AGPAT2 acts at the crossroads of lipid biosynthesis and DRP1-mediated ER morphogenesis

Domain: physics
Published: 2025-12-12
Editorial outcome: major_revision

Abstract
The morphology of the endoplasmic reticulum (ER), characterized by central sheets and peripheral tubules, is controlled by membrane-shaping proteins. However, the role of lipids in ER morphogenesis remains elusive, despite the ER being the major site for lipid synthesis. Here, by examining the role of eighteen phosphatidic acid (PA)-generating enzymes in ER morphology, we identify lysophosphatidic acid acyltransferase 2 (AGPAT2) as a critical factor in mouse and human cells. AGPAT2 produces PA in the glycerophospholipid/triacylglycerol biosynthesis pathway, and its mutations cause congenital generalized lipodystrophy. We find that AGPAT2-generated PA drives ER tubulation through gene knockout, 3D structural analysis by FIB-SEM, super-resolution microscopy, lipidomics, AlphaFold, and in vitro reconstitutions of ER tubulation and AGPAT2 activity. AGPAT2 interacts with and supplies PA to the PA-binding, dynamin-related GTPase, DRP1, which subsequently tubulates the ER in a manner independent of GTP hydrolysis and oligomerization, distinct from its function in mitochondrial division. Consistently, the reduction of PA levels by ectopic expression of a PA phosphatase, LIPIN1, transforms ER tubules into sheets. Our results reveal an unforeseen interplay between lipid biosynthesis and membrane organization in the ER.

Introduction
The ER establishes a unique morphology with sheets and tubules and plays crucial roles in cellular functions1–9. ER sheets, contiguous to the nuclear membrane, create rough ER which is essential for protein translation and post-translational modification. In contrast, ER tubules, which are involved in lipid synthesis and signaling, form dynamic platforms, that interact with other organelles such as the plasma membrane, mitochondria, endosomes, lipid droplets, and P-bodies10–18.

Previous studies have elucidated mechanisms that contribute to ER morphology through multiple ER-resident integral membrane proteins. These include reticulons and DP1/Yop1p, responsible for tubule formation; atlastin, which fuses ER tubules; lunapark/Lnp1p, crucial for the formation of three-way junctions in ER tubules; and Climp-63, which is involved in the formation of ER sheets19–22. However, in contrast to these protein factors, the role of lipids in ER morphology is unknown.

Dynamin-related protein 1 (DRP1) is a soluble protein and key member of the dynamin superfamily, known for severing the membranes of mitochondria and peroxisomes23–30. Beyond these organelles, DRP1 also localizes to the ER, where it contributes to the tubulation of the ER membrane31,32. The mechanism through which DRP1 induces ER tubulation differs significantly from its role in mitochondrial division. Mitochondrial division is governed by the oligomerization and GTP hydrolysis activities of DRP1. In contrast, ER tubulation does not depend on these activities31. Instead, an 18-amino-acid region in the variable domain (D-octadecapeptide) drives ER tubulation31. Therefore, DRP1 utilizes a unique mechanism for membrane deformation and ER tubulation. Notably, both DRP131 and D-octadecapeptide (Supplementary Fig. 1) preferentially bind to phosphatidic acid (PA). However, whether PA is required for ER morphogenesis remains unknown.

In this study, we performed a gene knockdown screen targeting enzymes responsible for PA production at the ER. Our data revealed that a specific lysophosphatidic acid acyltransferase, AGPAT2, cooperates with DRP1 to regulate ER morphology.

Results
AGPAT2 is required for ER morphogenesis
PA is produced through three major biosynthetic pathways: two phospholipases D (PLDs) generate PA from phosphatidylcholine (PC), and six lysophosphatidic acid acyltransferases (AGPATs) produce PA by adding a fatty acid to lysoPA from acyl-CoA (Fig. 1a)33. To test whether any of these enzymes are required for ER morphogenesis, we individually knocked down all 18 PA-producing enzymes in mouse embryonic fibroblasts (MEFs) using esiRNAs and siRNAs (Supplementary Fig. 2a). As a negative control, esiRNA targeting RLuc was used. ER morphology was assessed by laser confocal immunofluorescence microscopy of MEFs expressing an ER marker, GFP-SEC61β, using anti-GFP antibodies (Supplementary Fig. 2b). We previously reported that ER tubules constitute ~50% of the ER in wild-type (WT) MEFs31. In the initial screen, we set a threshold of approximately half the WT level as the criterion for a positive hit. This degree of reduction can be readily identified by visual inspection without quantification. Among the 18 enzymes tested, only knockdown of a lysophosphatidic acid acyltransferase (1-acyl-sn-glycerol-3-phosphate acyltransferase beta, AGPAT2) met this criterion in two experimental replicates (Supplementary Fig. 2b).

The role of AGPAT2 in ER morphology was further confirmed by shRNA knockdown (Fig. 1b, c, e), CRISPR/Cas9-mediated knockout (KO) in MEFs (Fig. 1f, g; and Supplementary Fig. 3), and esiRNA-mediated knockdown in human U2OS cells (Fig. 1h–j). Altered ER morphology in AGPAT2-KO cells was rescued by expressing AGPAT2-HA, demonstrating its specificity (Fig. 1f, g). To analyze the 3D ultrastructure of the peripheral ER, we performed focused ion beam scanning electron microscopy (FIB-SEM)34 on WT and AGPAT2-KO MEFs, along with DRP1-KO MEFs, in which ER tubulation is decreased31 (Supplementary Fig. 4). Consistent with the data from laser confocal immunofluorescence microscopy, 3D reconstruction of the ER structure demonstrated ER sheets at the cell periphery in AGPAT2-KO and DRP1-KO cells, in contrast to WT cells, which contained a network of tubules of the ER (Fig. 2a, b, and Supplementary Movies 1–3).

AGPAT1 is another lysophosphatidic acid acyltransferase closely related to AGPAT233. In the knockdown screen, AGPAT1 esiRNA did not affect ER morphology (Supplementary Fig. 2b). This result was confirmed using shRNA (Fig. 1b–d). In addition, further knockdown of AGPAT1 did not affect ER morphology in AGPAT2-KO MEFs (Supplementary Fig. 5a–c). Moreover, the altered ER morphology in AGPAT2-KO MEFs was not rescued by the expression of AGPAT1 (Supplementary Fig. 5d, e). Collectively, these data show that AGPAT1 is not involved in the regulation of ER morphology.

Since ER stress affects ER morphology35, we assessed ER stress by examining markers such as BIP and CHOP expression levels, as well as phosphorylation of eIF2α. The results show that ER stress is not elevated in AGPAT2-KO cells compared to WT cells (Fig. 3). As a positive control, we treated cells with tunicamycin and confirmed robust activation of the ER stress response (Fig. 3).

The topology of AGPAT2
AGPAT2 is an integral membrane protein with a single predicted transmembrane domain in the ER based on TmAlphaFold36,37 (Fig. 4a). To determine its topology, we expressed C-terminally FLAG-tagged AGPAT2 in MEFs (Fig. 4b). As a control, we co-expressed SEC61β, which is fused to HA at its C-terminus facing the ER lumen (Fig. 4b). We then performed laser confocal immunofluorescence microscopy with antibodies to FLAG and HA under different membrane permeabilization conditions using saponin (which selectively permeabilizes the plasma membrane) or Triton X-100 (which permeabilizes both the plasma and ER membranes). In the absence of detergents, we observed essentially no staining of AGPAT2-FLAG or SEC61β-HA (Fig. 4b–d). When the plasma membrane was permeabilized with saponin, we detected AGPAT2-FLAG but not SEC61β-HA (Fig. 4b–d). Conversely, when all membranes were permeabilized with Triton X-100, we detected both AGPAT2-FLAG and SEC61β-HA (Fig. 4b–d). These data suggest that the C-terminus of AGPAT2 faces the cytosol (Fig. 4e).

AGPAT2 is preferentially localized at ER tubules
ER-shaping proteins show preferential localization to specific ER subdomains. For instance, the ER tubulating protein, reticulon 4A (RTN4A), is enriched in ER tubules, while the ER sheet-forming protein Climp-63 is predominantly found in sheets19. To investigate the intra-ER localization of AGPAT2, we compared the localization of AGPAT2-FLAG with that of RTN4A-GFP in cells expressing mCherry-SEC61β, which is uniformly distributed in the ER membrane (Fig. 4f–i). We found that AGPAT2-FLAG was preferentially enriched in ER tubules (Fig. 4f, g), to a degree similar to RTN4A-GFP (Fig. 4h, i). This localization, together with the knockdown and knockout phenotypes showing decreased ER tubules, suggests that AGPAT2 plays a significant role in ER tubulation.

AGPAT2 functions in ER tubulation together with DRP1
Two ER tubulating proteins, DRP1 and the reticulon RTN4A, function through distinct mechanisms31. RTN4A induces ER tubulation by oligomerizing and inserting wedges into the ER membrane22, while DRP1 facilitates ER tubulation by inserting D-octadecapeptide in the variable domain31. To determine whether AGPAT2 works together with RTN4A or DRP1 in ER morphogenesis, we ectopically expressed AGPAT2-FLAG in WT, RTN4A-KO, and DRP1-KO cells (Fig. 5a, b). Complementing the data from AGPAT2 depletion and localization studies, the ectopic expression of AGPAT2 increased the levels of ER tubules in WT cells (Fig. 5a, b; WT). Similarly, in RTN4-KO cells, the overexpression of AGPAT2 led to increased ER tubules, indicating that RTN4A is not required for AGPAT2-induced ER tubulation (Fig. 5a, b; RTN4A-KO). In contrast, the overexpression of AGPAT2 in DRP1-KO cells did not induce ER tubulation (Fig. 5a, b; DRP1-KO). Therefore, AGPAT2’s ability to mediate ER tubulation depends on DRP1. Taken together, these data suggest that AGPAT2 and DRP1 collaboratively contribute to ER tubule formation within the same pathway. Further supporting this notion, DRP1 overexpression increased ER tubules in RTN4A-KO cells but not in AGPAT2-KO cells (Fig. 5c, d), and RTN4A overexpression induced ER tubulation in AGPAT2-KO cells (Fig. 5e, f).

To test whether the loss of AGPAT2 affects the expression levels of reticulons, we analyzed their protein abundance by Western blotting. Among the four reticulons in mice, RTN3 and RTN4 are highly expressed in MEFs, whereas RTN1 and RTN2 are expressed at very low levels or are undetectable (www.ebi.ac.uk/gxa/home). Therefore, we examined RTN3A and RTN4A levels in WT and AGPAT2-KO MEFs. We found that RTN3A levels remained unchanged between genotypes, while RTN4A levels were modestly increased in AGPAT2-KO MEFs (Supplementary Fig. 6). We speculate that this upregulation may represent a compensatory response to reduced ER tubules in AGPAT2-KO cells. We also examined the level of the sheet-forming protein Climp-63 and found it to be reduced in AGPAT2-KO MEFs (Supplementary Fig. 6), which may similarly reflect a compensatory mechanism.

In vitro reconstitution of AGPAT2- and DRP1-mediated ER tubulation
To further investigate the functional relationship between AGPAT2 and DRP1, we developed an in vitro reconstitution of ER tubulation using purified ER and recombinant DRP1 (Fig. 6). We isolated ER from WT, DRP1-KO, and AGPAT2-KO MEFs and incubated them with DRP1. To distinguish DRP1’s function in mitochondrial division, GTP was omitted from the reaction. Transmission electron microscopy revealed that purified WT ER contains both sheets and tubules and the addition of DRP1 did not alter these structures (Fig. 6a, b). In contrast, isolated DRP1-KO ER predominantly displayed sheet structures without tubules, and incubation with DRP1 induced tubulation of the membrane (Fig. 6a, b). As a negative control, recombinant GFP did not tubulate the ER (Fig. 6c). The intracellular concentration of DRP1 has been estimated to be 0.5 μM38. DRP1 significantly induced ER tubulation below this concentration (0.3 μM) in our assay (Fig. 6d, e). Similar to DRP1-KO ER, AGPAT2-KO ER primarily exhibited sheet morphology. However, in sharp contrast to DRP1-KO ER, DRP1 was unable to induce tubulation of AGPAT2-KO ER membranes (Fig. 6a, b). These data indicate that AGPAT2 is essential for DRP1 to tubulate the ER membrane in vitro.

