Molecular mechanisms of flotillin complexes in organizing membrane microdomains

Domain: neuroscience
Published: 2026-02-09
Editorial outcome: major_revision

Abstract
Flotillin-1 and flotillin-2 form hetero-oligomers to create flotillin membrane microdomains essential for endocytosis and protein sorting. However, the mechanisms of flotillin oligomerization and microdomain organization remain incompletely understood. Here, we present the cryo-EM structure of human flotillin complex, showing that flotillin-1 and -2 form a 44-mer, membrane attached, and dome-shaped structure that defines a 30-nm circular membrane domain. The cryo-ET data demonstrates that while attached to the cytoplasmic leaflet, flotillin complexes possess intrinsic structural plasticity in situ on the native membrane. Each flotillin complex may represent a fundamental unit of membrane microdomains, with their clustering enabling the formation of larger and more elaborate domains. We further reveal that phosphorylation at residues Y160 (flotillin-1) and Y163 (flotillin-2) may act as a molecular switch to modulate complex assembly, potentially regulating its function in endocytosis. These findings demonstrate the molecular mechanism of flotillin-mediated membrane segregation and microdomain formation, and suggest a previously unrecognized role of flotillin in sequestrating membrane proteins.

Introduction
Biological membranes are heterogenous in composition and due to the non-uniform distribution of lipids and proteins, are segregated into discrete nanoscale domains that function as dynamic platforms to orchestrate biomolecule interactions both within the membrane and between the membrane and intra/extracellular contents1,2. These functional membrane microdomains (FMMs), as basic organizational units of the membrane, are essential for many cellular processes3,4. FMMs are enriched in certain lipid species, such as sterols and sphingolipids and difference classes of proteins5,6. Mammalian flotillin-1 and flotillin-2 are two highly homologous proteins found in the membrane microdomains of plasma membrane7. They can be efficiently co-immunoprecipitated by each other and extensively colocalize, forming small puncta in the plasma membrane that correspond to flotillin membrane microdomains8. The membrane microdomains containing flotillin complexes are laterally mobile within the plasma membrane, and play crucial roles in various cellular processes, including endocytosis9–11, signaling12,13, protein trafficking13,14 and cytoskeleton remodeling15–17 (reviewed in refs. 18,19). The flotillin microdomains can be internalized into the cell with various cargos8,10,20–22 or secreted out of the cell23,24. For examples, Fyn kinase11 and RAB3123 respectively regulate the roles of flotillins in endocytosis and exosome secretion. And flotillin overexpression has been implicated in cell invasion and metastasis during tumorigenesis19,25. This overexpression disrupts membrane domain homeostasis, promoting plasma membrane invagination and endocytosis towards late endosomes, which in turn alters the trafficking of various cargos8, potentially contributing to oncogenic processes.

Although a scaffolding role of flotillins in FMM organization was proposed decades ago7,26, the physical basis of flotillin microdomain remains to be an enigma and it is more intriguing that how flotillin is involved in such a variety of different biological processes. Flotillin belongs to the SPFH (Stomatin, Prohibitin, Flotillin, and HflK/C) protein family, which shares a conserved structural organization that comprises an SPFH domain and a subsequent long α-helix (coiled-coil, CC domain)27. These long α-helices of both flotillin-1 and flotillin-2 were demonstrated to be important for their oligomerization28. Recent findings suggest that several representative SPFH family proteins can self-oligomerize and anchor to membranes, creating diverse membrane microdomains across various subcellular compartments27, such as stomatin complexes in the eukaryotic plasma membrane29–31, prohibitin (PHB) complexes in the inner mitochondrial membrane32–35, erlin complexes in the endoplasmic reticulum (ER)34,36, and HflK/C complexes in the prokaryotic plasma membrane37,38. Different from these typical SPFH proteins, another SPFH protein, bacterial flotillin (also known as FloA) contains a unique membrane-targeting region39, and was also demonstrated to play a role in organizing FMMs on bacterial membrane40,41. Given the conserved domain architecture of FloA, it likely employs a similar oligomerization mechanism to segregate bacterial membrane39,42. While most of these proteins possess transmembrane domains, mammalian flotillins lack such domains yet exhibit robust association with membranes. This association is facilitated by a combination of post translation modifications: S-palmitoylation of a few cysteine residues in their N-terminal domains and an N-terminal myristoylation specific for flotillin-215. In addition, a putative cholesterol recognition/interaction amino acid (CRAC) motif within their SPFH domain may also facilitate membrane association43. However, the structural basis for flotillin-1 and flotillin-2 oligomerization and the molecular mechanisms underlying the formation of flotillin membrane microdomains remain unknown.

In this work, we employ multi-modal imaging approaches to resolve the high-resolution structure of purified flotillin complex and the in situ architecture of the flotillin microdomain. Our results reveal that flotillin complexes exhibit intrinsic structural plasticity on membranes, where each complex may serve as a fundamental unit for microdomain formation. Clustering of these units enables the formation of larger and more elaborate domains. These findings elucidate the molecular basis of flotillin-mediated membrane segregation and uncover their role in membrane protein sequestration, opening avenues for the further exploration of their mechanisms in various cellular functions.

Results
Sample preparation and structural determination of the flotillin complex
Flotillin-1 and flotillin-2 were co-expressed in HEK293F cells, and a 3×FLAG tag was placed at the C-terminus of flotillin-1 for the complex purification from solubilized membrane fractions. To further increase sample homogeneity, glycerol density gradient centrifugation was performed and peak fractions were used for structural analysis (Supplementary Fig. 1a). The distribution of flotillin-1 and flotillin-2 in the glycerol fractions was concentrated in high molecular-weight fractions, and each fraction was examined using negative-staining electron microscopy (nsEM) (Supplementary Fig. 1b). The flotillin complex exhibited trapezoid-like shapes (side view) (Supplementary Fig. 1c) and round shapes (top/bottom view) in the nsEM images. From the nsEM images, some complexes were seen to attach to membrane vesicles with the lower bottom of the trapezoids (Supplementary Fig. 1d).

Subsequent cryo-electron microscopy (cryo-EM) analysis confirmed that flotillin complexes resembled a dome-like structure (Supplementary Fig. 2). However, a significant portion of particles exhibited irregularly deformations, and some classes from early 3D classification even show ruptures in the structures (Supplementary Fig. 2d, e). Due to the heterogeneity of the particles, the alignment of 2D particle images during single-particle cryo-EM 3D reconstruction was challenging and resulted in low-resolution density maps. To obtain a high-resolution refinement, we eliminated deformed particles during data processing. Only those particles without apparent deformations were included in the analysis. C22 symmetry was imposed during refinement, which resulted in a final density map of overall 3.57 Å resolution (Supplementary Figs. 2f, 3). Atomic models for flotillin-1 and flotillin-2 were built based on this map.

The structure of the flotillin complex shows that 22 copies of flotillin-1 and 22 copies of flotillin-2 are alternately arranged to form a dome-like structure, denoted as F1(1)–F1(22) and F2(1)–F2(22), respectively, giving rise to the C22 symmetry (Fig. 1). When viewed from the side, the structure of flotillin complex can be divided into three parts: the base region (~70 Å high), the wall region (~140 Å high), and the roof region (~175 Å wide) (Fig. 1c). The base region comprises the SPFH domains of flotillin-1 and flotillin-2, which are associated with the membrane (density of detergent micelles shown in Fig. 1c). The wall region of the structure is formed by a single layer of parallel-arranged helices, creating a helical barrel. The dome structure is tightly sealed in both the base region and wall region, with the inner leaflet of the plasma membrane serving as the bottom.

Recently, concurrent with our work, another study reported the structural analysis of flotillin complexes on isolated membrane vesicles44. Their results demonstrate the same overall architecture of the flotillin complex, indicating that the detergent-solubilized flotillin complex has retained its native structure in the membrane environment.

Structure of the flotillin complex
With approximately 48% sequence identity, flotillin-1 and flotillin-2 have nearly identical 3D structures (Fig. 2a, b). The structures of both proteins can be divided into five domains, including two SPFH domains (SPFH1 and SPFH2), two coiled-coil domains (CC1 and CC2), and a C-terminal domain (CTD), similar to the structure of the HflK/C complex37,38.

Flotillin-1 and flotillin-2 form two distinct heterodimeric interfaces, F1:F2 and F2:F1 (Fig. 2c). The SPFH1 domains of flotillin-1 and flotillin-2 interact with each other via the less conserved loop regions on their structurally conserved five-stranded core (Supplementary Figs. 4a–d and 5). The interfaces of adjacent SPFH2 domains are characterized by oppositely charged surface patches, which exhibit a clear charge complementarity (Supplementary Fig. 4e–h). These charged patch pairs dictate that flotillin-1 and flotillin-2 should be alternately arranged to form a complete complex. Following the SPFH domains, the CC1 domains of flotillin-1 and flotillin-2 both contain 189 residues, which are significantly longer than those of other SPFH family members (Supplementary Fig. 5), and they align perfectly to form a right-handed helical barrel and serve as the dome wall (Fig. 1). The interactions between parallel CC1 domains are evenly distributed along the entire helices, with no preference for either polar (Supplementary Fig. 6a–c) or hydrophobic interactions (Supplementary Fig. 6d–f). Besides, there are two small bends at the sites of Pi-helical (π) and 310-helical (η) motifs of CC1 domains (Fig. 2a, b), constituting the two hoops observed in the side view of the dome (Fig. 1c).

Following the CC1 domains, the CC2 domains and CTDs of both flotillins form the roof of the dome, primarily stabilized by hydrophobic interactions (Fig. 2d–f). The 44 CC2 domains (α6) converge towards the center, where 44 β-strands assemble into a β-barrel core. The subsequent sequences in the CTDs of flotillin-1 and flotillin-2 exhibit distinct structural characteristics, resembling the organization observed in the HflK/C complex37,38. In flotillin-1, the helices (α7) in the CTDs are oriented outwards from the center, with the C-terminus exposed to the cytoplasmic space. In contrast, the putative helices (α7) in the CTDs of flotillin-2 appear to be buried within the flotillin complex, although their exact conformation is not fully resolved in the map. This structural difference aligns with experimental observations, as attempts to purify the flotillin complex using a 3×FLAG tag at the C-terminus of flotillin-2 were unsuccessful, suggesting that the C-terminal region of flotillin-2 is inaccessible in the assembled complex. Similar to the HflK/C complex38, progressive truncation of the C-terminal sequences may disrupts dome structure formation (Supplementary Fig. 7).

Potential phosphorylation mediated regulation on the flotillin complex assembly
Y160 in flotillin-1 and Y163 in flotillin-2 are two known phosphorylation sites for Src family kinase Fyn6. These two residues are localized in the hinge regions between the SPFH2 and CC1 domains of flotillin-1 and -2, which might play a role in determining the orientation of the CC1 domain relative to the SPFH2 domain. Structure-wise, Y163 and Y160 are stabilized by multiple conserved hydrophilic and hydrophobic interactions across both flotillin homologs, in the F1(1):F2(1) and F2(1):F1(2) interfaces, respectively (Fig. 2g, h). On the one hand, polar interactions are established between the hydroxyl group of Y163 of F2(1) and R168 of F1(1) (Fig. 2g), and between the hydroxyl group of Y160 of F1(1) and Q171 of F2(1) (Fig. 2h), forming the solvent-exposed surfaces. On the other hand, the benzene rings Y163 and Y160 directly contribute to the formation of a hydrophobic core involving both inter- and intra-molecular interactions, including V172 of F1(1) with V65, L67, V120, L164, and L167 of F2(1) (Fig. 2g), and V175 of F1(1) with V61, I63, V117, L161, and L164 of F2(1) (Fig. 2h), respectively.

Next, we introduced mutations on Y160 of flotillin-1 and Y163 of flotillin-2 and examined their effects on the complex assembly. Since there is no phosphomimetic mutations for tyrosine, we introduced Y160E and Y163E to partially mimic the charge effect of phosphorylation, and found that they completely abolished the complex assembly (Fig. 2i, Supplementary Fig. 7). In contrast, neither the tyrosine mutations (Y160F of flotillin-1 and Y163F of flotillin-2) nor the double mutations (Y160F/R168A of flotillin-1 and Y163F/Q171A of flotillin-2) affected flotillin complex assembly (Fig. 2i, Supplementary Fig. 7). These results indicate that the stability of the Y160/Y163 participated hydrophobic core is essential for the formation of the flotillin complex. Very interestingly, it was previously shown that Y160F (flotillin-1) and Y163F (flotillin-2) mutations could block Fyn-regulated flotillin-mediated endocytosis11,17. In combination with our results, it appears to suggest that flotillin-mediated endocytosis might involve the disassembly of the dome structure.

