VPS13D interacts with VCP/p97 and negatively regulates endoplasmic reticulum–mitochondria interactions

VPS13D suppression results in extensive endoplasmic reticulum–mitochondria tethering

Given the role of VPS13D in determining mitochondrial morphology and function in disease, we explored whether it could affect the interactions between the ER and mitochondria, using live-cell confocal microscopy. Remarkably, small interfering RNA (siRNA)–mediated VPS13D suppression resulted in extensive ER–mitochondria tethering in U2OS cells by live-cell confocal microscopy (Figure 1A). The efficiency of siRNA-mediated VPS13D suppression was confirmed by Western blots (Supplemental Figure S1A) and quantitative PCR (qPCR; Supplemental Figure S1B). Increased interactions between the ER and mitochondria could be observed in both perinuclear and peripheral regions of cells (Figure 1, A–C). Our definition of perinuclear regions is described in Supplemental Figure S1C.

In addition, the extensive ER−mitochondria tethering induced by VPS13D suppression was quantitatively confirmed in a doxycycline (Dox)-inducible splitGFP-based ER−mitochondria MCS reporter U2OS cell line, which was a modified version of that used in a previous study (Cieri et al., 2018). Because the reconstitution of GFP (green fluorescent protein) was irreversible, a low concentration (0.1 μM) and a short incubation time (12 h) of DOX were applied to avoid artificial tethering between the ER and mitochondria. Under this condition, both the ER and mitochondria appeared to be morphologically normal, and the GFP signals were specifically localized to the ER–mitochondria intersections (Supplemental Figure S1D). More importantly, we found that VPS13D suppression substantially increased the fluorescence intensity of splitGFP at sites where mitochondria were tethered to the ER (Supplemental Figure S1E).

To gain structural details of hypertethered ER–mitochondria contacts, we examined 3D reconstructed projections of these contacts in a perinuclear region of VPS13D-suppressed U2OS cells. We observed that the mitochondria were almost completely circumscribed by multiple ER tubules (Figure 1D). In addition, we directly examined ER-mitochondrial MCSs in VPS13D-suppressed U2OS cells by transmission electron microscopy (TEM). The ER–mitochondria MCSs were defined as regions within ∼30-nm gaps between the ER and the outer mitochondrial membranes (OMM) under TEM (Csordas et al., 2006). Compared with the controls, VPS13D suppression caused substantial increases in both the number and length of ER–mitochondria contacts (Figure 1, E and F).

In addition, we examined whether VPS13D regulated contacts between the ER and other organelles, including the Golgi apparatus, peroxisomes, early endosomes, and late endosomes/lysosomes. We did not detect any substantial morphological changes in these ER-associated MCSs upon VPS13D suppression in U2OS cells (Supplemental Figure S2).

Given that VPS13A, another member of the human VPS13 family, was reported to be a lipid transporter at ER–mitochondria MCSs (Kumar et al., 2018), we tested whether the suppression of VPS13A could alter the ER–mitochondria interactions. The siRNAs-mediated suppression of VPS13A failed to increase the ER–mitochondrial interaction substantially by live-cell confocal microscopy (Supplemental Figure S3, A and B). In contrast, VPS13A and VPS13D double suppression led to an increase in ER–mitochondria interactions to an extent similar to that of VPS13D suppression alone (Supplemental Figure S3, A and B). The efficiency of siRNA-mediated silencing of VPS13A was confirmed by qPCR (Supplemental Figure S3C). These results indicated that the extensive associations between the ER and mitochondria were specific to VPS13D suppression and suggested that VPS13A was not required in the ER–mitochondria hypertethering, as the hypertethering phenotype still existed in the VPS13A/D double-knockdown condition.

In addition, we examined the localization of endogenous VPS13D relative to ER–mitochondria MCSs by immunofluorescence (IF) using a polyclonal antibody against VPS13D, which was precleared as previously described (Valverde et al., 2019). IF experiments using this precleared antibody showed that VPS13D was preferentially present at the intersections between the ER and mitochondria (Supplemental Figure S3D, Top panel); however, IF signals of endogenous VPS13D almost completely disappeared upon VPS13D suppression (Supplemental Figure S3D, Bottom panel). We found that a substantial portion of endogenous VPS13D (49%) was present or enriched at the sites of mitochondria tethering to the ER (Supplemental Figure S3E); meanwhile, 79% of ER–mitochondria junctions were marked by endogenous VPS13D (Supplemental Figure S3F). As a negative control, we rotated the VPS13D image 90° relative to the ER image and quantified the percentage of VPS13D puncta or foci at the ER contacts. We found that the percentage of 90°-rotated VPS13D at ER contacts is substantially reduced (Supplemental Figure S3E), suggesting that the presence of VPS13D at ER contacts is significant and not due to chance. Our results are in accordance with a recent study (Guillen-Samander et al., 2021).

