2026 年 51 巻 1 号 p. 81-92
Podocytes are terminally differentiated renal epithelial cells that play a crucial role in kidney filtration. Given this essential function, podocyte dysfunction results in kidney diseases known as podocytopathies. Previous studies have demonstrated that maintaining the activation–deactivation balance of mechanistic target of rapamycin complex 1 (mTORC1) is vital for podocyte function. Podocyte-specific knockout (KO) mouse models revealed that abnormal mTORC1 activation leads to severe podocytopathy. Therefore, elucidating the mechanism underlying mTORC1 activation in podocytes may contribute to the development of treatments for certain podocytopathies. In our previous study, we showed that macropinocytosis—large-scale endocytosis—is involved in the molecular mechanism of mTORC1 activation in podocytes. Growth factor (GF) stimulation induces circular dorsal ruffles (CDRs), which are large membrane protrusions on the dorsal surface of podocytes. CDRs serve as precursors to macropinocytosis, generating vesicles called macropinosomes, which transport extracellular nutrients to lysosomes, thereby activating mTORC1. These findings suggest that CDRs-derived macropinosomes modulate the mTORC1 pathway. In the present study, we investigated the molecular mechanism underlying macropinosome formation in podocytes, focusing on flotillin-1 (Flot1), a protein enriched in lipid microdomains. Imaging analysis revealed the localization of Flot1 at CDRs, and Flot1 depletion reduced macropinosome formation. Biochemical analysis further demonstrated impaired GF-stimulated mTORC1 activation in Flot1-KO cells, which exhibited slower growth than control cells. Notably, immuno-staining analysis showed that Flot1 is expressed specifically in podocytes but not in other renal cells. These findings indicate that Flot1 participates in the formation of CDRs-derived macropinosomes and contributes to macropinosome-dependent mTORC1 activation in podocytes.
Key words: Flot1, circular dorsal ruffles, macropinocytosis, mTORC1, podocytes
Graphical Abstract
The glomerular filtration barrier is essential for renal function because it prevents the loss of plasma proteins into urine (Pardawala et al., 2025). Podocytes, which are terminally differentiated renal epithelial cells, constitute a critical component of this barrier (Assady et al., 2017; Cunanan et al., 2025; Garg, 2018). These cells exhibit a complex structure characterized by interdigitating “foot processes” that maintain the slit diaphragm, a specialized cell-cell junction crucial for the filtration mechanism. Consequently, podocyte injury represents a convergent pathological mechanism underlying a range of proteinuric kidney diseases collectively termed podocytopathies, including minimal change disease, focal segmental glomerulosclerosis, and diabetic nephropathy (Kopp et al., 2020). Therefore, elucidating the signaling pathways that regulate podocyte homeostasis and function is of paramount importance.
The development of a podocyte-specific Cre gene has enabled the generation of podocyte-specific knockout (KO) mouse models, which have been instrumental in identifying key molecules involved in podocyte function (Moeller et al., 2003). For instance, the roles of two major protein quality control systems—the unfolded protein response (UPR) and endoplasmic reticulum-associated protein degradation (ERAD)—have been examined (Hassan et al., 2016; Kaufman et al., 2017; Yoshida et al., 2021). Deletion of the IRE1α/XBP1 pathway, which is central to the UPR, exerted minimal effects on renal function, with only moderate albuminuria observed after one year of age (Hassan et al., 2016; Kaufman et al., 2017). In contrast, deletion of Sel1L, a key component of ERAD, disrupted foot process structure, caused severe podocyte dysfunction, and resulted in mouse death within three to four weeks (Yoshida et al., 2021). These findings suggest that ERAD serves as the predominant protein quality control mechanism in podocytes. Regarding autophagy, depletion of Atg5, a critical autophagy-related protein, did not severely compromise podocyte integrity, and the KO mice survived (Hartleben et al., 2010). Notably, the activation of mechanistic target of rapamycin complex 1 (mTORC1) is tightly regulated in podocytes (Inoki et al., 2011). Depletion of TSC1, a negative regulator of mTORC1, led to mTORC1 hyperactivation, resulting in podocyte loss and podocytopathies; KO mice died within four weeks of age. Therefore, maintaining appropriate mTORC1 activity is essential for podocyte function. However, the molecular mechanisms by which podocytes regulate this activation remain poorly understood.
