH 7.4 at 4°C, and their concentrations determined by absorbance at 280 nm using calculated extinction coefficients from aa sequences. Dialyzed peptides were stored in small aliquots at −20°C.

Cellular experiments

Plasmid transfections were carried out using PolyFect (Qiagen, Germantown, MD, USA) or JetPrime (Polyplus, New York, NY, USA) reagents. For CHX chase experiments, approximately 24 hours following plasmid transfection, cells were treated with 50 μg/mL CHX in complete media for the indicated times. For lipid droplet experiments, cells were treated with 200 μM oleic acid (Millipore Sigma) for 16 hours, beginning 24 hours following transfection. For assessment of GFP^u stability, cells were cultured in DMEM supplemented with 10% FetalPlex animal serum complex (Gemini Bio, West Sacramento, CA, USA) and treated with 50 μM emetine for the indicated times to inhibit protein synthesis [40]. For experiments assessing basal degradation of HMGCR, expression of HMGCR was induced by culturing cells for 24 hours in LPDS media [55], which contains DMEM supplemented with 10% (v/v) fetal bovine lipoprotein-deficient serum (LPDS; d > 1.25), 100 μM sodium mevalonate, and 5 μM compactin. Cells were then treated with 50 μg/mL CHX in LPDS media for the indicated duration and cell lysates were harvested. LPDS was prepared by ultracentrifugation, as described [87]. For assessment of whole cell lysates, cells were lysed in modified RIPA buffer (1X PBS pH 7.4, 1% Triton X-100, 0.5% sodium deoxycholate) supplemented with 25 μM MG132 and cOmplete Protease Inhibitor Cocktail (Roche, Burlington, MA, USA). For co-immunoprecipitation with Cue1pTM-UBE2G2-FLAG, cells were treated with 25 μM MG132 for 4 hours, lysed in PBS (pH 7.4) with 1% digitonin and 40 μM MG132, and lysates immunoprecipitated with Anti-FLAG (M2) Magnetic Beads. For co-immunoprecipitation with FLAG-UBE2G2, cells were lysed in PBS (pH 7.4) with 0.5% Triton X-100, 0.1% deoxycholate and 40 μM MG132, and immunoprecipitated with Anti-FLAG (M2) magnetic beads. Clarified lysates or immunoprecipitates were denatured by heating in NuPAGE LDS Sample Buffer with β-mercaptoethanol at 70°C for 10 minutes, resolved on Bis-Tris polyacrylamide gels in NuPAGE MES or MOPS SDS running buffer and analyzed by western blotting. Proteins were detected using Clarity Western ECL (Bio-Rad, Hercules, CA, USA), Radiance Q (Azure Biosystems, Dublin, CA, USA), or SuperSignal Femto West (Thermo Scientific, Waltham, MA, USA) ECL reagents and a c280 Imager (Azure Biosystems) or autoradiography.

Membrane fractionation

Membrane fractionation experiments were previously described [23,55]. Twenty-four hours after transfection, cells were washed and resuspended in 0.25 M sucrose, 10 mM triethanolamine, supplemented with 25 μM MG132 and protease inhibitors, and then passed through a 27.5-gauge needle 15 times. Cell homogenates were centrifuged twice for 5 minutes at 1,000 × g to remove the nuclei, and the supernatants were spun twice for 30 minutes at 100,000 × g at 4°C in a Sorvall Discovery M150 ultracentrifuge using a S100 AT4-541 rotor to separate membrane (pellet) and cytosolic (supernatant) components. Cytosolic and membrane fractions were resolved on Bis-Tris polyacrylamide gels and analyzed by western blot.

Fluorescence microscopy

qPCR

RNA was extracted from HT1080 cells with TRIzol (Invitrogen) according to manufacturer’s instructions and reverse transcribed to cDNA using SuperScript III First-Strand Synthesis System for RT-PCR (Invitrogen). PCR reactions were performed on a CFX96 Real-Time PCR system using iQ SYBR Green Supermix (Bio-Rad). Transfected AUP1-FLAG constructs were amplified using the forward and reverse primers 5′-CCTGAAGACATCACCAAGGGA-3′ and 5′-GTGGTGCTCGAAGATCTTGTC-3′, respectively. UBE2G2 and HRD1/SYVN were amplified using RT² quantitative polymerase chain reaction (qPCR) Primer Assay (Qiagen) using NM_182668.2 and NM_172230, respectively. ER stress markers were amplified using the following primers (IDT): ATF-4: 5′-ATGGCCGGCTATGGATGAT-3′ and 5′-CGAAGTCAAACTCTTTCAGATCCATT-3′; BiP: 5′-TGTTCAACCAATTATCAGCAAACTCTTCTGCTGTATCCTCTTCACCAGT-3′ and 5′-ACTGGTGAAGAGGATACAGCAGAAGAGTTTGCTGATAATTGGTTGAACA-3′; CHOP: 5′-CTGCCTTTCACCTTGGAGAC-3′ and 5′-CGTTTCCTGGGGATGAGATA-3′; and XBP1-spliced: 5′-GAGTCCGCAGCAGGTG-3′ and 5′-GTGTCAGAGTCCATGGGA-3′. A total of 3 technical replicates for each condition were measured and analyzed by the ΔΔCt method using either β-actin or GAPDH as an internal control (QuantiTect primers QT01680476).

