SNX-mediated biogenesis of a plant-unique vesicle derived from the multivesicular body

Domain: biology
Published: 2026-03-24
Editorial outcome: major_revision

Abstract
Retrograde transport is central to endomembrane homeostasis, yet the identity and origin of plant retrograde carriers remain unresolved. Prevailing models propose that plant vacuolar sorting receptors (VSRs) recycle either from multivesicular bodies (MVBs) to the trans-Golgi network (TGN) or from the TGN to the Golgi apparatus and/or endoplasmic reticulum (ER). However, the ultrastructural features of plant retrograde transport carriers remain largely unresolved. Here, we show that plant retrograde transport is likely mediated by a previously unrecognized class of MVB-derived spherical vesicles. Using correlative light and electron microscopy and three-dimensional electron tomography, we identify a distinct population of ~30–50 nm spherical vesicles adjacent to MVBs, including nascent vesicles budding from the MVB limiting membrane in Arabidopsis root cells. Immunogold labeling shows that these vesicles are enriched in retromer components and VSRs, suggesting that they possibly function as retrograde transport carriers. To investigate their biogenesis, we perform cryo-electron microscopy and liposome tubulation assays, showing that Arabidopsis SNX1 generates shorter membrane tubules than its mammalian counterpart, consistent with reduced membrane affinity linked to differences in the amphipathic helix. Notably, the SNX1-SNX2 heterodimer produces heterogeneous structures, including spherical vesicles, recapitulating in vivo observations. Lastly, knockdown of SNX1 or SNX2 results in vacuolar mislocalization and increased degradation of GFP-VSR2, and defects in SNX1 and VPS29 inhibit formation of spherical vesicles adjacent to MVBs, resulting in embryonic lethality before the globular stage. Together, these findings establish MVB-derived spherical vesicles as plant retrograde carriers and reveal a distinct SNX-mediated mechanism underlying their formation.

Introduction
All eukaryotic cells possess a highly efficient endomembrane system that ensures the precise transport of biomacromolecules, such as proteins, to their designated locations, allowing them to carry out their functions effectively1. This system depends on tightly regulated anterograde and retrograde trafficking to deliver cargo and recycle components. Lysosomes in animal cells and vacuoles in plant and fungal cells function as central degradation compartments and play essential roles in nutrient recycling, signal transduction, membrane turnover, and the maintenance of cellular homeostasis. In animal cells, lysosome biogenesis requires the coordinated synthesis of lysosomal proteins and their trafficking through the endosomal system. Lysosomal hydrolases are synthesized in the endoplasmic reticulum (ER), modified in the Golgi apparatus with mannose 6-phosphate tags, and recognized by mannose 6-P receptor (MPR). The hydrolase-MPR complexes are then packaged at the trans-Golgi network (TGN). Lysosomal proteins are delivered either directly from the TGN to lysosomes via endosomes or indirectly via the plasma membrane (PM) through endocytosis. Proteins such as receptors that are not destined for lysosomes are recycled back to the PM or Golgi. In endocytosis, internalized membrane proteins can be directed to lysosomes or retrieved into tubulo-vesicular carriers for recycling—either directly back to the PM (fast recycling), indirectly via the endocytic recycling endosomes (slow recycling), or, in some cases, back to the secretory pathway2. Defects in these transport processes are associated with various neurodegenerative disorders3-5. In plants, retrograde transport is thought to be closely related to numerous physiological functions, including embryo development, plant morphogenesis, gravity response, and disease resistance6-16. However, the subcellular localization and formation mechanisms of retrograde transport carriers involved in recycling vacuolar sorting receptors (VSRs) in plants remain elusive, leading to two prevailing hypotheses15, 17. The classical hypothesis suggests that retrograde transport primarily mediates the retrieval of VSRs from multivesicular bodies (MVBs), also referred to as prevacuolar compartments (PVCs), back to the TGN18, 19. In this model, the recognition between VSRs and cargo proteins occurs at the TGN. Upon completion of sorting, VSRs and cargo proteins separate within the lumen of the MVB, after which VSRs are retransferred to the TGN to participate in the next round of cargo protein sorting20, 21. Conversely, a more recent hypothesis suggests that the recognition between VSRs and cargo proteins occurs early in the ER, with sorting completed in the TGN. Thus, retrograde transport mainly mediates the transport of VSRs from the TGN to the Golgi apparatus and/or the ER to engage in subsequent rounds of cargo protein sorting22, 23. In addition to the ongoing controversy surrounding their subcellular localization, the ultrastructure and formation mechanisms of plant retrograde transport carriers remain elusive23-25.

In yeast, retrograde transport is mediated by the retromer complex, a pentamer composed of a membrane-binding SNX subcomplex, formed by the heterodimer Vps5p and Vps17p, and a cargo-recognition VPS subcomplex comprising Vps26p, Vps29p, and Vps35p26. In animals, SNX1 and SNX2 are homologous to Vps5p, whereas SNX5 and SNX6 share a common ancestor with Vps17p and are closely related. The SNX protein family commonly features two specialized functional domains: a BAR (Bin/amphiphysin/Rvs) domain, which enables membrane curvature, and a PX (phox homology) domain, which binds to phosphoinositides. SNX1 and SNX2 interact with phosphatidylinositol 3-phosphate (PI(3)P) and phosphatidylinositol 3,5-bisphosphate (PtdIns(3,5)P2) through the PX domain, thereby directly anchoring to the membrane27. The VPS subcomplex consists of VPS26-VPS29-VPS35, where VPS35 interacts with VPS26 and VPS29 through its N-terminal and C-terminal domains, respectively, forming a horseshoe-shaped α-helix28. Unlike the stable heteropentameric retromer complex in yeast, the SNX-BAR heterodimers SNX1/2–SNX5/6 in animals can assemble into functionally distinct endosomal sorting complexes (ESCPE-1) that operate independently of the retromer core29. In Arabidopsis, homologs of the Vps5p and VPS proteins are present, comprising three genes encoding SNX (SNX1, SNX2a, and SNX2b), three encoding VPS35 (VPS35a, VPS35b, and VPS35c), two encoding VPS26 (VPS26a and VPS26b), and one encoding VPS29. SNX1 has been shown to be essential for the recycling of PIN2, BRI1, IRT1, PHT1, and SOS1 to the plasma membrane14, 20, 30-32. Recent studies indicate that the retrieval of VAMP727, a protein involved in vacuolar fusion in plants, from the vacuolar membrane depends on the SNX complex but not on the VPS complex33. In addition, VPS complex also participates in proper host invasion in rice blast34-36. Although the structure and function of retromer components in animals and yeast have been increasingly well characterized, the mechanism by which the retromer complex mediates retrograde transport in plants remains unclear37-41.

In this study, we combined correlative light and electron microscopy (CLEM), 3D electron tomography (ET), cryo-EM, and all-atom molecular dynamics simulations, along with cell biology and genetic approaches, to investigate plant retrograde transport in the model plant Arabidopsis thaliana, with the following major findings: (1) The CLEM, 3D ET, and immunogold transmission electron microscopy (TEM) analyses revealed distinct spherical vesicles (~30–50 nm in diameter) labeled by retromer markers and VSR proteins in Arabidopsis root cells, with nascent vesicles budding from MVBs in Arabidopsis root cells; (2) We applied cryo-EM to demonstrate that, when incubated with liposomes, Arabidopsis MVB-localized, BAR domain-containing SNX proteins generate significantly shorter tubules than mouse SNX1 in vitro; (3) The combined structural data and all-atom molecular dynamics simulations indicate that Arabidopsis SNX1 has a reduced membrane affinity compared to mouse SNX1, primarily due to differences in its AH domain, which in turn restricting the formation of longer tubular structures relative to those formed by mouse SNX1; (4) The Arabidopsis SNX1-SNX2 heterodimer produces a more heterogeneous mixture of irregular tubules and spherical vesicles when incubated with liposomes in vitro, closely mirroring the formation of the spherical vesicles near MVBs observed in vivo; and (5) We demonstrated that knockdown of SNX1 or SNX2 resulted in vacuolar mislocalization and increased degradation of GFP-VSR2 and impaired proper seed development and germination in plants. Disruptions in SNX1 and the core retromer component VPS29 impair formation of the spherical vesicles near MVBs, lead to the development of enlarged MVBs, and result in embryonic lethality before the globular stage. Taken together, our study provides evidence for the structural features and biogenesis of plant-unique vesicles derived from MVBs and mediated by SNX1 and SNX2.

Results
Plant retromer markers and VSRs localize to MVBs. To investigate plant retrograde transport, we first performed a systematic analysis of the subcellular localization of retromer markers and VSRs, which serve as the transport machinery and cargoes, respectively, for retrograde trafficking. Confocal imaging analysis of transgenic Arabidopsis root cells revealed that VSR1, the key retromer component VPS29 and SNX1, partially colocalized with MVBs labelled by RHA1/ARA7 (conventional members of the Arabidopsis RAB5 family) and RABG3f (a member of the RAB7 family) (Supplementary Fig. 1). However, these markers did not colocalize with the Golgi apparatus or the TGN (Supplementary Fig. 2). Similarly, in protoplasts, VSRs exclusively co-localizes with MVBs labeled by ARA7 and RABG3f, and do not overlap with other organelles such as the ER, Golgi apparatus, or TGN (Supplementary Fig. 3). Collectively, these findings support the hypothesis that retrograde transport may mediate the retrieval of VSRs from MVBs. Furthermore, we show that VSRs specifically colocalize with ARA7 and RABG3f, but not with other Rab GTPases (Supplementary Fig. 4), suggesting that RHA1/ARA7 and RABG3f may play a role in plant retrograde transport.

CLEM, 3D ET analyses, and immuno-EM reveal a plant-unique population of spherical vesicles (~30–50 nm in diameter), with nascent vesicles budding from MVBs. Given that plant retrograde transport likely to mediate the retrieval of VSRs from MVBs, we utilized CLEM and 3D ET to systematically identify and analyze potential structures around MVBs. Using CLEM, we first mapped the positions of all MVBs in a 200-nanometer root slice (Supplementary Fig. 5a). Through a systematic 3D ET analysis, we identified previously uncharacterized spherical vesicle structures, approximately 30 to 50 nm in diameter, located near MVBs (Fig. 1a-f, Supplementary Fig. 5b-t, Supplementary Fig. 6 and Supplementary Video 1). Occasionally, spherical vesicle structures are observed connected to MVBs (Fig. 1d-e, arrowhead), suggesting they may represent newly budded vesicles from the MVB membrane. In addition, we observed that the lumen of these spherical vesicles is more electron-transparent than the intraluminal vesicles (ILVs) of MVBs (Fig. 1f), indicating a lack of cytosolic cargo proteins. This preliminary observation suggests that these spherical vesicles are not plant secretory vesicles, which typically contain cargo proteins and exhibit higher electron density within their lumens under TEM, despite their similar sizes. Next, we applied immunogold labelling to identify the nature of these spherical vesicles. In wild type plants, VSR antibodies predominantly labelled the spherical vesicle and the limiting membrane of the MVB (Fig. 1g, 1j and Supplementary Fig. 7a). Interestingly, in transgenic plants expressing VPS29-GFP and SNX1-GFP, GFP antibodies primarily labelled the limiting membrane of the MVB, with a specific enrichment at the budding vesicle regions (Fig. 1h-i, 1k-l and Supplementary Fig. 7b-c). Taken together, all these results indicate that the newly identified spherical vesicles, with diameters ranging from approximately 30 to 50 nm, may serve in plant retrograde transport and are distinct from the tubular recycling endosomes described in animal systems.