Altogether, the previous and our observations indicate that phosphorylation at these tyrosines may destabilize the local hydrophobic cores, inducing a change on the orientation between the SPFH2 and CC1 domains that could function as a molecular switch to trigger the flotillin complex disassembly.

Flotillin complex mediated membrane compartmentalization
The SPFH1 domains mediate the membrane association of flotillins, with approximately half embedded in the detergent micelle (Fig. 3a–d). These domains are rich in aromatic residues, including F2, F3, F16 and F30 of flotillin-1, and W32, W34, W36 and W37 of flotillin-2 (Fig. 3d), contributing to the highly hydrophobic nature of the bottom rim of the flotillin complex (Fig. 3e). Notably, palmitoylated cystine residues (C34 of flotillin-1; C4, C19 and C20 of flotillin-2)15 and myristoylated glycine residue (G2) of flotillin-2 are all buried in membrane (Fig. 3d). Together, these features collectively ensure the stable and efficient membrane association of flotillins.

The 44 circularly arranged SPFH1 domains define a 30 nm-diameter membrane area on the cytosolic leaflet of the membrane (Fig. 3a, b). When all flotillin subunits are precisely aligned, they form a sealed dome that effectively compartmentalizes both the membrane and the perimembrane space, physically isolating it from the surrounding environment (Fig. 1). However, in addition to these well-aligned complexes, irregular deformations and structural ruptures were frequently observed during in vitro structural determination (Fig. 3f, Supplementary Fig. 2d–f). These observations suggest that the sealed dome may dynamically open and reseal, potentially regulating access to the segregated region.

In situ structure of the flotillin complex
To explore whether such deformations occur within a cellular context and to gain further insight into the potential functional roles of the flotillin complex, we developed a stable Jurkat T cell line co-expressing F1-EGFP and F2-mCherry for in situ studies. Flotillin complexes are known to exhibit a polarized distribution in T cells, localizing to caps, uropods, and the immune synapse during resting45, migrating16, and conjugated stages13, respectively. Consistently, our fluorescent confocal imaging results show that flotillin-1 and flotillin-2 were co-localized in caps or uropods of Jurkat T cells only when both flotillin homologs are co-expressed (Fig. 4a, Supplementary Fig. 8a–c), highlighting the necessity of hetero-oligomer formation for their function. Additionally, glycerol density gradient centrifugation assay confirmed that the fluorescent tags did not disrupt the assembly of flotillin complexes (Supplementary Fig. 8d).

Guided by the fluorescent signals of F1-EGFP and F2-mCherry, lamellae were prepared at regions enriched with flotillin complexes (Fig. 4b). Subsequent tomographic reconstructions revealed a cellular periphery with a clearly visible plasma membrane, consistent with the known localization of flotillin complexes in caps or uropods (Fig. 4c, d, Supplementary Movie 1). Notably, a significant number of hollow dome-shaped structures with heights of approximately 20 nm were observed attached to the cytosolic side of the plasma membrane (Fig. 4e), resembling our cryo-EM structure of the flotillin complex. To validate the identity of these structures, sub-tomogram averaging (STA) was performed. While the limited data size restricts the final map resolution and precludes definitive identification based solely on structure, the overall shape of the structure in conjunction with the correlative light and electron microscopy (CLEM) results strongly suggest their formation by the flotillin complex (Fig. 4f, g).

Individual subtomograms revealed additional electron densities within the complexes (Fig. 4e). These densities likely correspond to various partner proteins. However, they were absent in the averaged structure, suggesting a high heterogeneity of partner proteins with the domes. Additionally, a significant degree of structural variation was observed among the individual complexes (Supplementary Figs. 9a, b, 10a). This observation aligns with the findings from purified samples (Fig. 3f, Supplementary Figs. 2b–e, 9c, d), further supporting the notion that the flotillin complexes are highly dynamic in nature. This property might be essential for the flotillin complex’s cellular functions.

Organization model of flotillin membrane microdomains
The tomographic data offered an opportunity to investigate the spatial distribution and membrane preferences of flotillin complexes in situ. A majority of the observed flotillin complexes existed as individual units. Given the substantial membrane area encompassed by the dome-shaped complex, it is conceivable that the complex, along with its associated proteins, contribute to the organization of a specialized membrane microdomain. Notably, we also observed clusters of flotillin complexes on extracellular vesicles (Fig. 5a, Supplementary Movie 2), endosomes and plasma membrane (Fig. 5b, Supplementary Movie 3), which aligns with the presence of larger flotillin membrane microdomains implicated in exosome formation23 and endocytosis8,10,11.

The flotillin complexes were observed on membranes with varying curvature in situ. Most flotillin particles were located on relatively flat membranes (Supplementary Fig. 10a), and some appeared on convex or concave vesicle surfaces (Supplementary Fig. 10b, c). In purified samples, the averaged structure of non-vesicle-associated particles retained a concave-shaped membrane at the bottom (Supplementary Fig. 10d), while some complexes attached to the convex membrane surfaces of vesicles (Supplementary Fig. 10e). These observations indicate that flotillin complexes exhibit no specific preference for membrane curvature.

Since flotillin complexes also exhibit a clustering pattern, we sought to examine whether this behavior is related to the distribution of certain lipid species on the plasma membrane. Notably, flotillin was considered as a marker of lipid rafts7,26,27, which commonly refer to those highly dynamic, cholesterol and sphingolipids enriched microdomains (ranging from 10 to 200 nm) on the plasma membrane46. A model system to study the lipid driven phase separation property of plasma membrane is the giant plasma membrane vesicle (GPMV)47. Therefore, we prepared GPMVs from human Jurkat cells stably expressing tagged flotillin-1 and flotillin-2. Previously, flotillins were found to be enriched in the detergent-resistant membrane (DRM) fractions7,26, a sign usually suggesting a localization in the liquid-ordered (Lo) phase of plasma membrane. Unexpectedly, but consistent with a previous report on flotillin-248, our data indicate that both flotillin-1 and -2 predominantly partition into the liquid-disordered (Ld) phase in GPMV-based partitioning assay (Fig. 5c, Supplementary Fig. 11a). Similarly, tetraspanin-4 (T4), a member of the tetraspanin protein family that scaffolds tetraspanin-enriched microdomains49,50, colocalizes with flotillins in the vesicles within L929 cells (Supplementary Fig. 11b), and also partitions into the Ld phase (Supplementary Fig. 11c).

Furthermore, the use of GDN detergents during protein purification preserved portions of the membrane within the complex (Supplementary Fig. 1c), enabling mass spectrometry to identify endogenous lipids co-purified with flotillins (Supplementary Data 1). Comparative lipidomics analysis revealed a 1.8-fold enrichment of sphingosine (SPH) in the purified flotillin sample compared to the plasma membrane fraction, with no significant increase observed in other raft-associated lipids, such as cholesterol (CH) or other sphingolipids. This enrichment was accompanied by relative decreases in several other lipid species, including phosphatidylinositol (PI), lysophosphatidylcholine (LPC), ceramide (Cer), and triacylglycerol (TG) (Fig. 5d). This observation aligns with previous reports indicating that flotillins interact with sphingosine51. These findings, together with results of GPMV partitioning assay suggest that the flotillin complex does not have a strong preference for certain lipid-enriched membrane regions and its further clustering may not primarily dependent on raft-lipid mediated segregation mechanism3.

Here, we propose a model of the flotillin membrane microdomain (Fig. 5e): The membrane encompassed by the flotillin complex is laterally segregated as a basic unit of the microdomain (30 nm in diameter). Together with specifically associated sphingosine and proteins, the flotillin membrane microdomain could exert diverse functions in different cellular processes. Under certain conditions, or dependent on contextual factors, flotillin complexes could cluster to form larger membrane microdomains (over 200 nm in diameter), serving as a platform for signaling or inter-membrane trafficking.

Methods
Cloning, expression and purification of the flotillin complex
The coding DNA sequences for human flotillin-1 (UniProt ID: O75955) and flotillin-2 (UniProt ID: Q14254) were individually cloned into the pMLink vectors at the multiple cloning sites. Flotillin-1 contains a C-terminal 3×FLAG tag, and flotillin-2 has no tag or a C-terminal Strep tag. Using the LINK sequences in pMLink vectors58, after digestion with PacI and SwaI separately, the two genes in different pMLink vectors were cloned into the same pMLink vector via ligation-independent cloning (LIC) method59,60. To co-express both flotillins, the pMLink-flotillin-1 and pMLink-flotillin-2 vectors were combined in this way to generate the pMLink-flotillin-1/2 vector. Mutations and truncations of these genes were introduced individually into the pMLink-flotillin-1 and pMLink-flotillin-2 vectors prior to the combination.

HEK293F cells were cultured in SMM293T-II medium (Sino Biological Inc.) at 37°C under 5% CO2. When the cell density reached 1.5×106 cell/mL, the cells were transfected with the pMLink plasmid containing two flotillins. For one liter of cell culture, 1 mg of plasmid and 4 mg of polyethylenimine (Polysciences) were preincubated in 50 mL of fresh SMM293-TII medium for 30 min. The mixture was then added to the cell culture. After 48 h of incubation, 6 liters of the cell were harvested and resuspended in the prechilled Buffer A (40 mM Tris-HCl pH 7.4, 150 mM NaCl, 0.5 mM Mg(OAc)2, 10% glycerol) supplemented with 1% (v/v) protease inhibitor cocktail (Mei5bio). After sonication, the sample was subjected to centrifugation at 3000×g for 30 min to remove unbroken cells and large debris. Membranes were then pelleted by ultra-centrifugation at 50,000×rpm for 1 h (rotor 70 Ti, Beckman Coulter). All pelleted membranes were solubilized with Buffer B (Buffer A supplemented with 1% (w/v) glyco-diosgenin (GDN, Anatrace)) for 2 h. After centrifugation at 8000×g for 10 min, the supernatant was incubated with anti-FLAG M2 affinity resin (Sigma) at 4°C for 2 h. After rinsing the resin with wash Buffer C (Buffer A supplemented with 0.004% (w/v) GDN), the protein was eluted with Buffer D (Buffer C supplemented with 0.5 mg/ml 3×FLAG peptide) and concentrated to ~100 μL using a 100-kDa-cutoff spin concentrator (Millipore).

Then, the samples were loaded on a 20–50% glycerol density gradient and subjected to ultra-centrifugation at 100,000×g for 14 h (TLS-55 rotor, Beckman Coulter). All fractions of the glycerol gradient were collected manually in separate tubes, with 200 μL per fraction and analyzed with 10% SDS-polyacrylamide gel electrophoresis (SDS-PAGE). Fractions with highest homogeneity, as verified by negative staining electron microscopy, were pooled and dialyzed into Buffer E (40 mM Tris-HCl pH 7.4, 150 mM NaCl, 0.5 mM Mg(OAc)2, 5% glycerol, 0.004% (w/v) GDN). At last, the purified protein samples were concentrated to a desired concentration using a 100-kDa-cutoff spin concentrator (Millipore), and then stored at −80°C.

Glycerol gradient fractionation analysis of crude membrane extracts
To evaluate whether the designed mutations disrupt dome structure formation, we adapted the aforementioned protocol while omitting the affinity purification step. Briefly, HEK293F cells expressing wild-type (WT) or mutant flotillin constructs (individually or in combination) were harvested and resuspended in Buffer A supplemented with 1% (v/v) protease inhibitor cocktail (Mei5bio). Cell lysis was performed using a French press, followed by clarification centrifugation at 3000×g for 30 min. Membrane fractions were subsequently pelleted by ultracentrifugation at 45,000 rpm (70 Ti rotor, Beckman Coulter) for 1 h at 4°C. The membrane pellets were solubilized overnight at 4°C in Buffer F (Buffer A containing 1% (w/v) n-dodecyl-ß-D-maltoside (DDM, Anatrace)), followed by clearance centrifugation at 52,000 rpm (TLA-55 rotor, Beckman Coulter) for 25 min. The resulting supernatant was layered onto a 20–50% glycerol density gradient and subjected to ultra-centrifugation at 52,000 rpm (TLS-55 rotor, Beckman Coulter) for 5 h at 4°C. Gradient fractions (200 μL each) were manually collected and analyzed by SDS-PAGE and western blotting using anti-Flag antibody (Sigma-Aldrich, F3165) for flotillin-1-Flag and anti-Strep antibody (GenScript, A01732) for flotillin-2-Strep.