VPS13D negatively regulates the level of VAPB

To understand how VPS13D negatively regulates ER–mitochondria interactions, we examined the cellular distribution of the known tethering proteins involved in ER–mitochondria MCSs by IF. Among these tethers, we found that the ER-resident tethering protein VAPB, known for tethering the ER to mitochondria with its binding partner PTPIP51, was strongly enriched at sites around mitochondria (Figure 2A, Middle panel), presumably at ER–mitochondria MCSs, upon VPS13D suppression. To test whether this enrichment is specific for VAPB, we examined the distribution of sec61, a general ER membrane marker, in cells depleted of VPS13D. The sec61 protein is almost evenly distributed all over the ER, though we can still see enhanced ER–mitochondria interactions in VPS13D-suppressed U2OS cells (Figure 2A, Bottom panel). This result suggests that the enrichment of VAPB at ER contacts is specific upon VPS13D suppression. The antibody against VAPB used in the IF assays was validated by siRNA-mediated suppression (Figure 2B).

To further confirm the enhanced enrichment of VAPB at ER–mitochondria MCSs in VPS13D-suppressed cells, we performed cell fractionation to examine the levels of VAPB and PTPIP51 in crude mitochondrial fractions, which contained pure mitochondria and mitochondria-associated membranes (MAMs). The level of VAPB increased strongly in the crude mitochondrial fractions in VPS13D-suppressed cells (Figure 2C). In addition, consistent with our imaging results, the cellular fractionation assays showed that levels of calnexin, a marker for MAMs, increased in crude mitochondrial fractions upon VPS13D suppression (Figure 2C), suggesting enhanced ER–mitochondria interactions. The suppression of VPS13D also increased the level of PTPIP51 in the crude mitochondrial fractions, but to a lesser extent than that of VAPB (Figure 2C). To test whether the increase of VAPB at MAMs/mitochondria only mirrors the increase of VAPB in whole-cell lysate, we examined the ratio of the VAPB level in scrambled cells to that in VPS13D-suppressed cells. We found that the level of VAPB increased upon VPS13D suppression in whole-cell lysate (WCL), but the extent of the increase in VAPB level in MAMs/mitochondrial fractions is significantly higher than that in WCL (Figure 2, C and D).

Previous work showed that overexpression of VAPB increased ER–mitochondria associations in NSC34 cells by EM (Radu Stoica et al., 2014). Consistently, we showed that overexpression of VAPB alone or coexpression of VAPB and PTPIP51 substantially enhanced the interactions between the ER and mitochondria in the ER–mitochondria MCSs reporter cell line by both confocal microscopy (Supplemental Figure S4, A–C) and flow cytometry (Supplemental Figure S4, D–H). These results indicated that the expression of VAPB alone or the coexpression of VAPB and PTPIP51 was able to promote ER–mitochondria interactions. Therefore, we hypothesized that VPS13D controlled the extent of ER–mitochondria MCSs by regulating the level of VAPB. To prove this, we tested whether suppression of both VAPB and its binding partner PTPIP51 could rescue the extensive tethering between the ER and mitochondria resulting from VPS13D suppression. Consistent with previous studies (De Vos et al., 2012), double suppression of VAPB and PTPIP51 robustly reduced ER–mitochondria interactions in wild type (WT) U2OS cells. The efficiency of siRNAs-mediated suppression of VAPB and PTPIP51 was confirmed by Western blots (Figure 2E). More importantly, we found that the suppression of VAPB and PTPIP51 substantially rescued the ER–mitochondria hypertethering at both perinuclear and peripheral regions in VPS13D-suppressed cells (Figure 2, F and G). This result suggested that VPS13D negatively regulates ER–mitochondria MCSs by modulating the levels of VPAB and PTPIP51.