In liposome tubulation assays, Arabidopsis SNX1 induces the formation of shorter tubules compared to those generated by mouse SNX1. 3D ET analysis revealed that spherical vesicles may mediate plant retrograde transport (Fig. 1), a mechanism notably distinct from the tubular recycling endosomes observed in animal and yeast cells26, 42. However, the mechanism underlying this structural difference remains unclear. In animals and yeast, the formation of tubular recycling endosomes is primarily mediated by dimers of SNX1/2 and SNX5/6 (in animals) or Vps5p and Vps17p (in yeast)26. Thus, we speculate that SNX proteins are also likely involved in mediating the formation of spherical vesicles in plants. We begin with a detailed analysis of Arabidopsis SNX1 (AtSNX1). The protein-lipid overlay assay showed that AtSNX1 could bind to PI(3)P (Fig. 2a), consistent with its localization to MVBs, as MVB membranes are enriched in PI(3)P. Although AtSNX1 can bind to other phospholipids such as PI(4)P, it did not colocalize with PI(4)P reporters (Supplementary Fig. 8a), and its punctate localization remained largely unchanged following PI(4)P depletion induced by treatment with 50 μM phenylarsine oxide (PAO), a PI(4)K inhibitor (Supplementary Fig. 8b-f), supporting the notion that its in vivo function is primarily associated with PI(3)P-enriched compartments. We then generated liposomes containing the major phospholipids of organellar membranes along with PI(3)P. In liposome tubulation assays AtSNX1 can induce tubule formation upon the addition of PI(3)P, but not PI(4)P (Fig. 2b, Supplementary Fig. 8g-h). Quantitative liposome sedimentation assays showed that AtSNX1 binds more efficiently to liposomes containing PI(3)P than to liposomes either lacking PI(3)P or containing PI(4)P (Fig. 2c, Supplementary Fig. 8i). Interestingly, the tubules formed by AtSNX1 are significantly shorter than those generated by mouse SNX1 (MmSNX1) (Fig. 2b). To investigate the underlying cause, we further analyzed the structure of AtSNX1 on membrane tubules using cryo-EM. The cryo-EM image showed that the tubules were coated with AtSNX1 (Fig. 2d), with diameters ranging from approximately 30 to 50 nm (Fig. 2e). Two helical diffraction patterns were observed and indexed, referred to as Class I and Class II (Supplementary Fig. 9-10, Supplementary Fig. 11a). We determined the cryo-EM structures of SNX1-coated tubules with a resolution of 7.2 Å for Class Ⅰ and 8.7 Å for Class Ⅱ, as assessed by Fourier Shell Correlation (FSC0.143) (Supplementary Fig. 9 and Supplementary Table 1). The map revealed that Class Ⅰ tubules contain seven dimers per complete helical turn, with a tubule diameter of around 33 nm (Fig. 2e). Observation of the spiral coating structure of Class I tubules revealed the presence of distinct interaction interfaces. The first interaction interface, termed “Interface Ⅰ”, occurs along the same spiral line, where two adjacent SNX1 dimers interact through contacts between the α2 helix and α3 helix of the BAR domain (Fig. 2f, lower left). Another major protein-protein interaction interface, referred to as “Interface Ⅱ”, occurs between adjacent helical rows, facilitating the crosslinking of these parallel rows. The PX domain of the SNX1 dimer interacts with the α2 helix of the BAR domain from another dimer located in an adjacent helical row (Fig. 2f, lower middle). The residues potentially involved in tubule formation at these two interfaces are also highlighted (Supplementary Fig. 12). We also identified an interaction interface between SNX1 dimers, termed “Interface Ⅲ”, which is formed through the interaction of α helices between two BAR domains (Fig. 2f, lower right). Compared to Class Ⅰ tubules, the Class Ⅱ tubules formed by AtSNX1 exhibit a distinct helical symmetry, with six dimers per complete helical turn and a tubule diameter of around 30 nm (Supplementary Fig. 11b). The monomer (asymmetric unit) and dimer configurations are largely similar between Class I and Class II (Supplementary Fig. 13a). When comparing the structures of plant SNX1 with those of their homologs, these proteins adopt a similar overall architecture and display a comparable radius of curvature (~100 Å) (Supplementary Fig. 13a). However, at the level of higher-order scaffold organization, although Class Ⅰ and Class Ⅱ SNX1 dimers exhibit analogous stacking arrangements, the intersection angles of adjacent dimers at Interface Ⅰ show distinct discrepancies (Supplementary Fig. 11c and Supplementary Fig. 13b). Relative to Class Ⅰ, Class Ⅱ displays a smaller intersection angle, which reduces the overall radius of curvature and consequently decreases the number of dimers required to form a single helical turn. In addition, among the plant SNX1 and its homologs, the mouse SNX1 forms the smallest interaction interface at Interface I via the BAR domains while the fungi SNX1 forms the largest (Supplementary Fig. 13b). Notably, these differences in the homotypic BAR-BAR domain interactions does not affect the radius of curvature of the helical lattice. Taken together, our analysis shows that higher-order lattice packing plays an important role in regulating membrane curvature in plant SNX1.

The amphipathic helix (AH) of AtSNX1 is essential for membrane binding of AtSNX1. Consistent with the previously reported MmSNX1, we also observed a segment inserted into the membrane within the AtSNX1 dimer (Fig. 3a). Sequence alignment and helix prediction revealed that this segment forms an AH (Fig. 3b). Unlike MmSNX1, where the AH (280-297aa) is primarily located between the BAR (300-521aa) and PX (139-269aa) domains, the AH (157-176aa) of AtSNX1 is situated at the N-terminus of the BAR domain (160-402aa). The AH features polar or charged residues on one side and hydrophobic residues on the opposite side (Fig. 3c). The non-polar side of the AH inserts into the membrane (Fig. 3c), displacing lipid molecules and inducing curvature, which is essential for curvature-driven protein assembly43. We also investigated the function of the AH in AtSNX1 through mutational analysis. Using membrane-binding assays, we found that replacing residues 157 to 176aa of the AH with a flexible linker (GGGSGGGSGGGS, AH-mutant) significantly reduced AtSNX1’s ability to bind liposomes, thereby impairing its membrane tubulation capacity (Fig. 3d-f). These findings demonstrate that the AH is critical for both membrane binding and deformation by AtSNX1.

All-atom molecular dynamics simulations reveal that AtSNX1 has a lower binding affinity to membranes compared to MmSNX1. The length of tubules generated by SNX1 proteins is influenced by several factors, particularly the membrane-binding affinity of SNX1, which is determined by its AH and directly impacts tubule stability and length37. To further compare the membrane-binding affinities of AtSNX1 and MmSNX1, we performed all-atom molecular dynamics simulations. Two simulations were conducted, one using AtSNX1 and the other using MmSNX1 proteins, each with a timescale of 200 ns. The protein-membrane binding simulations revealed that, for both AtSNX1 and MmSNX1, the interaction with the membrane involved a gradual decrease in the distance between the protein molecules and the membrane, along with the insertion of the AH domain into the membrane (Fig. 4a-f, Supplementary Fig. 14a-f, 15a-b, 15d-e and Supplementary Video 2-3). At 200 ns, the simulations showed that the AH domain of MmSNX1 (right) was fully inserted into the membrane, whereas the AH domain of AtSNX1 was only partially inserted (Fig. 4b,c,e,f, Supplementary Fig. 14b,c,e,f, and Supplementary Fig. 15a-b, 15d-e).

To explore the underlying cause, we compared the cryo-EM structures of AtSNX1 and MmSNX1 and identified several potential factors: (1) differences in their AH tilt angles; notably, The AH domain of MmSNX1 aligns parallel to the membrane upon binding, suggesting a larger lateral contact area and a more stable interaction with the membrane, compared to that of AtSNX1 (Fig. 4g); (2) based on the pseudoatomic model built obtained by AlphaFold and flexible docking into our cryo-EM map, which is further supported by computational membrane interaction prediction, we found that the AH of AtSNX1 has three negatively charged residues (D) and two positively charged residues (K and R) on its polar side, giving it a net negative charge (Fig. 3c and Fig. 4h, left). In contrast, the polar side of the AH in MmSNX1 contains three positively charged residues (K) and one negatively charged residue (D), resulting in an overall positive charge (Fig. 4h, right)44. The cell membrane surface is negatively charged, leading us to speculate that the positively charged AH of MmSNX1 may insert more effectively into the membrane. In contrast, the negatively charged AH of AtSNX1 could experience repulsion from the membrane, potentially reducing its membrane-binding efficiency and impairing subsequent membrane deformation. To further validate these findings, we replaced the AH of MmSNX1 with that of AtSNX1 and evaluated its membrane remodeling ability. The liposome tubulation assay demonstrated that replacing the AH of MmSNX1 with that of AtSNX1 resulted in the formation of shorter tubules, similar to those produced by AtSNX1. This indicates that the AH domain is the key factor underlying the differences in membrane binding and remodeling abilities between MmSNX1 and AtSNX1, and therefore influences the stability and length of the formed tubules (Fig. 4i); (3) In addition, we calculated the interaction energy, Ei(interaction), defined as Ei = Ei(coul) (electrostatic interaction) + Ei(vdw) (van der Waals interaction), which typically indicates attractive forces in all-atom molecular dynamics simulations systems45. The results demonstrated that as the simulation time progressed, the interactions between AtSNX1/MmSNX1 and the membrane gradually strengthened, as indicated by the increasing absolute value of Ei (Supplementary Fig. 15c,f). Notably, the interaction between AtSNX1 and the membrane was relatively weaker than that of MmSNX1, indicating lower attractive forces between AtSNX1 and the membrane compared to those between MmSNX1 and the membrane (Supplementary Fig. 15c,f).

SNX1 and SNX2 jointly induce the formation of spherical vesicles in liposome tubulation assays. In Arabidopsis, six SNX-like proteins have been identified. Yeast two-hybrid (Y2H) experiments revealed that SNX1 interacts not only with itself but also with SNX2 (Fig. 5a), indicating that SNX1 can form a heterodimer with SNX2 to mediate vesicle formation. Consistently, the protein-lipid overlay assay showed that SNX2a, like SNX1, binds to PI(3)P, which is essential for its membrane-binding ability (Fig. 5b-c). To investigate the combined effect of SNX1 and SNX2a on membrane deformation, we purified the SNX1-SNX2a complex (Fig. 5d). The liposome tubulation assay showed that SNX2a alone was unable to deform the liposomes. However, the tubules formed by the SNX1-SNX2a complex were more distorted than those formed by SNX1 alone as revealed by negative-stain TEM (Fig. 5e). Detailed observation of the tubules using cryo-EM revealed that the SNX1-SNX2a heterodimer produces a more heterogeneous mix of irregular tubules and spherical vesicles in vitro (Fig. 5f), closely reflecting the formation of spherical vesicles near MVBs observed in vivo. This suggests that the combined action of SNX1 and SNX2 may play a key role in the formation of spherical vesicles near MVBs in plants. We attempted to analyze the structure of the SNX1-SNX2a complex on membrane tubules using cryo-EM. However, the pronounced irregularity of the spherical vesicles and tubules induced by the SNX1-SNX2a heterodimer hindered the application of iterative helical real-space reconstruction, making it impractical to obtain high-resolution structural information for the complex. To further investigate the membrane-binding properties of the SNX1-SNX2a complex, we performed all-atom molecular dynamics simulations. Protein-membrane binding simulations showed that the AH domain of the SNX1-SNX2a complex resists insertion into the membrane, as average AH‑membrane distance does not decrease over the course of the simulation (Supplementary Fig. 16 and Supplementary Video 4). We have thus proposed that SNX2a modulates the activity of SNX1 by altering the conformation or membrane interaction dynamics of the heterodimer, thereby fine-tuning the curvature generation process. These results indicate that the SNX1-SNX2a complex has an even lower membrane‑binding affinity than the Arabidopsis SNX1 homodimer, leading to the formation of shorter membrane tubules and spherical vesicles, as shown by liposome tubulation assays in vitro.

Knockdown of SNX1 or SNX2 resulted in vacuolar mislocalization and increased degradation of GFP-VSR2 and impaired proper seed development and germination in plants. To investigate the functional roles of SNX1 and SNX2 proteins in retrograde transport, we first examined whether their suppression affects VSR targeting. Notably, RNAi-mediated suppression of SNX1 or SNX2 expression in protoplasts resulted in its mislocalization of GFP-VSR2 to the vacuole (Fig. 6a). Subsequently, we performed protein immunoblotting and semi-quantitative analysis, which revealed that, compared to the control group, the degradation rate of GFP-VSR2 was significantly increased in protoplasts transformed with RNAi vectors targeting SNX1 or SNX2 (Fig. 6b and 6c). Similarly, we found that suppression of the core retromer protein VPS29 or the RAB7 activator MON1 by RNAi, as well as expression of dominant-negative mutants of RAB5 and RAB7 (ARA7S24N and RABG3f T22N), misdirected GFP-VSR2 to the vacuole for degradation as demonstrated by confocal microscopy, protein immunoblotting, and semi-quantitative analysis (Supplementary Fig. 17). These results indicate that SNX1 and SNX2, along with RAB5 (ARA7), RAB7 (RABG3f), and the retromer complex protein VPS29, are required for proper VSR targeting in plant cells.

Since VSRs are crucial for directing cargo proteins to protein storage vacuoles (PSVs) during seed development and mediating the targeting of newly synthesized proteinases to PSVs for seed germination46, we observed significant defects in the seeds of the snx1 mutant and the snx1/snx2a/snx2b triple mutant, as well as GEF mutants of RAB5 and RAB7, such as vps9a-2 and mon1/ccz1ab. In these mutants, protein secretion into the extracellular space (ECS) was evident (indicated by arrows), and PSVs were significantly smaller compared to those in the wild type (WT) (Fig. 6d-f and Supplementary Fig. 18). During seed germination, a delayed degradation of PSVs was also observed in the snx1 mutant and the snx1/snx2a/snx2b triple mutant (Fig. 6d, 6g and Supplementary Fig. 18). Taken together, SNX1 and SNX2, as well as RAB5 (ARA7), RAB7 (RABG3f), and the retromer complex are essential for proper VSR targeting, which play a critical role in seed development and germination in plants.