Stable cell line construction, GPMV preparation and microscopy imaging
For lentivirus packaging, HEK293T cells were plated in 10 cm dishes (4×106 cell/dish) and cultured in DMEM (Gibco), penicillin-streptomycin, and 10% fetal bovine serum (Sigma). A transfection mix was set up as follows: 10 µg transfer vector (pLVX-F1-EGFP, pLVX-F2-mCherry or pLVX-T4-BFP), 15 µg psPAX2 and 5 µg pVSVG in 0.5 mL Opti-MEM were combined with 60 µg PEI in 0.5 mL Opti-MEM (Thermo Fisher) and incubated for 20 min. Cell media was removed, and 9 mL Opti-MEM and 1 mL of transfection mix were added per dish. The transfection media was replaced after 6 h by plating media supplemented with 10% fetal bovine serum, and the supernatant was collected after additional 48 h. 10 mL lentivirus containing supernatant was concentrated to 1 mL using a 100-kDa-cutoff spin concentrator (Millipore) at 1500×g and freshly used for infection after being supplemented with equal volume of complete medium.

Jurkat cells were suspended cultured in RPMI 1640 (Gibco), penicillin-streptomycin, and 10% fetal bovine serum (Sigma) at 37°C under 5% CO2. When the cell density reached 0.25×106 cell/mL, the cells were infected with F1-EGFP and/or F2-mCherry lentivirus. Infected cells were sorted by Astrios EQ (Beckman Coulter) based on the corresponding fluorescence and cultured as normal. L929 cells stably expressing T4-BFP, F1-EGFP and F2-mCherry were also constructed in this way. To adhere Jurkat cells to dishes or cryo-EM grids, 50 µg/ml poly-D-lysine (Thermo Fisher) or 10 µg/ml fibronectin (Yeasen) solutions were added to the culture surface. After incubating 1 h at 37°C, the coating reagents were removed and the dishes were freshly used for Jurkat cell adhering.

GPMVs were isolated and imaged as described previously47,61. Briefly, cell membranes of Jurkat or L929 stable cell lines were labeled with 5 µg/ml fluorescent liquid-disordered phase or liquid-ordered phase marker for 20 min at 37°C. Then, cells were washed in GPMV buffer (10 mM HEPES, 150 mM NaCl, 2 mM CaCl2, pH 7.4) and incubated with GPMV buffer supplemented with 25 mM paraformaldehyde (PFA) and 2 mM dithiothreitol (DTT) (or 2 mM N-ethylmaleimide (NEM) alone) for 1 h at 37°C. The suspension was centrifuged at 100×g for 1 min, to pellet cell debris. The GPMV supernatant was transferred into a microcentrifuge tube by pipetting, and 400 µM deoxycholic acid (DCA) was added to increase phase separation temperature of GPMV membranes62,63. The GPMV suspension was left in the microcentrifuge tube for 20–30 min, and GPMVs would concentrate at the bottom of the tube. The sample was pipetted directly from the bottom of the tube for further microscopy experiments.

For confocal microscopy, cells or GPMVs were cultured or loaded in 35 mm Glass bottom dish (Cellvis), and imaged using Nikon A1R Si+ laser scanning microscope with 60× and 100× objective under oil and lasers exciting at 405 nm, 488 nm, 561 nm and 640 nm. To visualize polarized uropod or “cap” structure of Jurkat cells, fibronectin-coated dishes were used to promote cell adhesion. Polarization was induced by treatment with 100 ng/ml RANTES (MedChemExpress) for 4 h at 37°C/5% CO2 to stimulate migratory morphology. Nuclei were counterstained with Hoechst 33342 prior to live-cell imaging. To view proteins and lipids partitioning on GPMV membranes, the temperature was decreased below 15°C for GPMVs derived with PFA/DTT and below 5°C for the ones derived with NEM. Image analysis was conducted using NIS-Elements Viewer (Nikon) and ImageJ64.

Cryo-EM single particle sample preparation, image collection
The sample homogeneity was examined by negative staining electron microscopy. Four microliters of samples were applied onto copper grids and stained with 2% uranyl acetate. The grids were examined with an FEI Tecnai T20 TEM at 120 kV.

Cryo-EM grids were prepared with GraFutureTM RGO TEM Grid (GraFuture-RGO001A, Shuimu BioSciences Ltd., CN). The grids were glow-discharged for 10 s with a plasma cleaner prior to sample freezing. Three microliters of freshly prepared flotillin complex samples (~0.1 mg/mL) were applied onto the grids mounted in the chamber of an Thermo Fisher Vitrobot Mark IV at 16°C with 100% humidity (adsorption time 1 s, waiting time 60 s, and force −1), and flash-frozen in liquid ethane cooled by liquid nitrogen. Grids were screened on Thermo Fisher Talos Arctica (Gatan K2 Summit detector), operated at 200 kV. Images were collected with a 300 kV Thermo Fisher Titan Krios TEM (Gatan K3 summit detector, with GIF energy filter). The movies were acquired using EPU (Thermo Fisher Scientific) at a dose rate of 8.52 e–/s/Å2 and an exposure time of 4.81 s with 32 frames. The data was collected at the magnification of 64,000× corresponding to a physical pixel size of 1.37 Å/pixel and with defocus ranging from −1.6 to −2.2 μm. Two datasets of movie stacks (19,477 in total) were collected separately.

Cryo-EM single particle analysis (SPA)
The motion correction and electron-dose weighting of the micrographs were performed with MotionCor265 and the contrast transfer function (CTF) parameters were estimated with the program of Gctf66. Particle picking, classification and structure refinement were done with RELION 4.067.

The two datasets underwent separate processing, which is outlined in Supplementary Fig. 2. For the first batch of 6057 movie stacks, the two-dimensional (2D) class averages calculated from manually picked particles were used as the templates for particle autopicking. Then, the autopicked particles were subjected to several rounds of reference-free 2D classification. 2D class averages of the top views of the complex revealed the presence of 44 centrosymmetric dots along the external rim, suggesting a potential 44-mer form of oligomerization. Particles from good classes were combined and subjected to generate the initial model. After several rounds of 3D classification, the resulting classes without apparent structural deformation in each round were joined. Any duplicate particles in the combined data were removed to make the most of the data, as the particles tended to deform easily. These “classification-join-removing” cycles aimed to enrich the good particles as much as possible. Each round of 3D classification was carried out with C1 symmetry imposed. In the final round, the top view of the complex revealed a 44-mer form of oligomerization, which was consistent with the 2D class averages shown in Supplementary Fig. 3c.

With this prior knowledge, the second batch of 13,420 movie stacks was processed using a similar approach, and 3D classifications were performed with C11 symmetry imposed at the beginning. The most apparent deformation of the particles was observed at the bottom rim of the dome, where they changed from circular to elliptical shapes with varying eccentricities. By imposing high rotational symmetry during 3D classification, classes enriched with elliptical-shaped particles were characterized by features that were easier to observe, such as broken features and incorrect sizes of the bottom rim. Consequently, it facilitated the identification of the most homogeneous classes and the removal of deformed particles.

Alignment of particle images with the same chirality during 3D reconstruction is another challenge. When images with both the correct and inverse chirality coexist in the same map, the resulting resolution is low. To address this issue, a good reference map with a definite chirality was required. Based on the medium-quality C11-symmetric 3D density map, a preliminary 44-mer model was manually constructed and then converted to a map. The artificial map was used as the reference to refine the reconstructed map, resulting in a definite chirality and significant improvement in resolution. A final set of particles, containing 25,977 particles from two datasets, were pooled together and subjected to 3D refinement, resulting in a 3.57 Å (Gold-standard FSC 0.143) map with C22 symmetry imposed, a 6.32 Å (Gold-standard FSC 0.143) map with C1 symmetry imposed.

Model building and refinement
Flotillin-1 and flotillin-2 were modeled de novo using Coot68, partially based on the AlphaFold269,70 predicted monomer structures.

A poly-alanine model was built using Coot, and sequence substitution was performed using bulky side-chain residues as makers. Models of SPFH domain were generated by performing rigid body fitting of the AlphaFold2 predicted structures into the density map, followed by manual adjustments using Coot.

The 1/11 map, containing two flotillin-1 and two flotillin-2 subunits, was used for model refinement using real-space refinement (phenix.real_space_refine) in Phenix71. Model validation (Table S1) was calculated by MolProbity72. Figure preparation and structure analysis were performed with UCSF Chimera73.

Cryo-ET sample preparation, cryo-FIB milling, tomographic tilt series collection
The resuspended Jurkat stable cell line described above were loaded onto Quantifoil grids (R2/2, Au 200-mesh grid, Quantifoil Micro Tools, Germany), which were freshly glow discharged with a plasma cleaner before use. Excess liquid was blotted away from the back, and grids were plunge-vitrified with the Thermo Fisher Vitrobot Mark IV system. The vitrified samples were stored in liquid nitrogen until milling.

To prepare lamellae at the region of interest, cryo-FIB milling was performed following a previous protocol74. Briefly, cryo-fluorescence microscopy was performed with Thunder Imager EM Cryo CLEM (Leica) equipped with a 50×/0.9 NA objective to identify regions of interest. Then samples were loaded into the Thermo Fisher Aquilos 2 Cryo-FIB system using a cryo-transfer rod. To reduce the accumulation of electrons on samples and provide extra support while milling, sputtering was first performed on the samples with a 30 mA current for 15 s followed by organometallic platinum (Pt) deposition via GIS for 20 s. The fluorescence image was overlayed with the SEM images to locate the region of interest for milling. A 500 pA current was used for rough milling around the target region, and then the current was reduced gradually to 10 pA for fine milling, until the thickness of the lamellae reached ~200 nm.

The milled lamellae were reloaded into to a Thermo Scientific Titan Krios TEM equipped with a Falcon 4i camera and a Thermo Scientific Selectris X energy filter. Automatic tomographic tilt series acquisition was performed using Tomography 5.12.0 (Thermo Fisher Scientific) dose symmetrically75 from −56° to +56° (with respect to the pre-tilt angle) in 2° increments. 80 sets of images were acquired at a magnification of 42,000× (pixel size 3.00 Å) using dose fractionation mode. The defocus range was set from −3.0 to −5.0 μm. The total electron dose per tilt series was limited to 120 e–/Å2.

Tomogram reconstruction, annotation and rendering
We used the TOMOMAN script for batch tomogram reconstruction76. The collected frames were motion-corrected using MotionCor2 software65. Subsequent reconstruction was performed using IMOD software, where the tilt series were aligned by patch tracking, and then reconstructed using the weighted back projection method77. For visualization, the tomograms were downscaled using a binning factor of 4 and filtered to improve the contrast78.

Membrane segmentation was performed automatically using a CNN based MemBrain-seg software79, followed by manual polishing with Amira 2022.2 (Thermo Fisher Scientific). Actin in the tomograms was localized by the fiber tracking module in Amira (Thermo Fisher Scientific), where a reference cylinder with a radius of 3.5 nm and a length of 40 nm was used for template matching. The results were rendered using UCSF Chimera73.

Template matching and subtomogram averaging (STA)
We used the MATLAB (Mathworks) TOM toolbox80 to perform general preprocessing of the tomograms. Then the SPA structure of the flotillin complex obtained in our work was low-pass filtered as an initial template. Template matching was performed in the tomograms using the template above through PyTom81. In total, 218 flotillin complexes were selected from 20 tomograms. The subtomograms with a box size of 176 pixels3 (1× binned) were extracted using Warp78, then aligned in RELION v3.182, where C11 symmetry was applied. The model of flotillin complex was fitted into the STA map using rigid body docking within UCSF Chimera73. For visualization, the averaged structures were mapped back to the tomograms in their original locations and orientations, using the information from template matching and subtomogram averaging.

Comparative lipidomics analysis
Plasma membrane fractions were isolated following a previous publication83. In brief, HEK293F cells (10 mL) were pelleted and resuspended in 2 mL phosphate-buffered saline (PBS). The cells were lysed by 50 strokes in a Dounce homogenizer. Nuclei and cell debris were removed by centrifugation at 1000×g for 10 min. The resulting supernatant (2 mL) was layered onto 8 mL of 30% Percoll in PBS and centrifuged at 84,000×g for 30 min. The plasma membrane fraction was collected from the membrane band approximately halfway down the gradient.