mTORC1 is a key component of growth factor (GF)-induced signaling pathways. Over the past few decades, the molecular mechanisms underlying mTORC1 activation have been well characterized (Condon and Sabatini, 2019). Upon GF stimulation, the AKT pathway is activated, which in turn triggers the TSC–Rheb signaling cascade. The small GTPase Rheb subsequently activates mTORC1 directly at the lysosomal surface. The recruitment of mTORC1 to lysosomes is essential for its activation, and this process is regulated by Rag GTPases. When the lysosomal amino acid concentration reaches an optimal level, Rag GTPases are activated, thereby modulating the molecular mechanism responsible for mTORC1 recruitment to lysosomes. Thus, mTORC1 regulation under GF stimulation involves two distinct steps: (1) amino acid–dependent lysosomal recruitment and (2) AKT/TSC/Rheb pathway–dependent activation. To satisfy the biosynthetic demands of the first step, cells frequently utilize macropinocytosis, a large-scale, nonselective endocytic process responsible for internalizing extracellular solutes (Pacitto et al., 2017; Palm et al., 2015; Yoshida et al., 2015, 2018). GF stimulation induces macropinocytosis, generating vesicles called macropinosomes that deliver extracellular amino acids to lysosomes, thereby promoting mTORC1 recruitment. Hence, in certain cell types, macropinocytosis modulates mTORC1 activation by mediating the transport of extracellular nutrients (Swanson and Yoshida, 2019).
Macropinocytosis is an actin-driven endocytic process that begins with the formation of membrane ruffles and concludes with the fusion of plasma membrane protrusions (Stow et al., 2020; Swanson, 2008; Swanson and Araki, 2022). This process engulfs extracellular material into vesicles typically larger than 0.2 μm in diameter. Upon GF stimulation, dynamic membrane ruffles are induced, some of which fuse to generate macropinosomes. These macropinosomes subsequently detach from the plasma membrane and migrate into the cytosol, where they encounter and fuse with lysosomes, facilitating the transfer of extracellular solutes. As a precursor to macropinocytosis, certain cells form circular dorsal ruffles (CDRs) in response to GF stimulation (Hoon et al., 2012; Itoh and Hasegawa, 2013; Sun et al., 2022, 2025; Yan et al., 2024). CDRs are circular membrane ruffles that appear on the dorsal cell surface, gradually contract toward the center, and function to produce macropinosomes. Therefore, CDRs indirectly regulate mTORC1 activity through the macropinocytic process. Indeed, inhibition of CDRs formation has been shown to attenuate GF-induced mTORC1 activation by preventing macropinosome generation (Hua et al., 2023; Yoshida et al., 2018).
In our previous study, we demonstrated that CDRs are present in podocytes in vitro, ex vivo, and in vivo (Hua et al., 2023). Using biochemical and imaging analyses, we found that the GF-stimulated mTORC1 pathway in the podocyte cell line MPC5 and in isolated glomeruli depends on CDRs formation. Based on these observations, we propose that CDRs serve as signaling platforms for mTORC1 and related pathways in glomerular podocytes. We further suggest that elucidating the CDRs–macropinosome mechanism could provide an effective therapeutic strategy for podocytopathies.
Building upon this background, we investigated the molecular mechanisms underlying CDRs-derived macropinocytosis in the context of mTORC1 activation in podocytes, focusing on the role of Flot1, a core structural protein enriched in lipid microdomains (Bickel et al., 1997; Zhan et al., 2023). We found that Flot1 localizes to CDRs and that depletion of this protein impairs macropinosome formation. Moreover, both mTORC1 activation and cell growth rates were reduced in Flot1-KO cells compared with control cells. Notably, tissue staining of mouse kidneys revealed distinctive Flot1 expression in podocytes. Collectively, these findings suggest the existence of an operational Flot1–macropinocytosis–mTORC1 axis in podocytes, the dysregulation of which may contribute to pathological signaling. We propose that Flot1-mediated macropinocytosis provides a critical source of amino acids driving mTORC1 activation in podocytes and that Flot1 represents a promising therapeutic target.
Recombinant mouse epidermal growth factor (EGF; AF-315-09) and recombinant mouse platelet-derived growth factor (PDGF)-BB (AF-315-18) were purchased from PeproTech (Cranbury, NJ, USA). Anti-mTOR (#2983), anti-ERK (#4695), anti-pERK (#4376), anti-AKT (#9272), anti-pAKT (#4060), anti-pS6K (#9234), anti-S6K (#2708S), and anti-WAVE2 (#3659) antibodies were purchased from Cell Signaling Technology (Danvers, MA, USA). Anti-Flot1 (A6220), anti-Tubulin (AC012), and anti-GFP (AE012) antibodies were purchased from ABclonal (Wuhan, China). Anti-LAMP1 (65050-1-1g), anti-Abi1 (27387-1-AP), anti-Rin1 (16388-1-AP), and anti-Podocin (20384-1-AP) antibodies were purchased from Proteintech (Wuhan, China). Anti-Nephrin (ab216341) antibody was purchased from Abcam (Cambridge, UK). Goat anti-rabbit IgG (A21020) and goat anti-mouse IgG (A21010) secondary antibodies were purchased from Abbkine (Wuhan, China). EIPA (1154-25-2) was purchased from Tocris Bioscience (Bristol, UK).