We previously reported that DRP1’s GTPase activity and oligomerization are not required for ER tubulation in cells31. To test whether these activities are also dispensable in vitro, we used recombinant DRP1 harboring the K38A and G350D mutations, which impair GTP hydrolysis and oligomerization, respectively31. The mutant protein tubulated isolated DRP1-KO ER as effectively as the WT protein (Fig. 6f, g). It has been shown that tubules generated from liposomes and purified DRP1 measure ~60 nm in diameter and become constricted to ~40 nm upon GTP hydrolysis, a change proposed to represent a step in mitochondrial division39. Importantly, the width of purified ER tubules was ~30 nm, narrower than these reported values (Fig. 6h). Moreover, ER tubules induced by both WT and mutant DRP1 were also ~30 nm (Fig. 6h). These findings indicate that neither GTP hydrolysis nor oligomerization is required for DRP1-mediated ER tubulation in vitro, distinguishing this process from mitochondrial division.

To determine whether PA is responsible for the defect in DRP1-mediated tubulation of AGPAT2-KO ER, we added soluble 8:0 PA and DRP1 to in vitro reactions using AGPAT2-KO/DRP1-depleted ER. PA restored ER tubulation in a dose-dependent manner, but only when both PA and DRP1 were present (Supplementary Fig. 7a, b). In contrast, no such structures were observed in a mixture of DRP1 and PA alone, without ER, as a negative control (Supplementary Fig. 7c). These findings indicate that AGPAT2 is required to supply PA for DRP1-mediated ER membrane tubulation. It remains possible that 8:0 PA inserted into the membrane and stimulated DRP1-dependent tubulation, or alternatively, that PA activated DRP1 as a soluble lipid without membrane integration.

To ask if ER tubules in WT ER membranes are generated by endogenous DRP1, we incubated WT ER with antibodies against DRP1 or, as a negative control, TOM20. Incubation with the DRP1 antibody—but not the TOM20 antibody—led to the loss of ER tubules (Fig. 6i, j). Therefore, endogenous DRP1 is required to maintain ER tubular structures. We speculate that the antibody-mediated inhibition does not occur through direct interference with membrane-embedded regions of DRP1, but rather through binding to other domains, thereby altering its conformation to one less favorable for membrane association or deformation.

The catalytic activity of AGPAT2 is critical for ER tubulation
To determine whether the enzymatic activity of AGPAT2 is required for maintaining ER morphology, we introduced mutations in the conserved catalytic motifs of AGPAT2-HA and expressed the mutants in AGPAT2-KO MEFs33,40,41 (Fig. 7a–c). While WT AGPAT2 successfully rescued the ER morphology in the KO cells, all the mutants failed to do so (Fig. 7a–c). Hence, the PA-producing activity of AGPAT2 is critical for maintaining proper ER morphology. To assess the role of AGPAT2 in PA production, we developed a biochemical assay to measure the activity of PA production from lysoPA in vitro by incubating purified ER membranes with two substrates of AGPAT2: fluorescent lysoPA-TopFlour and acyl-CoA (Fig. 7d). We then analyzed the production of PA-TopFlour using thin-layer chromatography. Our results indicated that PA production is greatly decreased in AGPAT2-KO ER membranes compared to WT ER membranes, underscoring the significance of AGPAT2 in PA production within the ER (Fig. 7e, f).

To further investigate whether PA is critical for ER tubulation, we ectopically expressed WT HA-LIPIN1 in AGPAT2-KO MEFs that were also expressing AGPAT2-FLAG. LIPIN1 catalyzes the dephosphorylation of PA to DAG, thus its overexpression is expected to downregulate PA-mediated functions42 (Fig. 7g). Our results showed that the expression of WT HA-LIPIN1 significantly reduced the number of ER tubules (Fig. 7h, i; HA-LIPIN1). In contrast, a catalytically-inactive mutant of LIPIN1-HA did not influence the ER tubules42 (Fig. 7h, i; D712A). Thus, these data indicate that PA, produced by AGPAT2, plays a pivotal role in maintaining ER morphology.

We confirmed that LIPIN1 overexpression decreases PA levels using lipid mass spectrometry (Supplementary Fig. 8a). In contrast, DAG levels remained unchanged (Supplementary Fig. 8a). One possibility is that although LIPIN1 overexpression increases DAG production, the DAG is rapidly converted into downstream products such as triacylglycerols or other phospholipids. Alternatively, because DAG is 30–40 times more abundant than PA (Supplementary Fig. 8a), the conversion of PA to DAG may not be detectable upon LIPIN1 overexpression.

AGPAT2 and DRP1 interact
PA is predominantly synthesized in the ER and serves as a major precursor for the biosynthesis of phospholipids and triacylglycerol43–47. Remarkably, PA is dynamic, short-lived, and stealthy in nature—large amounts are produced and then rapidly transformed into other lipids or transported to mitochondria43–45,48. Consistent with the exceptionally transient nature of PA, only small amounts are detected in the ER by mass spectrometry (Supplementary Fig. 8b), and it is undetectable by the PA biosensor PASS at steady state45,49–52 (Supplementary Fig. 9). To address how this stealthy lipid is supplied to ER morphogenesis, we tested whether AGPAT2 interacts with DRP1 through co-immunoprecipitation. We found that AGPAT2-FLAG co-immunoprecipitated with endogenous DRP1 (Fig. 8a, b). Furthermore, the co-immunoprecipitation of endogenous AGPAT2 and DRP1 was also confirmed in human HEK293T cells without ectopic expression (Fig. 8c, d). In contrast, AGPAT1, a closely related enzyme to AGPAT2, showed less co-immunoprecipitation with DRP1 (Fig. 8e, f). These results suggest that AGPAT2 is in close proximity to DRP1.

The interaction with AGPAT2 and its generated PA does not seem to be required for the ER localization of DRP1. We examined the localization of DRP1 in WT and AGPAT2-KO MEFs using immunofluorescence super-resolution Airyscan microscopy with anti-DRP1 antibodies. In WT MEFs, 3D reconstruction of images revealed that DRP1 is associated with the ER marked by mCherry-SEC61β, displaying a snake-like pattern (Fig. 8g, h, and Supplementary Movie 4). In AGPAT2-KO cells, DRP1 also exhibited an association with the ER (Fig. 8i, j, Supplementary Movie 5). Furthermore, biochemical fractionation confirmed indistinguishable co-fractionation of DRP1 with the ER in both WT and AGPAT2-KO cells (Fig. 8k, l). These data suggest that AGPAT2 and PA are involved in the functions of DRP1 but not its localization.

PA modulates the AGPAT2-DRP1 interaction
To further examine the interaction of AGPAT2 with DRP1, we expressed three catalytically inactive mutants of AGPAT2-FLAG in AGPAT2-KO cells and performed co-immunoprecipitation with endogenous DRP1. All three mutants co-immunoprecipitated with DRP1 much less efficiently than WT AGPAT2-FLAG did (Fig. 9a, b). These data suggest that either the mutations affect the conformation of AGPAT2, disrupting its co-immunoprecipitation with DRP1, or that AGPAT2’s enzymatic activity (i.e., the production of PA) is important for the AGPAT2-DRP1 co-immunoprecipitation. To test these two models, we expressed the PA phosphatase LIPIN1 in AGPAT2-KO cells carrying WT AGPAT2-FLAG. Results showed that WT LIPIN1, but not the enzymatically inactive mutant LIPIN1 (D172A), decreased co-immunoprecipitation between WT AGPAT2-FLAG and DRP1 (Fig. 9c, d). These data suggest that AGPAT2-produced PA plays an important role in the interaction of AGPAT2 and DRP1.

To understand how PA affects the AGPAT2-DRP1 interaction, we first compared the co-immunoprecipitation of AGPAT2-FLAG with GFP fused to full-length DRP1 (GFP-FL) and DRP1 consisting of the GTPase and stalk domains, lacking the PA-binding variable domain53 (GFP-GS) (Fig. 9e–g). We used DRP1-KO HEK293T cells to avoid the contribution of endogenous DRP1. WT AGPAT2 similarly co-immunoprecipitated with both GFP-FL and GFP-GS (Fig. 9f, g). Consistent with these data, AlphaFold3 suggests that AGPAT2 interacts with DRP1 near the junction of the GTPase and stalk domains53 (Fig. 9h). Distinct from GFP-FL, which showed reduced co-immunoprecipitation with the mutant AGPAT2 (H98A, D103N), GFP-GS lacking the variable domain did not show reduced co-immunoprecipitation with the mutant (Fig. 9f, g). These data suggest that the variable domain inhibits the interaction of DRP1 with AGPAT2 when AGPAT2 does not produce PA.

We hypothesize that the GTPase and/or stalk domains interact with AGPAT2 while the variable domain regulate such interactions. Indeed, GFP-GTPase domain (GFP-G) or GFP-stalk domain alone (GFP-S) co-immunoprecipitated with AGPAT2 (Fig. 9i, j). Further supporting the inhibitory role of the variable domain, both GFP-G and GFP-S co-immunoprecipitated similarly with WT and the mutant AGPAT2 (H98A, D103N) (Fig. 9i, j). In contrast, the variable domain does not co-immunoprecipitate with AGPAT2 (Fig. 9k, l). These data suggest that the variable domain promotes the association of DRP1 with AGPAT2 when AGPAT2 produces PA, creating a robust mechanism by which PA enhances both the DRP1–AGPAT2 interaction and ER tubulation.

Since the variable domain regulates DRP1 oligomerization54, we investigated whether oligomerization contributes to PA-mediated regulation of the DRP1–AGPAT2 interaction. We performed a co-immunoprecipitation assay using the oligomerization-defective G350D mutant and found that DRP1 (G350D) interacts more strongly with enzymatically inactive AGPAT2 (H98A, D103N) than WT DRP1 (Supplementary Fig. 10). These results suggest that inhibition involves DRP1 oligomerization. Due to the unstructured nature of the variable domain, it is poorly resolved in existing structural models of DRP1, and how this domain interacts with or influences the GTPase and stalk domains remains largely unknown55–58. Nonetheless, oligomerization may facilitate intermolecular interactions within DRP1 oligomers such that the variable domain physically hinders interaction with AGPAT2. Alternatively, oligomerization may induce conformational changes in the GTPase or stalk domains that enhance their interaction with AGPAT2 in the absence of PA.

Finally, the interaction between DRP1 and AGPAT2 raised the question of whether AGPAT2 is required for ER tubulation because its loss depletes the majority of PA in the ER, or because AGPAT2 supplies PA directly to DRP1, thereby stimulating DRP1-mediated ER tubulation without altering overall PA levels. To test these possibilities, we performed lipidomics on purified ER membranes using shotgun mass spectrometry (Supplementary Fig. 8b)59–61. Our results revealed that the steady-state level of PA and other phospholipids, was comparable in ER purified from WT and AGPAT2-KO cells. These findings suggest that AGPAT2 does not regulate ER morphology by altering steady-state PA levels, but rather by supplying PA to DRP1 through their direct interaction.

Methods
Cells
WT, AGPAT2-KO, DRP1-KO31, and RTN4A-KO31 MEFs were grown in IMDM medium (12440061, Thermo Fisher Scientific) containing 10% FBS. WT, AGPAT2-KO, and DRP1-KO HEK293T cells were grown in DMEM media (10-017-CV, Corning) supplemented with 10% FBS. U2OS cells (HTB-96, ATCC) were cultured in DMEM medium containing 1 mM sodium pyruvate (11360-070, Gibco). KO cells were generated using the GeneArt CRISPR Nuclease Vector with OFP Reporter Kit (A21174, Invitrogen), following the manufacturer’s instructions. The target sequences for AGPAT2 (mouse: ACCGACCTTGGCGTAGAACC; human: AGGTACATGATGAGGCCCAC) and DRP1 (human: GCCTGTAGGTGATCAACCTA) were cloned into the vector and transfected into WT MEFs (for mouse AGPAT2) or HEK293T cells (for human AGPAT2 and DRP1). Based on the OFP fluorescent signal, transfected cells were sorted into 96-well plates as single cells at the Johns Hopkins Bloomberg Flow Cytometry and Immunology Core. The knockouts of AGPAT2 and DRP1 were screened and confirmed by DNA sequencing and/or Western blotting.