For lipid extraction, MTBE/methanol extraction method was used. Briefly, purified flotillin protein sample or plasma membrane fraction containing 0.5–2 µg protein was diluted with Buffer E or PBS to a final volume of 100 μL, respectively. The sample was mixed with 300 μL methanol and vortexed. Subsequently, 1 mL of MTBE containing 0.3 µg of each internal standard (LPC(16:0)-D49, LPE(17:0)-D5, PC(16:0-18:1)-D31, PE(16:0-18:1)-D31, PS(16:0-18:1)-D31 from Avanti Research, and TAG (15:0-18:1-15:0)-D7, DAG (15:0-18:1)-D7, Cer (d18:1-22:0)-D7, SM (18:1-18:1)-D9, Cholesterol-D7 from Alta Scientific) was added and vortexed again, followed by adding 250 µL of H2O to facilitate phase separation. The vortexing and resting process was repeated three times to ensure thorough extraction. The mixture was centrifuged at 8000 rpm for 15 min. The upper organic phase was carefully transferred into glass vials and dried under an argon flow in a fume hood. Aspirating precipitated protein in the interphase and the lower phase should be avoided during collection of the upper phase. Ensure the argon flow is gentle and avoid applying heat during the drying process. All steps above were performed at room temperature. The dried samples were stored at −80°C for subsequent LC-MS/MS analysis.

For quantitative analysis of general lipids, LC-MS/MS was conducted using an Orbitrap OE 240 mass spectrometer coupled with an Ultra-High Performance Liquid Chromatography (UHPLC) system (Thermo Fisher Scientific, USA). The liquid chromatography (LC) separation was performed on a Cortecs C18 column (2.1×100 mm, Waters, USA). The LC gradient utilized 60% acetonitrile with 5 mM ammonium acetate as mobile phase A and isopropanol/acetonitrile (v/v 9:1) as mobile phase B. Data-dependent MS/MS acquisition mode was employed, with an MS resolution of 60,000 and an MS/MS resolution of 15,000. The top 10 most intense precursor ions were selected for fragmentation. The mass range for MS acquisition was set to 240–2000 m/z in positive ion mode. Lipid identification was performed using LipidSearch 4.2 software (Thermo Fisher Scientific, USA). Lipids were identified by matching MS/MS spectra with the database in the software. Results with low MS/MS matching scores or inconsistent chromatographic behavior were manually reviewed and excluded to ensure the reliability of lipid identification.

For analysis of cholesterol, LC-MS/MS was conducted using a Q-Exactive HFX mass spectrometer coupled with a UHPLC system (Thermo Fisher Scientific, USA). A 2 µL aliquot of the supernatant was loaded to a Kinetex® Biphenyl column (2.1×150 mm, 2.6 µm, Phenomenex). Samples were eluted using acetonitrile containing 0.1% formic acid as the eluent, with a gradient from 80% to 99% within 6.5 min. The stationary phase was aqueous with 0.1% formic acid. Data acquisition was performed in positive ion mode with a mass range of 300–500 m/z using data-dependent MS/MS. Full scan and fragment spectra were collected at mass resolutions of 60,000 and 15,000, respectively. The source parameters were set as follows: spray voltage, 3200 V; capillary temperature, 320°C; heater temperature, 300°C; sheath gas flow rate, 35 Arb; auxiliary gas flow rate, 10 Arb. Data analysis was performed using Xcalibur 4.4 software (Thermo Fisher Scientific, USA). Cholesterol assignment was confirmed using a chemical standard based on retention time.

The molar amount of each lipid species was determined using corresponding internal lipid standards. For lipids without specific internal standards, calculations were based on similar lipid species: PI and PG were estimated using the PS standard, and SPH was estimated using the Cer standard. For every sample, the molar amounts of all lipid species, including cholesterol, were summed, and each lipid species was normalized to its mole percentage of the total lipid content.

Statistical tests
Statistical analyses were conducted using GraphPad Prism software. To evaluate the differences between two independent groups (purified flotillin protein sample and plasma membrane fraction sample) (Fig. 5d), unpaired two-sample t-tests were applied with Welch’s correction, which does not assume equal variances between groups. Each kind of lipid species was treated as an independent comparison, and multiple t-tests were performed without assuming consistent standard deviations across groups. All statistical tests were two-tailed, assumed a Gaussian distribution, and parametric methods were applied.