Cell culture and GF stimulationThe mouse podocyte cell line MPC-5 was purchased from Tongpai Biotechnology (Shanghai, China). Cells were cultured in RPMI-1640 basal medium (Gibco, Waltham, MA, USA; C11875500BT) supplemented with 10% fetal bovine serum (FBS; TransGen Biotech, Beijing, China), penicillin (Shanghai Yuanye Bio Technology, Shanghai, China; B25911), and streptomycin (Sangon Biotech, Shanghai, China; A610494-0050). To prevent mycoplasma contamination, cells were treated with prophylactic Plasmocin (InvivoGen, San Diego, CA, USA; ant-mpp) according to the manufacturer’s instructions. For GF stimulation, MPC-5 cells were serum-starved in low-glucose medium (Gibco, Waltham, MA, USA; C11885500BT) for 18 hours and subsequently stimulated with EGF.
Establishment of stable KO cell lines in MPC5Flot1-KO cell lines were generated using the CRISPR/Cas9 system delivered via the lentiCRISPRv2 vector (Addgene). Cells were selected with 5 μg/mL puromycin dihydrochloride (Solarbio, Beijing, China; P8230) for three days. Three guide RNAs targeting mouse Flot1 were designed as follows: 1#: 5'-CACCGGTTCCTGGGGAAGACAGAGG-3', 5'-AAACCCTCTGTCTTCCCCAGGAACC-3'; 2#: 5'-CACCGGCCGGAGAGTGTGGAGAGGC-3', 5'-AAACGCCTCTCCACACTCTCCGGCC-3'; 3#: 5'-CACCGTCCAGATGTACCAGGAGGCC-3', 5'-AAACGGCCTCCTGGTACATCTGGAC-3'.
Live-cell imagingLive-cell phase-contrast imaging was performed using a Zeiss Axio Observer Z1 microscope equipped with a 20× objective. Cells were cultured in 20 mm glass-bottom dishes (NEST, China; 801007) and maintained at 37°C in a humidified atmosphere containing 5% CO2 using a climate chamber (PECON, USA; 160-800489). Prior to GF stimulation, cells were cultured in low-glucose medium for 18 hours.
Immunofluorescence (IF) staining and imagingFor IF staining, cells were seeded onto 14-mm coverslips (Biosharp, Hefei, China; BS-14-RC) placed in 24-well plates (BIOFIL, Guangzhou, China; TCP010024). After incubation, cells were washed with phosphate-buffered saline (PBS; MeilunBio, Dalian, China; MA0015) and fixed with 4% paraformaldehyde (PFA; SparkJade, Shanghai, China; EE0001) in PBS (pH 7.5) for 30 minutes. Cells were then permeabilized with 0.1% Triton X-100 in PBS for 5 minutes and blocked with 5% bovine serum albumin in TBST (20 mM Tris, 150 mM NaCl, 0.1% Tween-20, pH 7.6) for 30 minutes. Subsequently, cells were incubated with primary antibodies for 3 hours at room temperature, followed by three 5-minute washes with TBST. They were then incubated with fluorescently labeled secondary antibodies for 2 hours and washed three additional times with TBST for 5 minutes each. Next, cells were stained with rhodamine–phalloidin (ABclonal, Wuhan, China; RM02835) for 1 hour to visualize F-actin. Finally, after a final set of TBST washes, the coverslips were mounted onto glass slides using an antifade mounting medium containing DAPI (Abcam, Cambridge, UK; ab104139) for nuclear counterstaining. Primary antibodies against Flot1, WAVE2, Abi1, Rin1, LAMP1, mTOR, Nephrin, and Podocin were used at a dilution of 1:50. Secondary antibodies were as follows: anti-rabbit IgG Alexa Fluor 488 (1:500; Abcam, Cambridge, UK; ab150077) and anti-rat IgG Alexa Fluor 594 (1:500; Abcam, Cambridge, UK; ab150160). Confocal images were acquired using a Leica TCS SP5 microscope equipped with a 63×/1.4 NA oil-immersion objective under the following excitation conditions: DAPI at 405 nm, FITC at 488 nm, and DsRed at 543 nm.