Defects in SNX1 and the core retromer complex protein VPS29 inhibit formation of spherical vesicles near MVBs, resulting in embryonic lethality before the globular stage. Although SNX1 and SNX2 proteins are essential for VSR targeting, single (snx1, snx2a, snx2b) and double mutants exhibit only mild growth defects (Supplementary Fig. 19a-e, 20a-c). The snx1/snx2a/snx2b triple mutant shows slightly more pronounced defects in overall growth but displays negligible phenotypic changes during embryonic development (Fig. 6h-i and Supplementary Fig. 19f, 20d). We speculate that the core retromer proteins may partially compensate for the functions of SNX1 and SNX2 in plant retrograde transport. To fully disrupt plant retrograde transport, we generated double mutants by crossing the snx1 mutants with the core retromer complex mutant vps29. The snx1-/-vps29+/- mutant exhibited significant defects during the seedling stage. Within the siliques of snx1-/-vps29+/-, seeds showed a high abortion rate (approximately 10.56%, n = 3548), which deviated from the expected theoretical value of 25%. Reciprocal crosses between wild type (Col-0) and snx1-/-vps29+/- revealed that fertility was compromised in the male but not in the female (Supplementary Fig. 21a). To further investigate the underlying cause, we performed Alexander staining to assess pollen viability. After staining, normal pollen appeared dark purple, while abnormal pollen was light green. In wild type and snx1-/-vps29+/-, 96.7 ± 1.3% and 94.7 ± 0.9% of the pollen, respectively, stained normally (Supplementary Fig. 21b-c). Statistical analysis revealed a slight decrease in pollen viability in snx1-/-vps29+/- compared to WT, indicating that the SNX1/VPS29 mutation had a minor effect on pollen viability (Supplementary Fig. 21c). Further analysis revealed that 4 hours after pollen germination in vitro, there was no significant difference in germination rate between WT and the snx1-/-vps29+/- mutant. However, significant shorter pollen tubes were observed in the snx1-/-vps29+/- mutant compared to WT (Supplementary Fig. 21b-c), indicating reduced competitive ability caused by the SNX1/VPS29 mutation. To determine the seed abortion period in snx1-/-vps29+/- mutant offspring, whole-mount clearing was performed. Wild-type seeds contained late heart-shaped embryos at 9 days after pollination (DAP). In contrast, embryos in snx1-/-vps29+/- seeds were arrested at the globular stage and exhibited abnormal cell division (Fig. 7a-d and Supplementary Fig. 20e-f). To further clarify the cause of embryo abnormalities, TEM analysis of embryos of snx1-/-vps29+/- were carried out. The results showed that embryos of snx1-/-vps29+/- exhibited enlarged MVBs and a reduced number of spherical vesicles near MVBs, compared to WT (Fig. 7e-f). In summary, defects in SNX1 and VPS29 inhibit formation of spherical vesicles near MVBs, resulting in embryonic lethality before the globular stage.

Methods
Plant materials and growth conditions. All mutants and transgenic plants were in the Columbia-0 (Col-0) background. The following have been previously described: vps2920, snx152, snx2a52, snx2b52, vps9a58, mon1-246, ccz1a59, ccz1b59, vps35a-1b-2c-17, vps35b-1c-17, GFP-VSR160, VPS29-GFP20, SNX1-GFP/Ch8, mRFP-ESYP32 (WAVE22)61, Ch-RAB5 (WAVE7)61, Ch-RAB7 (WAVE5)61, VHA-a1-mRFP62, mCIT-PHOSBP (P7Y)63. AtSNX1-Ch/mYongHong and MmSNX1-mYongHong were cloned into pCAMBIA1305 vector and generated by the floral dip method64. The primers used are listed in Supplementary Table 2. All seeds were surface sterilized with 75% (v/v) ethanol containing 0.05% Triton X-100 for 10 min, washed with ddH2O five times and then sown on 1/2 MS plates. The plates were kept at 4 °C in the dark for two days before being moved to a growth chamber (22 °C, 16 h light/8 h dark).

High-pressure freezing and freeze substitution. The general procedures for high-pressure freezing and freeze substitution were essentially performed as previously described65. Briefly, Five-day-old Arabidopsis root tips were excised and rapidly frozen using a high-pressure freezing apparatus (EM ICE, Leica). For ultrastructural analysis, freeze substitution was performed using 2% OsO₄ and 0.5% uranyl acetate in anhydrous acetone at −80 °C for 96 hours, followed by a gradual warming to −20 °C over a 24-hour period. The samples were incubated at −20 °C for 12 hours, then gradually warmed to 4 °C over a 4-hour period. Following three acetone rinses at room temperature, the roots were removed from the planchets and gradually infiltrated with increasing concentrations of Spurr resin (Ted Pella) over a 96-hour period. The sample polymerization was performed under vacuum at 70 °C for over 48 hours. For immunogold labeling analysis, high-pressure frozen root tips were transferred to the AFS (Leica Microsystems) and freeze-substituted in dry acetone containing 0.1% uranyl acetate at −85 °C. Stepwise infiltration with HM20, embedding, and UV polymerization were performed at −45 °C. For immunolabelling, the working concentration of rabbit anti-VSR and anti-GFP antibodies were 40 μg/ml. Gold particle-coupled secondary antibodies were diluted at a ratio of 1:50, followed by post-staining with aqueous uranyl acetate and lead citrate. Sections were examined using a Hitachi H-7650 transmission electron microscope equipped with a charge-coupled device camera, operating at 80 kV. (Hitachi High-Technologies; www.hitachi-hightech.com/jp/).

Electron tomography observations. The general procedures for performing electron tomography have been described previously66. In brief, 200-nm-thick sections were imaged using a Tecnai F20 electron microscope (FEI Company) operating at 200 kV. For each grid, a tilt image stack comprising 81 images was collected from +60° to −60° in 1.5° increments, followed by the acquisition of a second tilt image stack after rotating the grid by 90°. Dual-axis tomograms were reconstructed from pairs of image stacks using the Etomo program within the IMOD software package (version 4.9.7). The 15-nm gold particles on the surface of the sections were used as fiducial markers to align individual images in the tilt series. After electron tomography analysis, the IMOD software allows the display of an optical slice cut through the 3D image data in any desired orientation. For model generation, contours were manually drawn and meshed using the 3dmod program within the IMOD software package. To account for the thinning of plastic sections during electron imaging, the z-scale factor was calculated to be 1.6 for the root cell tomograms.

Correlative light and electron microscopy. Chose seedlings displaying strong fluorescence, then subject them to high-pressure freezing followed by freeze substitution at −80 °C for 24 hours in anhydrous acetone containing 0.25% glutaraldehyde and 0.1% uranyl acetate. The samples were then washed with precooled anhydrous acetone, embedded in graded HM20 resin (33%, 66%, and 100%), and polymerized under UV light for 18 hours at −45 °C. The 200 nm semi-thin sections were cut, collected on slot grids, and air-dried. Fluorescent images were initially captured using a confocal microscope, and the sample-loaded grids were subsequently examined using 3D ET.

Confocal microscopy and image processing. Confocal images were acquired using a Leica TCS SP8 confocal microscope. A 405 nm laser line was used to visualize PSV autofluorescence, and emission was collected between 440 and 490 nm. A 488 nm laser line was used to excite green fluorescence, and emissions for GFP were collected in the 500-550 nm range. A 514 nm laser line was used to excite red fluorescence, and emissions for mRFP or mCherry (Ch) were collected in the 600-650 nm range.

Cytological observation. Ovules were dissected from siliques and cleared with Hoyer's solution, following the previously described protocol67. Images of cleared embryos were captured using a Zeiss LSM980 laser scanning microscope equipped with differential interference contrast (DIC) optics.

Y2H assay. The full-length coding regions of the genes SNX1, SNX2a, SNX2b, SNX3, SNX4, and SNX5 were cloned into the pGADT7 vector to create fusions with the activating domain. The full-length coding sequence of the AtSNX1 and MmSNX1 genes were inserted into the pGBKT7 vector to create fusions with the GALL4 DNA-binding domain. The primers used are listed in Supplementary Table 2. The indicated plasmid combinations were used for the protein-protein interaction test according to the manufacturer’s instructions (Clontech, http://www.clontech.com/).

Phenotype characterization. In vitro pollen germination was conducted in a Petri dish containing solid PGM [1% low melting point agarose (w/v), 18% sucrose (w/v), 0.01% boric acid (w/v), 1 mM CaCl2, 1 mM Ca(NO3)2, and 1 mM MgSO4, adjusted to pH 7.5 using 0.2 M KOH] at 28°C, following a protocol modified from Boavida and McCormick (2007)68. Alexander staining (Solarbio, G3050) was performed to assess cytoplasmic viability, as described by Alexander (1969)69.

Chemical treatments. Oryzalin and PAO were prepared in DMSO and stocked at -20 ℃. Seedlings were cultured in 1/2 MS liquid medium containing 50 μM oryzalin for 6 hours or 50 μM PAO for 1.5-hour before being observed by confocal microscopy. For PAO treatment of Arabidopsis protoplasts, 50 μM PAO was injected into the Arabidopsis protoplasts, which expressed the fluorescent signal, 1 hour before being observed by confocal microscopy.

HeLa cells assay. AtSNX1 and MmSNX1 CDS were cloned into the plasmid pLV-IRES-EGFP to generate GFP-tagged forms and then infected HeLa cells with lentiviruses packaged using the pLV-IRES-AtSNX1-EGFP and pLV-IRES-MmSNX1-EGFP plasmids, respectively, to obtain HeLa-AtSNX1-GFP and HeLa-MmSNX1-GFP cell lines. Cells were fixed with 4% paraformaldehyde in PBS for 15 min. After washing with PBS for three times, coverslips were mounted in ProLongTM Gold Antifade Mountant with DAPI (Invitrogen). Cells were examined using a Leica TCS SP8 confocal microscope with 63× objective and 488 nm laser line. A total of 10 cells were examined in each condition.

Antibodies. The primary antibodies, anti-Myc (XMU-CRCE005), anti-GFP (Rockland, 600-401-215L), anti-His (Abbkin, ABT2050), anti-VPS29 (Agrisera, AS204401), anti-VSR (Agrisera, AS204407) were purchased from commercial companies. For AtSNX1 primary antibody, AtSNXl cDNA was cloned into the pET28a plasmid which incorporates an N-terminal histidine (His) tag into the expressed protein. Recombinant proteins were generated in Escherichia coli (Rosseta 2; Novagen) and purified using nickel agarose beads. Polyclonal antibodies were raised in rabbit.

Expression of recombinant protein, protein extraction. For the expression of MmSNX1, the full-length mouse SNX1 (NP_062701.2) was subcloned into the pGEX-6P-1 plasmid and expressed as glutathione S-transferase (GST)-tagged fusion proteins in Escherichia coli BL21 (DE3) cells. Cells were grown in Terrific Broth (TB) medium at 37 °C until reaching an optical density of 1.2 at 600 nm, then induced at 18 °C for 16 hours with 0.24 mM isopropyl-β-D-thiogalactoside (IPTG). Cells were harvested by centrifugation, resuspended in phosphate-buffered saline (PBS) buffer (140 mM NaCl, 2.7 mM KCl, 10 mM Na₂HPO₄, 1.8 mM KH₂PO₄, pH 7.4, and 10% glycerol), and lysed by sonication. After centrifugation at 18,000 rpm for 30 minutes, the supernatant was collected and incubated with glutathione-Sepharose 4B at 4 °C, followed by washing with PBS buffer. Following cleavage with Precision Protease (Beyotime) to remove the GST tag, the eluted proteins underwent further purification by size-exclusion chromatography on a Superdex 200 10/300 GL column (GE Healthcare) using an elution buffer of 50 mM HEPES (pH 7.4) and 150 mM NaCl. The target protein was collected and stored at -80 °C.

For the expression of full-length AtSNX1 and AtSNX2a, the full-length coding sequences of Arabidopsis SNX1 (AT5G06140) and Arabidopsis SNX2a (AT5G58440) were subcloned into the pET28a-His6-SUMO vector. Plasmids encoding AtSNX1 or AtSNX2a were transformed into LOBSTR cells, and protein expression was carried out as described for MmSNX1 above. For the purification of AtSNX1 and AtSNX2a, cell pellets were lysed by sonication in His-lysis buffer containing 50 mM Na₂HPO₄, 50 mM NaH₂PO₄ (pH 8.0), 400 mM NaCl, 20 mM imidazole, and 0.1% Tween 20. After clearing the bacterial lysates by centrifugation at 18,000 rpm for 30 minutes, the soluble fraction was incubated with Ni-NTA beads for 2 hours. The beads were then washed with His-wash buffer A (50 mM Na₂HPO₄, 50 mM NaH₂PO₄ (pH 8.0), 400 mM NaCl, 60 mM imidazole, and 0.1% Tween 20). After cleavage with ULP1 to remove the His-SUMO tag, the eluted proteins were further purified by size-exclusion chromatography using a Superose 6 Increase 10/300 GL column with an elution buffer containing 20 mM HEPES (pH 8.0) and 400 mM NaCl. The target protein was concentrated and stored at −80 °C. For AtSNX1-AtSNX2a complex expression, full-length AtSNX1 was subcloned into the pMAL-C6t-MBP vector, and full-length AtSNX2a was subcloned into the pET28a-His6-SUMO vector. Overexpressed AtSNX1 and AtSNX2a cell pellets were mixed and lysed together in MBP buffer containing 20 mM Tris-HCl (pH 7.4) and 200 mM NaCl. Then the AtSNX1-AtSNX2a complex was purified using Amylose resin. After cleavage with His-TEV to remove the His-MBP tag, the eluted fraction (SNX1-His-SUMO-SNX2a) was further loaded onto a Ni-NTA column and washed with buffer containing 20 mM HEPES (pH 7.4), 200 mM NaCl, and 40 mM imidazole to remove unbound samples. Following the removal of the His-SUMO tag by ULP1, the eluted protein was further purified by size-exclusion chromatography using a Superose 6 Increase 10/300 GL column with an elution buffer containing 20 mM HEPES (pH 7.4) and 200 mM NaCl. The purified target protein was concentrated and stored at −80 °C.