Statistics and reproducibility
All experiments were performed with appropriate replicates to ensure statistical validity. Data presented in Figs. 2i, 4a, b and Supplementary Fig. 7b are based on three independent experiments.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "This paper investigates the structural features of flotillin complex in model systems and in Jurkat cells. These complexes are associated with the formation of distinct membrane domains that are implicated in numerous membrane functions including endocytosis, mitochondrial function, and cell signaling. The study uses a suite of techniques including direct experimentation by negative stain electron microscopy, cryo-EM, and tomography, as well as contrast transfer function modeling for 3D reconstruction and de novo structural modeling using Coot (partially based on AlphaFold2).\nBased on this combined experimental and analytical approach, the study demonstrates that the structure of the flotillin complex is formed by the assembly of 22 repeats of alternate flotillin-1 and flotillin-2 proteins, resulting in a dome-like structure with a wide base (~ 30 Å) and a narrow roof region (~ 17 Å). These structures were observed in glycerol fractions, on vesicles, and in cells. Given that this major finding was already reported ~ a year ago with significantly higher resolution data and detailed model building algorithms (Fu and McKinnon, PNAS 2024), it raises major questions about the novelty of this paper. The authors acknowledge the other study, yet they fall short of distinguishing their own work from these prior discoveries. Instead, they list a series of critiques (such as the use of model vesicles from GPMVs, the use of Ca ions, etc.) that hardly affect the extracted structure (which as mentioned above agrees remarkably with the present findings). What is more, the authors ignore addressing the limitations of their own approach, such as the heavy utilization of detergents which could significantly compromise their studies of flotillin localization in liquid-ordered vs liquid-disordered membrane domains. On this point, the paper opens with an extensive discussion on the association of flotillin with raft-like domains only to indicate that their studies showed partitioning of flotillin to the disordered domains in GPMVs. These results seem to be questionable since the GPMVS studies had to be conducted at much lower temperatures (4 degrees) to observe phase separation. This is in stark contrast with other GPMV studies (e.g. Balakrishnan and Kenworthy, JACS 2024) which show Lo-Ld phase separation at room temperature. Is it possible that the introduction of detergent compromised the membrane composition and lateral segregation state? In fact, the data reported with PFA/DTT vs. NEM show major differences in the reported observations (Extended Figure 11). These findings also contradict recent studies reporting enrichment of flotillin in detergent-resistant membrane fractions typically associated with functional membrane microdomains (FMM) that resemble Lo raft-like domains (Ukleja et al., Nature Commun. 2024). The paper goes on to propose a functional mechanism based on the clustering of flotillin complexes observed in a few occurrences. However, there speculation is not founded on tangible evidence. The clustering of proteins on membrane surfaces is quite common, but unless a clear functional consequence is demonstrated it is hardly a convincing argument to state that clustering is somehow involved in function. Unfortunately, there are several such speculative, unsubstantiated statements that are made throughout the manuscript.\nBesides the comments above, the paper is very technique heavy with loose connections between the applied methods, obtained data, and contextualization of the obtained results relative to previous literature. For example, there are numerous citations of raft-related studies (not including flotillin) and an absence of more relevant, foundational, flotillin-focused studies. In addition, it is unclear whether some data are obtained by direct experimental observations or through data modeling. As a case in point, Extended Figure 2b shows cryo-EM images that marginally represent the extracted structure of the flotillin complex (same applies to Extended Figure 1). Also, the caption of Extended Figure 4a and 4c states that the transparent surfaces superimposed with the atomic structures are cyro-EM maps of SPFH1 and SPFH2. How? These structures are smaller than the reported resolution. This is not helped by the fact that several of the images are missing scale bars (e.g. Extended Figure 4 and Extended Figure 1c and 1d). These shortcomings compromise the potential impact and clarity of the manuscript, especially considering the quality of research published in Nature Communications.", "reviewer_2": "This manuscript describes the cryoEM structure of the flotillin1-flotillin2 complex in vitro and by cryoET in situ. Although the actual function of flotillin and related SPFH proteins is still somewhat mysterious, they are important for the generation of membrane domains and potentially endocytic vesicle structures. This work provides a framework for understanding how these proteins assemble and for future studies of their precise cellular interactions and functions. Notably the work does follow from a recent similar structure in PNAS by Fu and MacKinnon. This recent paper is properly cited and discussed in the current manuscript and I believe does not detract from the conclusions or significance. In addition there is another recent preprint in Biorxiv describing in situ structures of the endoplasmic reticulum (ER)-resident Erlin1/2 complex and the mitochondrial prohibitin (PHB1/2) complex by Gao and colleagues (https://doi.org/10.1101/2025.04.21.649849). Again, I don’t feel this detracts from this current study, and in fact highlights that there are a lot of current advancements being made in the field thanks to modern cryoEM developments.\nI thought the paper was really well written and worthy of publication without major changes. I have some minor queries and edits suggested below. For context I have expertise in structural biology of membrane associated protein complexes, but I am not specifically an expert in flotillins or cryoEM data processing.\nQueries.\n1. Line 94. During initial refinement what was the rationale for imposing C22 symmetry? Was this based on the 2D classes, or based on the previous findings of Fu and MacKinnon?\n2. Line 130. It is noted that several designed mutations did not fully prevent the formation of dome structures as assessed by glycerol gradients in Ext. Fig. 7. One thing I noticed if I am interpreting the blots correctly is that flotillin-2 seems to be able to form high molecular weight structures on its own without flotillin-1. Is this correct? If so, are these just aggregates, or have these been looked at by EM to verify if flotillin-2 can assemble as a homo-oligomeric domes?\n3. From line 177. It is stated that only when co-expressed do flotillin-1-GFP and mCherry-flotillin-2 localise to uropods (in Fig. 4 and Ext. Fig. 8). As I am not familiar with these cells or what uropods should look like it wasn’t entirely clear that the individual expressed proteins were mislocalised. For example, to my eye the localisation of flotillin-1-GFP doesn’t look much different on its own compared to the co-expressed and merged images. Could this be clarified in some way, for example by showing enlarged views in Ext Fig. 8 with labels or arrows pointing out the correct structures?\n4. Related to these same experiments and subsequent in situ tomography, could the authors include a brief comment as to what they think is happening with the mCherry-tag at the N-terminus of the flotillin-2 protein? As the C-terminus of the flotillin-1-GFP construct is likely free at the top of the dome structures this seems like it should not affect the structure at all (and from the sub-tomogram densities perhaps the low-threshold density at the top in Fig. 4g represents the diffuse localisation of the 22 copies of GFP?). But where does the N-terminal mCherry tag sit with respect to the membrane and is it possible that the presence of this tag is causing some of the particle heterogeneity that is mentioned on line 197?\nMinor points:\n5. Line 102. “associated to the membrane” should be “associated with the membrane”.\n6. Line 112. “similar to the structure of HflK/C complex” should be “similar to the structure of the HflK/C complex”."}, "author_response": "We sincerely thank both reviewers for their time and constructive feedback on our manuscript. While\nReviewer #1 raised concerns on the novelty and methodological approach of our study, we appreciate these\ncritiques as they have helped us strengthen key aspects of the work. Below, we address their concerns and\nhighlight key revisions:\n1. Novelty and independence of our work: We have clarified the concurrent yet independent nature of our\ndiscovery (preprint in bioRxiv, May 25, 2024) relative to Fu & McKinnon’s work (preprint in bioRxiv,\nMay 9, 2024; accepted by PNAS, May 30, 2024), emphasizing our first-reported in situ structural\nanalysis and our comprehensive mechanistic insights into how flotillin complexes organize membrane\nmicrodomains.\n2. Additional experimental support: New data (Figure 2i, Extended Data Figs. 7b, 8a-c, and Rebuttal\nFigs) now reinforce our findings on flotillin-1/2 oligomerization specificity and regulation.\n3. Methodological clarification: To clarify, we have provided more detailed explanations of our\nmethodological approaches (e.g., GPMV conditions, cryo-EM symmetry determination) both in this\nresponse and in the revised manuscript.\nWe are particularly grateful to Reviewer #2 for recognizing the technical rigor and biological relevance of\nour findings. Below we provide point-by-point responses to all critiques, with corresponding manuscript\nrevisions marked in the updated version.\nReviewer #1 (Remarks to the Author):\nThis paper investigates the structural features of flotillin complex in model systems and in Jurkat cells. These\ncomplexes are associated with the formation of distinct membrane domains that are implicated in numerous\nmembrane functions including endocytosis, mitochondrial function, and cell signaling. The study uses a suite\nof techniques including direct experimentation by negative stain electron microscopy, cryo-EM, and\ntomography, as well as contrast transfer function modeling for 3D reconstruction and de novo structural\nmodeling using Coot (partially based on AlphaFold2).\nBased on this combined experimental and analytical approach, the study demonstrates that the structure of\nthe flotillin complex is formed by the assembly of 22 repeats of alternate flotillin-1 and flotillin-2 proteins,\nresulting in a dome-like structure with a wide base (~ 30 Å) and a narrow roof region (~ 17 Å). These\nstructures were observed in glycerol fractions, on vesicles, and in cells. Given that this major finding was\nalready reported ~ a year ago with significantly higher resolution data and detailed model building\nalgorithms (Fu and McKinnon, PNAS 2024), it raises major questions about the novelty of this paper. The\nauthors acknowledge the other study, yet they fall short of distinguishing their own work from these prior\ndiscoveries.\nWe appreciate the reviewer’s comments and address the concerns below with clarifications:\n1) Concurrency of Findings: Our work was independently developed and first reported on bioRxiv (May\n25, 2024; https://doi.org/10.1101/2024.05.25.595881 ), merely 16 days after the cited PNAS study’s\npreprint submission (Fu & McKinnon, May 9, 2024; https://doi.org/10.1101/2024.05.09.593390 ). This\ntimeline underscores independence and concurrent development of our findings.\n2) Resolution Comparison: The resolution difference between their structure (3.5 Å) and ours (3.57 Å) is\nnegligible, in contrary to the comment of \"significantly higher\".\n3) Unit Correction: We note a typo in the reviewer’s comment: The dome dimensions are ~30 nm (base)\nand ~17 nm (roof), not Å as stated.\n4) Novelty and Functional Insights:\nOur study not only resolves the high-resolution structure of the flotillin complex in purified samples,\nbut also provides mechanistic insights into how flotillin complexes organize membrane microdomains,\nadvancing beyond the work of Fu & McKinnon. Key contributions include:\n(a) Phosphoregulation of complex assembly:\nMutagenesis data reveal that phosphomimetic mutations (Y160E in flotillin-1 and Y163E in flotillin-2)\nabolish flotillin complex assembly. This suggests that Fyn-regulated phosphorylation of these residues\n(Y160 in flotillin-1 and Y163 in flotillin-2) may serve as a physiological trigger for complex\ndisassembly.\n(b) Essential role of hetero-oligomerization:\nWe demonstrate that flotillin-1/2 hetero-oligomerization is functionally indispensable, as evidenced by\nits strict requirement for uropod localization in Jurkat cells upon co-expression.\n(c) Native structural dynamics revealed by cryo-ET:\nOur in situ cryo-electron tomography (cryo-ET) analysis resolves the native architecture of flotillin\ncomplexes in cellular membranes (Jurkat cells), uncovering their dynamic plasticity—including opening\nand resealing behaviors—that cannot be clearly defined in purified systems.\nCollectively, these discoveries have substantially advanced our understanding of the functional principles\ngoverning the formation and regulation of flotillin-mediated membrane microdomain in cellular contexts.\nInstead, they list a series of critiques (such as the use of model vesicles from GPMVs, the use of Ca ions,\netc.) that hardly affect the extracted structure (which as mentioned above agrees remarkably with the present\nfindings).\nWe appreciate the opportunity to clarify our position. The reviewer appears to have misinterpreted our\ndiscussion regarding the study by Fu & McKinnon:\n1) Agreement on the Structure: We have explicitly acknowledged that our flotillin complex structure from\npurified samples aligns with their findings. This consistency validates both studies’ structural models.\n2) Divergence on Functional Interpretations: Our in situ data challenge two specific interpretations in their\nwork:\n(a) Their proposed C-terminal roof-mediated membrane interaction (Fu & McKinnon, Fig. 5C) may\ndepend on two artifactual factors:\ni. Complementary Curvature: The convex membrane of vesicles and concave roof region of\nflotillin complexes exhibit complementary curvatures, but their natural electrostatic repulsion\n(both surfaces being negatively charged) should prevent spontaneous interaction (Rebuttal Fig.\n1).\nii. Ca²⁺ Neutralization: 2 mM Ca²⁺ neutralizes these repulsive charges, thereby bridging these\nsurfaces, creating false-positive binding that may not occur at cellular Ca²⁺ levels (1–10 μM).\nMoreover, the high vesicle concentrations used in their study might have further amplified this\ninteraction. This raises questions about whether the observed membrane association reflects a genuine\nbiological mechanism or a Ca²⁺-driven experimental artifact. (Discussion in our revised manuscripts,\nLines 296–304).\n(b) Their proposed flotillin-mediated membrane curvature generation model relies on the two\nartifactual orientations (inside-out and outside-out) of flotillin complexes in GPMV-derived vesicles. In\ncontrast, our findings support flotillin's intrinsic structural plasticity, indicating that these complexes\nadapt to pre-existing membrane curvature rather than actively generating it. Additionally, while they\nsuggest phosphorylation at Y160 (flotillin-1) and Y163 (flotillin-2) induces curvature while maintaining\ndome structure, we propose these PTMs instead drive flotillin complex disassembly (Figure 2i,\nExtended Figure 7b: iii–v), challenging their mechanistic interpretation (Discussion in our revised\nmanuscripts, Lines 304–309).\nOur goal is not to critique their methodology per se, but to rigorously evaluate how experimental conditions\n(e.g., Ca²⁺ levels, vesicle models) may bias functional interpretations. These distinctions are essential for\nadvancing the field’s understanding of flotillin’s biological role.\nRebuttal Fig. 1. Electrostatic surface potential of the roof region of the flotillin complex. The surface of\nresidues comprising the roof region (Flot1: 345–427 aa; Flot2: 348–428 aa) is colored by electrostatic\npotential (red: negative potential; blue: positive potential).