Plasmids and lentiviruses
Plasmids carrying GFP-SEC61β (15108) and RTN4A-GFP (61807) were obtained from Addgene. mCherry-SEC61β plasmids were constructed by replacing GFP with mCherry in the GFP-SEC61β plasmid. Plasmid transfection was carried out using Lipofectamine 3000 (Invitrogen). WT HA-LIPIN1 and the HA-LIPIN1(D712A) mutant were generated previously42. To construct AGPAT2 plasmids, AGPAT2 was PCR-amplified from cDNAs derived from MEFs and HEK293T cells and then cloned into either the pHR-SIN plasmid or the Clontech pEGFP plasmid, replacing the GFP gene. Mutations in the highly conserved amino acid residues in the catalytic motifs were introduced through site-directed mutagenesis using PCR with primers carrying specific mutations33,40,41. The WT and mutant alleles were confirmed by DNA sequencing of the plasmids. Epitopes, HA and FLAG, were fused to AGPAT2 by adding their sequences in PCR primers. To produce lentiviruses, the pHR-SIN plasmids carrying GFP-SEC61β, mCherry-SEC61β, AGPAT2-FLAG, AGPAT2-HA, AGPAT1-FLAG, RTN4A-GFP, and HA-LIPIN1 were co-transfected into HEK293T cells along with two other plasmids, pHR-CMV8.2ΔR and pCMV-VSVG, using Lipofectamine 2000 (Invitrogen)31. Two days after transfection, the supernatant containing the released viruses was collected from the transfected cells. The viruses were quick-frozen in liquid nitrogen and stored at -80°C. To generate shRNA lentiviruses, the following target sequences were cloned into pLKO.1: Scramble (CCTAAGGTTAAGTCGCCCTCG), AGPAT1 (CGTTCTGAGGAGCAATCTATC), and AGPAT2 (CCTGTGTCATCATCTCTAATC).

Knockdown screen
WT MEFs expressing GFP-SEC61β were individually transfected with 18 esiRNAs (6 AGPATs, 2 PLDs, 9 DGKs, 1 control RLuc) and one silencer select siRNA for Dgkz, as its esiRNA was not available, using Lipofectamine RNAiMAX (Invitrogen) according to the manufacturer’s instructions in two independent experiments. The cells were fixed with pre-warmed 4% paraformaldehyde in PBS for 20 min at room temperature and were permeabilized with PBS containing 0.1% Triton X-100 for 8 min at room temperature. Subsequently, the cells were incubated with anti-GFP antibodies to enhance the GFP-SEC61β signal, in PBS containing 0.5% BSA, followed by incubation with Alexa 488 anti-rabbit IgG (A21206, Invitrogen). The samples were examined using a Zeiss LSM800 laser scanning confocal microscope. In control cells (RLuc knockdown), ER tubules constituted ~50% of the ER network. Based on our previous work, we set a threshold of approximately half the control level as the criterion for a positive hit31. This level of reduction can be readily identified by visual inspection without quantification. Therefore, to efficiently screen many genes, we qualitatively assessed ER morphology without percentage quantification. In all knockdown conditions, ER tubule levels were comparable to those in control cells, except for AGPAT2 knockdown, which consistently showed a marked reduction in ER tubules in both experimental replicates.

qPCR
Cells were plated on 24-well plates. Total RNAs were purified from the cells using the RNeasy Mini Kit (74106, Qiagen) and reverse-transcribed using the ReadyScript cDNA Synthesis Mix (RDRT, Sigma-Aldrich). qPCR was performed using the QuantStudio 3 Real-Time PCR System (Applied Biosystems) with PowerUp SYBR Green Master Mix (A25741, Thermo Fisher Scientific). GAPDH was used as the reference gene. The following primers were used:
Mouse AGPAT1: CCCTACGTGGTTGTGTCCAA and AGTAGCTCACGCTTGGCAAT
Mouse AGPAT2: CCAAGGTCGGTCTCTACTGC and GCGAAGGCATACACGTACTTG
Human AGPAT2: GACGGTGGAGAACATGAGCATC and GGAGACGATGACACAGGGAC
Mouse GAPDH: AAGGTCATCCCAGAGCTGAA and CTGCTTCACCACCTTCTTGA
Human GAPDH: CATGAGAAGTATGACAACAGCCT and AGTCCTTCCACGATACCAAAGT

Western blotting
Proteins were separated using SDS-PAGE and transferred onto PVDF membranes. The primary antibodies used were: DRP1 (1:2000, 611113, BD Biosciences), TIM23 (1:1000, 611223, BD Biosciences), HA (1:10000, NB600-362, Novus Biologicals), RTN4 (1:1000, ab47085, Abcam), RTN3 (1:1000, 12055-2-AP, Proteintech), GAPDH (1:10000, MA5-15738, Thermo Fisher Scientific), tubulin (1:1000, 2125, Cell Signaling Technology), FLAG (1:2000, F7425, Sigma-Aldrich), AGPAT2 (1:1000, 14937S, Cell Signaling Technology), Climp-63 (1:100, sc-393544, Santa Cruz Biotechnology), BiP (1:1000, 3183, Cell Signaling Technology), eIF2α (1:1000, 2103, Cell Signaling Technology), phospho-eIF2α (1:1000, 3597, Cell Signaling Technology), CHOP (1:1000, 2895, Cell Signaling Technology), GFP31,72 and actin (1:1000, sc-47778, Santa Cruz Biotechnology). Immunocomplexes were visualized using fluorescently-labeled secondary antibodies, including Alexa Fluor 488 donkey anti-mouse IgG (1:2,000, A21202, Invitrogen), Alexa Fluor 488 donkey anti-rabbit IgG (1:2,000, A21206, Invitrogen), Alexa Fluor 488 chicken anti-goat IgG (1:2,000, A21467, Invitrogen), Alexa Fluor 647 donkey anti-mouse IgG (1:2,000, A31571, Invitrogen), Alexa Fluor 647 donkey anti-rabbit IgG (1:2,000, A31573, Invitrogen), and Alexa Fluor 647 donkey anti-goat IgG (1:2,000, A21447, Invitrogen), and detected with a Typhoon Molecular Imager (Amersham). To quantify band intensity, bands were boxed and mean values were measured using NIH Fiji software31. Background intensity was also measured in the corresponding region in a negative control lane and subtracted from the band intensity.

Lipid dot blot assay
Membrane lipid strips (P-6002, Echelon) were rehydrated and blocked in PBS containing 3% fatty acid free BSA (A3070, Sigma) and 1% Tween-20 for 1 hour at room temperature42. After washing in PBS containing 0.05% Tween-20 (PBST), the membranes were incubated with 1 μg/ml GFP (Ag33633, Proteintech) or GFP-D-octadecapeptide in 2 ml of binding buffer (100 mM NaCl and 10 mM sodium phosphate, pH 7.0) overnight at 4°C with gentle mixing. The membranes were washed in PBST three times for 10 min and incubated with anti-GFP antibodies in PBST containing 3% BSA for 1 h at room temperature. The immunocomplexes were visualized using fluorescently labeled secondary antibodies.

Liposome flotation assay
DOPC (850457), DPPG (840455), DPPS (840037), and rhodamine-DPPE (810158) were purchased from Avanti Polar Lipids. DPPA (L-4116), DPCL (L-C160), PI(4)P (P-4016), and PI(4,5)P2 (P-4516) were obtained from Echelon Biosciences. Lipids were mixed at a molar ratio of 84:1:15 for POPC:rhodamine-DPPE:variable lipid(s)42. The lipids were dried under a stream of nitrogen gas for 5 minutes, followed by further drying in a SpeedVac overnight. The dried lipid film was resuspended to a final concentration of 2 mM in 150 mM KCl and 20 mM MES (pH 7.0), vortexed for 1 h, and subjected to five freeze-thaw cycles using liquid nitrogen and a 42°C heat block. Unilamellar liposomes were generated by extrusion through a 400 nm polycarbonate membrane (21 passes)42. Liposomes (1 mM total lipid) were incubated with 1 μM GFP or GFP-D-octadecapeptide in a final volume of 200 μl at 4°C for 1 h with gentle mixing. The mixture was diluted in 1.728 M sucrose and 20 mM MES (pH 7.0) to a final volume of 1.25 ml and placed at the bottom of an ultracentrifuge tube. A sucrose step gradient was formed by overlaying 2.9 ml of 1.25 M sucrose in 20 mM MES (pH 7.0), followed by 0.85 ml of 0.25 M sucrose in the same buffer. The samples were centrifuged at 55,000 × g for 2 h at 4°C using an SW55 Ti rotor (Beckman Coulter). Two 2.5-ml fractions were collected from the top. Liposomes were recovered in the upper fractions42. Both upper and lower fractions were analyzed by Western blotting using anti-GFP antibodies.

Immunofluorescence microscopy
Cells were fixed with pre-warmed 4% paraformaldehyde in PBS for 20 min at room temperature in 8-well chambered cover glasses42. The cells were then permeabilized with PBS containing 0.1% Triton X-100 for 8 min at room temperature. Alternatively, to selectively permeabilize the plasma membrane, cells were treated with PBS containing 5% BSA and 0.1% saponin for 30 min at room temperature. Following permeabilization, cells were washed with PBS containing 1% BSA. They were then incubated with primary antibodies in PBS with 0.5% BSA, followed by incubation with the appropriate secondary antibodies. The primary antibodies used were: DRP1 (1:300, 611113, BD Biosciences), PDH subunit E2/E3bp (1:300, ab110333, Abcam), FLAG (1:300, 66008-3-Ig, Proteintech), GFP (to enhance GFP-SEC61β)31,72, mCherry (1:10000, M11217, Invitrogen, to enhance mCherry-SEC61β), and HA (1:300, NB600-362, Novus Biologicals, to detect AGPAT2-HA and HA-LIPIN1). Immunocomplexes were visualized using fluorescently-labeled secondary antibodies, including Alexa Fluor 488 donkey anti-rabbit IgG (1:400, A21206, Invitrogen), Alexa Fluor 568 donkey anti-mouse IgG (1:400, A10037, Invitrogen), Alexa Fluor 568 donkey anti-goat IgG (1:400, A11057, Invitrogen), and Alexa Fluor 568 donkey anti-rat IgG (1:400, ab175475, Abcam). The samples were examined using a Zeiss LSM800 laser scanning confocal microscope73.

To quantify ER morphology, we measured the area of the cell occupied by the ER—whether composed of tubules or sheets—using the ER marker SEC61β fused to GFP31. The total ER area was defined as the combined area occupied by ER tubules and sheets. The area of ER tubules was calculated by subtracting the sheet area from the total ER area. To determine the relative abundance of tubular ER, the tubule area was divided by the total ER area, yielding the percentage of the area occupied by the tubular form (Tubular ER%). This measurement reflects the spatial coverage of the ER tubule network within the cell, rather than the membrane surface area of individual tubules. All measurements were performed using NIH Fiji software.

For quantifying the distribution of mCherry-SEC61β, AGPAT2-FLAG, and RTN4A-GFP in the ER, the relative fluorescence intensity of the signal from these proteins in the area covered by ER tubules was measured in comparison to the total fluorescence intensity. For super-resolution imaging, the samples were viewed using a Zeiss LSM880-Airyscan FAST Super-resolution Single-point, laser scanning confocal microscope. Image analysis was performed using Zen and Imaris software31.

Focused ion beam scanning electron microscopy (FIB-SEM)
WT, DRP1-KO, and AGPAT2-KO MEFs were pre-fixed in 2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer (pH 7.4, Electron Microscopy Sciences) at Johns Hopkins University and then shipped to the University of Michigan’s Microscopy Core34 overnight at ice temperature. The cells were subjected to post-fixation in a mixture of 1% osmium tetroxide (OsO₄) and 1% potassium ferrocyanide in 0.1 M sodium cacodylate buffer for 15 min at ice temperature. The fixed cells were then contrasted with 1% thiocarbohydrazide, 1% OsO₄, and 1% uranyl acetate, each for 5 min at room temperature. Cells were further treated with Walton’s lead aspartate for 10 min at 60°C. The cells were dehydrated in an ascending ethanol series (10%, 30%, 50%, 70%, 80%, 95%, and 100%) and embedded in Durcupan resin (Electron Microscopy Sciences) in beam capsules. After thermal polymerization at 70°C for 48 hours, the glass coverslip was removed by liquid nitrogen, and the cell monolayer was transferred to the surface of the resin block. The samples were serially sectioned from the side of the resin block close to the coverslip using a Leica EM ARTOS 3D ultramicrotome (Leica Microsystems) with a diamond knife (DiATOME). 600 μm × 600 μm sections with 1 μm thickness were obtained. Six to seven serial sections were manually transferred to a 10 × 10 mm Si wafer by submerging the Si wafer as 3 quarters as into the diamond knife trough filled with water and inclined facing the diamond knife edge. The serial sections were anchored to the horizontal line between the Si chip and water. Then water in the diamond knife trough was very slowly drained until the sections were attached on the Si wafer and the sections were dried on a heat block at 45°C for 3 h. During these processes, the order of the serial sections was maintained. The serial sections on the Si wafer were coated with a 5 nm thick carbon layer using a Leica EM ACE600 high vacuum coater (Leica Microsystems). The first or second section from each group (1-2 μm from the bottom of the cells, where the ER is usually observed) was observed using an EVO 15 LaB6 SEM (Carl Zeiss USA) at 10 keV with a high-definition backscattered detector to identify regions containing the ER in these sections. The ER was imaged at 1000 × magnification, with ~10 nm resolution and a 6144 × 4608-pixel output prior to FIB-SEM analysis.