Western blottingFor western blot analysis, cells were lysed on ice for 10 minutes in CHAPS buffer (40 mM HEPES, pH 7.5; 120 mM NaCl; 1 mM EDTA; 10 mM sodium pyrophosphate; 10 mM sodium glycerophosphate; 1.5 mM Na3VO4; 0.3% CHAPS; and a protease inhibitor cocktail). The lysates were centrifuged at 12,000 × g for 10 minutes at 4°C, and the resulting supernatant was collected, mixed with SDS loading buffer, and boiled at 100°C for 5 minutes to ensure complete protein denaturation. Samples were then subjected to SDS–PAGE, transferred onto polyvinylidene difluoride membranes (Cytiva, Marlborough, MA, USA; 10600023), and blocked with 5% nonfat milk in TBST. The membranes were incubated with primary antibodies (1:1000 dilution) followed by horseradish peroxidas-conjugated secondary antibodies (1:5000 dilution).
FDx70 uptake assayMPC-5 cells expressing scrambled control (Scr) or Flot1-KO constructs were seeded onto coverslips and serum-starved in low-glucose DMEM for 18 hours. Cells were then treated with EGF and fluorescein isothiocyanate–dextran (FDx70; 0.5 mg/mL) (Thermo Fisher Scientific, Waltham, MA, USA; D1823) for 15, 30, or 60 minutes at 37°C. After fixation with 4% PFA in PBS (pH 7.5) for 30 minutes and washing with Dulbecco’s phosphate-buffered saline (Gibco, Waltham, MA, USA; 14190144), the coverslips were mounted for imaging. All images were acquired using a Zeiss Axio Observer Z1 microscope, with at least ten fields captured per sample. Cell counts and macropinosome numbers (FDx70-positive vesicles) were quantified from phase-contrast and fluorescence images, respectively. FDx70 fluorescence intensity was analyzed using ImageJ software (NIH, Bethesda, MD, USA).
Cell viability assayCells were seeded into 96-well plates at a density of 2,000 cells per well and allowed to adhere overnight. For each experiment, cells were pretreated with the indicated agents for 30 minutes, after which cell viability was assessed using the Cell Counting Kit-8 (CCK-8; Dojindo, Kumamoto, Japan; CK04). Specifically, the culture medium was replaced with CCK-8 working solution (serum-free DMEM mixed with CCK-8 reagent at a 9:1 ratio), and the plates were incubated in the dark for 2 hours. Absorbance at 450 nm was measured using a microplate reader (Agilent BioTek Cytation 5, Winooski, VT, USA). Each condition was tested in triplicate, and mean absorbance values were used for data analysis and graphical representation.
Our previous study delineated the process of GF-induced CDRs formation in podocytes (Hua et al., 2023). To visualize the membrane dynamics of CDRs in MPC5 cells, we performed live-cell imaging to monitor their formation following EGF stimulation (Fig. 1). The CDRs appeared and gradually contracted inward toward the center, eventually closing into a single macropinosome (Fig. 1A). Interestingly, some CDRs were also observed to close and give rise to two distinct macropinosomes (Fig. 1B). To further confirm this phenomenon, cells were treated with another growth factor, PDGF, which also induced the generation of multiple macropinosomes (Supplementary Fig. 1).
Heterogeneity in macropinosome generation in MPC5 cells
Representative live-cell phase-contrast images of epidermal growth factor (EGF)-stimulated MPC5 cells illustrating the formation of macropinosomes from circular dorsal ruffles (CDRs) following membrane closure. Red and yellow arrows indicate the locations of CDRs and macropinosomes, respectively. Upon EGF stimulation, CDRs in MPC5 cells developed into either a single (A) or multiple (B) macropinosomes, demonstrating heterogeneity in macropinosome formation. Times after addition of EGF are indicated at top right (minutes : seconds).
To elucidate the molecular mechanism underlying macropinosome formation from CDRs, we validated the presence of these structures by examining the expression of known CDRs-associated proteins. WAVE2 and Abi1 were previously reported to localize at CDRs (Dubielecka et al., 2011; Suetsugu et al., 2003). Consistently, IF analysis confirmed strong localization of WAVE2 (Fig. 2A) and Abi1 (Fig. 2B) at F-actin–rich ring structures, verifying their identity as CDRs. We next examined whether the lipid raft–associated protein Flot1 participates in this process. Notably, Flot1 was prominently recruited to CDRs (Fig. 2C), suggesting its potential involvement in CDRs formation. To exclude nonspecific staining, Rin1—a protein not implicated in CDRs—was used as a negative control. As expected, Rin1 showed no enrichment at these structures (Fig. 2D), confirming the specific localization of Flot1, WAVE2, and Abi1 to CDRs. To further verify antibody specificity, a GFP-tagged Flot1 plasmid was constructed for independent validation. Western blot analysis confirmed correct expression of the GFP–Flot1 fusion protein (Fig. 2E; Supplementary Fig. 2). The colocalization of GFP–Flot1 with antibody-labeled endogenous Flot1 further validated the reliability of the anti-Flot1 antibody (Fig. 2F). Consistent with these findings, GFP–Flot1 specifically localized to CDRs (Fig. 2G). Line-scan analysis of fluorescence intensity profiles revealed a high degree of overlap between GFP–Flot1 and F-actin signals, with coinciding peak distributions that confirm their colocalization (Fig. 2H). Collectively, these findings identify Flot1 as a novel CDRs-associated protein and suggest a potential functional role for Flot1 in CDRs formation.