VPS13D interacts with p97 and is required for the stability of p97

Next, we investigated how VPS13D negatively regulates the level of VAPB at ER–mitochondrial MCSs. AAA-ATPase valosin-containing protein (VCP/p97) reportedly plays essential roles in ubiquitin-dependent protein degradation, dynamics, and homeostasis (Ye et al., 2001). p97 acts downstream of ubiquitin signaling events and utilizes the energy from ATP hydrolysis to extract its substrate proteins from membrane structures or multiprotein complexes (Ye et al., 2001). We found that endogenous VPS13D interacted with p97 by coimmunoprecipitation assays (Supplemental Figure S5A). Furthermore, we investigated which domains of VPS13D were responsible for interacting with p97 via GFP-trap assays. Our bioinformatics analysis predicted that VPS13D consists of an N-terminal lipid transfer domain, a UBA domain followed by a VPS13 adaptor-binding (VAB) domain, and a VPS13_C domain at its C-terminus (Figure 3A). The GFP-trap assays indicated that both the UBA domain and the VAB domain were capable of interacting with p97 (Figure 3B). In contrast, the C-terminal region of VPS13D (residues 3791–4363) failed to interact with p97, supporting the conclusion that VPS13D-p97 interaction is specific. p97 was predicted to harbor an N domain at the N-terminus and a D1/D2 domain at its C-terminus (Stolz et al., 2011; Supplemental Figure S5B, Top panel). Our GFP-trap assays showed that neither the N domain alone nor the D1/D2 domain alone was able to interact with VPS13D (Supplemental Figure S5B, Bottom panel), suggesting that both of these two domains of p97 were required for interaction with VPS13D.

The UBA domain of VPS13D was reported to bind to K63-, but not K48-linked ubiquitin chains, as determined by in vitro binding assays (Anding et al., 2018). We examined whether the VAB domain of VPS13D interacted with ubiquitin via IF. The IF results showed that VAB-GFP puncta were associated with both K63- and K48-linked ubiquitin chains in cells, with a preferential interaction with K63-linked ubiquitin (Supplemental Figure S5C). In addition, our GFP-trap assays confirmed the interactions between VAB-GFP and K63- and K48-linked ubiquitin chains (Supplemental Figure S5D). To further validate the VAB domain−ubiquitin interactions, GFP-trap assays using harsh wash buffers were performed to wash out proteins that were noncovalently bound to VAB-GFP. These results showed that neither K63- nor K48-linked ubiquitin was detectable (Supplemental Figure S5E), indicating that the VAB domain interacted with ubiquitin chains in a noncovalent manner and also that the VAB domain might not be conjugated by ubiquitin. In addition, our GFP-trap assays showed that p97 interacted with both K48- and K63-linked ubiquitin chains (Supplemental Figure S5F). Taken together, these lines of evidence indicated that the UBA domain and the VAB domain of VPS13D were independently associated with p97 and ubiquitin, suggesting that VPS13D, p97, and ubiquitin might form a ternary protein complex.

Previous studies demonstrated that p97 promoted the degradation of VAPB mutants (Papiani et al., 2012; Baron et al., 2014), but whether p97 could interact with and facilitate the degradation of WT-VAPB was unclear. Therefore, we examined the interactions between p97 and VAPB by GFP-trap assays, and we found that p97-GFP indeed interacted with endogenous VAPB (Supplemental Figure S5G). Because our results showed that VPS13D interacted with p97 and p97 also interacted with VAPB, we hypothesized that VPS13D might cooperate with the protein extractor p97 to dissociate the ER-resident tether VAPB from ER membranes, and consequently control the ER–mitochondria MCSs. To test this idea, we examined the effects of p97 on the degradation of WT-VAPB or two disease mutant variants of VAPB, VAPB-P56S and VAPB-T46I (Chen et al., 2010), both of which were implicated in ALS. We found that overexpression of p97 substantially decreased the level of WT-VAPB, similarly to that of VAPB-P56S (Figure 3, C and D). One striking difference was that p97 overexpression failed to reduce the level of another VAPB mutant, VAPB-T46I (Figure 3, C and D), and this result was further confirmed in other cell lines, including HEK293, COS7, and HeLa cells (Supplemental Figure S5H), suggesting a new mechanism underlying the pathology of VAPB-T46I in ALS. To test whether VAPB-T46I is still sensitive to p97-mediated repression, but starts from a much higher level than the WT in the absence of p97 overexpression, we examined the level of VAPB and its pathogenic mutants in the absence of p97 overexpression in U2OS cells. We found that there is no substantial difference in the levels of VAPB-GFP, VAPB-GFP-T46I, and VAPB-GFP-P56S (Supplemental Figure S5I).