For FIB-SEM analysis, samples were imaged using a FEI Helios Nanolab 650 Dual Beam system equipped with a gas injection system for platinum deposition and milling, at the Michigan Center for Materials Characterization. Using back-scattered electron imaging at 10 kV accelerating voltage, we located the same region of interest containing the ER. Once the region of interest was found, a platinum rectangle was deposited on a 6 × 8 μm region at 10 kV and 0.8 nA, followed by a protective deposition of carbon at 30 kV and 0.23 nA. Rough trenches around the region were cut using 30 kV and 9.3 nA beam current. The region of interest was lift-out and transferred to a new Si wafer using a micromanipulator (Omniprobe 200). The sample face was polished using 2.5 nA and 0.79 nA beam currents. The selected block was sequentially cut and imaged to obtain an image stack for 3D reconstruction using AutoSlice and View G3 V1.3 software. FIB cuts were made using 30 kV accelerating voltage and 80 pA beam current, while image of the block face was done using the SEM at 2 kV, set in backscatter mode. The field of view of the SEM and the slice thickness were set to produce 5 nm cubic voxels. The dimensions of the imaged blocks (width × height × thickness) were 8.1 × 6.8 × 0.4 μm for WT cells, 8.0 × 7.0 × 0.5 μm for DRP1-KO cells, and 8.4 × 6.4 × 0.5 μm for AGPAT2-KO cells. The resulting stack of images was aligned and reconstructed using Avizo v.9.3 (Thermo Fisher Scientific) and Imaris 9.3 (Oxford Instruments). Manual segmentation was performed to determine the boundaries of the ER features contained in the image stack dataset34.

Isolation of the ER
Cells were cultured, washed with ice-cold PBS, and placed on ice. Homogenization buffer [10 mM HEPES-KOH (pH 7.4) containing 0.22 M mannitol, 0.07 M sucrose, and protease inhibitors (11836170001, Roche)] was added, and cells were collected using a cell scraper31. The cells were washed twice for 5 min each at 4°C. To isolate the ER and mitochondria, cells were homogenized with 10 strokes using a syringe fitted with a 27-gauge needle in the same buffer. The post nuclear supernatant was obtained by centrifugation at 800 g for 5 min and then for 10 min at 4°C. The supernatant was further separated into an ER-enriched supernatant and a mitochondria-enriched pellet by centrifugation at 8000 g for 10 min at 4°C. The ER-enriched supernatant was further clarified by centrifugation at 17,400 g for 20 min at 4°C. Finally, the supernatant was separated into the ER pellet and the cytosolic supernatant by ultracentrifugation at 100,000 g for 1 h at 4°C31. To isolate the ER for in vitro AGPAT2 activity assays, cells were homogenized with 30 strokes using a PYREX homogenizer (7725T-5) and purified as described above.

Lipidomics
Lipid species were profiled using a multidimensional mass spectrometry-based shotgun lipidomics approach74. The protein concentration of each isolated ER sample was quantified using the Pierce BCA Protein Assay Kit (23225, Thermo Scientific). Bovine serum albumin was used as the standard. An adequate amount of each sample (equivalent to 0.1 mg protein) was transferred into a disposable glass culture test tube. A premixed solution of lipid internal standards (IS), including di14:1 choline glycerophospholipid (PC), di16:1 ethanolamine glycerophospholipid (PE), di15:0 phosphatidylglycerol (Na) (PG), di14:0 phosphatidylserine (Na) (PS), di14:0 phosphatidic acid (Na) (PA), and 17:0 lysophosphatidic acid (LPA), was added prior to lipid extraction for quantification of the targeted lipid species75. Lipid extraction was performed using a modified Bligh and Dyer procedure74. The organic and aqueous phases were collected separately into new glass tubes.

The aqueous phase was used to extract LPA. Briefly, an equal volume of 4% formic acid in methanol was added to the aqueous phase and vortexed for 30 s. A Hybrid SPE cartridge was preconditioned twice with 2 ml of methanol. The aqueous solution was then loaded onto the cartridge to recover and enrich LPA species from biological matrices. The SPE column was washed twice with 2 ml of methanol, and the lipids trapped in the column were eluted with 4 ml of 20% ammonium methanol solution and dried under a nitrogen stream76. The lipid extract for LPA analysis was resuspended at 100 μl/mg of protein in methanol and stored at -20°C until use.

The organic phase from the Bligh and Dyer extraction was used to isolate other lipids. To minimize oxidation and preserve lipid integrity, the lipid extracts were flushed with nitrogen gas, sealed with caps, and stored at -80°C. Before lipidomic analysis, the extracts were resuspended in a chloroform:methanol mixture (1:1, v/v) at 500 μL/mg of protein. A 10 μl aliquot of each extract was used for derivatization of PE and DAG, respectively77,78. All samples were analyzed within one month of extraction.

For shotgun lipidomics, individual lipid extracts were further diluted to a final concentration of ~500 fmol total lipids per μL for mass spectrometric analysis. Data acquisition for PC, PE, and LPA was performed on a TSQ Altis triple quadrupole mass spectrometer. Data acquisition for PS, PG, PA, and PI was performed on a Q-Exactive hybrid quadrupole-Orbitrap system (Thermo Fisher Scientific). An automated nanospray ion source (TriVersa NanoMate, Advion Bioscience) was used for sample introduction59.

Data were processed following standard principles of shotgun lipidomics, including ion peak selection, baseline correction, and isotope deconvolution61,75,79. Internal standards were applied to correct for extraction efficiency and matrix effects. Minimal batch variation was observed due to normalization procedures. Final lipid levels were expressed as nmol per mg of protein. Details on the linear dynamic range, signal-to-noise threshold, and quality control procedures followed previously established protocols59,74.

DRP1 expression and purification
Rosetta 2 (DE3) pLysS competent cells (Novagen) were transfected with pET15b vectors carrying DRP1 and were grown in LB containing ampicillin and chloramphenicol overnight at 37°C42. 1 ml of the culture was diluted into 1 L of the same growth medium and continued to grow for 3–5 hours at 37°C. The culture was cooled down on ice, and 0.1 mM of IPTG was added. The expression of His6-DRP1 (mouse isoform 3, Uniprot: Q8K1M6-3)80,81, His6-DRP1 (K38A, G350D), and His6-GFP-D-octadecapeptide was induced overnight at 16°C. The bacteria were washed twice with PBS by centrifugation at 5000 × g for 15 min at 4°C in a JLA 8.1000 rotor and harvested and froze at -80°C. The frozen cell pellets were resuspended in 40 ml of lysis buffer (10 mM imidazole, 1 mM MgCl2, 500 mM NaCl, 2 mM β-mercaptoethanol, 20 mM Hepes, pH 7.4) and sonicated on ice for 10 × 5 sec at setting 5 and then 10 × 5 sec at setting 2 using a Fisher Scientific Sonic dismembrator model 100. The cell homogenate was centrifuged at 1935 × g for 15 min at 4°C in JA-20X, and the supernatant was further clarified at 20,100 × g for 15 min at 4°C in JA-20X. After being passed through a membrane filter with a pore size of 0.45 μm, the lysate was incubated with 2 ml of pre-washed 50% Ni-NTA beads (His-Bind Resin, Novagen) overnight at 4°C and placed in a 15-ml column (Poly-prep chromatography column, Bio-Rad). The beads were washed with 3 ml of lysis buffer, 15 ml of lysis buffer, 3 ml of wash buffer (40 mM imidazole, 1 mM MgCl2, 500 mM NaCl, 2 mM β-mercaptoethanol, 20 mM Hepes, pH 7.4), and 15 ml of wash buffer. The His6-tagged recombinant proteins were eluted from the column using 3 × 0.9 ml of elution buffer (250 mM imidazole, 1 mM MgCl2, 500 mM NaCl, 2 mM β-mercaptoethanol, 20 mM Hepes, pH 7.4) and collected it into nine 300-μl fractions. After confirming the purification of proteins using SDS-PAGE and Coomassie Brilliant Blue staining of each fraction, the peak fractions (typically fractions #3 and 4) were collected and diluted into 15 ml of lysis buffer without imidazole. The residual imidazole was removed by passing the solution three times through an Amicon Ultra Centrifugation Filter (50 kDa MWCO) according to the manufacturer’s instructions. The purified proteins were mixed with final 20% DMSO, snap frozen in liquid nitrogen, and stored at -80°C.

In vitro ER tubulation assay
Freshly purified ER (7.5 μg of proteins in 50 μl) was incubated with or without recombinant DRP1 protein and 8:0 PA (830842, Avanti) in 150 mM KCl and 20 mM MES (pH 7.0) at 23°C for 3 h. We selected 8:0 PA based on preliminary experiments comparing the solubility of PA species with different acyl chain lengths (8:0 and 10:0) under our assay conditions, in which 8:0 PA remained soluble while 10:0 PA (830843, Avanti) precipitated out of solution. As a negative control, GFP protein (Ag33633, Proteintech) was used. For negative-stain electron microscopy, 20 μl of the samples were placed on EM grids (CF400-CU-UL, Electron Microscopy Sciences) that had been glow-discharged using a Quorum GloQube-D (Laughton)31. After an incubation period of 2 min, the grids were washed three times with 150 mM KCl and 20 mM MES (pH 7.0), stained with 0.1% uranyl acetate and 0.04% Tylose for 30 seconds twice, and then air-dried. The samples were examined using a Hitachi 7600 transmission electron microscope31. The length of tubules was analyzed using NIH Fiji software. Small vesicles with an area less than 0.05 μm2 were excluded from the analysis.

To test the effect of antibodies on ER tubules, freshly purified ER (7.5 μg of protein in 50 μl) was incubated with 12.5 μg/ml of TOM20 antibody (612278, BD Biosciences) or DRP1 antibody (611113, BD Biosciences) in a buffer containing 150 mM KCl and 20 mM MES (pH 7.0) at room temperature for 3 h. Electron microscopy sample preparation, negative staining, and length analysis were performed as described above.

In vitro AGPAT2 activity assay
Freshly isolated ER (30 μg of proteins) was incubated with 50 μM 18:0 acyl-CoA (870718, Avanti) and 10 μM lysoPA-TopFluor (810280, Avanti) in 200 μl of 100 mM Tris-HCl (pH 7.4) containing 1 mg/ml of fatty acid-free bovine serum albumin (A7030, Sigma) for 10 min at 37°C. Lipids were extracted using 200 μl of chloroform/methanol (2:1, v/v) at room temperature for 1 h with gentle agitation. The extracted lipid samples were applied to a TLC plate (60805, Sigma) and developed in a solvent system of chloroform/ethanol/water in a ratio of 65:25:4 (v/v/v). NBD-PA (810174P, Avanti) was used as lipid markers. After development, the TLC plates were dried and the lipid spots were visualized and scanned using a Typhoon imager system.

Immunoprecipitation
HEK293T cells were cultured in 10-cm dishes and transfected with various plasmids using Lipofectamine 300042. Two days after transfection, the cells were washed with ice-cold PBS, placed on ice, and lysed with 0.5 ml of ice-cold 1x lysis buffer [20 mM Tris-HCl pH 7.5, 100 mM KCl, 1% Triton X-100, protease inhibitor cocktail (11836170001, Roche)]. The samples were gently homogenized by pipetting on ice for 15 min. The homogenates were clarified by centrifugation at 14,500 × g for 5 min at 4°C in a microcentrifuge twice. A total of 500 μl of the supernatants was then incubated with either 10 μl of Anti-FLAG M2 Affinity Gel (A2220, Sigma), GFP-Trap (ChromoTek), or 10 μl of Protein A-Sepharose 4B (101041, Life Technologies) combined with 5 μl of DRP1 antibody (110-55288, Novus Biologicals) for 3 h at 4°C. The beads were washed three times with 1x lysis buffer. The clarified cell lysates and immunoprecipitated fractions were analyzed by SDS-PAGE and Western blotting using antibodies specific for DRP1 (611113, BD), FLAG (F7425, Sigma), AGPAT2 (14937S, Cell Signaling), and GAPDH (MA5-15738, Thermo Fisher Scientific). Immunocomplexes were visualized using fluorescent-labeled secondary antibodies and detected using a Typhoon Molecular Imager (Amersham). To quantify band intensity, bands were boxed and mean values were measured using NIH Fiji software31. Background intensity was also measured in the corresponding region in a negative control lane and subtracted from the band intensity.