Both endogenous and exogenous Flot1 localize to CDRs in MPC5 cells
(A–D) Representative confocal IF images of MPC5 cells stimulated with EGF for 5 minutes. WAVE2 (green) and Abi1 (green) were used as CDRs markers and colocalized with F-actin (red), confirming their localization at CDRs (A and B, respectively). Flot1 (green) was also observed at CDRs (C). Rin1 (green), a protein unrelated to CDRs, served as a negative control for the staining and showed no enrichment at these sites (D). (E) Expression of GFP-tagged Flot1 was confirmed by immunoblotting. The green asterisk denotes exogenous GFP-Flot1, and the yellow asterisk denotes endogenous Flot1. (F) GFP-Flot1 expressed in MPC5 cells was visualized to validate the specificity of the Flot1 antibody. GFP fluorescence (green) and antibody-labeled endogenous Flot1 (red) signals showed strong colocalization. (G) Confocal images showing GFP-Flot1 (green) in MPC5 cells with and without EGF stimulation. GFP-Flot1 was recruited to CDRs identified by F-actin (red). (H) Line-scan analysis of F-actin and GFP-Flot1 fluorescence intensity along the dotted line in (G) demonstrated a high degree of signal overlap, confirming their colocalization at CDRs.
To investigate the role of Flot1 in CDRs formation, we generated a Flot1- KO MPC5 cell line. Western blot analysis confirmed the complete loss of Flot1 expression (Fig. 3A; Supplementary Fig. 3). By comparing KO and Scr cells, we examined whether Flot1 depletion affects EGF-induced CDRs formation in MPC5 cells. Phase-contrast microscopy revealed that CDRs were still induced in Flot1-KO podocytes following EGF stimulation (Fig. 3B). The average diameter of CDRs remained unchanged in the absence of Flot1 (Fig. 3C). A time-course analysis showed no significant difference in CDRs numbers between Flot1-KO and control cells, with CDRs formation peaking at 5 minutes after stimulation and dissipating by 30 minutes (Fig. 3D). Collectively, these results indicate that Flot1 is not essential for CDRs formation. We therefore hypothesized that Flot1 might function in later stages of macropinocytosis. To test this, we quantified macropinosome numbers up to 60 minutes after EGF stimulation. No difference was detected between the groups up to 15 minutes; however, macropinosome numbers in Flot1-KO cells were lower than those in Scr controls at 30 minutes and showed a significant reduction at 1 hour post-stimulation (Fig. 3E–F). These findings suggest that Flot1 is required for efficient macropinosome formation and may play a crucial role in nutrient uptake in podocytes.
Depletion of Flot1 reduces the generation of multiple macropinosomes from CDRs
(A) Establishment and validation of Flot1-knockout (KO) MPC5 cells. Successful Flot1 depletion was confirmed by western blot analysis. Tubulin, AKT, and ERK were used as loading controls, and pAKT was detected to verify the viability of KO cells. Scr: scramble control cells. (B–D) Flot1 depletion does not affect EGF-induced CDRs formation. Representative phase-contrast micrographs of Scr and Flot1-KO cells were captured 5 minutes after EGF stimulation. Red arrows indicate the locations of CDRs (B). Quantification of CDRs size in Scr (n = 258) and Flot1-KO (n = 260) cells showed no significant difference. The y-axis represents arbitrary units (AU), with the average CDRs size in Scr cells normalized to 1.0 (C). Time-course analysis of CDRs numbers at 5, 10, 15, 30, and 60 minutes post-EGF stimulation showed no significant difference between groups. Data represent the mean ± SD of three independent experiments, with >500 cells analyzed per time point (D). (E–F) Flot1 depletion reduces the generation of macropinosomes. A macropinocytosis assay using fluorescent dextran (FDx) was performed in Scr and Flot1-KO cells at 15, 30, and 60 minutes after EGF stimulation. Representative images are shown (E), with red arrows highlighting macropinosomes. Quantification revealed a significant decrease in FDx uptake at 60 minutes in Flot1-KO cells compared with Scr controls. Data represent three independent experiments, with >500 cells analyzed per time point (F). Statistical analysis was performed using an unpaired two-tailed t-test (C–D, F). **p<0.01.