Next, we investigated whether VPS13D suppression affected p97. We initially examined the cellular distributions of endogenous p97 upon VPS13D suppression by IF using an anti-p97 antibody (Supplemental Figure S6A), which was validated by siRNA-mediated suppression (Supplemental Figure S6A, bottom panel). In control cells, p97 was mainly cytosolic, with a small fraction of p97 localizing to ER–mitochondria intersections in form of the punctae-like structures (Supplemental Figure S6A, top panel). We found a significant decrease in the fluorescence of endogenous p97 protein upon VPS13D suppression (Supplemental Figure S6A, middle panel), suggesting a correlation between VPS13D and p97 level. Therefore, we sought to explore whether VPS13D suppression affected the level of p97. Indeed, we found that the level of endogenous p97 was substantially reduced in response to VPS13D suppression (Figure 3, E and F). In addition, we explored whether VPS13D suppression affected the p97-VAPB interactions. Our GFP-trap assays showed that a significantly smaller amount of VAPB was coimmunoprecipatated with p97-GFP in response to VPS13D suppression than in control cells (Figure 3G). We also noted that the level of p97-GFP was remarkedly reduced in VPS13D-depelted cells (Figure 3, E and F), which may account for the smaller amount of VAPB coimmunoprecipitated by p97-GFP.

Given the correlation between VPS13D suppression and the reduced level of p97 in cells, we tested whether VPS13D was required for p97 protein stability by cycloheximide (CHX) chase assays. In control cells, endogenous p97 was stable, with a small but significant decrease over time, and p97 was readily detected after 30 h treatment of CHX (Figure 3, H and I). In contrast, the initial level of endogenous p97 was substantially lower, and p97 was almost undetectable after 17 h treatment of CHX in VPS13D-depleted cells (Figure 3, H and I), suggesting that p97 stability was compromised in response to VPS13D suppression.

To further confirm the role of p97 in the VPS13D-mediated regula­tion of ER–mitochondria interactions, we examined ER−mitochondria interactions in cells in which p97 was inhibited by its allosteric inhibitor NMS873, using live-cell confocal microscopy. We found that p97 inhibition resulted in extensive ER–mitochondria tethering in both perinuclear and periphery regions in a dosage-dependent manner (Figure 3, J and K), which mimicked the phenotype induced by the suppression of VPS13D.

In addition, we examined the ER–mitochondria interactions in p97 siRNA–treated U2OS cells. We found that siRNA-mediated p97 suppression significantly enhances the ER–-mitochondria interactions (Figure 3, L–O), consistent with the p97 inhibitor results (Figure 3, J and K). In addition, we examined the ER–mitochondria interactions in response to p97 overexpression in VPS13D-suppressed U2OS cells. Cells with VPS13D suppression and GFP vector expression substantially increase the ER–mitochondria connections (Figure 3N, Top panel; Figure 3O). Expression of p97-GFP moderately decreases, but does not completely rescue, the ER–mitochondria interaction in both perinuclear and peripheral regions in VPS13D-suppressed cells (Figure 3N, Bottom panel; Figure 3O), suggesting that, in addition to the level of p97, unknown pathways may also be responsible for the ER–mitochondria hypertethering phenotype observed in VPS13D-suppressed cells.

VPS13D suppression perturbs mitochondrial morphology, mitochondrial distribution, and mtDNA synthesis

Next, we investigated whether the extensive ER–mitochondria tethering resulting from VPS13D suppression affected mitochondrial dynamics and functions. We tracked the dynamics of the ER–mitochondria interactions in either control or VPS13D-suppressed U2OS cells. Time-lapse image analysis demonstrated that mitochondrial motility was dramatically impaired in VPS13D-suppressed cells compared with that in controls (Figure 4A). Also, mitochondrial length was significantly reduced (Figure 4B), in agreement with a previous study in Drosophila (Anding et al., 2018). In addition, we found that the majority of VPS13D-suppressed cells exhibited defects in mitochondrial distribution, in which mitochondria were clustered in perinuclear regions (Figure 4C), suggesting that VPS13D is required for mitochondrial transport.

We further explored whether these defects in mitochondrial morphology and distribution resulting from VPS13D suppression can be rescued by VAPB/PTPIP51 double knockdown or p97 overexpression. We found that both of these defects in mitochondrial perimeter and distribution are substantially rescued upon the simultaneous suppression of VAPB and PTPIP51 (Figure 4, B and C). Interestingly, p97 overexpression can significantly rescue, but not completely restore, these two defects (Figure 4, B and C), consistent with its effects on the ER-mitochondrial hyper-tethering phenotype.