In vitro E2 binding assay

In vitro translated ³⁵S methionine-labeled E2s were generated using the E. coli T7-S30 Extract System (Promega) and excess ³⁵S methionine was removed by desalting twice with Zeba Spin Desalting Columns (Thermo Scientific). GST fusion proteins were quantified by Coomassie blue staining. Moreover, 0.1 μg of GST fusion protein (approximately 3 pmol) was prebound to GS 4B (GE Healthcare) and then incubated with approximately 10⁵ cpm of ³⁵S methionine-labeled E2 (approximately 0.2 pmol) in 200 μl Binding Buffer (25 mM Tris-HCl, pH 7.4, 50 mM NaCl, 5 mM DTT, 0.5% IGEPAL CA-630) overnight at 4°C. Beads were extensively washed in Binding Buffer, resolved on Bis/Tris polyacrylamide gels under reducing conditions, dried and visualized on a Typhoon FLA 7000 (GE Healthcare). Binding was quantified using ImageQuant Software.

In vitro ubiquitination assay

GST-Cys-HA-ubiquitin was expressed in E. coli BL21 (DE3), purified with GS 4B, and incubated with excess Cy5 maleimide (Amersham PA15131) at room temperature for 1 hour. After removing excess dye by extensive washing, Cy5-HA-ubiquitin was released from the beads with PreScission Protease (GE Healthcare) overnight at 4°C, leaving the N-terminal sequence (GPLGS) prior to the labeled Cys. Cy5-labeled ubiquitin was quantified with Coomassie staining. Ubiquitination reactions were performed with 1 μM of E3 immobilized on GS 4B in 50 μl reaction volume. The reaction mixture contained approximately 100 nM E1, 2 μM UBE2G2, 8 μM Cy5-labeled ubiquitin, and 5 μM peptide in reaction buffer (50 mM Tris-HCl pH 7.4, 5 mM MgCl₂, 20 mM ATP, 1 mM DTT, and 0.1% Triton X-100). Reactions were carried out at 37°C for 90 minutes with continuous shaking, quenched with 4X NuPAGE LDS Sample Buffer (Thermo Fisher), resolved on Bis/Tris polyacrylamide gels under reducing conditions, and visualized on a Typhoon FLA 7000.

In vitro deubiquitination

HT1080 cells were lysed in lysis buffer (50 mM Tris-HCl pH 7.4, 120 mM NaCl, 1% Triton X-100) supplemented with 10 mM iodoacetamide, protease inhibitors, and 40 μM MG132. Lysate was clarified by centrifugation at 1,000 x g at 4°C for 10 minutes. Ubiquitin conjugates were enriched by binding with TUBE1 and TUBE2 agarose (UM401, UM402, respectively; Life Sensors, Malvern, PA, USA) on a rotating platform for 4 hours at 4°C. Control agarose (UM400) was used as negative control. After washing 3× with lysis buffer (20× bed volume), the beads were treated with 5 mM DTT in 0.1% Triton X-100 buffer (50 mM Tris-HCl pH 7.4, 100 mM NaCl, 0.1% Triton X-100) for 15 minutes and washed 2x with buffer containing 1 mM DTT. The beads were then treated with buffer alone or a DUB cocktail (DB599, Life Sensors) at 37°C for 4 hours with constant agitation. The beads were washed 3× with 0.1% Triton X-100 buffer and bound proteins were eluted with 2X sample buffer.

Protein expression for crystallization

The pETDuet-GST vector was used to co-express GST-AUP1(379 to 410) and UBE2G2 in E. coli BL21(DE3) cells (Invitrogen). Briefly, cells were induced at OD₆₀₀ 0.8 with 0.15 mM IPTG overnight at 25°C to express both proteins. Cells were lysed with an APV-2000 homogenizer in binding buffer (50 mM Tris-HCl pH 7.5, 150 mM NaCl, 1 mM DTT), and the homogenate was centrifuged at 27,000 x g for 20 minutes to remove bacterial cell wall. The supernatant was collected and incubated with GS 4B for 2 hours at 4°C to isolate GST-AUP1(379 to 410) and the associated UBE2G2. The beads were then collected by centrifugation and extensively washed with binding buffer. The GST tag was removed through on-column cleavage with PreScission Protease overnight at 4°C. After purification, the complex containing AUP1(379 to 410) and UBE2G2 was immediately used for crystallization.