\nWhat is more, the authors ignore addressing the limitations of their own approach, such as the heavy\nutilization of detergents which could significantly compromise their studies of flotillin localization in liquid-\nordered vs liquid-disordered membrane domains.\nOn this point, the paper opens with an extensive discussion on the association of flotillin with raft-like\ndomains only to indicate that their studies showed partitioning of flotillin to the disordered domains in\nGPMVs.\nWe appreciate the opportunity to clarify our experimental rationale. Our study demonstrates that flotillin’s\ndome-like structure spatially compartmentalizes membranes, prompting us to investigate whether this\ncompartmentalization also involves raft-like domain formation (raft lipid partitioning).\nHistorically, flotillins were named for their presence in detergent-resistant membranes (DRMs) (Bickel et al.,\nJ. Biol. Chem. 1997), which were often incorrectly used as an equivalent term of lipid rafts in some\nliteratures. However, DRMs are defined by biochemical fractionation and do not strictly correspond to the\nconceptual definition of lipid raft domains in physiological contexts. Thus, while flotillins have been widely\nused as raft markers, their actual property and association with rafts in native membranes remain an open\nquestion.\nUsing the well-established GPMV method (Sezgin et al., Nat. Protoc. 2012) and comparative lipidomics,\nwe\nobserved that flotillin preferentially localizes to disordered domains and does not co-enrich with the most\nabundant species of raft lipids. These findings suggest that the flotillin-mediated membrane\ncompartmentalization is likely driven by its intrinsic dome-like structural architecture, rather than the self-\nassociation property of raft lipids.\nRegarding the reviewer’s comment on the \"heavy utilization of detergents\" in our GPMV experiments,\nwe\nacknowledge the concern and will provide detailed methodological clarifications in our subsequent response.\nThese results seem to be questionable since the GPMVS studies had to be conducted at much lower\ntemperatures (4 degrees) to observe phase separation. This is in stark contrast with other GPMV studies (e.g.\nBalakrishnan and Kenworthy, JACS 2024) which show Lo-Ld phase separation at room temperature.\nWe appreciate the reviewer’s concern regarding the temperature conditions in our GPMV experiments.\nHowever, the methodology we employed is well-established and carefully optimized to minimize artifacts\nwhile enabling the observation of phase separation.\nAs noted in the foundational GPMV protocol (Sezgin et al., Nat. Protoc. 2012), the use of PFA/DTT—\ncommon vesiculants in other studies—introduces significant perturbations, including (i) nonspecific cross-\nlinking of lipids and proteins by the aldehyde; (ii) cleavage of protein disulfides and thioesters, leading to\ndepalmitoylation; (iii) specific coupling of phosphatidylethanolamines to proteins. These effects artificially\nelevate the miscibility transition temperature by ~15°C, enabling phase separation at room\ntemperature but at the cost of substantial membrane perturbation (Box 2, Sezgin et al., Nat. Protoc.\n2012). The study cited by the reviewer (Balakrishnan & Kenworthy, JACS 2024) further illustrates this point:\ntheir use of lipid peroxidation dramatically enhances demixing but also strongly perturbs membrane\norganization, even displacing canonical raft markers (e.g., GPI-anchored proteins and CTxB) into disordered\ndomains.\nRebuttal Fig. 2. Phase partitioning of flotillins in GPMVs. GPMVs were labeled with different dyes\nfollowing isolation from the Jurkat cells expressing Flot1-EGFP (a), Flot2-mCherry (b), both Flot1-EGFP\nand Flot2-mCherry (c). Phase separation into the Lo and Ld phases was induced by cooling, with FAST DiI\nand FAST DiO dye marking the Ld phase, and CTxB (Cholera Toxin Subunit B (Recombinant), Alexa\nFluor™ 647, Invitrogen) marking the Lo phase.\nBy contrast, we opted for a milder approach using N-ethyl maleimide (NEM), which avoids cross-linking\nartifacts but requires lower temperatures (<5°C) to observe phase coexistence (Box 2, Sezgin et al., Nat.\nProtoc. 2012). To improve phase separation under these conditions, we supplemented with 400 µM\ndeoxycholic acid (DCA), a detergent concentration rigorously validated in prior work (Zhou et al., J. Biol.\nChem. 2013; Levental et al., Biophys. J. 2016). Importantly, our conditions preserved the expected\npartitioning of CTxB into Lo domains (Rebuttal Fig. 2), confirming minimal perturbation compared\nto lipid peroxidation-based methods in the study cited by the reviewer (Balakrishnan & Kenworthy,\nJACS 2024). We acknowledge that GPMV methodologies vary across studies, but our protocol prioritizes\nphysiological relevance by avoiding aggressive perturbations that compromise membrane integrity.\nFinally, our observation of flotillin enrichment in Ld domains aligns with earlier findings (Fig. 6B,\nSengupta et al., Biochim Biophys Acta. 2008), further supporting the validity of our experimental approach.\nIs it possible that the introduction of detergent compromised the membrane composition and lateral\nsegregation state?\nWe acknowledge the reviewer’s valid concern regarding potential detergent-induced perturbations to\nmembrane composition and lateral organization. Indeed, any GPMV formation protocol may inherently\nperturb native membrane organization to some extent through vesiculation-induced component\nreorganization, with protein asymmetry generally preserved but lipid asymmetry partially disrupted (Sezgin,\nBBA Biomembranes 2022).\nWe further note that GPMVs, while imperfect, remain the most experimentally tractable model for studying\nmacroscopic membrane domains. As highlighted by the study cited by the reviewer (Balakrishnan &\nKenworthy, JACS 2024), mammalian plasma membranes are tuned near a miscibility critical point, making\nphysiological nanodomains challenging to resolve in live cells. GPMVs amplify these nanoscopic\nfluctuations into microscopically observable phases below the miscibility transition temperature, providing\nunique insights into underlying principles of membrane organization.\nTo minimize potential artifacts, our experimental conditions were carefully optimized. The 400 µM\ndeoxycholic acid (DCA) concentration used is well below its critical micelle concentration (CMC ≈ 5 mM)\nand has been previously validated to facilitate phase separation without compromising protein partitioning\n(Zhou et al., J. Biol. Chem. 2013; Levental et al., Biophys. J. 2016). Importantly, as established earlier, our\nconditions preserved the expected Lo domain partitioning of CTxB (Rebuttal Fig. 2), indicating the\nmaintenance of raft-like organization.\nIn fact, the data reported with PFA/DTT vs. NEM show major differences in the reported observations\n(Extended Figure 11).\nThe observed differences between PFA/DTT and NEM treatments are indeed expected. As described in the\nExtended Figure 11 legend, PFA/DTT induces depalmitoylation of flotillins due to DTT-mediated cleavage\nof thioester bonds, causing flotillins (which lack transmembrane domains) to dissociate from the membrane\nand localize to the GPMV lumen. In contrast, NEM preserves flotillin palmitoylation and membrane\nassociation while minimizing artifacts, making it the preferred method for studying their partitioning in\nGPMVs.\nThese findings also contradict recent studies reporting enrichment of flotillin in detergent-resistant\nmembrane fractions typically associated with functional membrane microdomains (FMM) that resemble Lo\nraft-like domains (Ukleja et al., Nature Commun. 2024).\nOur findings are consistent with previous reports demonstrating flotillin partitioning into the Ld phase of\nGPMVs (Sengupta et al., Biochim Biophys Acta. 2008).\nThe study cited by the reviewer (Ukleja et al., Nat. Commun. 2024) examines FloA, a bacterial flotillin-like\nprotein from Staphylococcus aureus, which differs fundamentally from mammalian flotillins. FloA contains\nan additional N-terminal transmembrane helix, a divergent SPFH domain, and a much shorter coiled-coil\nregion, making direct comparisons to mammalian flotillins biologically irrelevant. Thus, the reviewer’s\nreference does not contradict our observations, as these proteins belong to the same superfamily but exhibit\ndistinct structural and functional properties.\nThe paper goes on to propose a functional mechanism based on the clustering of flotillin complexes observed\nin a few occurrences. However, there speculation is not founded on tangible evidence. The clustering of\nproteins on membrane surfaces is quite common, but unless a clear functional consequence is demonstrated\nit is hardly a convincing argument to state that clustering is somehow involved in function. Unfortunately,\nthere are several such speculative, unsubstantiated statements that are made throughout the manuscript.\nOur study employs multi-modal imaging approaches to investigate the organization mechanism of flotillin\nmembrane microdomains. We demonstrate that individual flotillin complexes exhibit a dome-like structure\nthat physically compartmentalizes the membrane, forming discrete microdomains (~30 nm in diameter),\nas\nrevealed by both in vitro and in situ structural analyses. These well-defined microdomains then serve as\nfundamental units that can laterally cluster, creating higher-order membrane microdomain organization,\nwhich is mechanistically distinct from generic protein clustering.\nImportantly, flotillin microdomains of similar dimensions have been reported elsewhere (John Abousawan et\nal., J Cell Sci. 2023; Gregory M.I. Redpath et al., Nat. Commun. 2019) and shown to serve as platforms for\nrecruiting client proteins to regulate processes like endocytosis and signaling. Thus, our proposal of\nfunctional clustering is grounded in direct structural evidence and prior literatures, rather than pure\nspeculation.\nBesides the comments above, the paper is very technique heavy with loose connections between the applied\nmethods, obtained data, and contextualization of the obtained results relative to previous literature.\nOur study employs a carefully designed, multi-modal approach where each technique logically builds upon\nprior findings to investigate flotillin membrane microdomain organization. The experimental workflow is as\nfollows:\n1) We first resolved the high-resolution structure of the flotillin complex using cryo-EM SPA, revealing its\ndome-like architecture, which spatially compartmentalizes membranes into ~30 nm microdomains.\n2) Leveraging this distinct structural signature, we then performed in situ cryo-ET to directly visualize\nflotillin complexes in native membranes, confirming their dome-like structure and observed them to\ncluster into higher-order microdomains.\n3) Given flotillin’s historical association with lipid rafts, we tested this hypothesis using GPMV assays and\ncomparative lipidomics. Contrary to expectations, flotillin localized to the Ld (non-raft) phase in\nGPMVs and showed no specific enrichment of the most abundant species of raft lipids except\nsphingosine—results consistent with prior studies (Sengupta et al., Biochim Biophys Acta. 2008; Riento,\nK. et al., PLoS One, 2018).\nFor example, there are numerous citations of raft-related studies (not including flotillin) and an absence of\nmore relevant, foundational, flotillin-focused studies.\nWe appreciate the reviewer’s feedback regarding literature citations and would like to clarify our rationale.\n1) Comprehensive citation of flotillin-focused studies\nOur manuscript extensively cites foundational and functional studies on mammalian flotillins, including:\n Discovery of flotillins (Ref. 3)\n Roles in endocytosis (Ref. 5) and its regulation by Fyn kinase (Ref. 9)\n Roles in signaling (Ref. 7, 8), protein trafficking (Ref. 8, 9), and cytoskeleton remodeling (Ref. 10–\n12)\n Functions in tumorigenesis, invasion, and metastasis (Ref. 14, 15)\n Flotillin-1/-2 coassembly and membrane microdomain formation (Ref. 17).\n2) Bacterial FloA is different from mammalian flotillin\nThe reviewer appears to refer to studies on FloA from Daniel Lopez lab (Ukleja et al., Nat. Commun.,\n2024 (previously cited by the reviewer); Esther Garcı´a-Ferna´ ndez et al., Cell, 2017; Gudrun Koch et\nal., Cell Chem. Biol., 2017), which is a bacterial flotillin homolog from Staphylococcus aureus. FloA is\nfunctionally distinct from mammalian flotillins. Our manuscript focuses on literatures relevant to\nmammalian flotillin biology.\n3) DRM is not equal to lipid rafts\nWhile flotillins are frequently described as raft markers due to their presence in detergent-resistant\nmembranes (DRMs), it is important to note that DRMs are experimentally defined by biochemical\nfractionation that do not necessarily correlate with physiologically relevant raft domains. Our GPMV\nanalysis and comparative lipidomics data demonstrate that flotillin-mediated microdomains are not\nequivalent to lipid rafts. To our knowledge, only Sengupta et al. (BBA, 2008) has used GPMV to study\nflotillin partitioning in coexisting Lo-Ld phases, with their results consistent with ours.\nIn addition, it is unclear whether some data are obtained by direct experimental observations or through data\nmodeling. As a case in point, Extended Figure 2b shows cryo-EM images that marginally represent the\nextracted structure of the flotillin complex (same applies to Extended Figure 1).\nAll structural data presented in this study were obtained through direct experimental observations. The high-\nresolution density map of the flotillin complex was determined by cryo-EM single-particle analysis (SPA),\nand its in situ architecture and distribution were visualized using cryo-ET within the native cellular context.\nAs for data modeling, we're not sure what the reviewer meant by “data modeling”. We built the atomic model\nbased on the high-resolution cryo-EM density map, partially with the assistance of AlphaFold2.\nExtended Figure 2b displays a raw cryo-EM image of purified flotillin sample, where individual complexes\ncan be clearly distinguished and show similar orientations to those depicted in Extended Figure 2c. Similarly,\nExtended Figure 1b–d presents negative-stain EM images of the complex.\nAlso, the caption of Extended Figure 4a and 4c states that the transparent surfaces superimposed with the\natomic structures are cyro-EM maps of SPFH1 and SPFH2. How? These structures are smaller than the\nreported resolution.\nThe cryo-EM maps of SPFH1 and SPFH2 domains were determined at ~4.5 Å resolution, which, while\ninsufficient for precise side-chain modeling, provides reliable backbone tracing. These maps showed strong\nagreement with the AlphaFold2-predicted structures of individual flotillin monomers. As detailed in the\nMethods section, the SPFH domain models were generated by rigid-body fitting of the AlphaFold2-predicted\nstructures into the cryo-EM density, followed by manual refinement in Coot.\nTo reflect the fact that this region was not resolved at atomic resolution, we have revised Extended Data\nFigure 4a-d by removing the distance labelling. Nevertheless, based on (1) the reliable backbone tracing,\n(2)\nstructural conservation of SPFH1 domains across the SPFH protein family, and (3) consistency with both our\ndata and the flotillin complex structure reported by Fu & McKinnon, we propose potential interactions\nbetween the loop regions of adjacent SPFH1 domains in Flot1 and Flot2. We have accordingly modified the\ntext to reflect this interpretation more precisely.\nThis is not helped by the fact that several of the images are missing scale bars (e.g. Extended Figure 4 and\nExtended Figure 1c and 1d).\nWe appreciate the reviewer's attention to details. Regarding Extended Figure 4, since it depicts a molecular\nmodel rather than experimental microscopy data, scale bars are not applicable. For Extended Figure 1c and\n1d, the scale bars are indeed present and positioned in the bottom right corner of each respective panel.\nThese shortcomings compromise the potential impact and clarity of the manuscript, especially considering\nthe quality of research published in Nature Communications.