Previous studies demonstrated that mtDNA synthesis was spatially and temporally regulated by ER–mitochondria MCSs (Murley et al., 2013; Lewis et al., 2016). Therefore, we sought to explore whether VPS13D regulated mtDNA synthesis. We performed siRNA-mediated suppression of VPS13D in a U2OS cell line stably expressing polG2-GFP, a marker of replicating mtDNA (Young et al., 2015). We found that replicating mtDNA localized to mitochondria at a density of 0.28 μm^–1 in control cells, with some polG2-GFP puncta preferentially localizing at the tips of mitochondria (Figure 4D, Top panel). In contrast, VPS13D suppression reduced the density of replicating mtDNA by a factor of 4 and strongly increased the size of replicating mtDNA, suggesting aggregation of mitochondrial nucleoids (Figure 4D, Bottom panel). In addition, the aggregated replicating mtDNA failed to localize to the tips of mitochondria in VPS13D-suppressed cells (Figure 4D, Bottom panel), suggesting defects in synthesis and distribution of replicating mtDNA. In addition, suppression of dynamin-related protein 1 (Drp1) or mitochondrial fission factor (Mff) significantly decreased the number of replicating mtDNA and caused the aggregation of replicating mtDNA (Figure 4E; Supplemental Figure S7, A–D), resembling the phenotype observed in VPS13D-suppressed cells. Suppression of MFN2 also significantly altered the density and size of replicating mtDNA, but to a lesser extent than that resulting from the suppression of Drp1 or Mff (Figure 4E; Supplemental Figure S6, E and F). Therefore, our results suggest that VPS13D suppression might lead to defects in mtDNA synthesis and distribution through affecting mitochondrial dynamics.

Plasmids and siRNA oligonucleotides

mCherry-mito7 (55102), mitoBFP (49151), p97-GFP (23971), pRK5-HA-ubiquitin-WT (17608), pRK5-HA-ubiquitin-K48 (17605), and pRK5-HA-ubiquitin-K63 (17606) were purchased from Addgene. eBFP-peroxisome was previously described (Ji et al., 2017). ER-tagRFP, ER-mGFP, polG2-GFP, mStrawberry-Rab4b, mStrawberry-Rab7A, CFP-VAPB, and PTPIP51-TagBFP2 were gifts from Henry N. Higgs (Dartmouth Geisel School of Medicine). Lamp1-OFP and B4GALT1-GFP was purchased from Sinobiological. ACSL3-Halo was generated by cutting the ACSL3 ORF from pEFIRES-P-ACSL3-mCherry (Addgene 87158) and pasting it into Halo-N1 vector, which was generated by replacing GFP with Halo in mGFP-N1 (Addgene 54767). The construct for quantification of the extent of ER–mitochondria contacts was generated by inserting the mitochondrial targeting sequence fused in frame with GFP_1–10 (tom20-GFP_1–10, a gift from Tito Calì at the University of Padua) and the ER targeting sequence of Sac1 fused in frame with GFP₁₁ (Sac1-long linker-GFP₁₁, a gift from Tito Calì at the University of Padua) into two MCS sites of pCW57-MCS1-P2A-MCS2 (Addgene 89180) under control of a doxycycline-inducible promoter. The Lentivirus packaging plasmids psPAX (12260) and pMD2. G (12259) were bought from Addgene. All of the truncated mutants of VPS13D or p97 used in this study were generated by PCR using human VPS13D-myc-flag (Origene, RC205601) or p97-GFP (Addgene, 23971) as templates and cloning them into mGFP-N1 (Addgene 54767), mGFP-C1 (Addgene 54579), or the 3*flag vector, generated by replacing GFP with a 3*flag tag in mGFP-N1 (Addgene 54767). All of constructs used in this study were generated by using a ClonExpress II One Step Cloning kit (Vazyme, C112-01). Oligonucleotides for human VPS13D siRNAs were synthesized by RiboBIO against target sequences 5′-GCAGAAGGATCCTTAAAGA-3′ (siRNA#1); 5′-GTAGCACGTTTGACATGAA-3′ (siRNA#2); and 5′-CAAAGAGCCTCTACTATGA-3′ (siRNA#3). Oligonucleotides for VPS13A were against sequences 5′-GGATGGGACTGGAAATCAA-3′ (siRNA#1); 5′-GCAGCTACATTCCTCTTAA-3′ (siRNA#2); and 5′-GGATAGAGCTTATGATTCA-3′ (siRNA#3). Oligonucleotides for p97 were against sequences 5′-GAAUAGAGUUGUUCGGAAU-3′ (siRNA#1); and 5′-GGAGGUAGAUAUUGGAAUU-3′ (siRNA#2). Oligonucleotides for Drp1 were against sequences 5′-GGAACAAAGTATCTTGCTA-3′ (siRNA#1); 5′-GAAGGGTTATTCCAGTCAA-3′ (siRNA#2); and 5′-GGTTGGAGATGGTGTTCAA-3′ (siRNA#3). Oligonucleotides for Mff were synthesized by RiboBIO against sequences 5′-GGAGAGGATTGTTGTAGCA-3′ (siRNA#1); 5′-GTACTTACGCTGAGTGAAA-3′ (siRNA#2); and 5′-GTAGCTTTCTGGCTGCTTA-3′ (siRNA#3). Oligonucleotides for Mfn2 were against sequences 5′-GGCCAAACATCTTCATCCT-3′ (siRNA#1); 5′-CGGTTCGACTCATCATGGA-3′ (siRNA#2); and 5′-GCGAGGAAATGCGTGAAGA-3′ (siRNA#3). Oligonucleotides for VAPB were against sequences 5′-CCAACAGCGGAATCATCGA-3′ (siRNA#1); 5′-ACAGCACCACGTAGGTACT-3′ (siRNA#2); and 5′-CCGATGTTGTCACCACCAA-3′ (siRNA#3). Oligonucleotides for PTPIP51 were against sequences 5′-GAAGCTAGATGGTGGATGA-3′ (siRNA#1); 5′-GGATGAGAGTGCTGACTGT-3′ (siRNA#2); and 5′-GCGGGACTCTGACAAAGAA-3′ (siRNA#3). As a control, scrambled siRNA (Negative Control), 5′-CGUUAAUCGCGUAUAAUACGCGUAT-3′ (RiboBIO) was used.