For the purification of His-tagged AtSNX1 and His-tagged AtSNX2a, the procedure was the same as that for full-length AtSNX1, except for slight variations in buffer compositions. For lysis and Ni-NTA affinity purification, His-lysis buffer (50 mM PBS buffer, pH 7.4, 20 mM imidazole, 0.1% Tween 20) was used. Unbound samples were removed with 60 mM imidazole, and the target protein was eluted with buffer containing 250 mM imidazole. The elution protein was collected and stored at -80℃ for lipid binding assay.

Protein-lipid overlay assay. The nitrocellulose membranes contained different lipids (PIP StripsTM membranes; P23751) used in protein-lipid overlay assay were purchased from ThermoFisher. The nitrocellulose membranes were incubated for 2 hours at room temperature in TBST buffer (Tris-buffered saline with Tween 20) containing 3% (w/v) fatty-acid-free BSA. After three washes with TBST buffer, the membranes were incubated at 4 °C for 4 hours in TBST containing the purified His-tag protein at a final concentration of 3-5 μg/mL. After washing the membranes four times with TBST buffer, they were incubated at room temperature for 2 hours with anti-His tag antibody (1:2000, Abbkine) in TBST buffer. This was followed by three washes with TBST and a 1-hour incubation at room temperature with the secondary antibody (1:4000, Beyotime). Finally, the membranes were washed three times with TBST.

Liposome preparation. All lipids were purchased from Avanti Polar Lipids. Stock solutions of lipids were prepared at 10 mg/mL in chloroform for 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC), 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine (DOPE), 1,2-dioleoyl-sn- glycero-3-phosphor-1-serine (DOPS), or dissolved in chloroform at 0.2 mg/mL for 1,2-dioleoyl-sn-glycero-3-phospho-(1’-myo-inositol-3’-phosphate) (PtdIns(3)P) and 1,2-dioleoyl-sn-glycero-3-phospho-(1'-myo-inositol-4'-phosphate) (PtdIns(4)P). Lipid molar ratios used for liposome preparation are 45% DOPC, 30% DOPE, 20% DOPS, and 5% PtdIns(3)P or PtdIns(4)P). Lipid mixtures were dried under nitrogen gas and kept under vacuum overnight. The dry lipid mixtures were resuspended in 50 mM HEPES (pH 7.4) and 150 mM NaCl for at least 30 minutes at 37 °C, then vortexed gently to obtain a homogeneous hydrated solution. The solution was extruded through 0.4 μm membrane filters to produce liposomes with a diameter of 400 nm.

Liposome-protein co-sedimentation assay. For liposome-protein co-sedimentation assay, the 400 nm liposomes (0.4 mg/mL) and protein (0.2 mg/mL) were incubated for 30 min at 20℃ before ultracentrifugation at 100,000×g for 30 min and then the supernatants and pellets were subjected to sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) analysis.

Liposome tubulation assay and Negative-Stain EM. Protein (0.4 mg/mL) was incubated with liposomes (0.2 mg/mL) for 4 hours at 4 °C. The mixture was then applied to a glow-discharged, carbon-coated EM grid and stained with uranyl acetate. The EM grids were examined using a Hitachi H-7650 electron microscope operating at 80 kV.

Cryo-EM sample preparation and data acquisition. Four microliters of the sample (protein concentration: 3 mg/mL, liposome concentration: 1.5 mg/mL) were loaded onto a Quantifoil Au grid (R2/2, 300 mesh), followed by plunge-freezing in liquid ethane using a Vitrobot Mark IV (Thermo Fisher) at 4 °C and 100% humidity. Movies were recorded using a Titan Krios G4 300 kV cryo-TEM system equipped with a Falcon 4i direct electron detector and a Selectris X energy filter (Thermo Fisher). Cryo-EM data was acquired automatically with EPU v3.4.0.5704REL in EER mode, with a total dose of ~50 e-/Å2 and targeted defocus ranging from -1.0 to -2.5 μm, slit width of 10 eV and a pixel size of 0.92 Å (×130,000 magnification). A total of 6,062 movies across 2 imaging sessions were collected.

Cryo-EM data processing. All processing was performed in CryoSPARC v.4.4.170. Micrographs were aligned with patch motion correction, and the contrast transfer function (CTF) was estimated using patch CTF estimation. A total of 1,914 particles were manually picked using the manual picker tool from 58 randomly selected movies. Particle cleaning was performed using 2D classification, resulting in the selection of 1,896 particles for Topaz training71. After Topaz picking, 487,808 particles were obtained and further aligned through iterative 2D classifications. During 2D classification, multiple 2D classes corresponding to tubes with different diameters were obtained from Topaz-picked particles (Supplementary Fig. 9a). Following asymmetric helical refinement and extensive symmetry searches in CryoSPARC, only two particle subsets reproducibly converged into stable consensus maps (highlighted by black and green boxes in Supplementary Fig. 9a). Other classes either failed to converge or yielded low-quality maps (highlighted by red and salmon boxes) and were therefore excluded from further analysis. Two populations with distinct diameters (Class Ⅰ: 49,380 particles; Class Ⅱ: 20,728 particles) were manually grouped, and initial 3D volumes were generated using asymmetric helical reconstruction. These volumes were subjected to symmetric search utility in CryoSPARC to determine twist and rise parameters. The symmetry parameters were applied in subsequent rounds of helical refinement. The final resolutions for the two maps were 7.2 Å for Class Ⅰ and 8.7 Å for Class Ⅱ, respectively. We attempted helical reconstruction in RELION 5 using BLUSH regularization and also tested symmetry expansion followed by subunit classification. However, these approaches did not improve the resolution relative to our final maps and were therefore not pursued further.

Model building. To assist with model building, Alphafold 372 was used to predict the dimer structure of plant SNX1 dimer. The dimer was rigid fitted into the Class I and II density maps in ChimeraX73, followed by automatic molecular dynamics flexible fitting (MDFF) in Namdinator74. The final helical assemblies were obtained by docking multiple identical copies of the dimer model followed by manual adjustment in COOT75 and real space refinement in PHENIX76. Residues 1-23 in the PX domain of the final model were removed due to the absence of map density. After flexible model refinement, the subcomplex was docked into the two maps to reconstruct the intact scaffold structures of the two populations in ChimeraX. Figures were generated using ChimeraX.

All-atom molecular dynamics simulations. The GROMACS software was used for all-atom molecular dynamics simulations77. The CHAIRMM-GUI website (https://charmm-gui.org/) was utilized to select the CHARMM36 force field for generating system input files78, 79. The cryo-EM structure of MmSNX1 (PDB: 7D6D) was retrieved from the Protein Data Bank, while the AtSNX1 cryo-EM structure was determined in this study37. The structure of AtSNX2a was predicted using the AlphaFold Server (powered by AlphaFold3; https://alphafoldserver.com)72. AtSNX2a-related information was retrieved from Uniport (Uniport ID: Q8L5Z7). Given that the N-terminus of AtSNX2a harbors a long intrinsically disordered region, only the amino acid sequence corresponding to residues 157–587 was used for structure prediction (parameters: protein entity, single copy, 10 replicate predictions). Each prediction run generated five models, and the top-ranked model (rank_0, highest confidence score) was selected as the representative structure. Predicted template modeling (pTM) scores were then compared, and the model with the highest pTM score (0.75) was selected for subsequent molecular simulations (Supplementary Fig. 23a). Additional predicted structures of AtSNX2a are shown in Supplementary Fig. 23b-e. While the overall folds of these models are highly conserved, the disordered region linking the PX and BAR domains is poorly resolved by AlphaFold3, leading to notable structural variability in this region (Supplementary Fig. 23f). The bilayer membrane system was composed of 45% DOPC, 30% DOPE, 20% DOPS, and 5% PI(3)P. The system was solvated using the TIP3P water model, and Na⁺ and Cl⁻ ions were added to maintain charge neutrality. Using VMD, the protein was vertically positioned approximately 4.5 Å above the bilayer with its AH domain oriented toward the bilayer; overlapping solvent molecules were then removed to obtain the initial structure. The Particle-mesh Ewald (PME) method was utilized to handle electrostatic interactions. Electrostatic interactions were handled via the Particle-mesh Ewald (PME) method. Following the CHARMM-GUI workflow, energy minimization was first performed with the steepest descent (steep) algorithm for a maximum of 5,000 steps. Subsequently, a 5-step system pre-equilibration was conducted for a total duration of 1,875 ps, during which restraints were applied to the protein and lipid bilayer prior to the production run. The cutoff distance for Coulomb interactions and van der Waals radii was set to 1 nm, and the truncation value for non-bonded interactions was 10 Å. The Nose-Hoover thermostat was used to maintain the simulation temperature at 300 K, and the Parrinello-Rahman barostat was employed to control the pressure at 1 bar under semiisotropic conditions. A 200 ns molecular dynamics simulation was carried out using GROMACS 2023 software. Statistical charts related to the molecular simulation data were generated using OriginPro. Protein and membrane visualizations were created using VMD and PyMOL.

Statistics and reproducibility. Data represent mean ± SEM. All experiments were performed at least three independent replicates. Analyses used GraphPad Prism 10: two-tailed unpaired t-tests and paired t-tests (two groups). Significance was defined as *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001, while “ns” indicates no significance (P > 0.05). Biological replicates are specified in figure legends. All schematic diagrams and cartoons presented in this study were originally created by our team using Adobe Illustrator (version 2021).