\nHaving addressed the reviewer’s concerns point-by-point in our previous responses, we note that certain key\naspects of our study appear to have been misunderstood by the reviewer—particularly regarding (1) our\nGPMV methodology, (2) the distinctions between bacterial FloA and mammalian flotillin, and (3) the\ndifferences between DRM and lipid rafts. Most critically, the reviewer incorrectly assumes that our major\nfindings were preempted by Fu & McKinnon (PNAS 2024). These misunderstandings have unfortunately\ncast doubt on the novelty and scientific rigor of our work. We hope our detailed clarifications have resolved\nthese concerns.\nWe emphasize that our work was posted on bioRxiv concurrently with Fu & McKinnon’s work, achieves\ncomparable resolution, and importantly provides comprehensive mechanistic insights into how flotillin\ncomplexes organize membrane microdomains. Our study presents the first in situ structural analysis of\nflotillin complexes, and reveals their intrinsic structural plasticity and dynamic opening/resealing\nmechanisms for partner protein encapsulation. And we proposed a novel paradigm wherein flotillin\ncomplexes autonomously organize membrane microdomains through their distinctive dome-like architecture.\nThese advances underscore the novelty and significance of our work in elucidating the molecular\nmechanisms of flotillin complexes in organizing membrane microdomains.\nReviewer #2 (Remarks to the Author):\nThis manuscript describes the cryoEM structure of the flotillin1-flotillin2 complex in vitro and by cryoET in\nsitu. Although the actual function of flotillin and related SPFH proteins is still somewhat mysterious, they are\nimportant for the generation of membrane domains and potentially endocytic vesicle structures. This work\nprovides a framework for understanding how these proteins assemble and for future studies of their precise\ncellular interactions and functions. Notably the work does follow from a recent similar structure in PNAS by\nFu and MacKinnon. This recent paper is properly cited and discussed in the current manuscript and I believe\ndoes not detract from the conclusions or significance. In addition there is another recent preprint in Biorxiv\ndescribing in situ structures of the endoplasmic reticulum (ER)-resident Erlin1/2 complex and the\nmitochondrial prohibitin (PHB1/2) complex by Gao and colleagues\n(https://doi.org/10.1101/2025.04.21.649849). Again, I don’t feel this detracts from this current study, and in\nfact highlights that there are a lot of current advancements being made in the field thanks to modern cryoEM\ndevelopments.\nI thought the paper was really well written and worthy of publication without major changes. I have some\nminor queries and edits suggested below. For context I have expertise in structural biology of membrane\nassociated protein complexes, but I am not specifically an expert in flotillins or cryoEM data processing.\nWe thank the reviewer for the positive feedback. We hope that this updated version of the manuscript,\nespecially our improved gradient separation of all WT and mutant flotillins and properly labelled images will\naddress these concerns.\nQueries.\n1. Line 94. During initial refinement what was the rationale for imposing C22 symmetry? Was this based on\nthe 2D classes, or based on the previous findings of Fu and MacKinnon?\nWe thank the reviewer for raising this important point. To clarify, our work and the study by Fu and\nMacKinnon were developed independently and shared publicly within a month (our preprint: May 25, 2024.\nhttps://doi.org/10.1101/2024.05.25.595881 ; Fu and MacKinnon’s preprint: May 9, 2024.\nhttps://doi.org/10.1101/2024.05.09.593390). Thus, our and their conclusions were reached concurrently and\nindependently, without prior influence from each other.\nRegarding the imposition of C22 symmetry, our decision was based on two key observations:\n1) 2D class analysis (Extended Data Fig. 2c, bottom-left panel) revealed a ring-like arrangement of 44\ndiscernible density dots.\n2) 3D classification with C1 symmetry further confirmed this arrangement, showing 44 helices at the roof\nregion of the dome-like structure (Extended Data Fig. 2f, blue density map in Dataset 1).\nWe systematically tested multiple symmetries (C4, C11, C22, C44 and others) and found that C22 yielded\nthe highest resolution, and most importantly, allowed an unambiguous assignment of flotillin-1 and flotillin-2\nwithin the complex. The computational details of this refinement process are provided in the Methods section\n(Lines 535–544).\n2. Line 130. It is noted that several designed mutations did not fully prevent the formation of dome structures\nas assessed by glycerol gradients in Ext. Fig. 7. One thing I noticed if I am interpreting the blots correctly is\nthat flotillin-2 seems to be able to form high molecular weight structures on its own without flotillin-1. Is this\ncorrect? If so, are these just aggregates, or have these been looked at by EM to verify if flotillin-2 can\nassemble as a homo-oligomeric domes?\nWe thank the reviewer for their insightful observation regarding flotillin behavior in glycerol gradient assays.\nTo provide clearer characterization of flotillin distribution across low/high molecular weight (MW) fractions,\nwe repeated gradient separations for all WT and mutant constructs under more stringent conditions\n(including a prior ultracentrifugation to remove potential large membrane fragments carrying heterogeneous\naggregates) (Lines 453–467). The updated results are presented in Figure 2i and a revised Extended Data\nFigure 7, replacing the previous version.\nOur optimized experimental conditions reveal that while flotillin-1 (but not flotillin-2) can form some high-\nMW species when expressed alone (lanes 7-9 in revised Extended Data Fig. 7b: i), multiple lines of evidence\ndemonstrate these do not represent functional homo-oligomeric domes:\n1) EM characterization: Anti-Flag affinity purification followed by negative-stain EM shows that while\nflotillin-1 appears in both low- and high-MW fractions in crude membrane extracts (Rebuttal Fig. 3a),\nthe purified flotillin-1 protein predominantly localizes to low-MW fractions (Rebuttal Fig. 3b).\nImportantly, EM analysis of high-MW fractions reveals only amorphous aggregates without any dome-\nlike architecture (Rebuttal Fig. 3c).\n2) Endogenous interactions: The observed high-MW species may reflect heterocomplex formation with\nendogenous flotillins rather than true homo-oligomers. Specifically: (a) flotillin-1-Flag could associate\nwith endogenous flotillin-2 (Extended Data Fig. 7b:i, lanes 7-9), and (b) trace amounts of flotillin-2-\nStrep in high-MW fractions (Extended Data Fig. 7b:ii, lane 8) might complex with endogenous flotillin-\n1.\n3) Structural constraints: Our cryo-EM structure demonstrates that the native dome architecture requires\nprecise geometric arrangements (Rebuttal Fig. 4). The dome roof features precisely 22 flotillin-1 C-\nterminal domains (CTDs) arranged in a C22-symmetric ring layered atop 44 CC2 domains, representing\nthe maximal capacity for flotillin-1 incorporation and leaving no space for additional CTDs from\nflotillin-1, as the binding interfaces are already fully occupied. While flotillin-1 may form alternative\nhigh-MW assemblies through self-association, these would necessarily adopt non-native configurations\nthat fail to meet the stringent geometric requirements for proper dome formation, resulting in non-\nfunctional aggregates rather than the biologically relevant architecture.\n4) Evolutionary conservation: Flotillin paralogs exhibit mutual stabilization, as demonstrated by multiple\nstudies showing that the absence of one paralog leads to degradation of the other (Hoehne et al., Mol.\nCell Neurosci. 2005; Solis et al., Biochem. J. 2007; Jang et al., J. Cell Sci. 2015). This co-dependence\nunderscores the biological requirement for both flotillin-1 and flotillin-2 in forming functional\ncomplexes.\nIn conclusion, while flotillin-1 may form some high-MW species, the integrated evidence confirms that\nnative-like dome assembly strictly requires both flotillin paralogs.\nRebuttal Fig. 3. Analysis of flotillin-1 homo-oligomeric assemblies. a, Glycerol gradient fractionation\nprofile of solubilized crude membrane extracts from HEK293F cells overexpressing flotillin-1-Flag. b,\nGradient profile of anti-Flag affinity-purified flotillin-1 from the same membrane preparation shown in (a).\nFractionation was performed as described in Methods (Lines 453–467). c, Representative negative-stain EM\nimage of fraction 7 from (b). Scale bar, 100 nm.\nRebuttal Fig. 4. Top view of the flotillin complex. The model is rendered as helical tubes, with flotillin-1\nsubunits colored cyan and flotillin-2 subunits colored yellow.\n3. From line 177. It is stated that only when co-expressed do flotillin-1-GFP and mCherry-flotillin-2 localise\nto uropods (in Fig. 4 and Ext. Fig. 8). As I am not familiar with these cells or what uropods should look like\nit wasn’t entirely clear that the individual expressed proteins were mislocalised. For example, to my eye the\nlocalisation of flotillin-1-GFP doesn’t look much different on its own compared to the co-expressed and\nmerged images. Could this be clarified in some way, for example by showing enlarged views in Ext Fig. 8\nwith labels or arrows pointing out the correct structures?\nWe thank the reviewer for this suggestion and sincerely apologize for the lack of clarity in our original\npresentation of the flotillin localization data.\nTo address this concern, we have provided representative images that clearly demonstrate the difference\nbetween proper uropod localization (which only occurs upon co-expression of both proteins) and the\nmislocalization observed when either protein is expressed alone. In the revised Extended Data Fig. 8a–c, we\nhave included enlarged views with clear annotations and dashed lines to highlight the uropod structures\nwhere the co-expressed proteins localize, as well as the uniform plasma membrane distribution seen with\nindividually expressed constructs.\n4. Related to these same experiments and subsequent in situ tomography, could the authors include a brief\ncomment as to what they think is happening with the mCherry-tag at the N-terminus of the flotillin-2\nprotein? As the C-terminus of the flotillin-1-GFP construct is likely free at the top of the dome structures this\nseems like it should not affect the structure at all (and from the sub-tomogram densities perhaps the low-\nthreshold density at the top in Fig. 4g represents the diffuse localisation of the 22 copies of GFP?). But where\ndoes the N-terminal mCherry tag sit with respect to the membrane and is it possible that the presence of this\ntag is causing some of the particle heterogeneity that is mentioned on line 197?\nThank you for pointing this out. We sincerely apologize for not clearly stating the tag localization in the\noriginal manuscript. Both the GFP-tag (flotillin-1) and mCherry-tag (flotillin-2) are fused to the C-terminus\nof their respective proteins. These tags remain at the top of the dome structures, likely accounting for the\nlow-threshold density observed in Fig. 4g. Importantly, the particle heterogeneity observed in our in situ\ntomograms (Extended Data Fig. 9a–b, 10a)—primarily localized to the wall and base regions of the dome—\nis not attributable to the mCherry tag, as its C-terminal position does not interfere with the assembly of the\nroof region.\nRegarding the reviewer’s specific question about N-terminal mCherry tagging of flotillin-2, we agree that\nthis would disrupt membrane association. Flotillin-2 lacks a transmembrane domain and instead depends on\nN-terminal myristoylation (at G2) followed by palmitoylation (at C4, C19 and C20) for stable membrane\nbinding. An N-terminal mCherry tag would: (1) Prevent Met1 cleavage, blocking the exposure of G2 for\nmyristoylation; (2) Consequently abolish subsequent palmitoylations (akin to the G2A mutant (Carolin\nNEUMANN-GIESEN et al., Biochem. J. 2004)), severely impairing membrane localization and dome\nassembly.\nTo validate flotillin-2 membrane targeting, we compared Flot2WT and Flot2G2A (both co-expressed with\nFlot1WT) using glycerol gradient assays (Rebuttal Fig. 5). Flot2WT localized to membrane fractions (S2) and\nhigh-MW fractions (#7-9), while Flot2G2A shifted to soluble fractions (S1) with only trace amounts in low-\nMW fractions (#2-4). These results demonstrate that intact Gly2 myristoylation is essential for flotillin-2\nmembrane targeting and dome complex formation. The N-terminal mCherry tag would block Met1 cleavage\nand Gly2 myristoylation, disrupting Flot2 membrane association.\nRebuttal Fig. 5. Analysis of dome structure formation in wild-type and G2A mutant flotillin-2. Glycerol\ngradient fractionation was performed as described in Methods (Lines 453–467). Whole cell lysate (WCL)\nwas obtained as the supernatant after initial clarification centrifugation following French press lysis. S1\nfraction represents the supernatant after pelleting the membrane fraction by ultracentrifugation, primarily\ncontaining soluble proteins. S2 fraction corresponds to the solubilized membrane fraction, obtained after\npelleting unsolubilized membranes via clearance ultracentrifugation, and served as the input for glycerol\ngradient assays.\nMinor points:\n5. Line 102. “associated to the membrane” should be “associated with the membrane”.\nThis has been corrected. Thank you for pointing it out.\n6. Line 112. “similar to the structure of HflK/C complex” should be “similar to the structure of the HflK/C\ncomplex”.\nThis has been corrected. Thank you for pointing it out."}, {"round": 2, "reviewer_comments": {"reviewer_1": "The authors have addressed most of the concerns raised about the previous version. As a result, the paper is significantly improved. I particularly appreciate the author’s clarification regarding the novelty of this study and the additional measurements that validate their findings. In fact, it would be great if more of the clarifications provided in the response were integrated in the main manuscript or provided as further supporting evidence in the SI (e.g. response figure 2). This said, there are still some salient points that need to be addressed to further support the statements made and to improve the readability of the paper. Please find the recommendations below for further details.\n• The introduction defines microdomains and lipid rafts as synonymous structures. While it is understood that these terminologies are used interchangeably in some fields, it is important to make a clear distinction here since this is related to a core finding of this study. As the authors stated, lipid rafts are small domains in the range of 10’s to 100’s of nm. However, microdomains are typically in the micron-scale regime (like those investigated in GPMVs). The authors should clearly define both terms and use them accurately.\n• The paper introduces flotillin as lipid raft biomarkers (lines 42-43, 59) only to demonstrate later that the mammalian homologs studied here preferentially associate with the Lo phase of the lipid membrane. To avoid confusion, the authors are advised to provide a clearer discussion of this and briefly summarize their findings at the end of the introduction to help set the stage. This will significantly elevate the impact of the results and will help contextualize them compared to current knowledge. This would be also a great opportunity to distinguish between the behavior of mammalian flotillin and non-mammalian flotillin-like structures.\n• Since the structure of flotillin has been already published, please provide a reference earlier in the paper. In fact, the agreement between the current findings and earlier results from Fu & MacKinnon provides further support for this paper. This will also help readers in distinguishing the present in-situ study from the earlier work performed on plasma membrane vesicles.\n• Figure 4 is a key figure in this paper. The use of tomography for 3D reconstruction of in situ flotillin complexes is unique. But further information is needed to support the 3D rendering shown in panel d. For example, only one flotillin complex (#1) can be seen in panel c. This could be due to the fact that only one tomographic slice is shown. If applicable, the authors should show the rest of the slices in the SI and clearly label the location of the structures shown in panel e. If not, they should thoroughly explain how the 3D rendition was obtained from this single EM image.