Antibodies and reagents

Anti-GFP (1:1000, 50430-2-AP, ProteinTech), anti-Mff (17090-1-AP; ProteinTech), anti-Drp1 (#8570; CST), anti-Mfn2 (12186-1-AP; ProteinTech), anti-PTPIP51 (20641-1-AP; ProteinTech), anti-calnexin (10427-1-AP; ProteinTech), anti-COXIV (11242-1-AP; ProteinTech), anti-flag (F1804; Sigma), and anti-p97/VCP (10736-1-AP; ProteinTech) were used at 1:1000 dilutions for Western blots. Anti-VAPB (14477-1-AP; ProteinTech) was used at 1:1000 for Western blots and 1:200 for immunofluorescence. Anti-tom20 (sc-17764; Santa Cruz Biotechnology), anti-tubulin (11224-1-AP; ProteinTech), and anti-actin (60008-1-Ig, ProteinTech) were used at 1:5000 dilutions for Western blots. Anti-VPS13D (A304-691A; BETHY Laboratories) was used at 1:50 dilutions for coimmunoprecipitation and 1:100 for Western blots. The reagents NMS873 (TargetMol, T1853) and BODIPY 493/503 (Thermo Fisher, D3922) were used. Antibiotics such as G418 (10131027), puromycin (A1113803), hygromycin B (10687010), and blasticidin (A1113903) were obtained from Thermo Fisher (U.S.).

Cell culture, transfection, and stable lines

Human osteosarcoma U2OS cells (ATCC), human embryonic kidney HEK293 cells (ATCC), African green monkey kidney fibroblast-like COS7 cell line (ATCC), and human cervical cancer HeLa cells (ATCC) were grown in DMEM (Invitrogen) supplemented with 10% fetal bovine serum (Life Technologies) and 1% penicillin/streptomycin. All of the cell lines used in this study are free of mycoplasma contamination. For transfection, cells were seeded at 4 × 10⁵ cells per well in a six-well dish ∼16 h before transfection. Plasmid transfections were performed in OPTI-MEM (Invitrogen) with 2 µl Lipofectamine 2000 per well for 6 h, followed by trypsinization and replating onto glass-bottomed confocal dishes at ∼3.5 × 10⁵ cells per well. Cells were imaged in live-cell medium (DMEM with 10%FBS and 20 mM HEPES no P/S) ∼16–24 h after transfection. For all transfection experiments in this study, the following amounts of DNA were used per 3.5-cm well (individually or combined for cotransfection): 500 ng for mito-BFP, eBFP2-peroxisome, mCherry-mito7, B4GALT1-GFP, ACSL3-Halo, and Lamp1-OFP; 1000 ng for ER-tagRFP, ER-mGFP, and p97-GFP; 300 ng for mStrawberry-Rab4b and mStrawberry-Rab7A. For siRNA transfections, cells were plated on 3.5-cm dishes with density 30–40%, and 2 µl lipofectamine RNAimax (Invitrogen) and 50 ng siRNA were used per well. At 48 h after transfection, a second round of transfection was performed with 50 ng siRNAs. Cells were analyzed 24 h post–second transfection for suppression. For the generation of the stable Dox-inducible ER–mitochondria MCSs reporter U2OS cell line: day 1: 293T cells were seeded into 10-cm dish with ∼3 × 10⁶ cells 18 h before transfection; day 2: transfections were performed with 4 μg PCW-Tom20-GFP_1-10-P2A-Sac1-long linker-GFP₁₁, 3 μg psPAX2, and 2 μg pMD2.G; day 3: U2OS cells were seeded into 10-cm dishes with 2 × 10⁶ cells; day 4: the supernatant from transfected HEK293 cells was collected 48 h post–initial transfection, followed by filtering through a syringe with 0.22-μm filters, and 1 ml filtered HEK293 supernatant was added to U2OS cells; day 5: U2OS cells were grown in tetracycline-free medium containing G418 (2 μg/ml), followed by verification by imaging and Western blots. To make a polG2-GFP U2OS cell line, we transfected U2OS cells with polG2-GFP construct by lipofectamine 2000. At 24 h after transfection, cells were grown in G418 (1 μg/ml)-containing growth medium for another week, followed by fluorescence-activated cell sorting (FACS).