Statistical and Reproducibility
Statistical analyses were conducted using GraphPad Prism 10. Sample sizes were not predetermined by statistical methods, and experiments were not blinded. No data were excluded from the analyses. The experiments were not randomized, and the investigators were not blinded to allocation during either the experiment or the outcome assessment.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "This is an interesting manuscript that identifies AGPAT2 that generates phosphatidic acid (PA) from lyso-PA (LPA) as a direct effector of DRP1-mediated ER membrane tubulation, independent of DRP1’s self-assembly and GTPase activities. The tour de force cell biology experiments are impressive and comprehensive with little to comment on their execution, which is top-notch. However, the design and interpretation of several experiments leave a lot to be desired as outlined below. Given these drawbacks, the manuscript in its current form is not suitable for Nature Communications and its expected rigor.\n1. In Fig. 2i, via FIB-SEM experiments, the authors show that in DRP1 KO MEFs, ER tubules are entirely lost and are transformed into sheets, but this stands in direct contrast/conflict with past experiments (Friedman et al., Science 1 Sep 2011 Vol 334, Issue 6054 pp. 358-362 DOI: 10.1126/science.1207385) that clearly show that DRP1 knockdown does not disrupt ER-mitochondria contact sites, the cells still displaying pre-constricted ER tubule-mitochondria contacts also containing the DRP1 receptor Mff. Are the authors looking specifically at ER-mitochondria contact sites here? If not, they must, and also explain why these contacts persist even in the absence of DRP1 in previous experiments. Resolution of these conflicting data is an absolute must.\n2. In Fig. 4a, b, the authors show “that purified WT ER contains both sheets and tubules and the addition of DRP1 did not alter these structures”, but do not show these tubules are indeed decorated/generated by endogenous DRP1. Are they? What evidence exists that the ER tubules isolated from WT cells are indeed generated by endogenous DRP1 decoration? Moreover, how exactly was extrinsic PA (in the form of soluble, non-physiological short chain 8:0 PA) added to the ER sheets isolated from AGPAT2-KO+shDRP1 cells, and how was the efficiency of 8:0 PA incorporation into the native ER membranes determined? Could DRP1 be clustering 8:0 PA in solution to create adherent (sticky) tubes instead of tubulating the ER membrane vesicles themselves? What else do these tubes contain, besides DRP1? An ER membrane marker imaged via correlative light microscopy might provide the necessary distinction.\n3. I also have immediate and pressing concerns with the in vitro reconstitution experiments and the conclusions drawn, including that: (i) that DRP1 tubulates ER membranes in an oligomerization- and GTPase activity-independent manner, (ii) that an 18-residue sequence in the VD called Octadecapeptide constitutes an alpha-helix and an amphipathic helix one at that, and (iii) that this helix selectively binds PA over other lipids to remodel the ER membrane – none of which are supported by the data presented in this study, or from the published literature.\nTo demonstrate point (i) the authors should use a combined oligomerization- and GTPase-dead version of DRP1. Mutations R403C, G350D, or G363D have been shown to impair DRP1 self-assembly beyond dimers in solution, while mutations K38A or S39N ablate GTPase activity. The authors should use these two sets of mutations in combination to demonstrate full-length DRP1 rid of both activities still tubulates isolated ER membranes, or for that matter also PA-containing liposomes, specifically in vitro (with or without added GTP). Given DRP1’s interactions with various binding partners in the intracellular milieu, in vivo experiments alone (as performed in Adachi et al., Molecular Cell, 2020) do not suffice.\nTo demonstrate point (ii), the authors should use CD to demonstrate that this peptide indeed forms an alpha-helix in solution under physiological conditions. Moreover, helical wheel analysis of this sequence shows no clear amphipathicity (sidedness and demarcation of the polar and nonpolar residue faces with a strong and distinct hydrophobic moment). Without any supporting data, this sequence's description as an amphipathic helix should be removed here and moving forward. This does not fit the profile of a well-defined amphipathic helix.\nTo demonstrate point (iii) the authors should show that this stretch selectively binds PA over other negatively charged lipids such as PS, PG, CL, and PIPs. If so, what is the molecular basis of this selectivity? I do not think this has been established.\n4. In Fig. 7, the authors further claim that the variable domain inhibits the interaction of DRP1 with AGPAT2 when AGPAT2 does not produce PA. Given that AGPAT2 is predicted to interact with the GTPase domain-stalk interface at the opposite end of the molecule, how does this occur? Moreover, DRP1 rid of the variable domain has been shown to prematurely form fibril-like oligomers in solution, while the full-length remains mostly an assembly auto-inhibited dimer-tetramer. Could the differential co-IP be a reflection of the different oligomerization states of the GFP-FL and GFP-GS variants of DRP1? This part of the manuscript is ambiguous and mostly speculative with no consideration of the structural biology of the DRP1 molecule or the topology of the contacts when embedded in the same membrane bilayer.", "reviewer_2": "In this manuscript, the authors investigate the factors that contribute to ER morphology. In addition to the known proteins that shape ER sheets and tubules, the authors show that membrane phospholipid PA generated by AGPAT2 drives ER tubulation. The authors propose that the PA generated by AGPAT2 is used by DRP1, to generate membrane curvature. While, the observation is interesting, how does a phospholipid that constitutes less than 5% of the membrane phospholipid regulate the ER morphology by is unclear. Authors claim to report previously undetermined role of lipids in ER morphogenesis. It has been reported before that mutants in yeast such as spo7, nem1, pah1 and opi1 that lead to increase membrane phospholipids have more ER sheets than ER tubules. Other factors that can affect ER morphology is ER stress. Authors don’t address these alternative possibilities. While the manuscript proposes an interesting model, some of the data presentation and interpretations raise concerns about the conclusions.\n1. Provide quantification of % tubular ER for Figure 1B.\n2. Figure 2: Throughout the manuscript, the authors measure “tubular ER %”. It is unclear from the results and materials and methods sections how this percentage is being calculated as it seems that only one ER marker (SEC61B) is being used. SEC61B is used as a general ER marker and localizes to both ER sheets and tubules (as can be seen in microscopy images in Figure 1). A better explanation showing how this percentage is calculated should be given as this ratio is used in multiple figures.\n3. Need more evidence to show AGPAT2 plays a role in ER morphology. Overexpression of RTN4A in AGPAT2 KO cells to check if the ER tubules can be restored supporting Figure 2B.\n4. Does AGPAT2-KO have increased ER stress which can alter ER morphology?\n5. Is the expression of reticulons and reticulon-like proteins normal in AGPAT2?\n6. What is the ER membrane lipid composition in AGPAT2?\n7. Figure 4 is not convincing. Unable to see actual ER tubules and sheets due to magenta highlights.\n8. Can AGPAT2 addition in DRP1 KO and AGPAT2 KO cells increase tubulation in isolated ER?\n9. Is AGPAT2-KO ER membrane completely devoid of PA? As, addition of external PA is tubulating the ER membrane in Figure 4.\n10. Measure PA and DAG to support 5i.\n11. Quantification of western blots would be helpful, especially when authors claim that there is an increase or decrease in protein levels.\n12. Does lipin OE lead to increased ER sheets?\n13. Inference from figure 6i not convincing. CLIMP63 seen in mitochondria.\nMinor-\n1. Figure 5G: should read “AGPAT2” instead of “AGPTA2” in figure", "reviewer_3": "In this paper, the authors investigate how lipid biosynthesis influences the three-dimensional organization of the ER membrane.\nMajor comments:\nThe EM work done both using TEM and SEM is not described well . It is not clear how the samples were prepared and therefore it is challenging to evaluate the interpretation of the results.\nPlease elaborate on how you perform sample prep for both TEM and SEM in an way that will allow the reader to repeat your experiment.\nMoreover- The graphs in the entire paper are too small to be seen, the data can not be understood that way.\nSpecific Commets:\nPage 3 line 61: The full name of PA, \"phosphatidic acid,\" is missing from the main text\nPage 4 line 70 : The full name of PC, is missing from the main text\nline 82: Figure 1b is not presented well; the result is not clearly evident. Please mark the AGPAT2 micrograph differently, and ensure the titles are capitalized as written in the text: AGPAT2 instead of agpat2.\nFig2i:\nThe 3D representation of the segmented surface data should be displayed alongside the data itself—which means it should be presented alongside at least one slice through the volume. Moreover, a scale cannot be added to the figure if the surface is presented in a perspective view. Please provide a scale on the slice, and for the surface, specify the FOV dimensions.\nMore importantly: What is the field of view of the imaged block? Is it identical for all three samples? I’m a bit confused by your materials and methods—are you using block-face imaging or array tomography? Please provide more information regarding the sample preparation.\nMaterial and method section:\nLine 387: Please provide more details. Was the cell monolayer embedded in resin? Which resin? what are the steps of fixation and embedding used? How did you manage to obtain 500 nm thick slices through a cell monolayer? Which microtome did you use? How were the 500 nm sections organized and lifted on the wafer? manually?\nLine392: What do you mean platinum rectangle was deposited by SEM? do you mean by GIS system installed at the SEM?\nLine 395: What is the thickness of the final lamella obtain after the polish?\nLine 400: What are the overall dimensions of the imaged block?"}, "author_response": "RESPONSES TO REVIEWERS' COMMENTS\nReviewer #1\nThis is an interesting manuscript that identifies AGPAT2 that generates phosphatidic acid (PA) from\nlyso-PA (LPA) as a direct effector of DRP1-mediated ER membrane tubulation, independent of DRP1’s\nself-assembly and GTPase activities. The tour de force cell biology experiments are impressive and\ncomprehensive with little to comment on their execution, which is top-notch. However, the design and\ninterpretation of several experiments leave a lot to be desired as outlined below. Given these drawbacks,\nthe manuscript in its current form is not suitable for Nature Communications and its expected rigor.\nRESPONSE 1: We sincerely appreciate the reviewer’s positive and encouraging feedback. We are also\ngrateful for the constructive and helpful comments that have significantly contributed to the\nimprovement of our manuscript. As detailed below, all comments have been thoroughly addressed\nduring the revision process. Changes in the text are shown in red font in the revised manuscript.\n1. In Fig. 2i, via FIB-SEM experiments, the authors show that in DRP1 KO MEFs, ER tubules are\nentirely lost and are transformed into sheets, but this stands in direct contrast/conflict with past\nexperiments (Friedman et al., Science 1 Sep 2011 Vol 334, Issue 6054 pp. 358-362 DOI:\n10.1126/science.1207385) that clearly show that DRP1 knockdown does not disrupt ER-mitochondria\ncontact sites, the cells still displaying pre-constricted ER tubule-mitochondria contacts also containing the\nDRP1 receptor Mff. Are the authors looking specifically at ER-mitochondria contact sites here? If not,\nthey must, and also explain why these contacts persist even in the absence of DRP1 in previous\nexperiments. Resolution of these conflicting data is an absolute must.\nRESPONSE 2: Thank you for bringing up these important points. Due to the limited volume that can be\nanalyzed by FIB-SEM, we focused our efforts on visualizing the 3D architecture of peripheral ER,\nrather than ER–mitochondria contact sites. Quantitative laser confocal immunofluorescence microscopy\nrevealed that although ER tubules are reduced in Drp1-KO MEFs, they are still present—likely\nmaintained through Drp1-independent mechanisms (Figure 5a, b)—consistent with our previous\nfindings (Adachi et al., Mol. Cell, 2018). Regarding contact sites, we previously observed pre-constricted\nER tubule–mitochondria contacts in Drp1-KO MEFs (Adachi et al., Mol. Cell, 2018). These observations\nare in agreement with the study by Friedman et al. (2011), which showed that mitochondrial\nconstriction can occur at ER contact sites independently of Drp1.\n2. In Fig. 4a, b, the authors show “that purified WT ER contains both sheets and tubules and the addition\nof DRP1 did not alter these structures”, but do not show these tubules are indeed decorated/generated by\nendogenous DRP1. Are they? What evidence exists that the ER tubules isolated from WT cells are indeed\ngenerated by endogenous DRP1 decoration?