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "In this manuscript, the authors identified a plant-specific membrane trafficking pathway between MVB and recycling endosomes. Using advanced light microscopy and electron microscopy techniques, in combination with structural biology and biochemical assays, they found that plant SNX1 localises to small vesicles around the MVB, and its function differs from that of mouse SNX1, which generates longer tubules in vitro. Ultimately, the authors demonstrated that the function of SNX is involved in embryo biogenesis in Arabidopsis. The study is interesting; however, one of the primary concerns is that the biological relevance of SNX1-regulated spherical recycling endosomes remains highly speculative. Although the authors have put considerable effort into studying the structure and biochemical activities of SNX1, its contribution to the manuscript is unclear to me.\nMajor issues:\n1. Please provide quantification data on the immuno-gold labeling (Line 161-164).\n2. Why is plant SNX different from animal SNX? Can MmSNX1 complement the Arabidopsis mutant phenotype? What functional insights can we gain from the results related to “Arabidopsis SNX1 induces the formation of shorter tubules compared to those generated by mouse SNX1” and “AtSNX1 has a lower binding affinity to membranes compared to MmSNX1”? Please provide further explanation or, ideally, some in vivo experimental data demonstrating that this specific feature of plant SNX1 is critical for its function.\n3. The genetic study does not appear very convincing to me. The authors should concentrate on analysing the snx1/snx2a/snx2b triple mutant and conducting genetic complementation assays using different versions of SNX. Knocking out SNX in the vps29 mutant background does not seem very relevant.\n4. The phenotype studies in Figure 6d-i are unclear; some quantification is needed for the developmental studies. What are the embryo phenotypes in the snx1/snx2a/snx2b triple mutant?\n5. Why are different mutants used in Figure 6f-I? The authors should explain why a specific mutant was chosen for that experiment. I would use one or two representative mutants for all studies, not radomly chosen. In general, some of the results in Figure 6 are confusing.", "reviewer_2": "The paper by Li et al. describes analysis of retrieval processes from multivesicular endosomes in Arabidopsis cells. While their findings are interesting, I have to say that the authors have made a very serious and hazardous mistake in the definition of “recycling endosomes (RE).” The authors appear to use this term to indicate retrograde carriers from the MVE, involving retromer and/or sorting-nexin complexes. This is wrong. RE was originally defined in mammalian cells as a compartment fulfilling slow processes of endocytic recycling from and back to the plasma membrane and has been widely used as such for a variety of eukaryotes in the field. The best-accepted RE markers are Rab11, Arf6 and transferrin in mammals and Ypt31/32 and Rcy1 in yeast. One can of course propose a change of term definition, but in that case, extreme care should be taken to rationalize it, especially when the definition has been long accepted and widely used. Using the term RE for retromer/SNX-dependent carriers is not acceptable to me, because it will cause very serious confusion in the membrane trafficking field. The authors’ finding that the compartment they are looking at does not overlap with RABA1c (one of Rab11 in plants) (suppl. Fig. 4) supports my view that they are distinct, although it should also be noted that the Rab11 group has been extensively diversified in plants.\nThe CLEM and immune-EM analysis is well done and the presence of 30-50 nm vesicles around the MVE looks convincing. The formation of these vesicles may well be formed from MVE. The important point that the authors like to claim is probably their structures, not long tubules but spherical vesicles. I have to point out here that whether they are tubules or vesicles is not easy to determine by conventional thin-section electron microscopy. The results of in vitro tubulation assays are interesting in that sense. The MD simulation analysis also appears to support the structural difference of these retrieval compartments. However, I am still concerned about a possibility that tubulation can be stabilized by interaction with cytoskeletons and motor proteins, which could cause differences between species.\nI would like to add one more critique. While colocalization analysis gives us very important clues to understand possible interactions between components, but we must remember that the results only tell us about “steady-state” locations. They are important but we should not ignore “transient” interactions when we are looking at traffic events transporting cargo from somewhere to elsewhere.", "reviewer_3": "Li et al. focused on the molecular structure and functions of SNX proteins in plants and reported that plant recycling endosomes (REs) are unique spherical vesicles approximately 30–50 nm in diameter, the formation of which is regulated by the SNX1-SNX2 heterodimer. Data obtained using correlative light and electron microscopy, 3D electron tomography, immunogold transmission electron microscopy, and MD simulation were of high quality and interesting. The authors also performed a liposome tubulation assay in vitro and found that Arabidopsis SNX1 generates significantly shorter tubules compared to mouse SNX1. They also showed that Arabidopsis SNX1 and SNX2 mediate the recycling of vacuolar sorting receptors (VSRs), a process that is essential for proper seed development and germination in plants. They further showed that disruptions in Arabidopsis SNX1 and the core retromer component VPS29 impair REs formation, leading to the development of enlarged MVBs. Based on these results, the authors claim to provide the first evidence for the localization and structure, as well as the formation mechanism, of plant REs mediated by SNX1 and SNX2. However, I feel that the definition of “REs” is misleading in the field of plant cell biology and a major revision is required before publication.\nThe detailed comments are below.\n1.My biggest concern lies with the authors' usage of the term \"recycling endosome/RE.\" RE usually refers to compartmental functions in the recycling pathway of endocytosed cargo to the plasma membrane, which are often identified by the localization of Rab11 in mammalian cells. In this manuscript, the authors call “retromer vesicles” formed at MVBs as REs, but to my knowledge, those retromer-dependent compartments are not referred to as REs. The authors cited multiple studies on retromer-dependent structures in animals and yeast; however, none of them referred to these structures as REs. Using a widely accepted term for a newly identified structure based on the authors’ definition is misleading and likely to cause confusion in the field. Although it may not be intentional, it appears that by using the established term \"RE,\" the authors are trying to increase the perceived impact of their work by claiming the discovery of a RE in plants, whose existence has been unclear. I strongly recommend that the authors reconsider this terminology.\n2.For the immunoelectron microscopy (Fig. 1g–i), more microscopic images and quantitative data are required. Fig. 1h shows only one gold particle on the bud, which is not sufficiently convincing.\n3.Fig. 2a shows that AtSNX1 binds much more strongly to PI(4)P than to PI(3)P, which is inconsistent with its subcellular localization. I strongly suggest that the authors examined the effect of PI(4)P on AtSNX1 expression.\n4.Page 10, line 276 “Since SNX2a and SNX2b function redundantly, …”: I cannot agree with this sentence. The functional redundancy of these SNX2s was not clearly demonstrated in this paper (#43).\n5.Page 10, line 288. The authors suggested that the combined action of SNX1 and SNX2 may play a key role in the formation of spherical structures. Although this suggestion was supported by an in vitro tubulation assay, the explanation of how SNX1 and SNX2 function was insufficient. A more detailed discussion of the molecular functions is required.\n6.Fig. 6a–c: The fluorescence signal in the RNAi-SNX lines shown in Figure 6a appears stronger than that in the control, and the Western blotting results in Figure 6b also indicate that the total amounts of GFP-VSR2 and GFP are higher in the RNAi lines. Therefore, it is not convincing to conclude from these data that SNX knockdown impairs VSR2 recycling. Since the authors already have the mutant lines, the mislocalization and degradation of VSR2 should be checked in the mutant plants, not only in protoplasts.\n7. The authors should specify which RAB5 and RAB7 were used for the localization and dominant-negative mutant expression analyses. In particular, conventional and plant-specific RAB5 are known to localize and function differently.\n8.The involvement of RABs was not mentioned in the discussion, and the authors’ interpretation of the related data was not clear.\n9.Although the authors acknowledge the technical difficulty of resolving the SNX1–SNX2 complex structure by cryo-EM in the Discussion, this limitation is not sufficiently reflected in the interpretation of their comparative analysis between Arabidopsis and mouse SNX1. Given that SNX1 and SNX2 have non-overlapping functions, as shown in this study and in previous work (Pourcher et al., 2010, Plant Cell), and that SNX2 requires SNX1 for proper endosomal localization, it would be important to clearly state how the inability to resolve the SNX1–SNX2 complex structure affects the interpretation and significance of the structural comparisons. A more explicit discussion of this point in both the Introduction and Discussion could help clarify the rationale and impact of the present findings.\n10.Most of this discussion is a repetition of what has already been described in the results. In contrast, the authors describe their hypothesis in the last section of the Discussion (page 15), but in a very strong way. Please provide a careful discussion by clarifying what is known and the authors’ proposition."}, "author_response": "Thank you for your constructive comments and your time in reviewing the MS.\nMajor issues:\n1. Please provide quantification data on the immuno-gold labeling (Line 161-164).\nResponse: We have now included additional representative immunoelectron microscopy images and provided quantification data on the immuno-gold labeling to further support the findings shown in Fig. 1h, 1j and Supplementary Fig. 7 with proper description. The results indicate that VSRs are predominantly localized to the retrieval vesicles (RVs) and the limiting membrane of the MVB (Fig. 1g, 1j and Supplementary Fig. 7a), while VPS29-GFP and SNX1-GFP primarily label the limiting membrane of the MVB, with a specific enrichment at the budding vesicle regions (Fig. 1h-i, 1k-l and Supplementary Fig. 7b-c).\n2. Why is plant SNX different from animal SNX? Can MmSNX1 complement the Arabidopsis mutant phenotype? What functional insights can we gain from the results related to “Arabidopsis SNX1 induces the formation of shorter tubules compared to those generated by mouse SNX1” and “AtSNX1 has a lower binding affinity to membranes compared to MmSNX1”? Please provide further explanation or, ideally, some in vivo experimental data demonstrating that this specific feature of plant SNX1 is critical for its function.\nResponse: AtSNX1 and MmSNX1 share only 28.6% amino acid sequence similarity. MmSNX1 does not interact with AtSNX1 or AtSNX2 in Y2H assays, unlike AtSNX1, which interacts with both AtSNX1 and AtSNX2 (Supplementary Fig. 20a). In vitro liposome tubulation assays further demonstrated that replacing the amphipathic helix (AH) of MmSNX1 with that of AtSNX1 led to the formation of shorter tubules, resembling those induced by AtSNX1 (Fig. 4i). These findings suggest that the primary difference between AtSNX1 and MmSNX1 lies in their AH domains, which lead to distinct membrane binding and remodeling behaviors due to structural and electrostatic differences (Fig. 3 and 4a-h).\nAtSNX1 exhibits lower membrane-binding affinity and consequently generates shorter tubules than MmSNX1 in vitro (Fig. 2b and Fig. 3). In vivo, AtSNX1 expression results in a punctate localization pattern, whereas MmSNX1 forms elongated tubular structures in Arabidopsis protoplasts, roots, and HeLa cells (Supplementary Fig. 20b-g). We thus propose that AtSNX1 drives the formation of spherical structures due to its limited ability to bind to and stabilize tubules, thereby preventing the formation of elongated tubular structures.\nInterestingly, expression of MmSNX1 fails to rescue the vacuolar degradation defect of VSRs caused by RNAi-mediated knockdown of AtSNX1 (Supplementary Fig. 20j). This finding indicates that, although MmSNX1 can assemble long, stable tubular retrieval vesicles in Arabidopsis protoplasts, such extended carriers are probably poorly suited to the spatial and physiological demands of plant cells. We have also generated transgenic plants to assess whether MmSNX1 can complement the Arabidopsis snx1 mutant phenotype. However, due to the limited three-month revision timeline, these results will be presented in a future study\nWe have now incorporated the above results into the new Supplementary Fig. 20 and integrated the corresponding discussions in the revised MS (page 15, lines 434-454, highlighted in red).\n3. The genetic study does not appear very convincing to me. The authors should concentrate on analysing the snx1/snx2a/snx2b triple mutant and conducting genetic complementation assays using different versions of SNX. Knocking out SNX in the vps29 mutant background does not seem very relevant.\nResponse: As recommended, we have now focused our genetic analysis on the snx1/snx2a/snx2b triple mutant. However, the snx1/snx2a/snx2b triple mutant exhibited only mild defects in plant growth and negligible phenotypic changes during embryonic development (Fig. 6h-i and Supplementary Fig. 17f and 18d). These findings led us to hypothesize that the VPS subcomplex of the retromer may partially compensate for the loss of SNX function. Therefore, we believe that analyzing SNX mutants in the vps subcomplex mutant background (e.g., vps29 mutant) remains relevant, as it may help uncover potential functional redundancy or compensatory mechanisms among retromer complex components. To maintain clarity and focus, we limited our analysis to the snx1-/-vps29+/- mutant (Figure 7). We also performed genetic complementation assays with various SNX constructs. However, due to the limited three-month revision timeline, these results will be included in a future study.\nWe have now incorporated the above results into the new Supplementary Fig. 17-18 and integrated the corresponding discussions in the revised MS (pages 12, lines 351-358, highlighted in red).\n4. The phenotype studies in Figure 6d-i are unclear; some quantification is needed for the developmental studies. What are the embryo phenotypes in the snx1/snx2a/snx2b triple mutant?\nResponse: We have now incorporated the quantification data for the developmental phenotypes in the revised Figure 6d-i. The snx1/snx2a/snx2b triple mutant exhibited a negligible phenotype during embryonic development (Supplementary Fig. 18d).\n5. Why are different mutants used in Figure 6f-I? The authors should explain why a specific mutant was chosen for that experiment. I would use one or two representative mutants for all studies, not radomly chosen. In general, some of the results in Figure 6 are confusing.\nResponse: Since the snx1/snx2a/snx2b triple mutant exhibits only mild growth defects and negligible phenotypic changes during embryonic development (Fig. 6h-i and Supplementary Fig. 17f and 18d), we hypothesized that the VPS subcomplex of the retromer may partially compensate for the loss of SNX function. In response to the reviewer’s suggestion, we have now streamlined our genetic analysis to improve clarity and consistency. Specifically, we now focus on the snx1/snx2a/snx2b triple mutant in Figure 6 and the snx1-/-vps29+/- mutant in Figure 7 to more effectively assess functional interactions within the retromer complex.