Macropinosomes serve as an auxiliary nutrient delivery mechanism that facilitates mTORC1 activation (Yoshida et al., 2018). Because fewer macropinosomes were generated in Flot1- KO cells, we hypothesized that Flot1 depletion would impair mTORC1 activation. To evaluate the functional consequences of this reduction on downstream nutrient signaling, we measured phosphorylated S6 kinase (pS6K) levels, a well-established downstream readout of mTORC1 activity. Compared with Scr cells, Flot1-KO cells exhibited a significant decrease in pS6K phosphorylation at 5, 10, and 60 minutes after EGF stimulation (Fig. 4A–B; Supplementary Fig. 4). In contrast, activation of the ERK pathway—serving as a control for growth factor signaling—remained unchanged under the same conditions (Fig. 4A and C). Previous studies have demonstrated that nutrients internalized via macropinosomes are delivered to lysosomes through vesicular fusion, promoting mTORC1 recruitment and activation (Palm et al., 2015; Swanson and Yoshida, 2019; Yoshida et al., 2015). To quantify the extent of mTORC1 activation, we examined the colocalization of mTOR with LAMP1, a lysosomal marker, using IF. Under basal conditions without EGF stimulation, both Scr and Flot1-KO cells displayed minimal colocalization. Following EGF treatment for 5 and 15 minutes, Scr cells exhibited pronounced perinuclear lysosomal clustering and markedly increased mTOR–LAMP1 colocalization. In contrast, Flot1-KO cells showed a significant reduction in mTOR–LAMP1 overlap at both time points (Fig. 4D–E). Together, these results indicate that the reduction in macropinosome formation in Flot1-deficient cells attenuates mTORC1 activation, likely due to a diminished supply of macropinosome-derived nutrients.
Depletion of Flot1 attenuates EGF-stimulated mTORC1 activation and cell growth in MPC5 cells
(A–C) Flot1 KO specifically attenuates mTORC1 signaling, as indicated by the decreased pS6K/S6K, without affecting ERK signaling (pERK/ERK), in response to EGF stimulation. Representative western blot results are shown in (A). Quantification of the average pS6K/S6K and pERK/ERK ratios from three independent experiments is presented in (B) and (C), respectively. Statistical analysis was performed using an unpaired two-tailed t-test. **p<0.05, *p<0.01. (D–E) Representative confocal images of Scr and Flot1-KO cells stained for lysosome-associated membrane protein 1 (LAMP1, red) and mTOR (green) following EGF stimulation. In Scr cells, LAMP1 and mTOR showed strong perinuclear colocalization at 15 minutes (D), whereas Flot1-KO cells displayed markedly reduced colocalization at both 5 and 15 minutes (E). (F–G) Cell viability assays using CCK-8 were performed under two conditions: inhibition of macropinocytosis by EIPA (F) and Flot1 depletion (G). Following EGF stimulation, EIPA treatment significantly reduced cell viability at 48 and 72 hours compared with DMSO-treated controls. Similarly, Flot1-KO cells exhibited a progressive reduction in viability from 24 to 96 hours compared with Scr controls. Statistical analysis was performed using an unpaired two-tailed t-test. ****p<0.0001, **p<0.01.
Since mTORC1 is a central regulator of cell growth, we investigated whether impaired macropinocytosis resulting from Flot1 depletion affects cell survival. First, we assessed the effect of pharmacological inhibition of macropinocytosis using EIPA, a well-established macropinocytosis inhibitor (Commisso et al., 2013; Koivusalo et al., 2010; Swanson and Watts, 1995). Cells were treated with EIPA or dimethyl sulfoxide (DMSO) as a control, and viability was measured at 48 and 72 hours. EIPA treatment significantly reduced cell viability at both time points (Fig. 4F), indicating an essential role for macropinocytosis in supporting cell growth. Next, we compared the growth of Scr and Flot1-KO cells under identical conditions. Cell viability was measured at 24, 48, 72, and 96 hours. Flot1-KO cells exhibited significantly lower viability than Scr cells at 24 hours, with the difference becoming more pronounced by 96 hours (Fig. 4G). Together, these results demonstrate that Flot1 functions as a critical molecular hub linking macropinocytic nutrient uptake to mTORC1 signaling, thereby promoting cell survival.