\nRESPONSE 3: We agree with the reviewer that determining whether the tubules are generated by\nendogenous DRP1 is a critical point. To address this, we conducted new experiments during the revision\nin which we incubated WT ER with antibodies against DRP1 or, as a negative control, TOM20. We found\nthat incubation with the DRP1 antibody, but not the TOM20 antibody, resulted in the loss of ER tubules\n(Figure 6h, i), suggesting that endogenous DRP1 is required to maintain ER tubular structures. We also\nattempted to examine the localization of endogenous DRP1 on isolated ER tubules using gold-\nconjugated DRP1 antibodies. However, as noted above, the DRP1 antibody disrupts ER tubules, making\nit technically challenging to localize DRP1 on the ER membrane under current conditions. We plan to\npursue this interesting direction in future studies.\nadded to the ER sheets isolated from AGPAT2-KO+shDRP1 cells, and how was the efficiency of 8:0 PA\nincorporation into the native ER membranes determined?\nRESPONSE 4: We thank the reviewer for this insightful question. In this experiment, we added 8:0 PA to\nfreshly purified ER in a buffer containing 150 mM KCl and 20 mM MES (pH 7.0). We selected 8:0 PA\nbased on preliminary experiments comparing the solubility of PA species with different acyl chain\nlengths (8:0 and 10:0) under our assay conditions. This step was critical, as we aimed to avoid using\ndetergents, organic solvents, or BSA as lipid carriers—particularly because BSA induced aggregation of\nER membranes in our assay. Under these conditions, 8:0 PA remained soluble, whereas 10:0 PA\nprecipitated out of solution. Therefore, we used 8:0 PA to assess its effect on DRP1-dependent ER\ntubulation. Due to the technical challenges in accurately measuring PA incorporation into membranes,\nwe did not assess incorporation efficiency. Accordingly, we have revised the manuscript to more\ncarefully describe the results of this experiment and expanded the discussion to include alternative\ninterpretations. For instance, it is possible that PA interacts with DRP1 without integrating into the\nmembrane, thereby enhancing DRP1’s ability to tubulate the ER membrane. We included a control\nexperiment showing that PA alone does not induce ER tubulation. Additionally, we performed a second\ncontrol demonstrating that DRP1 and PA do not form tubules (RESPONSE 5). We have moved these data\nto Supplementary Figure 7 to improve clarity and focus.\nCould DRP1 be clustering 8:0 PA in solution to create adherent (sticky) tubes instead of tubulating the\nER membrane vesicles themselves? What else do these tubes contain, besides DRP1? An ER membrane\nmarker imaged via correlative light microscopy might provide the necessary distinction.\nRESPONSE 5: To address this concern, we performed a new experiment during the revision. To\ndetermine whether PA and DRP1 alone can generate tubular structures under our experimental\nconditions, we examined DRP1 alone and a mixture of DRP1 and PA using electron microscopy. Our new\nresults show that neither condition results in tubule formation (Supplementary Figure 7c), supporting\nour original conclusion that the observed tubules originate from the ER membrane. Regarding the\ncontents of ER tubules, we agree this is an interesting question. However, we believe it falls beyond the\nscope of the current study.\n3. I also have immediate and pressing concerns with the in vitro reconstitution experiments and the\nconclusions drawn, including that:\n(i) that DRP1 tubulates ER membranes in an oligomerization- and GTPase activity-independent manner,\n(ii) that an 18-residue sequence in the VD called Octadecapeptide constitutes an alpha-helix and an\namphipathic helix one at that, and (iii) that this helix selectively binds PA over other lipids to remodel the\nER membrane – none of which are supported by the data presented in this study, or from the published\nliterature.\nRESPONSE 6: Thank you for the insightful comments. We have addressed each point in detail below.\nTo demonstrate point (i) the authors should use a combined oligomerization- and GTPase-dead version of\nDRP1. Mutations R403C, G350D, or G363D have been shown to impair DRP1 self-assembly beyond\ndimers in solution, while mutations K38A or S39N ablate GTPase activity. The authors should use these\ntwo sets of mutations in combination to demonstrate full-length DRP1 rid of both activities still tubulates\nisolated ER membranes, or for that matter also PA-containing liposomes, specifically in vitro (with or\nwithout added GTP). Given DRP1’s interactions with various binding partners in the intracellular milieu,\n2\nin vivo experiments alone (as performed in Adachi et al., Molecular Cell, 2020) do not suffice.\nRESPONSE 7: To address this concern, we performed the suggested experiment during the revision. We\npurified DRP1 carrying both an oligomerization-defective mutation (G350D) and a GTPase-defective\nmutation (K38A) and tested it in our in vitro ER tubulation assay. Our new data show that DRP1\n(G350D/K38A) tubulates the ER similarly to WT DRP1 (Figure 6f, g), substantiating our original conclusion\nthat these two activities are dispensable for ER tubulation.\nTo demonstrate point (ii), the authors should use CD to demonstrate that this peptide indeed forms an\nalpha-helix in solution under physiological conditions. Moreover, helical wheel analysis of this sequence\nshows no clear amphipathicity (sidedness and demarcation of the polar and nonpolar residue faces with a\nstrong and distinct hydrophobic moment). Without any supporting data, this sequence's description as an\namphipathic helix should be removed here and moving forward. This does not fit the profile of a well-\ndefined amphipathic helix.\nRESPONSE 8: As suggested, we examined the peptide's secondary structure using CD spectrometry in\ncollaboration with Dr. Doug Barrick (Department of Biophysics, Johns Hopkins University) and found no\nevidence of an alpha helix in solution (200 µM peptide, 20 mM MES, pH 7.0, 150 mM KCl). Since the\npeptides interact with PA, which is functionally important, we are interested in further analyzing their\nstructure under different conditions—such as in the presence of PA or membranes—in future studies.\nHowever, we believe this is beyond the scope of the current study. As recommended by the reviewer,\nwe have removed the description of the peptide as a helix in the revised manuscript.\nTo demonstrate point (iii) the authors should show that this stretch selectively binds PA over other\nnegatively charged lipids such as PS, PG, CL, and PIPs. If so, what is the molecular basis of this\nselectivity? I do not think this has been established.\nRESPONSE 9: As recommended, we have expanded our lipid-binding analysis of the peptide in the\nrevised manuscript. We performed a lipid dot blot assay using GFP fused to the peptide and full-length\nDRP1, as we have previously done (Adachi et al., Mol Cell, 2016). Our new results show that while DRP1\nbinds to several lipids—including PA, CL, and PIPs—the peptide binds more specifically to PA. Notably,\nthe peptide did not bind to other negatively charged lipids such as PS, PG, CL, or PIPs (Supplementary\nFigure 1), indicating its high specificity for PA.\n4. In Fig. 7, the authors further claim that the variable domain inhibits the interaction of DRP1 with\nAGPAT2 when AGPAT2 does not produce PA. Given that AGPAT2 is predicted to interact with the\nGTPase domain-stalk interface at the opposite end of the molecule, how does this occur? Moreover,\nDRP1 rid of the variable domain has been shown to prematurely form fibril-like oligomers in solution,\nwhile the full-length remains mostly an assembly auto-inhibited dimer-tetramer. Could the differential co-\nIP be a reflection of the different oligomerization states of the GFP-FL and GFP-GS variants of DRP1?\nThis part of the manuscript is ambiguous and mostly speculative with no consideration of the structural\nbiology of the DRP1 molecule or the topology of the contacts when embedded in the same membrane\nbilayer.\nRESPONSE 10: Thank you for the insightful comments on the DRP1 structure and assembly. To address\nthe key question raised by the reviewer regarding the role of DRP1 oligomerization in the DRP1–AGPAT2\ninteraction, we performed new experiments using the oligomerization-defective G350D mutant. Our\nresults show that oligomerization-defective DRP1 (G350D) interacts more strongly with enzymatically\ninactive AGPAT2 than WT DRP1 in co-immunoprecipitation assays (Supplementary Figure 10). These\n3\ndata suggest that the inhibition involves DRP1 oligomerization. Due to the unstructured nature of the\nvariable domain, it is not well resolved in existing structural models of DRP1, and how this domain\ninteracts with or influences the GTPase and stalk domains remains largely unknown. Nonetheless,\noligomerization may facilitate intermolecular interactions within DRP1 oligomers such that the variable\ndomain physically hinders interaction with AGPAT2. Alternatively, oligomerization may induce\nconformational changes in the GTPase or stalk domains that enhance their interaction with AGPAT2 in\nthe presence of PA. We have added the new data and corresponding discussion to the revised\nmanuscript.\nReviewer #2:\nIn this manuscript, the authors investigate the factors that contribute to ER morphology. In addition to the\nknown proteins that shape ER sheets and tubules, the authors show that membrane phospholipid PA\ngenerated by AGPAT2 drives ER tubulation. The authors propose that the PA generated by AGPAT2 is\nused by DRP1, to generate membrane curvature. While, the observation is interesting, how does a\nphospholipid that constitutes less than 5% of the membrane phospholipid regulate the ER morphology by\nis unclear. Authors claim to report previously undetermined role of lipids in ER morphogenesis. It has\nbeen reported before that mutants in yeast such as spo7, nem1, pah1 and opi1 that lead to increase\nmembrane phospholipids have more ER sheets than ER tubules. Other factors that can affect ER\nmorphology is ER stress. Authors don’t address these alternative possibilities. While the manuscript\nproposes an interesting model, some of the data presentation and interpretations raise concerns about the\nconclusions.\nRESPONSE 1: We genuinely appreciate the reviewer’s supportive and insightful comments on our\nmanuscript. We thoroughly addressed all the comments during the revision process and substantially\nrevised our study by conducting new experiments, as detailed below. Changes in the text are indicated\nin red font in the revised manuscript.\nMore specifically, regarding alternative explanations for our data, we first tested the role of ER stress\nduring the revision (RESPONSE 4) and found no induction of ER stress in AGPAT2-KO cells. Second, we\nagree that reduced lipid biosynthesis due to the absence of AGPAT2 could contribute to changes in ER\nmorphology, consistent with previous studies. To address this point, as requested, we performed\nlipidomic analysis and found similar levels of PA and other phospholipids in WT and AGPAT2-KO ER.\nTherefore, the increase in ER tubules is not simply due to PA depletion in AGPAT2-KO cells (RESPONSE\n7). Third, our in vitro assays distinguish between lipid biosynthesis and ER morphogenesis, as these\nassays do not involve lipid production (Figure 6). We have discussed these possibilities in the revised\nmanuscript. Thank you for your insightful suggestions. We have also cited key studies on lipid\nbiosynthesis and ER morphology in the budding yeast Saccharomyces cerevisiae.\n1. Provide quantification of % tubular ER for Figure 1B.\nRESPONSE 2: Thank you for this important suggestion. In our knockdown screen, we targeted genes\nencoding 18 ER-localized PA-producing enzymes and a control (RLuc) using RNAi in two independent\nexperiments. In control MEFs (RLuc knockdown), ER tubules constituted approximately 50% of the ER\nnetwork. Based on our previous work on Drp1 (Adachi et al., Mol Cell, 2020), which showed that Drp1\nloss reduced ER tubules to approximately 15%, we set a threshold of approximately half the control level\nas the criterion for a positive hit. This level of reduction can be readily identified by visual inspection\nwithout quantification. Therefore, to efficiently screen many genes, we qualitatively assessed ER\nmorphology without percentage quantification. In all knockdown conditions, ER tubule levels were\n4\ncomparable to those in control cells, except for AGPAT2 knockdown, which consistently showed a\nmarked reduction in ER tubules in both experimental replicates. We then validated the effects of\nAGPAT2 deficiency using shRNA-mediated knockdown and CRISPR-based knockout approaches. In these\nfollow-up experiments, we quantified ER tubules. In the revised manuscript, we have moved the screen\ndata to Supplementary Figure 2 and expanded the description of the screen in the Methods section.\n2. Figure 2: Throughout the manuscript, the authors measure “tubular ER %”. It is unclear from the\nresults and materials and methods sections how this percentage is being calculated as it seems that only\none ER marker (SEC61B) is being used. SEC61B is used as a general ER marker and localizes to both\nER sheets and tubules (as can be seen in microscopy images in Figure 1). A better explanation showing\nhow this percentage is calculated should be given as this ratio is used in multiple figures.