\nReviewer #2 (Remarks to the Author):\nThe paper by Li et al. describes analysis of retrieval processes from multivesicular endosomes in Arabidopsis cells. While their findings are interesting, I have to say that the authors have made a very serious and hazardous mistake in the definition of “recycling endosomes (RE).” The authors appear to use this term to indicate retrograde carriers from the MVE, involving retromer and/or sorting-nexin complexes. This is wrong. RE was originally defined in mammalian cells as a compartment fulfilling slow processes of endocytic recycling from and back to the plasma membrane and has been widely used as such for a variety of eukaryotes in the field. The best-accepted RE markers are Rab11, Arf6 and transferrin in mammals and Ypt31/32 and Rcy1 in yeast. One can of course propose a change of term definition, but in that case, extreme care should be taken to rationalize it, especially when the definition has been long accepted and widely used. Using the term RE for retromer/SNX-dependent carriers is not acceptable to me, because it will cause very serious confusion in the membrane trafficking field. The authors’ finding that the compartment they are looking at does not overlap with RABA1c (one of Rab11 in plants) (suppl. Fig. 4) supports my view that they are distinct, although it should also be noted that the Rab11 group has been extensively diversified in plants.\nResponse: Thank you for your constructive comments and your time in reviewing the MS. To avoid confusion with the well-established definition of recycling endosomes (REs), primarily associated with recycling processes to the plasma membrane (PM) in animal cells, we have adopted the term \"retrieval vesicle (RV)\" to accurately describe the novel structure identified in our study. Specifically, we observed a distinct spherical membrane compartment approximately 30-50 nm in diameter adjacent to MVBs in Arabidopsis root cells, with nascent structures actively budding from the MVB surface. These compartments mediate the retrieval of specific cargo proteins, such as VSRs. We have now clearly defined and consistently used the term RV throughout the revised manuscript to avoid potential confusion in the membrane trafficking field.\nThe CLEM and immune-EM analysis is well done and the presence of 30-50 nm vesicles around the MVE looks convincing. The formation of these vesicles may well be formed from MVE. The important point that the authors like to claim is probably their structures, not long tubules but spherical vesicles. I have to point out here that whether they are tubules or vesicles is not easy to determine by conventional thin-section electron microscopy. The results of in vitro tubulation assays are interesting in that sense. The MD simulation analysis also appears to support the structural difference of these retrieval compartments. However, I am still concerned about a possibility that tubulation can be stabilized by interaction with cytoskeletons and motor proteins, which could cause differences between species.\nResponse: We agree that distinguishing vesicles from tubules is challenging using conventional thin-section transmission electron microscopy (TEM). In our study, we employed electron tomography on 200-nm-thick sections, enabling the reconstruction of their 3D architecture. This analysis confirmed the spherical nature of the vesicles (approximately 30-50 nm in diameter) budding from MVBs. We have now added a series of sequential images to illustrate the structure of spherical RVs from top to bottom (Supplementary Fig. 6).\nAdditionally, we appreciate your insightful comment regarding the potential role of cytoskeletal elements and motor proteins in stabilizing membrane tubulation—a mechanism well documented in animal cells and one that may underlie the structural differences we observe1,2. Consistent with this, disrupting microtubules with 50 μM oryzalin for 6 h significantly reduced the length of MmSNX1-labeled tubules in Arabidopsis roots (Supplementary Fig. 20g–i), supporting microtubule-dependent stabilization of tubulation. Moreover, two recent studies in Arabidopsis further underscore a potential functional connection between endosomal and autophagosomal dynamics and the cytoskeleton. One study demonstrated that CLASP interacts with SNX1 to link microtubules to auxin transport through PIN2 recycling3. Another study revealed that SNX proteins interact with ATG8 and CLASP to facilitate autophagosome movement along microtubules4. However, whether microtubules contribute to the formation of RVs in Arabidopsis remains unclear and represents a promising direction for future research.\nWe have now incorporated the above results into the new Supplementary Fig. 6, Supplementary Fig. 20g-i and integrated the corresponding discussions in the revised MS (page 6, lines 160-162; pages 15-16, lines 455-467, highlighted in red).\nReferences:\n1. Delevoye, C. et al. BLOC-1 Brings Together the Actin and Microtubule Cytoskeletons to Generate Recycling Endosomes. Current biology : CB 26, 1-13 (2016).\n2. Delevoye, C. et al. Recycling endosome tubule morphogenesis from sorting endosomes requires the kinesin motor KIF13A. Cell reports 6, 445-454 (2014).\n3. Ambrose, C. et al. CLASP interacts with sorting nexin 1 to link microtubules and auxin transport via PIN2 recycling in Arabidopsis thaliana. Developmental cell 24, 649-659 (2013).\n4. Liao, Y. et al. The plant retromer components SNXs bind to ATG8 and CLASP to mediate autophagosome movement along microtubules. Mol. Plant 18, 416-436 (2025).\nI would like to add one more critique. While colocalization analysis gives us very important clues to understand possible interactions between components, but we must remember that the results only tell us about “steady-state” locations. They are important but we should not ignore “transient” interactions when we are looking at traffic events transporting cargo from somewhere to elsewhere.\nResponse: Thank you for raising this important point. We fully agree that colocalization analyses primarily reflect steady-state localization patterns and might not capture transient or dynamic interactions involved in cargo transport. We have attempted to visualize the dynamic behavior of the newly identified retrieval vesicles; however, due to their small size (~30–50 nm in diameter) and rapid movement, it is extremely challenging to capture their dynamics and transient interactions with other organelles using conventional confocal microscopy. In future work, we aim to employ advanced live-cell super-resolution imaging techniques, such as lattice light-sheet microscopy or instant structured illumination microscopy (iSIM), to resolve these transient events more effectively and thereby gain deeper insights into the dynamics of vesicle-mediated trafficking.\nWe have now incorporated the above discussions in the revised MS (page 14, lines 405-413, highlighted in red).\nReviewer #3 (Remarks to the Author):\nLi et al. focused on the molecular structure and functions of SNX proteins in plants and reported that plant recycling endosomes (REs) are unique spherical vesicles approximately 30–50 nm in diameter, the formation of which is regulated by the SNX1-SNX2 heterodimer. Data obtained using correlative light and electron microscopy, 3D electron tomography, immunogold transmission electron microscopy, and MD simulation were of high quality and interesting. The authors also performed a liposome tubulation assay in vitro and found that Arabidopsis SNX1 generates significantly shorter tubules compared to mouse SNX1. They also showed that Arabidopsis SNX1 and SNX2 mediate the recycling of vacuolar sorting receptors (VSRs), a process that is essential for proper seed development and germination in plants. They further showed that disruptions in Arabidopsis SNX1 and the core retromer component VPS29 impair REs formation, leading to the development of enlarged MVBs. Based on these results, the authors claim to provide the first evidence for the localization and structure, as well as the formation mechanism, of plant REs mediated by SNX1 and SNX2. However, I feel that the definition of “REs” is misleading in the field of plant cell biology and a major revision is required before publication.\nResponse: Thank you very much for your positive and constructive comments and your time in reviewing the MS.\nThe detailed comments are below.\n1.My biggest concern lies with the authors' usage of the term \"recycling endosome/RE.\" RE usually refers to compartmental functions in the recycling pathway of endocytosed cargo to the plasma membrane, which are often identified by the localization of Rab11 in mammalian cells. In this manuscript, the authors call “retromer vesicles” formed at MVBs as REs, but to my knowledge, those retromer-dependent compartments are not referred to as REs. The authors cited multiple studies on retromer-dependent structures in animals and yeast; however, none of them referred to these structures as REs. Using a widely accepted term for a newly identified structure based on the authors’ definition is misleading and likely to cause confusion in the field. Although it may not be intentional, it appears that by using the established term \"RE,\" the authors are trying to increase the perceived impact of their work by claiming the discovery of a RE in plants, whose existence has been unclear. I strongly recommend that the authors reconsider this terminology.\nResponse: We appreciate the reviewer’s thoughtful comment regarding the terminology used to describe the structure identified in our study. We fully acknowledge that in mammalian systems, the term \"recycling endosome (RE)\" traditionally refers to compartments involved in the recycling of endocytosed cargo back to the plasma membrane (PM), often marked by Rab11 localization. We also recognize that the retromer-associated structures we observed in plants, which originate from MVBs, do not correspond to REs as defined in animal systems.\nTo avoid confusion and prevent any misinterpretation regarding the functional or evolutionary equivalence of these structures, we have now revised the manuscript to replace the term \"recycling endosome (RE)\" with \"retrieval vesicle (RV)\" throughout. This term better reflects the role of these vesicles in the retrograde trafficking of VSRs and is consistent with their observed origin and function in plant cells. Importantly, our intention was not to overstate the impact of our findings but to highlight a potentially plant-specific mechanism of cargo retrieval from MVBs, distinct from classical RE pathways in animal cells.\nWe have also revised the Introduction and Discussion sections to clarify this distinction and explicitly state the differences between animal REs and the plant RVs described in this study.\n2.For the immunoelectron microscopy (Fig. 1g–i), more microscopic images and quantitative data are required. Fig. 1h shows only one gold particle on the bud, which is not sufficiently convincing.\nResponse: We have now included additional representative immunoelectron microscopy images and provided quantification data to further support the findings shown in Fig. 1h, 1j and Supplementary Fig. 7. The results indicate that VSRs are predominantly localized to the retrieval vesicles (RVs) and the limiting membrane of the MVB (Fig. 1g, 1j and Supplementary Fig. 7a), while VPS29-GFP and SNX1-GFP primarily label the limiting membrane of the MVB, with a specific enrichment at the budding vesicle regions (Fig. 1h-i, 1k-l and Supplementary Fig. 7b-c).\n3.Fig. 2a shows that AtSNX1 binds much more strongly to PI(4)P than to PI(3)P, which is inconsistent with its subcellular localization. I strongly suggest that the authors examined the effect of PI(4)P on AtSNX1 expression.\nResponse: We thank the reviewer for this insightful comment. We agree that the strong binding of AtSNX1 to PI(4)P observed in Fig. 2a appears inconsistent with its predominant localization to PI(3)P-enriched MVBs. To address the reviewer’s comments, we conducted additional experiments to examine the colocalization between AtSNX1 and PI(4)P reporters, as well as the effect of PI(4)P depletion on AtSNX1 localization and expression, by treating cells with phenylarsine oxide (PAO), a known PI(4)-kinase inhibitor. The results showed that AtSNX1 did not colocalize with PI(4)P markers (Supplementary Fig. 8a), and its localization remained largely unchanged upon PI(4)P depletion (Supplementary Fig. 8b-f), supporting the notion that its in vivo function is primarily associated with PI(3)P-enriched compartments. In addition, although AtSNX1 is capable of binding to liposomes in the presence of PI(4)P, it is unable to induce tubular structure formation under these conditions in vitro (Supplementary Fig. 8g-i).\nWe have now incorporated the above results into the new Supplementary Fig. 8 and included the corresponding descriptions in the revised MS (page 7, lines 191-202, highlighted in red).\n4.Page 10, line 276 “Since SNX2a and SNX2b function redundantly, …”: I cannot agree with this sentence. The functional redundancy of these SNX2s was not clearly demonstrated in this paper (#43).\nResponse: We appreciate the reviewer’s comment and agree that the functional redundancy of SNX2a and SNX2b was neither sufficiently demonstrated in this study nor supported by previous publications. Accordingly, we have removed the sentence to avoid overinterpretation.\n5.Page 10, line 288. The authors suggested that the combined action of SNX1 and SNX2 may play a key role in the formation of spherical structures. Although this suggestion was supported by an in vitro tubulation assay, the explanation of how SNX1 and SNX2 function was insufficient. A more detailed discussion of the molecular functions is required.\nResponse: To gain deeper insight into the functional interplay between SNX1 and SNX2, we have performed all‑atom molecular dynamics simulations. Simulation results show that SNX1 and SNX2a form a heterodimer; however, the amphipathic‑helix (AH) domain of the complex resists membrane insertion, as the average AH–membrane does not decrease over the course of the simulation (Supplementary Fig. 14a-c and Supplementary Video 4). We have thus proposed that SNX2a modulates the activity of SNX1 by altering the conformation or membrane interaction dynamics of the heterodimer, thereby fine-tuning the curvature generation process.\nWe have now incorporated the above results into the new Supplementary Fig. 14 and integrated the corresponding discussions in the revised MS (pages 10-11, lines 305-320, highlighted in red).\n6.Fig. 6a–c: The fluorescence signal in the RNAi-SNX lines shown in Figure 6a appears stronger than that in the control, and the Western blotting results in Figure 6b also indicate that the total amounts of GFP-VSR2 and GFP are higher in the RNAi lines. Therefore, it is not convincing to conclude from these data that SNX knockdown impairs VSR2 recycling. Since the authors already have the mutant lines, the mislocalization and degradation of VSR2 should be checked in the mutant plants, not only in protoplasts.\nResponse: To address the concern, we repeated the protoplast experiments under conditions that ensure comparable expression levels of GFP-VSR2 between control and RNAi-treated cells. The updated data confirm that RNAi-mediated knockdown of SNX proteins leads to increased GFP signal within vacuoles (Fig. 6a-b).\nAdditionally, in response to the reviewer’s suggestion, we are currently generating transgenic Arabidopsis lines stably expressing GFP-VSR2 in various snx mutant backgrounds, including the snx1/snx2a/snx2b triple mutant. These experiments are ongoing and will be presented in a future study.