Live imaging by confocal microscopy

Cells were grown on glass-bottomed confocal dishes. Confocal dishes were loaded into a laser scanning confocal microscope (LSM780, Zeiss, Germany) equipped with multiple excitation lasers (405, 458, 488, 514, 561, and 633 nm) and a spectral fluorescence GaAsP array detector. Cells were imaged with the 63 × 1.4-NA iPlan-Apochromat 63× oil objective using the 405-nm laser for BFP, 488-nm for GFP, and 561-nm for mStrawberry, OFP, tagRFP, or mCherry. To track the dynamics of mitochondria and ER, cells on confocal dishes were loaded into a spinning-disk confocal microscope (Olympus IX83, Japan) equipped with 100×/1.4 oil (WD 013 mm, DIC slider) objective, multiple lasers (405, 488, 561, and 640 nm) with corresponding filters, and EMCCD and scMOS cameras.

Antibody preclearing

For endogenous VPS13D IF, anti-VPS13D antibody was first cleared according to the protocol described in (Valverde et al., 2019). Briefly, VPS13D-depleted HEK293 cells were fixed in 4% paraformaldehyde (PFA) for 15 min, rinsed with phosphate-buffered saline (PBS), quenched in 50 mM NH₄Cl in PBS, washed twice with PBS, permeabilized with PBX (PBS containing 0.1% Triton X-100) for 10 min, and scraped in PBX with 1% Triton X-100. Antibodies were added to the fixed cells at the final concentration used for IF (1:100) and mixed overnight by rotation at 4°C. The mixture was then centrifuged at 17,000 × g for 20 min at 4°C, and the supernatant containing cleared antibodies was used for IF.

Quantitative RT-PCR for detecting the mRNA level of VPS13D and VPS13A

U2OS cells were transfected with either scrambled, VPS13D, or VPS13A siRNAs. At 4 d after transfection, RNA was isolated with Trizol (Thermo Fisher) according to the instructions of the manufacturer. cDNA was reverse transcribed using RverTra Ace (TRT-101, TOYOBO) according to the directions of the manufacturer. The cDNA was analyzed using quantitative PCR with qPCR Mix (QPS-201, TOYOBO) using the following primers: VPS13D-1: 5′-GTGACTGACAACAGATACGAGCCA-3′; 5′-CGACCAACCCATGTAACCCA-3′; VPS13D-2: 5′-TCGATATGCAGCACCAGATAAA-3′; 5′-CTGGTGGGAGTGAAGAG­AAATAA-3′; VPS13D-3: 5′-GGATTTGCCTCAGATGGAGTTAC-3′; 5′-TCCAGGACGTAGTGATGGCT-3′; VPS13A-1: 5′-CGGTGCTATGGCTAAG-3′; 5′-CCTCCACGAGTGATGC-3′; VPS13A-2: 5′-TGGGCTTATGCTATACATGGCG-3′; 5′-CCAAAGACACGAGAAGTTCACC-3′; VPS13A-3: 5′-TGTGATGAATCGCCGATCTGA-3′; 5′-CCATGTATAGCATAAGCCCACCA-3′.

Immunofluorescence staining

Cells were fixed with 4% PFA (Sigma) in PBS for 10 min at room temperature. After being washed three times with PBS, cells were permeabilized with 0.1% Triton X-100 in PBS for 15 min on ice. Cells were then washed three times with PBS, blocked with 0.5% BSA in PBS for 1 h, incubated with primary antibodies in diluted blocking buffer overnight, and washed three times with PBS. Secondary antibodies were applied for 1 h at room temperature. After being washed three times with PBS, samples were mounted on Vectashield (H-1000; Vector Laboratories).