\n• Similarly, Figure 5 is the basis of another key finding in this paper, i.e. the clustering of flotillin complexes. However, the 3D rendering shown in the right panels of Figure 5a and 5b are not fully supported by the EM images shown in the left panels. For instance, only one trapezoidal flotillin complex can be clearly identified within the box in left panel a. How were the rest of the complexes identified? On this point, please note that the boundaries of the box in Figure 5a do not match across the left and right panels, which is a cause of confusion. This should be fixed.\n• The same comment applied to panel b. It is almost impossible to identify the flotillin complexes on the vesicle (purple) and plasma membrane (yellow) from the presented EM image. If further images were taken on different tomographic slices, these should be shared in the SI. Since this is used as a demonstration of how flotillin clustering is involved in exosome formation (lines 225-228), it is crucial that the statement is accurately supported by data.\n• Assuming there are additional data to support flotillin clustering, it is important to revisit the discussion presented in lines 255-260. This clustering could be a mere outcome of statistical distribution of flotillin complexes – the EM images also show isolated complexes. Therefore, it should be made clear that the model presented in Figure 5 and in the text is a hypothesis rather than a fact. For example, please replace “we present a new model” by “we propose a new model”. Can you also provide a clear rationale for how this model informs the flotillin microdomain structures, given that flotillin complexes were predominantly associated with the Lo phase of GPMV membranes?\n• Finally, I recommend that the authors consider a more objective discussion in lines 294-311. For example, it is advisable to avoid statements like “our in situ studies raised questions about certain aspects of their conclusions” or “this discrepancy may arise from the non-physiological conditions in their study”. Instead, please consider the following: “our in situ studies reveal different behavior within cellular systems” or “these differences can be rationalized by different conditions of the two studies”. I further recommend that the authors rewrite the last sentence of this paragraph (lines 309-311) to read: “These different interpretations highlight importance of in situ studies in capturing the native behavior of flotillin complexes”.", "reviewer_2": "I have read the Authors responses, and examined the updated manuscript and I am happy that they have addressed all of my concerns."}, "author_response": "We sincerely thank Reviewer #1 for his/her thorough evaluation and constructive feedbacks on our revised\nmanuscript.\nIn the revision, we have improved the clarity, terminology, and data presentation of the manuscript—\nparticularly through the addition of supplementary tomographic movies (Movies S1–S3) that provide a direct\nvisualization of the tomographic datasets underlying Figures 4–5.\nReviewer #1 (Remarks to the Author):\nThe authors have addressed most of the concerns raised about the previous version. As a result, the paper is\nsignificantly improved. I particularly appreciate the author’s clarification regarding the novelty of this study\nand the additional measurements that validate their findings. In fact, it would be great if more of the\nclarifications provided in the response were integrated in the main manuscript or provided as further\nsupporting evidence in the SI (e.g. response figure 2). This said, there are still some salient points that need\nto be addressed to further support the statements made and to improve the readability of the paper. Please\nfind the recommendations below for further details.\n• The introduction defines microdomains and lipid rafts as synonymous structures. While it is understood that these\nterminologies are used interchangeably in some fields, it is important to make a clear distinction here since this is related to\na core finding of this study. As the authors stated, lipid rafts are small domains in the range of 10’s to 100’s of nm. However,\nmicrodomains are typically in the micron-scale regime (like those investigated in GPMVs). The authors should clearly define\nboth terms and use them accurately.\nWe thank the reviewer for this insightful comment. We have revised the manuscript, and ensured consistent\nterminology throughout. Additionally, we have incorporated Response Figure 2 into the Extended Data\nFigure 11 as one piece of supporting evidence. These modifications have improved the clarity of our\nmanuscript.\n• The paper introduces flotillin as lipid raft biomarkers (lines 42-43, 59) only to demonstrate later that the mammalian\nhomologs studied here preferentially associate with the Lo phase of the lipid membrane. To avoid confusion, the authors are\nadvised to provide a clearer discussion of this and briefly summarize their findings at the end of the introduction to help set\nthe stage. This will significantly elevate the impact of the results and will help contextualize them compared to current\nknowledge. This would be also a great opportunity to distinguish between the behavior of mammalian flotillin and non-\nmammalian flotillin-like structures.\nWe thank the reviewer for their suggestion. To address this concern, we now introduce flotillins as lipid raft\nbiomarkers specifically within the context of the GPMV assay, where we provide a clearer discussion and\nsummary of our findings (lines 252–278). This adjustment enhances the logical flow of the narrative and\nbetter contextualizes our results relative to current knowledge. Additionally, we have distinguished between\nmammalian flotillin and non-mammalian flotillin-like structures to eliminate potential ambiguity (lines 68–\n73). These modifications improve manuscript clarity.\n• Since the structure of flotillin has been already published, please provide a reference earlier in the paper. In fact, the\nagreement between the current findings and earlier results from Fu & MacKinnon provides further support for this paper. This\nwill also help readers in distinguishing the present in-situ study from the earlier work performed on plasma membrane\nvesicles.\nWe appreciate this constructive suggestion. We have now incorporated the reference to Fu & MacKinnon's\nwork earlier in the manuscript (lines 120–123). The revised text now explicitly delineates the key distinctions\nbetween our findings and prior research (lines 288–293, 326–342), enhancing clarity for readers. This\nmodification better positions our study within the existing literature while highlighting its novel\ncontributions.\n• Figure 4 is a key figure in this paper. The use of tomography for 3D reconstruction of in situ flotillin complexes is unique.\nBut further information is needed to support the 3D rendering shown in panel d. For example, only one flotillin complex (#1)\ncan be seen in panel c. This could be due to the fact that only one tomographic slice is shown. If applicable, the authors\nshould show the rest of the slices in the SI and clearly label the location of the structures shown in panel e. If not, they should\nthoroughly explain how the 3D rendition was obtained from this single EM image.\nWe acknowledge the reviewer's concern regarding Figure 4. Indeed, only one flotillin complex (#1) is visible\nin panel c because this represents just a single tomographic slice from the complete dataset. To address this\npoint comprehensively, we have prepared a supplementary movie (Movie S1) that sequentially displays all\ntomographic slices of this tomogram, with the 3D reconstruction shown in panel d superimposed for\nreference. This movie clearly labels the spatial positions of all structures featured in panel e. We have also\nincluded more information on the procedure to get such 3D rendering in the Methods session.\n• Similarly, Figure 5 is the basis of another key finding in this paper, i.e. the clustering of flotillin complexes. However, the\n3D rendering shown in the right panels of Figure 5a and 5b are not fully supported by the EM images shown in the left panels.\nFor instance, only one trapezoidal flotillin complex can be clearly identified within the box in left panel a. How were the rest\nof the complexes identified? On this point, please note that the boundaries of the box in Figure 5a do not match across the\nleft and right panels, which is a cause of confusion. This should be fixed.\nWe appreciate the reviewer's careful examination of Figure 5. Similar to Figure 4c–d, we have prepared a\nsupplementary movie (Movie S2) to facilitate visualization of the tomographic dataset. Regarding the box\nboundary mismatch between panels, we have corrected a minor scaling discrepancy in the right panel.\nWe would like to clarify that the apparent difference in the number of visible flotillin complexes between left\nand right panels stems from their fundamentally different representations: the left panel displays a single 2D\ntomographic slice, showing only complexes present in that specific plane, while the right panel presents the\nfull 3D reconstruction containing complexes from all slices. This explains why fewer complexes are visible\nin the 2D view.\nAll flotillin complexes have been carefully annotated in Movie S2, which provides a comprehensive\nvisualization of their spatial distribution throughout the tomogram. This supplemental material would\naddress the reviewer's concerns about flotillin complex identification and localization.\n• The same comment applied to panel b. It is almost impossible to identify the flotillin complexes on the vesicle (purple) and\nplasma membrane (yellow) from the presented EM image. If further images were taken on different tomographic slices, these\nshould be shared in the SI. Since this is used as a demonstration of how flotillin clustering is involved in exosome formation\n(lines 225-228), it is crucial that the statement is accurately supported by data.\nWe thank the reviewer for the suggestion. Similarly, we have prepared a supplementary movie (Movie S3).\nThis movie clearly annotates the spatial distribution of flotillin complexes on both the vesicle (purple) and\nplasma membrane (yellow) across all tomographic slices. This supplemental material would address the\nreviewer's concerns regarding data representation and interpretation.\n• Assuming there are additional data to support flotillin clustering, it is important to revisit the discussion presented in lines\n255-260. This clustering could be a mere outcome of statistical distribution of flotillin complexes – the EM images also show\nisolated complexes. Therefore, it should be made clear that the model presented in Figure 5 and in the text is a hypothesis\nrather than a fact. For example, please replace “we present a new model” by “we propose a new model”. Can you also\nprovide a clear rationale for how this model informs the flotillin microdomain structures, given that flotillin complexes were\npredominantly associated with the Lo phase of GPMV membranes?\nWe appreciate the reviewer's insightful comment regarding the interpretation of flotillin clustering. We have\nrevised the text to more carefully present our model as a hypothesis.\nConcerning the structural basis of flotillin microdomains, our observations suggest that the intrinsic dome-\nlike architecture of individual flotillin complex (~30 nm in diameter) mediates membrane segregation,\neffectively acting as a physical container that compartmentalizes the membrane into nanoscale domains . Our\nin situ cryo-ET data (Movies S2–S3) further reveal that these complexes could cluster into larger (~200 nm)\nmicrodomains on vesicles and the plasma membrane.\nTo explore the mechanistic basis, we considered whether raft domains (typically 10–200 nm) might induce\nclustering of flotillin cages. While flotillins are widely used as raft markers, their actual association with rafts\nin native membranes remains unclear. Given the technical challenges in detecting nanoscopic rafts in living\nmembranes, we employed GPMVs—an established model for studying membrane phase behavior (Sezgin et\nal., Nat. Protoc. 2012; Balakrishnan & Kenworthy, JACS 2024). Surprisingly, flotillins preferentially\nlocalized to the Ld phase rather than the Lo phase in GPMVs, suggesting that lipid-driven partition of the Lo\nphase is not the cause for the observed clustering of flotillins in Figure 5c and Extended Data Figure 11.\nOf course, we are also aware that GPMVs are merely a model system for studying plasma membrane phase\nseparation. The vesiculation process may introduce artifacts such as disrupting native membrane leaflet\nasymmetry, whose impact on phase separation remains unclear, and altering dynamic non-equilibrium states\nessential to living cell membranes. While GPMVs provide insights, they cannot conclusively determine raft\norganization or molecular domain preferences in living cells (Sezgin et al., Nat. Protoc. 2012).\nWe speculate this clustering of flotillin likely depends on contextual factors including local lipid composition\nand interactions with other membrane/non-membrane components (e.g., cytoskeleton).\n• Finally, I recommend that the authors consider a more objective discussion in lines 294-311. For example, it is advisable\nto avoid statements like “our in situ studies raised questions about certain aspects of their conclusions” or “this discrepancy\nmay arise from the non-physiological conditions in their study”. Instead, please consider the following: “our in situ studies\nreveal different behavior within cellular systems” or “these differences can be rationalized by different conditions of the two\nstudies”. I further recommend that the authors rewrite the last sentence of this paragraph (lines 309-311) to read: “These\ndifferent interpretations highlight importance of in situ studies in capturing the native behavior of flotillin complexes”.\nWe have revised the discussion to adopt a more objective tone as suggested. We appreciate this constructive\nsuggestion."}, {"round": 3, "reviewer_comments": {"reviewer_1": "The authors have satisfactorily addressed the comments and concerns related to the previous version of the manuscript. The paper reads significantly better now and provides readers with a much more balanced discussion and contextual significance. I particularly appreciate the revisions made to the introduction, data interpretation, and comparison to previous literature.\nIn addition, the new videos compiled from the tomographic slices now provide the evidence needed for the 3D reconstruction of flotillin localization and distribution in different membrane types within the studied cells. For further clarity, I strongly recommend that a reference to the movies is provided in the legends of Figures 4 and 5.", "reviewer_2": "I believe the authors have answered my queries in the previous review. From their responses to Reviewer #1 I also believe they have reasonably answered those queries as well."}, "author_response": "We are grateful to the reviewers for their thorough evaluation and constructive feedback, which have\nsignificantly enhanced the quality of our work.\nReviewer #1 (Remarks to the Author):\nThe authors have satisfactorily addressed the comments and concerns related to the previous version of the\nmanuscript. The paper reads significantly better now and provides readers with a much more balanced\ndiscussion and contextual significance. I particularly appreciate the revisions made to the introduction, data\ninterpretation, and comparison to previous literature.\nIn addition, the new videos compiled from the tomographic slices now provide the evidence needed for the\n3D reconstruction of flotillin localization and distribution in different membrane types within the studied\ncells. For further clarity, I strongly recommend that a reference to the movies is provided in the legends of\nFigures 4 and 5.\nWe thank the reviewer for their comment. In this revision, we have followed the suggestions by adding\nreferences to the corresponding supplementary tomographic movies (Supplementary Movies 1–3) in the\nlegends of Figures 4 and 5. These modifications have improved the clarity of our manuscript.\nReviewer #2 (Remarks to the Author):\nI believe the authors have answered my queries in the previous review. From their responses to Reviewer #1\nI also believe they have reasonably answered those queries as well.\nThank you for your thoughtful review and for acknowledging our responses to the previous comments. We\nsincerely appreciate your time and constructive feedback."}]

Decision letter