\n7. The authors should specify which RAB5 and RAB7 were used for the localization and dominant-negative mutant expression analyses. In particular, conventional and plant-specific RAB5 are known to localize and function differently.\nResponse: We confirm that the RAB5 proteins used in our localization analyses are the conventional forms ARA7 and RHA1, while the RAB7 protein used in both the localization and dominant-negative mutant analyses refers specifically to RABG3f. We have now explicitly clarified this information in the Results section of the revised manuscript.\n8.The involvement of RABs was not mentioned in the discussion, and the authors’ interpretation of the related data was not clear.\nResponse: Thank you for pointing this out. We have now revised the Discussion section to include a more detailed interpretation of the data related to RABs. Specifically, based on the data and literature, we propose the following sequences: (i) RHA1/ARA7 is initially recruited to MVBs; (ii) RHA1/ARA7 then recruits the MON1–CCZ1 complex, which activates RABG3f; and (iii) RABG3f then serves as a key coordinator, recruiting downstream effectors—including the retromer component VPS35—to initiate RV formation. This revision clarifies the involvement of RABs and provides a more comprehensive interpretation of their contribution to the trafficking pathway we describe.\nWe have now integrated the above discussions in the revised MS (pages 14-15, lines 415-433 highlighted in red).\n9.Although the authors acknowledge the technical difficulty of resolving the SNX1–SNX2 complex structure by cryo-EM in the Discussion, this limitation is not sufficiently reflected in the interpretation of their comparative analysis between Arabidopsis and mouse SNX1. Given that SNX1 and SNX2 have non-overlapping functions, as shown in this study and in previous work (Pourcher et al., 2010, Plant Cell), and that SNX2 requires SNX1 for proper endosomal localization, it would be important to clearly state how the inability to resolve the SNX1–SNX2 complex structure affects the interpretation and significance of the structural comparisons. A more explicit discussion of this point in both the Introduction and Discussion could help clarify the rationale and impact of the present findings.\nResponse: Given the difficulty in obtaining structural details of the SNX1-SNX2a complex on membrane tubules through cryo-EM, we conducted all-atom molecular dynamics simulations to gain further insight into its membrane interaction properties. The simulation results revealed that the AH domain of the SNX1–SNX2a complex resists insertion into the membrane, as the average AH‑membrane distance does not decrease over the course of the simulation (Supplementary Fig. 14a-c and Supplementary Video 4). We propose that SNX2a modulates SNX1 activity by altering the conformation or membrane interaction dynamics of the heterodimer, thereby fine-tuning membrane curvature generation. Consequently, the SNX1–SNX2a complex exhibits lower membrane-binding affinity than the Arabidopsis SNX1 homodimer, resulting in the formation of shorter tubules and spherical vesicles in vitro, as demonstrated by liposome tubulation assays (Fig. 5e–f).\nWe have now integrated the above results and discussions in the revised MS (page 16, lines 472-485 highlighted in red).\n10.Most of this discussion is a repetition of what has already been described in the results. In contrast, the authors describe their hypothesis in the last section of the Discussion (page 15), but in a very strong way. Please provide a careful discussion by clarifying what is known and the authors’ proposition.\nResponse: We have now carefully revised the Discussion section to minimize redundancy with the Results section. In addition, we have adjusted the wording in the final section to more explicitly distinguish between current knowledge in the field and our proposed model, ensuring our conclusions are appropriately balanced and supported by evidence."}, {"round": 2, "reviewer_comments": {"reviewer_1": "The authors have addressed most of my previous concerns; the additional quantification and phenotype studies have further strengthened the main conclusion and the biological relevance of the unique feature of SNX in plants. Due to the weak phenotype of the snx1/snx2a/snx2b triple mutant, the authors were unable to provide additional genetic evidence; however, the explanation and rationale for using alternative mutant lines seem reasonable. I would encourage the authors to include this explanation in the discussion and provide an alternative hypothesis if possible.\nA few minor issues:\n1. The arrows on Figure 7e are too small; please enlarge them.\n2. Line 753, please provide further information on AlphaFold prediction (I assume it is AlphaFold 2), including the reference, parameters used and the number of repeats.", "reviewer_2": "I am satisfied by the revisions made.", "reviewer_3": "The revised manuscript addresses some of the requested experiments. Nevertheless, an important issue remains insufficiently clarified: the definition of “retrieval vesicles (RVs)” is still not convincing. Retrograde transport is a fundamental process in intracellular trafficking, and more rigorous evidence is required to define its carriers. At present, the data provided do not clearly demonstrate that SNX or retromer are reliable markers of RVs. While it is well established that SNX and retromer participate in recycling from MVBs, this does not automatically justify defining the vesicles they label as RVs. A more cautious description, such as “SNX- or retromer-associated vesicles,” would be more appropriate. Alternatively, the manuscript would be strengthened if it explicitly presented this work as a first attempt to propose a definition of plant RVs.\n1. In the Introduction (Page 4, line 97), the expression “recycling endosome form endosomes” is confusing and scientifically imprecise. Since “endosome” is a general term encompassing several distinct compartments (recycling, early, late, etc.), such wording can be misleading. I recommend rewriting this section with greater accuracy and with reference to current knowledge of membrane trafficking in eukaryotes, including yeast and animal systems.\n2. The authors argue that because VSRs are delivered to the vacuolar lumen in snx mutants, VSRs must be direct cargo of SNX. This interpretation seems somewhat overstated, and alternative explanations should be discussed. In particular, consideration of possible ESCRT-dependent MVB sorting would be important. Furthermore, the statement in the Abstract—“we demonstrated that SNX1 and SNX2 mediate the recycling of VSRs”—appears stronger than what the current data can support. The conclusions would be more convincing if they were presented in a more cautious manner, acknowledging possible indirect or secondary effects.", "reviewer_4": "The study by Li et al. employs a multipronged approach to characterize the plant-specific membrane trafficking pathway. The study is extensive and interesting but needs a few additional updates to meet the quality required for Nature Communications. As I am particularly asked to review the cryo-EM structural data, I have the carefully looked into this section and have following concerns regarding them which need to be addressed before its publication.\n1. While I agree that the map resolution is not sufficient for high-quality models, at 7–8 Å the map still shows clear secondary-structure features (e.g., helical tubes). However, the current models are not built appropriately; for instance, the two helices between residues 180–295 are not correctly placed in the observed electron density maps of SNX1_7-fold_intact. I recommend further improving the model qualities, including removing regions where no clear electron density is present (unless explicitly stated as hybrid model based on EM / Predication).\n2. In addition, the current figures do not illustrate how well the model fits the map, nor do they adequately present the cryo-EM data-processing workflow. I advise authors use one asymmetric unit and show map-model fitting from different orientations.\n3. Key details of cryo-EM processing workflow including the total number of particles after 2D classification and the number of particles used for each 3D classification jobs must be clearly reported. The authors should also provide additional representative 2D classes to give better overview of particle used for 3D classification rather than only two. It is puzzling for general readers why only two 3D classes or symmetries were used for refinement, and why additional classes or symmetries were not tried or explored.\n4. Based on the cryo-EM map, it appears that the resolution of some reconstructions could be improved. Did the authors attempt to use RELION, which is widely applied for helical reconstructions? Were alternative refinement strategies (e.g., Blush, symmetry expansion followed by subunit classification) tested to improve map quality? If so, these should be briefly described in the manuscript.\n5. Do the authors find any differences in overall conformation of AtSNX1 complexes between the classes with 1 or 2? This should be included in the supplementary figures.\n6. Additionally, the MD simulation experiments performed are not sufficient to draw proper conclusions. The key problem is the insufficient timescale of the experiments and lack of experimental replicates. I advise the authors to extend the simulation time to at least 150–400 ns, with one or more additional replicates for each experimental condition.\nMinor comments:\nI find that not all the EM micrograph images have scale bars. Do these images share the same scale bar as neighboring images? If so, this needs to be clarified in the figure legend."}, "author_response": "Reviewer #1 (Remarks to the Author):\nThe authors have addressed most of my previous concerns; the additional quantification and phenotype studies have further strengthened the main conclusion and the biological relevance of the unique feature of SNX in plants. Due to the weak phenotype of the snx1/snx2a/snx2b triple mutant, the authors were unable to provide additional genetic evidence; however, the explanation and rationale for using alternative mutant lines seem reasonable. I would encourage the authors to include this explanation in the discussion and provide an alternative hypothesis if possible.\nResponse: Thank you very much for your positive and constructive comments and your time in reviewing the MS.\nAs suggested, we have now incorporated the explanation into the Discussion in the revised MS (pages 18-19, lines 517-529, highlighted in red). Specifically, we added the following text:\n“Although SNX1 and SNX2 appear to play a central role in plant retrograde transport, the snx1/snx2a/snx2b triple mutant exhibited only mild defects in plant growth and negligible phenotypic changes during embryonic development (Fig. 6h-i and Supplementary Fig. 19f and 20d). These findings led us to hypothesize that the VPS subcomplex of the retromer may partially compensate for the loss of SNX function. Therefore, we generated double mutants by crossing snx1 with the core retromer mutant vps29. The resulting snx1⁻/⁻ vps29⁺/⁻ plants exhibited pronounced defects during seedling development. Ultrastructural analysis by TEM further revealed that embryos of snx1⁻/⁻ vps29⁺/⁻ contained enlarged MVBs and a markedly reduced number of spherical vesicles in the vicinity of MVBs compared with wild type (Fig. 7e–f). Collectively, these results suggest that the VPS subcomplex of the retromer may provide functional redundancy or engage compensatory mechanisms in response to the loss of SNX function. However, the underlying mechanisms remain unclear, and alternative explanations cannot be excluded.”\nA few minor issues:\n1. The arrows on Figure 7e are too small; please enlarge them.\nResponse: Done.\n2. Line 753, please provide further information on AlphaFold prediction (I assume it is AlphaFold 2), including the reference, parameters used and the number of repeats.\nResponse: We have now added detailed methodological information on the AlphaFold prediction (page 28, lines 833-845, highlighted in red), together with a new Figure S23, including the reference, parameters, and number of replicate predictions, as follows:\n“The structure of AtSNX2a was predicted using the AlphaFold Server (powered by AlphaFold3; https://alphafoldserver.com) (Abramson et al., 2024). AtSNX2a-related information was retrieved from Uniport (Uniport ID: QBL5Z7). Given that the N-terminus of AtSNX2A harbors a long intrinsically disordered region, only the amino acid sequence corresponding to residues 157–587 was used for structure prediction (parameters: protein entity, single copy, 10 replicate predictions). Each prediction run generated five models, and the top-ranked model (rank_0, highest confidence score) was selected as the representative structure. Predicted template modeling (pTM) scores were then compared, and the model with the highest pTM score (0.75) was selected for subsequent molecular simulations (Supplementary Fig. 23a). Additional predicted structures of AtSNX2a are shown in Supplementary Fig. 23b-e. While the overall folds of these models are highly conserved, the disordered region linking the PX and BAR domains is poorly resolved by AlphaFold3, leading to notable structural variability in this region (Supplementary Fig. 23f).”\nReference:\nAbramson, J. et al. Accurate structure prediction of biomolecular interactions with AlphaFold 3. Nature 630, 493-500 (2024).\nReviewer #2 (Remarks to the Author)\nI am satisfied by the revisions made.\nResponse: Thank you very much.\nReviewer #3 (Remarks to the Author)\nThe revised manuscript addresses some of the requested experiments. Nevertheless, an important issue remains insufficiently clarified: the definition of “retrieval vesicles (RVs)” is still not convincing. Retrograde transport is a fundamental process in intracellular trafficking, and more rigorous evidence is required to define its carriers. At present, the data provided do not clearly demonstrate that SNX or retromer are reliable markers of RVs. While it is well established that SNX and retromer participate in recycling from MVBs, this does not automatically justify defining the vesicles they label as RVs. A more cautious description, such as “SNX- or retromer-associated vesicles,” would be more appropriate. Alternatively, the manuscript would be strengthened if it explicitly presented this work as a first attempt to propose a definition of plant RVs.\nResponse: Thank you for your constructive comments. We agree that a rigorous definition of “retrieval vesicles” requires more evidence. In accordance with the reviewer’s suggestion and to ensure a more cautious and accurate interpretation, we have revised the manuscript throughout to replace the term “retrieval vesicles (RVs)” with more descriptive terminology, “spherical vesicles near MVBs” where appropriate. We believe this change more accurately reflects the scope of our data, improves the clarity and rigor of the manuscript, and avoids overinterpretation. We have also added a dedicated discussion addressing the nature of these spherical vesicles near MVBs and acknowledge that further evidence will be required to fully understand their identity and function (page 19, lines 536-540, highlighted in red). Specifically, we have added the following text:\n“The nature of the spherical vesicles observed near MVBs in plant cells. Although we identified a plant-specific vesicle population that appears to originate from MVBs, is labeled by retromer and VSR proteins, and whose formation depends on retromer components, additional evidence is required to demonstrate that these vesicles traffic to the TGN and function as bona fide retrieval carriers.”\n1. In the Introduction (Page 4, line 97), the expression “recycling endosome form endosomes” is confusing and scientifically imprecise. Since “endosome” is a general term encompassing several distinct compartments (recycling, early, late, etc.), such wording can be misleading. I recommend rewriting this section with greater accuracy and with reference to current knowledge of membrane trafficking in eukaryotes, including yeast and animal systems.