Crystallization, data collection, and structure determination

Crystals of the UBE2G2:G2BR^AUP1 complex were grown by sitting drop vapor diffusion at 19 ± 1°C. Each droplet contained 0.6-μl protein solution (15 mg/ml in 150 mM NaCl and 50 mM Tris-HCl, pH 7.5) and 0.2-μl reservoir solution (30% PEG 4000, 0.2 M NH₄Ac, and 0.1 M NaAc, pH 4.6), and the premixed 0.8 μl droplets were equilibrated against 60 μl reservoir solution. The rod-shaped crystals grew to full size (0.08 mm × 0.08 mm × 0.3 mm) in 3 days. Crystals were flash-cooled in cold nitrogen stream. X-ray diffraction data were collected from a single crystal on a MARMOSAIC 325 detector at the BL9-2 beamline of the Stanford Synchrotron Radiation Lightsource (SSRL) and processed using the HKL-3000 program suite [88]. Data statistics are presented in Table 1. The UBE2G2:G2BR^AUP1 structure was determined by molecular replacement using phenix.automr of the PHENIX program suite [89]. The crystal structure of the UBE2G2:G2BR^gp78 (PDB entry 3H8K) [17] was used as the search model after solvent molecules were removed. The structure was refined with phenix.refine of PHENIX, and the model building and adjustment was done with COOT [90]. About 1,000 reflections were randomly selected for cross-validation. Water molecules were included at the last stage of the refinement on the basis of difference electron density (Fo-Fc, above 3σ) and verified with omit maps. The refined structure was validated using the PROCHECK [91] and WHATIF [92] programs. Figures were generated with PyMol (DeLano Scientific, South San Francisco, CA, USA). Refinement statistics of the crystal structure are also presented in Table 1.

NMR

UBE2G2 and gp78 RING used in titration experiments were expressed and purified as previously described [24]. G2BR^gp78 and G2BR^AUP1 peptides were solubilized and dialyzed against 50 mM Tris-Cl pH 7.0, 0.5 mM TCEP, 0.05% sodium azide. NMR samples of G2BR^gp78:UBE2G2 and G2BR^AUP1:UBE2G2 were prepared in 50 mM Tris-Cl pH 7.0, 0.5 mM TCEP, 100 μM zinc sulfate, and 0.05% sodium azide. NMR spectra were collected on Bruker AVIII spectrometers operating at 800 or 850 MHz using cryogenic TCI probes. The ¹H-¹⁵N HSQC spectra of ¹⁵N-labeled UBE2G2 at 150 μM were collected in the absence and presence of gp78 RING at concentrations ranging between 0 and 450 μM. The changes in chemical shift for each peak in the 2D spectrum upon gp78 RING binding was calculated using the following CSP equation: (1)

The CSPs calculated from Eq 1 were plotted versus the molar ratio of gp78 RING to UBE2G2:G2BR, and the data were fit using the following equation to calculate the dissociation constant (K_d), where x is the ligand-to-protein ratio: (2)

ITC

ITC measurements were performed with purified UBE2G2 [17] and G2BR^gp78 or G2BR^AUP1 peptides (GenScript, Piscataway, NJ, USA). Samples were extensively dialyzed against PBS (pH 7.4) at 4°C, and final concentrations were determined by absorbance at 280 nm. UBE2G2:G2BR interactions were studied using an iTC200 calorimeter (Malvern/MicroCal) at 25°C. A typical experiment included injection of 18 aliquots (2.1 μL each) of 30 μM G2BR into a solution of 3 μM UBE2G2 in the cell (volume 200 μL) at a stirring speed of 750 RPM. An additional set of injections was run in a separate experiment with buffer in the cell instead of the protein solution. These blank experiment data were subtracted from the main ligand-into-protein experiment data. K_d values were determined from integrated binding isotherms using the “One set of sites” model in Origin 7.0—based Malvern/MicroCal data analysis software.

Fluorescence affinity measurements

FP experiments were performed by incubating purified UBE2G2 with 10 nM FITC-labeled G2BR peptide (Peptide 2.0) for 15 minutes at 25°C in 50 mM Tris-HCl pH 7.5, 1 mM DTT, 0.1 mg/ml BSA. Fluorescence anisotropy at 490-nm excitation and 515-nm emission were measured for each titration point in triplicate in Corning 384 well round bottom black polystyrene plates at 25°C with a Spectra Max M5 Microplate Reader (Molecular Devices, San Jose, CA, USA). MST was performed with titration of purified UBE2G2 with FITC-labeled G2BR peptide (Peptide 2.0) in Monolith NT.115 (NanoTemper Technologies, München, Germany). Measurements were repeated at least twice with biological triplicates. Data analysis was carried out using Prism software (GraphPad, San Diego, CA, USA) and fitted by nonlinear least squares assuming 1:1 stoichiometry to estimate the K_d.