Image analysis

All image analysis and processing were performed using ImageJ (National Institutes of Health). Colocalization-based analysis of ER–mitochondria contacts was performed by a colocalization plugin (imageJ, NIH) with the following settings: ratio (0–100%): 50; threshold channel 1 (0–255): 50; threshold channel 2 (0–255): 50; display value (0–255): 255. ER–mitochondria intersection sites were automatically identified by the colocalization plugin, with white pixels representing potential ER–mitochondria contact sites. For mitochondrial perimeter measurement, an imageJ macro named the mitochondrial morphology macro (Dagda et al., 2008) was used to measure the perimeter of mitochondria in regions of interest (ROIs) where mitochondria are well resolved. For the measurement of polG2-GFP–labeled replicating mitochondrial DNA density in mitochondria, two steps were performed. Step1: measure the number of polG2-GFP puncta by cell counter (imageJ plugin, NIH) in ROIs; step2: measure the mitochondrial perimeters of corresponding ROIs. The density of replicating mitochondrial DNA was calculated as the number of polG2-GFP puncta divided by the perimeter of mitochondria in a ROI.

Differential centrifugation

Differential centrifugation was a modified version of the protocol previously described (Ji et al., 2017). Cells were harvested from 2 × 10 cm² dishes at 90% confluency. The following steps were conducted at 4°C or on ice. Cells were washed with precold PBS twice and homogenized in isolation buffer (225 mM mannitol, 75 mM sucrose, 0.1 mM EGTA, 30 mM Tris-HCl pH7.4) by Dounce (Wheaton Dura-Grind). The homogenate was centrifuged twice for 10 min at 600 × g to remove nuclei and debris. The resulting supernatant was centrifuged three times for 15 min at 13,000 × g to obtain crude mitochondria. Western blotting was performed using rabbit anti-VAPB (1:1000, 14477-1-AP, ProteinTech), rabbit anti-PTPIP51 (1:1000, 20641-1-AP, ProteinTech), rabbit anti-p97 (1:1000, 10736-1-AP, ProteinTech), rabbit anti-calnexin (1:500, 10427-2-AP, ProteinTech), anti-Lamp1 (1:1000; sc-20011;Santa Cruz Biotechnology), anti-Rab7A (1:1000; A12784; ABclonal), anti-HSP60 (1:1000; 15282-1-AP; ProteinTech), and rabbit anti-COXIV (1:1000, 11242-1-AP, ProteinTech) antibodies.

Electron microscopy

Scrambled or VPS13D siRNA–treated U2OS cells were fixed with 2.5% glutaraldehyde in 0.1 M phosphate buffer, pH 7.4 for 2 h at room temperature. After being washed three times with 0.1 M phosphate buffer, cells were scraped and collected with 0.1 M phosphate buffer followed by centrifugation at 3000 rpm. The pellet was resuspended in PBS (0.1 M) and centrifuged at 3000 rpm for 10 min. This step was repeated three times. The samples were postfixed with precold 1% OsO4 in 0.1 M phosphate buffer for 2–3 h at 4°C, followed by rinsing three times with PBS for 20 min. The samples were dehydrated in graded ethanol (50%, 70%, 85%, 90%, 95%, 2 × 100%) for 15 min for each condition. The penetrations were performed in the order acetone-epoxy (2:1); acetone-epoxy (1:1); epoxy. Each round of penetration was performed at 37°C for 12 h. The samples were embedded in epoxy resin using standard protocols (Kumar et al., 2014). Sections parallel to the cellular monolayer were obtained using a Leica EM UC7 with a thickness of 60–100 nm and examined under a Tecnai G2 20 TWIN (FEI) with accelerating voltage 200 kV. Mitochondria and ER were identified based on their respective morphologies and were traced by hand. The extension length of ER–mitochondria contact regions was measured using the “measure” tool in ImageJ (NIH)s. Mitochondria with ER within 30 nm at any point on the mitochondrial circumference were identified as contact sites.

Green fluorescent protein–trap assay

GFP-trap assays performed according to the manufacturer’s protocol (GFP-trap agarose beads, ChromoTek) were used for detection of protein–protein interactions. The 5% input was used in GFP traps unless otherwise indicated.

Statistical analysis

All statistical analyses and p-value determinations were performed in GraphPad Prism6. To determine p-values, ordinary one-way ANOVA with Turkey’s multiple comparisons test was performed among multiple groups and a two-tailed unpaired Student’s t test was performed between two groups. Chi-squared analysis followed by Fisher’s exact tests was performed in Figure 4C and Supplemental Figure S3C.

Data availability

All the data and relevant materials, including constructs and primers, that support the findings of this study are available from the corresponding author upon reasonable request.