\nResponse: We appreciate the reviewer’s insightful comment. We have carefully revised this section of the Introduction to improve clarity and scientific accuracy, incorporating current understanding from yeast and animal systems in the revised manuscript (page 5, lines 101-105; pages 5-6, lines 113-126; highlighted in red). Specifically, we have added the following text:\n“In yeast, retrograde transport is mediated by the retromer complex, a pentamer composed of a membrane-binding SNX subcomplex, formed by the heterodimer Vps5p and Vps17p, and a cargo-recognition VPS subcomplex comprising Vps26p, Vps29p, and Vps35p26. In animals, SNX1 and SNX2 are homologous to Vps5p, whereas SNX5 and SNX6 share a common ancestor with Vps17p and are closely related.”\n“The VPS subcomplex consists of VPS26-VPS29-VPS35, where VPS35 interacts with VPS26 and VPS29 through its N-terminal and C-terminal domains, respectively, forming a horseshoe-shaped α-helix28. Unlike the stable heteropentameric retromer complex in yeast, the SNX-BAR heterodimers SNX1/2–SNX5/6 in animals can assemble into functionally distinct endosomal sorting complexes (ESCPE-1) that operate independently of the retromer core29. In Arabidopsis, homologs of the Vps5p and VPS proteins are present, comprising three genes encoding SNX (SNX1, SNX2a, and SNX2b), three encoding VPS35 (VPS35a, VPS35b, and VPS35c), two encoding VPS26 (VPS26a and VPS26b), and one encoding VPS29. SNX1 has been shown to be essential for the recycling of PIN2, BRI1, IRT1, PHT1, and SOS1 to the plasma membrane14, 20, 30-32. Recent studies indicate that the retrieval of VAMP727, a protein involved in vacuolar fusion in plants, from the vacuolar membrane depends on the SNX complex but not on the VPS complex33. In addition, VPS complex also participates in proper host invasion in rice blast34-36. Although the structure and function of retromer components in animals and yeast have been increasingly well characterized, the mechanism by which the retromer complex mediates retrograde transport in plants remains unclear37-41.”\n2. The authors argue that because VSRs are delivered to the vacuolar lumen in snx mutants, VSRs must be direct cargo of SNX. This interpretation seems somewhat overstated, and alternative explanations should be discussed. In particular, consideration of possible ESCRT-dependent MVB sorting would be important. Furthermore, the statement in the Abstract—“we demonstrated that SNX1 and SNX2 mediate the recycling of VSRs”—appears stronger than what the current data can support. The conclusions would be more convincing if they were presented in a more cautious manner, acknowledging possible indirect or secondary effects.\nResponse: We agree with the reviewer that our current data do not demonstrate that VSRs are direct cargo of SNX. Accordingly, we have revised the manuscript (page 13, lines 357-363, highlighted in red) to avoid overstatement and now describe our findings more accurately as “Notably, RNAi-mediated suppression of SNX1 or SNX2 expression in protoplasts resulted in its mislocalization of GFP-VSR2 to the vacuole (Fig. 6a). Subsequently, we performed protein immunoblotting and semi-quantitative analysis, which revealed that, compared to the control group, the degradation rate of GFP-VSR2 was significantly increased in protoplasts transformed with RNAi vectors targeting SNX1 or SNX2 (Fig. 6b and 6c).”\nIn addition, we have expanded the Discussion to more carefully consider the relationship between SNX function and VSR recycling, and we now explicitly acknowledge alternative interpretations of the observed phenotype. Specifically, we discuss the possibility that enhanced VSR degradation may result from indirect or secondary consequences of SNX perturbation, and we also consider the potential contribution of ESCRT-dependent MVB sorting to VSR degradation. The manuscript has been revised accordingly (pages 19-20, lines 540-560, highlighted in red), as follows:\n“An additional unresolved issue concerns the relationship between SNX function and VSR recycling. While knockdown of SNX1 or SNX2 resulted in vacuolar mislocalization and enhanced degradation of GFP–VSR2, it remains unclear whether VSRs constitute direct cargo of SNX proteins. Previous studies have shown that Arabidopsis VSR1 can bind VPS35 as revealed by immunoprecipitation assays15, and that in yeast, the VSR homolog, Vps10p receptor interacts directly with retromer via Vps35p51. However, the molecular relationship between SNX proteins and VPS35 in Arabidopsis, and whether SNXs directly participate in VSR recognition, remain unresolved. Alternatively, the observed phenotype may arise from indirect or secondary consequences of SNX perturbation. For example, loss of SNX function could broadly disrupt endosomal membrane organization, phosphoinositide composition, or retromer recruitment, thereby indirectly impairing VSR recycling20, 52. Furthermore, impaired recycling of other regulatory factors or cargo adaptors may secondarily compromise VSR localization, rather than VSRs being directly recognized by SNX proteins themselves. In addition, degradation of membrane proteins residing on the limiting membrane of MVBs, such as VSRs, generally requires their sorting into intraluminal vesicles (ILVs) through ESCRT-dependent mechanisms53. However, the mechanism by which VSRs become missorted into ILVs in the absence of SNX function remains unknown. Addressing these questions will be important for elucidating the precise molecular pathway underlying VSR recycling and retromer-mediated trafficking in plant cells.”\nAs suggested, we have also revised the Abstract to adopt a more cautious conclusion, replacing the original statement with: “Lastly, knockdown of SNX1 or SNX2 resulted in vacuolar mislocalization and increased degradation of GFP-VSR2.”\nReviewer #4 (Remarks to the Author):\nThe study by Li et al. employs a multipronged approach to characterize the plant-specific membrane trafficking pathway. The study is extensive and interesting but needs a few additional updates to meet the quality required for Nature Communications. As I am particularly asked to review the cryo-EM structural data, I have the carefully looked into this section and have following concerns regarding them which need to be addressed before its publication.\nResponse: Thank you for your constructive comments and your time in reviewing the MS.\n1. While I agree that the map resolution is not sufficient for high-quality models, at 7–8 Å the map still shows clear secondary-structure features (e.g., helical tubes). However, the current models are not built appropriately; for instance, the two helices between residues 180–295 are not correctly placed in the observed electron density maps of SNX1_7-fold_intact. I recommend further improving the model qualities, including removing regions where no clear electron density is present (unless explicitly stated as hybrid model based on EM / Predication).\nResponse: We thank the reviewer for this comment. We have now improved the model to achieve better agreement with the density map. The updated data have been incorporated into the revised Fig. 2e-f, Fig. 3a-c, and Supplementary Figs. 9-12. Please also refer to the updated validation report for details. A description of the revised model refinement has been added to the Methods section (Model building, page 28, lines 818-826, highlighted in red). The refined overall model and the asymmetric unit fitted into the map are shown in Fig. R1.\nFig. R1. Cryo-EM maps of two AtSNX1 classes and fitted models.\na, Different views of the cryo-EM map of AtSNX1 class I with the corresponding dimer fitted into the density. b, Different views of the cryo-EM map of AtSNX1 class II with the corresponding dimer fitted into the density. c, Asymmetric units of AtSNX1 class I and class II shown in different orientations.\n2. In addition, the current figures do not illustrate how well the model fits the map, nor do they adequately present the cryo-EM data-processing workflow. I advise authors use one asymmetric unit and show map-model fitting from different orientations.\nResponse: To enable a better assessment of map–model consistency, we have added additional figure panels showing both a dimer and a single asymmetric unit fitted into the map from multiple orientations (Fig. R1). In addition, the cryo-EM data-processing workflow is now presented in detail in a new Supplementary Fig. 9a, as described in the following response.\n3. Key details of cryo-EM processing workflow including the total number of particles after 2D classification and the number of particles used for each 3D classification jobs must be clearly reported. The authors should also provide additional representative 2D classes to give better overview of particle used for 3D classification rather than only two. It is puzzling for general readers why only two 3D classes or symmetries were used for refinement, and why additional classes or symmetries were not tried or explored.\nResponse: We have now explicitly reported the particle numbers at each stage of cryo-EM data processing, as shown in the new Supplementary Fig. 9a. During 2D classification, multiple 2D class averages corresponding to tubes with different diameters were observed. Accordingly, we performed asymmetric helical refinements and extensive symmetry searches in CryoSPARC to explore alternative structural solutions. Despite these efforts, only two particle subsets consistently converged to stable consensus maps, which are presented in the manuscript. Other classes either failed to converge or yielded maps of insufficient quality and were therefore excluded from further analysis. These details have been added to the Methods section (Cryo-EM data processing, pages 27–28, lines 801–808, highlighted in red).\nSupplementary Fig. 9a. Image processing of AtSNX1. a, Flowchart of imaging processing. The colored boxes of 2D classes represent the particle subsets used for the following asymmetric helical refinement.\n4. Based on the cryo-EM map, it appears that the resolution of some reconstructions could be improved. Did the authors attempt to use RELION, which is widely applied for helical reconstructions? Were alternative refinement strategies (e.g., Blush, symmetry expansion followed by subunit classification) tested to improve map quality? If so, these should be briefly described in the manuscript.\nResponse: We thank the reviewer for the suggestion. In attempt to employ RELION to further improve the map resolution, we re-extracted the final set of particles and then performed the BLUSH regularization-enable helical refinement in RELION by following the standard pipeline or by using the helical twist and rise parameters previously obtained from CryoSPARC (Fig. R2 a-c), with or without symmetry enforcement. Unfortunately, we were unable to obtain reconstructions that are of better quality than those obtained from CryoSPARC. Notably, CryoSPARC has become a widely adopted and reliable platform for helical reconstruction (Schäfer et al., 2025). In accordance with the reviewer’s suggestion, we have further conducted focus refinement after symmetry expansion and signal subtraction in CryoSPARC. Our local resolution analysis reveals that there is no substantial improvement in map quality for the majority of the masked region (Fig. R3). We therefore did not include the analyses in the manuscript and the corresponding discussion is found in the Methods Section (Cryo-EM data processing, page 28, lines 811-814, highlighted in red).\nReference: Schäfer, J.-H., O’Neill, R. T., Donnelly, J. P., Powers, E. T., Kelly, J. W., & Lander, G. C. (2025). Helical Reconstruction of Amyloids in cryoSPARC. bioRxiv, 2025.2010.2003.680389. doi:10.1101/2025.10.03.680389\nFig R2. Representative density maps from RELION 5 with BLUSH regularization after performing homogenous reconstruction only in Class3D (a), helical reconstruction with (b) or without symmetry enforcement (c) by using extracted particles exported from CryoSPARC. Maps obtained from helical reconstruction with (d) or without symmetry enforcement (e) after re-extraction and normalization in RELION5.\nFig R3. Local resolution estimation of AtSNX1 dimer from Class I (a) and Class II (b) after local refinement on symmetry expanded particle stack in CryoSPARC (left), compared to the local estimation of AtSNX1 tubes (right).\n5. Do the authors find any differences in overall conformation of AtSNX1 complexes between the classes with 1 or 2? This should be included in the supplementary figures.\nResponse: Upon comparing the asymmetric units from the 2 classes by overlaying, we found that the monomer (asymmetric unit) and dimer configurations are largely similar between Class I and Class II. However, there are significant differences at the level of higher-order scaffold organization, that is, at the dimer-dimer interfaces. We have now included the model comparisons between the 2 classes in Supplementary Fig. 13. The monomer from different classes reveal only subtle movements of the helices connecting the PX and BAR domains (new Supplementary Fig. 13a). In helical lattices, Class II exhibits a shorter radius of curvature, associated with a smaller intersecting angle between neighboring dimers (new Supplementary Fig.13b). As a result, fewer asymmetric units are required per helical turn in Class II when compared with Class I.\nThe above discussion of the conformational differences between the SNX1 classes has been included in the revised manuscript (pages 9–10, lines 239–256, highlighted in red).\nSupplementary Fig. 13. Structural comparisons between plant SNX1 and its homologs reveal the molecular basis for the distinct membrane remodeling behavior in plants. a, Superimposition of the BAR domains of the dimers of plant SNX1 and its homologs: Mouse SNX1 7-fold (PDB accession code 7D6E) and Fungi Vps5 (PDB accession code 6H7W). b, Dimer-dimer interactions revealed by structural alignment of the dimer of dimers of SNX1 with its homologs, shown by superimposing of the upper latitudinal interface (Interface I) the scaffold while keeping the position and orientation of the adjacent dimer fixed (indicated by arrow). The radius of curvature was measured by the 2D projection of the dimer of dimers and the curvature length was the arc length of the 2D projection of the dimer of dimers. The intersecting angle was measured by the 2D projection of the dimer on the upper layer as shown in the figure.\n6. Additionally, the MD simulation experiments performed are not sufficient to draw proper conclusions. The key problem is the insufficient timescale of the experiments and lack of experimental replicates. I advise the authors to extend the simulation time to at least 150–400 ns, with one or more additional replicates for each experimental condition.\nResponse: Using the updated AtSNX1 structures, we extended the MD simulation time to 200 ns and conducted two independent replicate simulations for each experimental condition to enhance the robustness and reproducibility of the results. The extended simulations yielded consistent trajectories and supported the same overall conclusions as our original analyses, thereby strengthening the reliability of our findings.\nThe revised results and updated analyses have been incorporated into the revised Figure 4a-f, Supplementary Fig. 14-16.\nMinor comments:\nI find that not all the EM micrograph images have scale bars. Do these images share the same scale bar as neighboring images? If so, this needs to be clarified in the figure legend.\nResponse: Scale bars have now been added to all EM micrograph images."}, {"round": 3, "reviewer_comments": {"reviewer_3": "I felt that the revised manuscript was substantially improved, and that my concerns were sincerely addressed.", "reviewer_4": "The authors have addressed all concerns, and the manuscript is suitable for publication."}, "author_response": ""}]

Decision letter