\nRESPONSE 3: We appreciate this helpful suggestion regarding the quantification method. To assess ER\nmorphology, we measured the area of the cell occupied by the ER—whether composed of tubules or\nsheets—using the ER marker SEC61β fused to GFP. The total ER area was defined as the combined area\nof ER tubules and sheets. We then calculated the relative area of ER tubules by dividing the area\noccupied by tubules by the total ER area. All measurements were performed using NIH Fiji software. In\nthe revised manuscript, we have included this expanded description of the ER morphology\nquantification in the Methods section.\n3. Need more evidence to show AGPAT2 plays a role in ER morphology. Overexpression of RTN4A in\nAGPAT2 KO cells to check if the ER tubules can be restored supporting Figure 2B.\nRESPONSE 4: We agree with the reviewer on the importance of providing additional evidence for the\nrole of AGPAT2 in ER morphology. As suggested, we overexpressed RTN4A in AGPAT2-KO cells. Our new\nresults demonstrate that RTN4A overexpression induces ER tubulation in AGPAT2-KO cells (Figure 5e, f).\nThese findings further support our conclusion that AGPAT2 and RTN4A regulate ER tubulation through\ndistinct mechanisms.\n4. Does AGPAT2-KO have increased ER stress which can alter ER morphology?\nRESPONSE 5: Thank you for this insightful question. In our revision, we assessed ER stress by examining\nER stress markers, including BiP and CHOP expression levels, as well as phosphorylation of eIF2α (Figure\n3). Our new results show that ER stress is not elevated in AGPAT2-KO cells compared to WT cells. As a\npositive control, we treated cells with tunicamycin and confirmed robust activation of the ER stress\nresponse.\n5. Is the expression of reticulons and reticulon-like proteins normal in AGPAT2?\nRESPONSE 6: As suggested, we analyzed the protein levels of reticulons during the revision. Among the\nfour reticulons in mice, RTN3 and RTN4 are highly expressed in MEFs, whereas RTN1 and RTN2 are\nexpressed at very low levels or are undetectable (www.ebi.ac.uk/gxa/home). Therefore, we examined\nRTN3A and RTN4A levels in WT and AGPAT2-KO MEFs. Our new results show that RTN3A levels remain\nunchanged between genotypes, while RTN4A levels are modestly increased in AGPAT2-KO MEFs\n(Supplementary Figure 6). We speculate that this upregulation may represent a compensatory response\nto reduced ER tubules in AGPAT2-KO cells. We also examined the level of the sheet-forming protein\nClimp63 and found it to be reduced in AGPAT2-KO MEFs (Supplementary Figure 6), which may similarly\nreflect a compensatory mechanism. It will be interesting to test in future studies whether restoring\nRTN4A or Climp63 levels affects ER morphology in AGPAT2-KO cells.\n5\n6. What is the ER membrane lipid composition in AGPAT2?\nRESPONSE 7: To address this valuable question, we isolated ER fractions from WT and AGPAT2-KO MEFs\nand performed lipidomic analysis. We found no significant differences in the levels of PA or other\nphospholipids—including LPA, PG, PI, PS, PC, PE, and DAG—between the two cell types (Supplementary\nFigure 8b). These results indicate that the loss of AGPAT2 does not alter the steady-state levels of PA.\nWe speculate that the presence of multiple PA-generating enzymes at the ER may compensate for the\nloss of AGPAT2, maintaining overall PA content. These findings suggest that AGPAT2 does not regulate\nER morphology by modulating steady-state PA levels (e.g., by depleting PA in the ER), but rather by\nsupplying PA to Drp1 through the AGPAT2-Drp1 interaction.\n7. Figure 4 is not convincing. Unable to see actual ER tubules and sheets due to magenta highlights.\nRESPONSE 8: We appreciate the reviewer’s feedback. To improve clarity and better visualize ER tubules,\nwe have removed the magenta highlights in the revised Figure 6 and Supplementary Figure 7. We\nbelieve the updated images now more clearly display the ER tubules and sheets.\n8. Can AGPAT2 addition in DRP1 KO and AGPAT2 KO cells increase tubulation in isolated ER?\nRESPONSE 9: This is an interesting point. However, AGPAT2 is an integral membrane protein and, under\nour assay conditions, cannot be readily added to isolated ER in vitro.\n9. Is AGPAT2-KO ER membrane completely devoid of PA? As, addition of external PA is tubulating the\nER membrane in Figure 4.\nRESPONSE 10: Thank you for this insightful question. During the revision, we performed quantitative\nlipid mass spectrometry on isolated ER from WT and AGPAT2-deficient cells and found similar levels of\nPA, as described in RESPONSE 7. These results support the presence of multiple PA-producing enzymes\nand argue against the possibility that AGPAT2 is required for ER tubulation solely through maintaining\noverall PA levels in the ER. Instead, the new data, together with our original findings, suggest that\nAGPAT2 is critical for Drp1-dependent ER tubulation by interacting with Drp1 and potentially providing a\nlocalized source of this low-abundance phospholipid. In our in vitro assay, the addition of exogenous PA\nmay create an excess relative to steady-state conditions, thereby bypassing the otherwise necessary\nAGPAT2–DRP1 interaction. We have added this discussion to the revised manuscript.\n10. Measure PA and DAG to support 5i.\nRESPONSE 11: As suggested, we measured PA and DAG levels in LIPIN1-overexpressing cells during the\nrevision process using lipid mass spectrometry of isolated ER. As shown in the original Figure 5i (now\nFigure 7i in the revised manuscript), we compared AGPAT2-KO cells reconstituted with AGPAT2 (control)\nto AGPAT2-KO cells co-expressing AGPAT2 and LIPIN1. The new results show that ectopic expression of\nLIPIN1 significantly reduces PA levels while DAG levels remain unchanged (Supplementary Figure 8a).\nThese findings support our conclusion that decreased PA levels lead to reduced ER tubulation.\nRegarding the unchanged DAG levels, we speculate that although LIPIN1 overexpression increases DAG\nproduction, the DAG may be rapidly converted into downstream products such as triacylglycerols or\nother phospholipids. Alternatively, because DAG is 30–40 times more abundant than PA (Supplementary\nFig. 8a), the conversion of PA to DAG may not be detectable upon LIPIN1 overexpression.\n6\n11. Quantification of western blots would be helpful, especially when authors claim that there is an\nincrease or decrease in protein levels.\nRESPONSE 12: Thank you for this important suggestion. In the revised manuscript, we have included\nquantification of the Western blot data to support our findings.\n12. Does lipin OE lead to increased ER sheets?\nRESPONSE 13: This experiment was originally presented in Figure 5h and 5i of the initial manuscript\n(now Figure 7h and 7i in the revised version). In this analysis, we compared AGPAT2-KO cells\nreconstituted with AGPAT2 (control) to AGPAT2-KO cells co-expressing AGPAT2 and LIPIN1. The results\nshow an increase in ER sheets upon LIPIN1 overexpression.\n13. Inference from figure 6i not convincing. CLIMP63 seen in mitochondria.\nRESPONSE 14: The primary goal of the experiment in Figure 6i (Figure 8k in the revised manuscript) was\nto assess DRP1 levels in the ER fraction. To this end, we optimized our subcellular fractionation protocol\nto minimize mitochondrial contamination in the ER fraction. While the reviewer is correct that some ER\nproteins may appear in the mitochondrial fraction, the key point is that our ER fraction is highly enriched\nand devoid of mitochondrial contamination, as demonstrated by the absence of the mitochondrial\nmarker TIM23. Therefore, the data reliably support our original conclusion that DRP1 levels in the ER\nfraction are comparable between WT and AGPAT2-KO cells.\nMinor-\n1. Figure 5G: should read “AGPAT2” instead of “AGPTA2” in figure\nRESPONSE 15: Thank you for pointing out the typo. It has been corrected.\nReviewer #3:\nIn this paper, the authors investigate how lipid biosynthesis influences the three-dimensional organization\nof the ER membrane.\nRESPONSE 1: We are grateful to this reviewer for carefully reading the manuscript and providing many\nvital suggestions, which immensely helped improve our revised manuscript. We have addressed all the\ncomments and incorporated the recommendations outlined below. Changes in the text are shown in red\nfont in the revised manuscript.\nMajor comments:\nThe EM work done both using TEM and SEM is not described well . It is not clear how the samples were\nprepared and therefore it is challenging to evaluate the interpretation of the results.\nPlease elaborate on how you perform sample prep for both TEM and SEM in an way that will allow the\nreader to repeat your experiment.\nRESPONSE 2: We agree with the reviewer that the original manuscript lacked sufficient detail regarding\nthe FIB-SEM methodology. During the revision process, we expanded the Methods section to include a\n7\nmore comprehensive description of the sample preparation and imaging procedures. Additionally, we\nadded a new figure illustrating each step of the FIB-SEM workflow to enhance clarity (Supplementary\nFigure 4). We also clarified in the revised manuscript that only SEM was performed in this study.\nMoreover- The graphs in the entire paper are too small to be seen, the data can not be understood that\nway.\nRESPONSE 3: As recommended, we have enlarged all graphs throughout the revised manuscript to\nimprove readability and ensure the data are clearly presented.\nSpecific Commets:\nPage 3 line 61: The full name of PA, \"phosphatidic acid,\" is missing from the main text\nRESPONSE 4: Thank you for this important suggestion. We have added the full name of PA to the revised\nmanuscript.\nPage 4 line 70 : The full name of PC, is missing from the main text\nRESPONSE 5: We have included the full name of PC in the revised manuscript.\nline 82: Figure 1b is not presented well; the result is not clearly evident. Please mark the AGPAT2\nmicrograph differently, and ensure the titles are capitalized as written in the text: AGPAT2 instead of\nagpat2.\nRESPONSE 6: Thank you for these important suggestions. In the revised manuscript, we have highlighted\nthe AGPAT2 image using red font in the new Supplementary Figure 2 and have capitalized all enzyme\nnames as recommended.\nFig2i:\nThe 3D representation of the segmented surface data should be displayed alongside the data itself—which\nmeans it should be presented alongside at least one slice through the volume. Moreover, a scale cannot be\nadded to the figure if the surface is presented in a perspective view. Please provide a scale on the slice,\nand for the surface, specify the FOV dimensions.\nRESPONSE 7: We agree with the reviewer on these important suggestions and have made the following\nchanges in the revised manuscript: (1) added one individual image for each cell type, each with a scale\nbar (Figure 2a); (2) removed scale bars from the reconstructed images for clarity (Figure 2b); and (3)\nprovided the field of view dimensions—8.1 × 6.8 × 0.4 µm for WT cells, 8.0 × 7.0 × 0.5 µm for DRP1-KO\ncells, and 8.4 × 6.4 × 0.5 µm for AGPAT2-KO cells (Figure 2a and its legend).\nMore importantly: What is the field of view of the imaged block? Is it identical for all three samples?\nI’m a bit confused by your materials and methods—are you using block-face imaging or array tomography?\nPlease provide more information regarding the sample preparation.\nRESPONSE 8: The sizes of the imaged blocks are similar but not identical across the three samples, as\nnoted in RESPONSE 7. This information has been added to the revised manuscript (Figure 2a and its\nlegend). Regarding the imaging method used to generate the 3D datasets, we exclusively used FIB-SEM.\nWe have corrected the Methods section accordingly to clarify this point. Additionally, as described in\nRESPONSE 2, we have included a new figure (Supplementary Figure 4) that outlines the sample\n8\npreparation steps and further confirms that FIB-SEM was the technique used in this study.\nMaterial and method section:\nLine 387: Please provide more details. Was the cell monolayer embedded in resin? Which resin? what\nare the steps of fixation and embedding used? How did you manage to obtain 500 nm thick slices through\na cell monolayer? Which microtome did you use? How were the 500 nm sections organized and lifted on\nthe wafer? manually?\nRESPONSE 9: We are grateful for these critical questions. In the revised manuscript, we have added the\nfollowing detailed description of the sample preparation procedure:\nWT, DRP1-KO, and AGPAT2-KO cells were cultured as monolayers on glass coverslips and pre-fixed in\n2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer (pH 7.4) overnight at ice temperature. Post-\nfixation was performed with a mixture of 1% osmium tetroxide (OsO ) and 1% potassium ferrocyanide in\n4\n0.1 M sodium cacodylate buffer for 15 minutes at ice temperature. The cells were further contrasted\nwith 1% thiocarbohydrazide, 1% OsO , and 1% uranyl acetate, each applied for 5 minutes at room\n4\ntemperature.