Flot1 is uniquely expressed in podocytes in vivoTo further elucidate the role of Flot1 in podocyte function, we performed IF staining of mouse kidney tissue for in vivo analysis. Podocin and Nephrin are well-established podocyte markers, both localized at the plasma membrane and used to delineate podocyte boundaries (Boute et al., 2000; Kestila et al., 1998; Yoshida et al., 2021). However, simultaneous detection of Flot1 with these membrane-associated markers cannot be applied due to the shared animal source of each antibody. In a previous study, we demonstrated that mTOR and LAMP1 form a unique TOR–autophagy spatial coupling compartment (TASCC) in podocytes, with strong LAMP1 expression specifically observed in these cells (Narita et al., 2011). Consistent with this, our IF staining recapitulated the characteristic LAMP1 pattern, showing diffuse cytoplasmic distribution and distinct perinuclear puncta within Podocin- and Nephrin-positive podocytes (Fig. 5A–B; Supplementary Fig. 5A–B). These findings indicate that LAMP1 can serve as a reliable complementary marker for identifying podocytes in tissue sections. We next examined the subcellular localization of Flot1 within glomeruli. A strong spatial correlation between LAMP1 and Flot1 signals was frequently observed, suggesting that Flot1 is uniquely expressed in podocytes but not in other renal cells (Fig. 5C; Supplementary Fig. 5C). However, due to experimental limitations, the subcellular location of the Flot1 signal could not be confirmed (Fig. 5C; Supplementary Fig. 5C). Collectively, these observations suggest that Flot1 is uniquely expressed in podocytes compared with other renal cell types and may contribute to maintaining podocyte-specific membrane organization.
Unique expression of Flot1 and the proposed model of its function in podocytes in vivo
(A–B) Representative confocal images of mouse kidney tissue co-stained for LAMP1 (red) and podocyte markers podocin (A, green) or nephrin (B, green). Strong LAPM1 expression was observed in podocytes, confirming its utility as a podocyte marker. (C) Representative confocal images of mouse kidney tissue co-stained for LAMP1 (red) and Flot1 (green). The Flot1 signal was robustly detected in podocytes, indicating its preferential expression in these cells. (D) Proposed model of Flot1 function in podocytes. GF stimulation activates receptor proteins and induces CDRs. Flot1 is recruited to these CDRs, which subsequently generate multiple macropinosomes (MPs) where Flot1 participates in the maturation process. These macropinosomes deliver extracellular nutrients to lysosomes, facilitating mTORC1 activation at the lysosomal membrane. The mTORC1 pathway regulates cell growth and the structural maintenance of podocyte foot processes. Therefore, Flot1 depletion reduces macropinosome abundance, leading to attenuated mTORC1 signaling and compromised podocyte function. Flot1 interacts with Flot2 to form the Flot1/Flot2 heterodimer, which stabilizes each other. Because of this, Flot2’s involvement is also proposed.
In this study, we identified a previously unrecognized role of Flot1 in podocytes. Our findings demonstrated that Flot1 localizes to CDRs and contributes to the regulation of their closing process. Depletion of Flot1 markedly reduced the number of macropinosomes generated from CDRs (Fig. 2 and 3). Western blot analysis further revealed that Flot1-KO cells exhibited decreased mTORC1 activation in response to GFs (Fig. 4A–B). Similarly, IF staining showed reduced colocalization of mTOR with LAMP1 in Flot1-KO cells (Fig. 4D–E), and these cells displayed slower growth rates than the controls (Fig. 4G). Taken together, these findings support a model in which Flot1 modulates GF-induced mTORC1 activation through macropinocytosis. Following GF stimulation, CDRs are induced to generate multiple macropinosomes. Flot1 is recruited to these CDRs, where it facilitates macropinosome maturation and closure, although it is not essential for their initial formation. The resulting macropinosomes deliver extracellular nutrients to lysosomes, where they promote mTORC1 activation (Fig. 5D).
The precise molecular mechanism by which Flot1 regulates CDRs-derived macropinosome formation remains to be elucidated. Flot1 forms a heterotetrameric complex with Flotillin-2 (Flot2), stabilizing each other. Together, they localize to lipid rafts on the cytoplasmic side of the plasma membrane. (Gauthier-Rouviere et al., 2020). Therefore, our current data are insufficient to conclude that Flot1 directly regulates macropinocytosis in podocytes. Rather, it is reasonable to hypothesize that the Flot1/Flot2 heterodimer itself is involved in the process (Fig. 5D). In fact, a recent study showed that depletion of Flot2 in podocytes worsens lipopolysaccharide (LPS)-induced kidney injury in vivo (Yu et al., 2023). Additionally, the distribution pattern of the actin cytoskeleton was disturbed in Flot2 KO podocytes (Yu et al., 2023). Meanwhile, a pioneering study demonstrated that Flot1 depletion inhibits clathrin-independent endocytosis (Glebov et al., 2006). Subsequent studies have referred to flotillin-dependent endocytosis as “upregulated flotillin-induced trafficking” (UFIT), and its physiological roles have been extensively investigated (Gauthier-Rouviere et al., 2020). UFIT has been reported in multiple cell types, suggesting a possible mechanistic link between UFIT and CDRs-mediated macropinocytosis in podocytes.
In a previous paper, we demonstrated that mTORC1 plays a critical role in maintaining foot-process structure, although the mechanism remains unknown (Inoki et al., 2011). For this project, we generated a podocyte-specific TSC2 KO mouse. Since TSC2 is a negative regulator of mTORC1 (Inoki and Guan, 2009), mTORC1 becomes “hyper-activated” in the podocytes of the KO mouse. Scanning electron microscopy (SEM) observations clearly showed that the foot processes of the knockout KO mouse were dramatically disturbed at 4 weeks, resulting in death within 14 weeks due to a malfunctioning filtration mechanism. Meanwhile, in the current project, our data strongly suggest that Flot1 is one of the key molecules in the mTORC1 activation mechanism in podocytes. Therefore, we speculate that Flot1 is involved in foot process formation through the mTORC1 pathway (Fig. 5D).
Tissue staining revealed unique Flot1 expression in podocytes compared to other renal cells in vivo (Fig. 5C; Supplementary Fig. 5C). This finding suggests that the proposed model applies not only to MPC5 cells, a podocyte-derived cell line, but also to podocytes on the glomerular surface. Although our tissue staining data is insufficient to conclude that Flot1 is enriched along the periphery of podocytes and foot processes, we hypothesize that the protein is located at the plasma membrane, similar to Podocin and Nehprin, for two reasons. First, IF data showed Flot1 at CDRs in MPC5 cells (Fig. 2C), which was confirmed by an alternative experimental method using GFP-Flot1 (Fig. 2G–H). Second, Flot1 has been established as a scaffold protein of lipid rafts. Studies have shown that the protein is primarily located at the plasma membrane (Zhan et al., 2023). In this case, one might propose that Flot1 is located at the plasma membrane, where it modulates the macropinosome-mTORC1 axis and contributes to podocyte function as part of the kidney filtration mechanism (Fig. 5D).
In summary, our study demonstrates that Flot1 plays a critical role in GF-stimulated activation of the mTORC1 pathway, which is essential for maintaining podocyte function. Based on these findings, we propose that Flot1 may serve as a promising therapeutic target for specific types of podocytopathies. Future investigations should include the generation and characterization of podocyte-specific Flot1 knockout mouse models to validate this hypothesis.
This work was supported by the Frontiers Science Center for Cell Responses grant from Nankai University (C029205001) and the Shenzhen Science and Technology Program (JCYJ20210324120813037) to S.Y.; the joint research program of the Research Center for GLOBAL and LOCAL Infectious Diseases, Oita University (2025A02) to H.M. and S.Y.; the Astellas Foundation for Research on Metabolic Disorders (2023A3542) to H.M.; the JSPS Bilateral Program (JPJSBP120257417) to H.M.; the National Natural Science Foundation of China (81970654), the Shenzhen High-level Hospital Construction Fund (XJ2022019601, 24250G1036), and the Shenzhen Science and Technology Program (JCYJ20220530162816036) to Y.F.
Conflict of Interest StatementThe authors declare that this study was conducted in the absence of any commercial or financial relationships that could be construed as potential conflicts of interest.
Data Availability StatementThe datasets presented in this study and the original data are available from the corresponding author upon request. All the original western blot images are shown as the Supplemental-Figures.
Author Contribution StatementLY and YX designed and performed the experiments, with the support of LC, HY, and XW. SY conceived the study, designed the experiments, and wrote the manuscript, with the support of YF and HM.
Ethics Approval and Consent to ParticipateEthics approval is not required in this study.
Patient Consent for PublicationPatient consent for publication is not required in this study.
Cell Counting Kit-8
CDRscircular dorsal ruffles
EGFepidermal growth factor
ERADendoplasmic reticulum-associated protein degradation
FBSfetal bovine serum
FDxfluorescein isothiocyanate–dextran
Flot1flotllin-1
Flot2flotllin-2
GFgrowth factor
IFimmunofluorescence
KOknockout
LPSlipopolysaccharide
mTORC1mechanistic target of rapamycin complex 1
PDGFplatelet-derived growth factor
PFAparaformaldehyde
pS6Kphosphorylated S6 kinase
SEMScanning electron microscopy
TASCCTOR–autophagy spatial coupling compartment
UFITupregulated flotillin-induced trafficking
UPRunfolded protein response