\nFollowing this, samples were treated with Walton’s lead aspartate for 10 minutes at 60°C, dehydrated\nthrough a graded ethanol series (10%, 30%, 50%, 70%, 80%, 95%, and 100%), and embedded in\nDurcupan resin using beam capsules. Thermal polymerization was carried out at 70°C for 48 hours. After\npolymerization, the glass coverslip was removed by immersion in liquid nitrogen, and the embedded cell\nmonolayer was transferred to the surface of the resin block.\nSerial sectioning was performed from the resin block face adjacent to the coverslip using a Leica EM\nARTOS 3D ultramicrotome (Leica Microsystems) equipped with a DiATOME diamond knife. Sections\nmeasuring 600 µm × 600 µm with a thickness of 1 µm were collected. Six to seven serial sections were\nmanually transferred onto a 10 × 10 mm Si wafer. This was achieved by partially submerging the wafer\ninto the water-filled trough of the diamond knife and aligning it with the knife edge. The sections were\ncollected at the interface between the wafer and the water surface. Water was then slowly drained\nfrom the trough, allowing the sections to adhere to the wafer surface. The wafer was subsequently dried\non a heat block at 45°C for 3 hours. The serial order of the sections was preserved throughout the\nprocess.\nLine392: What do you mean platinum rectangle was deposited by SEM? do you mean by GIS system\ninstalled at the SEM?\nRESPONSE 10: Yes, we used the FEI Helios Nanolab 650 DualBeam FIB-SEM, which is equipped with a gas\ninjection system (GIS) for platinum deposition and milling. We have clarified this point in the revised\nmanuscript.\nLine 395: What is the thickness of the final lamella obtain after the polish?\nRESPONSE 11: The final lamella thickness after polishing was 0.4 µm for WT cells and 0.5 µm for both\nDRP1-KO and AGPAT2-KO cells. This information has been added to the Methods section and Figure 2a.\nLine 400: What are the overall dimensions of the imaged block?\nRESPONSE 12: As noted in RESPONSE 7, the dimensions of the imaged blocks (width × height ×\n9\nthickness) are as follows: 8.1 × 6.8 × 0.4 µm for WT cells, 8.0 × 7.0 × 0.5 µm for DRP1-KO cells, and 8.4 ×\n6.4 × 0.5 µm for AGPAT2-KO cells.\n10"}, {"round": 2, "reviewer_comments": {"reviewer_1": "Although the authors have attempted to address the many queries previously raised, several of the serious concerns remain and are consigned by the authors to future studies, which severely undermine the claim that Drp1 tubulates ER membranes in a PA/AGPAT2-dependent manner, independent of its self-assembly and GTPase activites. The major concerns that have not been addressed are:\n(i) Evidence that full-length Drp1 rid of BOTH its self-assembly (G350D) and GTPase activities (K38A) can tubulate PA-containing liposomes is conspicuously absent. No data for WT is shown either. This is absolutely essential for demonstrating that full-length Drp1 possesses assembly-independent membrane remodeling activities with a distinct, resolvable mechanism.\n(ii) Evidence that endogenous Drp1 decorates isolated native ER tubes is not directly and unequivocally demonstrated. It is moreover unclear how antibodies targeting Drp1 embedded in pre-formed ER tubes purportedly in an unassembled and GTPase activity-impaired basal state disrupt ER tubulation.\n(iii) From the data presented, it is plausible that Drp1 independent of its self-assembly and GTPase activities promotes the assembly of another protein factor that tubulates ER membranes in a PA-dependent manner, instead of tubulating it by itself. This is not considered or addressed.\n(iv) The significance of PA-dependent, Drp1-decorated ER tubes, if not essential for the formation of ER-mitochondrial contact sites or contact-dependent mitochondrial constriction as suggested in the authors’ response, is not elaborated in the discussion. Adachi et al. 2020 suggested that Drp1-dependent ER tubulation is essential for ER-mitochondria contact site formation and ER tubule-dependent mitochondrial constriction and division.\n(v) The dot-blot assay presented in Supp Fig. 1 to address the lipid specificity of the D-octapeptide is weak and not convincing. This must be addressed using liposomes (lipid bilayers) in suspension.", "reviewer_2": "How was the relative area occupied by ER tubules calculated? Elaborate in methods how area specifically occupied by ER tubules was measured.\nThe authors have addressed all other concerns and I think new experimental additions have greatly improved the paper.", "reviewer_3": "The authors have gone to great lengths to address all the comments and concerns I've raised during the review. The manuscript at its current state is well presented and written in a clear and informative manner, especially regarding the EM part which is the focus of most of my comments."}, "author_response": "RESPONSES TO REVIEWERS' COMMENTS\nReviewer #1\nAlthough the authors have attempted to address the many queries previously raised, several of the serious\nconcerns remain and are consigned by the authors to future studies, which severely undermine the claim\nthat Drp1 tubulates ER membranes in a PA/AGPAT2-dependent manner, independent of its self-assembly\nand GTPase activites. The major concerns that have not been addressed are:\n(i) Evidence that full-length Drp1 rid of BOTH its self-assembly (G350D) and GTPase activities (K38A)\ncan tubulate PA-containing liposomes is conspicuously absent. No data for WT is shown either. This is\nabsolutely essential for demonstrating that full-length Drp1 possesses assembly-independent membrane\nremodeling activities with a distinct, resolvable mechanism.\nRESPONSE 1:\n[TEXT REDACTED]\n(ii) Evidence that endogenous Drp1 decorates isolated native ER tubes is not directly and unequivocally\ndemonstrated. It is moreover unclear how antibodies targeting Drp1 embedded in pre-formed ER tubes\npurportedly in an unassembled and GTPase activity-impaired basal state disrupt ER tubulation.\nRESPONSE 2: Our data show that Drp1 is associated with both ER tubules and sheets in cells (Fig. 8g),\nand that isolated ER fractions contain Drp1 (Fig. 8k). Furthermore, incubation with anti-Drp1\nantibodies—but not control anti-Tom20 antibodies—abolished ER tubules in vitro (Fig. 6j, k). These\nresults strongly support the conclusion that Drp1 is critical for the maintenance of ER tubules in vitro.\nWe speculate that antibody-mediated inhibition does not occur through direct interference with\nmembrane-embedded regions of Drp1, but rather by binding to other domains of Drp1, thereby altering\nits conformation to one less favorable for membrane association or deformation. We believe that a\nmore detailed investigation of Drp1’s specific association with ER tubules is beyond the scope of the\ncurrent study.\n(iii) From the data presented, it is plausible that Drp1 independent of its self-assembly and GTPase\nactivities promotes the assembly of another protein factor that tubulates ER membranes in a PA-\ndependent manner, instead of tubulating it by itself. This is not considered or addressed.\nRESPONSE 3: We agree with the reviewer that additional ER protein(s) may cooperate with Drp1 in ER\ntubulation, and that Drp1’s role could be to stimulate a separate tubulation factor. While this model\ncannot be excluded, our new data described in RESPONSE 1 support the notion that Drp1 is sufficient to\ndrive membrane tubulation through a PA-dependent, but GTP- and oligomerization-independent\nmechanism.\n(iv) The significance of PA-dependent, Drp1-decorated ER tubes, if not essential for the formation of ER-\nmitochondrial contact sites or contact-dependent mitochondrial constriction as suggested in the authors’\nresponse, is not elaborated in the discussion. Adachi et al. 2020 suggested that Drp1-dependent ER\ntubulation is essential for ER-mitochondria contact site formation and ER tubule-dependent mitochondrial\nconstriction and division.\nRESPONSE 4: We appreciate the insightful comments. As the reviewer correctly pointed out, we have\npreviously suggested that Drp1-dependent ER tubulation may play an important role in ER–\nmitochondria contact sites and mitochondrial division. However, we did not investigate the role of Drp1-\ndependent ER tubules in these processes in the current study.\n(v) The dot-blot assay presented in Supp Fig. 1 to address the lipid specificity of the D-octapeptide is\nweak and not convincing. This must be addressed using liposomes (lipid bilayers) in suspension.\nRESPONSE 5: Thank you for the important suggestion. As requested, we tested the lipid-binding\nspecificity of the D-octadecapeptide using liposomes. New results show that the D-octadecapeptide\nspecifically binds to liposomes containing PA, but not those containing PS, cardiolipin, PI(4,5)P₂ or PI4P\n(Supplementary Fig. 1b, c), confirming the results of the lipid dot blot assay (Supplementary Fig. 1a).\nReviewer #2\nHow was the relative area occupied by ER tubules calculated? Elaborate in methods how area specifically\noccupied by ER tubules was measured.\nThe authors have addressed all other concerns and I think new experimental additions have greatly\nimproved the paper.\nRESPONSE 5: Thank you for the positive comment and helpful suggestion. We have expanded the\ndescription of how we measured the area occupied by ER tubules in the revised Methods section."}, {"round": 3, "reviewer_comments": {"reviewer_1": "The authors have attempted to address my remaining concerns through additional experiments. However, a few nagging concerns remain on their latest experimental design and data interpretation. These will have to be addressed, at least in word, in the text.\n1.\n[TEXT REDACTED]\n2.\n[TEXT REDACTED]\n3.\n[TEXT REDACTED]\n4.\n[TEXT REDACTED]\n5.\n[TEXT REDACTED]\nCited references\n1. Francy, C. A., Alvarez, F. J., Zhou, L., Ramachandran, R., and Mears, J. A. (2015) The Mechanoenzymatic Core of\nDynamin-related Protein 1 Comprises the Minimal Machinery Required for Membrane Constriction J Biol Chem 290, 11692-\n11703\n2. Macdonald, P. J., Stepanyants, N., Mehrotra, N., Mears, J. A., Qi, X., Sesaki, H. et al. (2014) A dimeric equilibrium\nintermediate nucleates Drp1 reassembly on mitochondrial membranes for fission Mol Biol Cell 25, 1905-1915\n3. Ugarte-Uribe, B., Muller, H. M., Otsuki, M., Nickel, W., and Garcia-Saez, A. J. (2014) Dynamin-related protein 1 (Drp1)\npromotes structural intermediates of membrane division J Biol Chem 289, 30645-30656\n4. Lu, B., Kennedy, B., Clinton, R. W., Wang, E. J., McHugh, D., Stepanyants, N. et al. (2018) Steric interference from\nintrinsically disordered regions controls dynamin-related protein 1 self-assembly during mitochondrial fission Sci Rep 8,\n10879", "reviewer_2": "Authors have expanded their explanation for how the ER was quantified. I have no more comments."}, "author_response": "RESPONSES TO REVIEWERS' COMMENTS\nReviewer #1:\nThe authors have attempted to address my remaining concerns through additional experiments. However,\na few nagging concerns remain on their latest experimental design and data interpretation. These will\nhave to be addressed, at least in word, in the text.\n1.\n[TEXT REDACTED]\nRESPONSE 1:\n[TEXT REDACTED]\n2.\n[TEXT REDACTED]\nRESPONSE 2:\n[TEXT REDACTED]\n3.\n[TEXT REDACTED]\nRESPONSE 3:\n[TEXT REDACTED]\n4.\n[TEXT REDACTED]\nRESPONSE 4:\n[TEXT REDACTED]\n5.\n[TEXT REDACTED]\nRESPONSE 5:\n[TEXT REDACTED]\nCited references\n1. Francy, C. A., Alvarez, F. J., Zhou, L., Ramachandran, R., and Mears, J. A. (2015) The\nMechanoenzymatic Core of Dynamin-related Protein 1 Comprises the Minimal Machinery Required for\nMembrane Constriction J Biol Chem 290, 11692-11703\n2. Macdonald, P. J., Stepanyants, N., Mehrotra, N., Mears, J. A., Qi, X., Sesaki, H. et al. (2014) A dimeric\nequilibrium intermediate nucleates Drp1 reassembly on mitochondrial membranes for fission Mol Biol\nCell 25, 1905-1915\n3. Ugarte-Uribe, B., Muller, H. M., Otsuki, M., Nickel, W., and Garcia-Saez, A. J. (2014) Dynamin-\nrelated protein 1 (Drp1) promotes structural intermediates of membrane division J Biol Chem 289, 30645-\n30656\n4. Lu, B., Kennedy, B., Clinton, R. W., Wang, E. J., McHugh, D., Stepanyants, N. et al. (2018) Steric\ninterference from intrinsically disordered regions controls dynamin-related protein 1 self-assembly during\nmitochondrial fission Sci Rep 8, 10879"}, {"round": 4, "reviewer_comments": {"reviewer_1": "The authors have decided to remove [TEXT REDACTED] altogether and instead discuss alternative interpretations in the text. I'm okay with that. Quantification of ER tubule diameter (apparently induced by Drp1) however should be included in the revised manuscript and should be mentioned as being narrower to what was described for Drp1 previously. The Drp1 isoform used in this study (isoform 3?) should also be explicitly mentioned. As for the K38A mutant, it can bind GTP but cannot hydrolyze it. Therefore, adding GTP to the reaction to induce GTP binding-dependent conformational changes might help address the requirements for the reported PA-dependent membrane tubulation. This does not have to be done now given the above revisions. I have no further concerns."}, "author_response": "RESPONSE: Thank you for your comments. As requested, we have added and discussed the data on the\nwidth of ER tubules generated by DRP1 in the revised manuscript (Fig. 6h). We have also provided\ninformation on the specific DRP1 isoform used in the current study."}]

Decision letter