KIF1C facilitates retrograde transport of lysosomes through Hook3 and dynein (original) (raw)

Introduction

Lysosomes are dynamic cellular organelles integral to various biological processes, including autophagy, endocytosis, and phagocytosis1,2,3,4. Given the shared characteristics among lysosomes, late endosomes, and endolysosomes, such as the presence of identical membrane proteins like lysosome-associated membrane protein 1 (LAMP1), this article will refer to these collectively as lysosomes. Notably, the quantity, morphology, and spatial distribution of lysosomes can vary significantly depending on the cell type and physiological context5,6,7,8. Typically, lysosomes are predominantly located in the perinuclear region, often described as the perinuclear cloud, yet their positioning within a cell remains dynamic, influenced by several factors including nutrient availability, growth factors, cytosolic pH, oxidative stress, and infection4,9,10. For instance, nutrient deprivation or the absence of growth factors triggers lysosome clustering in the perinuclear area to promote efficient fusion with autophagosomes, facilitating the degradation and recycling of cellular materials11. Conversely, when nutrients are reintroduced, lysosomes migrate toward the plasma membrane, enhancing lysosome-localized mTORC1 signaling and thereby stimulating gene expression for protein synthesis12.

Lysosomes are transported bi-directionally along microtubules by the cytoplasmic dynein-1 (hereafter referred to as dynein), in association with dynactin, and several members of the kinesin family9,10. Dynein drives retrograde movement (towards the microtubule minus-end), while kinesins mediate anterograde movement (towards the microtubule plus-end). The minus-ends of microtubules are generally located near the centrosome, close to the nucleus, and the plus-ends are directed towards the cell periphery. Therefore, dynein transports lysosomes from the periphery to the cell center, and kinesins distribute them throughout the cytoplasm or position them at the cell periphery.

Motor proteins are linked to organelle membranes via adapters, often effectors of the Rab or Arf-like (Arl) small GTP-binding proteins13. A critical player in this process is Arl8, located on lysosomal membranes, which recruits motor proteins to facilitate directional transport. Specifically, Arl8 engages the kinesin-1 family member KIF5B via its effector SKIP, promoting anterograde movement of lysosomes14. Additionally, Arl8 directly attracts kinesin-3 family members KIF1A and KIF1Bβ to lysosomes, further supporting their anterograde transport15. For retrograde transport, Arl8 utilizes effectors like RUFY (RUN and FYVE domain-containing protein) 3/4 and DENND6A to facilitate dynein-driven movement. RUFY3/4 can bind dynein directly or indirectly through the dynein-activating adapter JIP416,17. Similarly, DENND6A activates Rab34, which then recruits dynein via the dynein-activating adapter RILP (Rab interacting lysosomal protein)18. In addition to small GTP-binding proteins and their effectors, the lysosomal membrane protein TMEM55B recruits dynein to lysosomes by associating with JIP4, thereby mediating the anterograde transport of lysosomes19.

KIF1C, like KIF1A and KIF1B, is a member of the kinesin-3 family20. It is autoinhibited by intramolecular interactions between its stalk and motor domains, which can be relieved upon binding of Protein Tyrosine Phosphatase D1 (PTPD1) or the dynein-activating adapter Hook321. Notably, Hook3 can simultaneously interact with KIF1C and dynein, scaffolding the interaction between these opposite-polarity motors and enabling bidirectional motility along microtubules22. KIF1C is ubiquitously expressed and involved in vesicle transport in neurons23, MHC class II antigen presentation in myeloid cells24, and α5β1-integrin transport in RPE1 and U2OS cells25. Additionally, it plays roles in regulating Golgi structure26, podosome dynamics27,28,29, and invadopodia formation30. Recent studies further highlight its involvement in mRNA transport31,32,33. Despite its broad involvement in cellular functions, the role of KIF1C in lysosomal positioning has been unclear. We show here that KIF1C facilitates dynein-mediated retrograde transport of lysosomes, which is responsible for their positioning in the perinuclear region. The motor activity of KIF1C is not essential for the perinuclear positioning of lysosomes, instead, it plays a negative regulatory role in this process. KIF1C facilitates the recruitment of the dynein-dynactin-Hook3 complex to lysosome-associated RUFY3, thereby modulating lysosomal distribution, which is critical for effective autophagic and endocytic degradation processes. Our study elucidates an unconventional function of KIF1C, activating dynein-driven lysosomal transport independent of its motor function, thus providing valuable insights into the complex regulation of organelle positioning.

Results

KIF1C regulates lysosomal distribution

To explore the function of KIF1C in regulating lysosomal positioning, we developed KIF1C knockout (KO) U2OS cell lines (Fig. 1A). Immunoblotting confirmed that control and _KIF1C_-KO cells express comparable levels of LAMP1, a major component of the lysosomal membrane used as a lysosomal marker (Fig. 1A). Subsequent immunostaining of LAMP1 revealed that in control cells, lysosomes were broadly distributed throughout the cell, with a marked concentration in the perinuclear region (Fig. 1B). In contrast, _KIF1C_-KO cells displayed a more uniform distribution of lysosomes, with a reduced concentration around the perinuclear region (Fig. 1B). We confirmed that total LAMP1 intensity per cell was comparable between control and _KIF1C_-KO cells (Fig. 1B). To quantify the distribution of lysosomes, we measured the LAMP1 intensity in the perinuclear (0–5 µm from the nuclear rim) and peripheral ( > 15 µm from the nuclear rim) regions relative to that in entire cell (Fig. 1C). As shown in Fig. 1D, _KIF1C_-KO cells had significantly lower LAMP1 intensity in the perinuclear region and higher intensity in the peripheral regions compared to control cells, indicating that loss of KIF1C causes the dispersion of lysosomes throughout the cell. Similar results were observed in _KIF1C_-KO HeLaS3 cells (Fig. S1), confirming the role of KIF1C across different cell types. Furthermore, the altered distribution of lysosomes was effectively reversed by reintroducing KIF1C into _KIF1C_-KO U2OS (Fig. 1E–G) and HeLaS3 cells (Fig. S1B,C), reinforcing the apparent contribution of KIF1C to lysosomal positioning. In contrast, we found no significant changes in the distribution of early endosomes (Fig. S2A,B) and mitochondria (Fig. S2C,D) between control and _KIF1C_-KO cells. Additionally, loss of KIF1C also did not affect the overall structure of the Golgi apparatus, which is typically positioned near the nucleus as a compact, stacked structure in a microtubule-dependent manner34,35 (Fig. S2E,F). These findings emphasize that KIF1C selectively influences lysosomal distribution without affecting the general organelle architecture within cells.

Fig. 1: KIF1C induces perinuclear positioning of lysosomes independently of its motor activity.

figure 1

A Representative immunoblots showing the levels of KIF1C and LAMP1 in control and _KIF1C_-KO (#1 and #2) U2OS cells. B Representative fluorescence images of control and _KIF1C_-KO (#1 and #2) U2OS cells stained with anti-LAMP1 antibody and DAPI. Scale bars, 10 μm. The values below each panel represent the relative fluorescence intensities of LAMP1 per cell, presented as the mean ± SD of three independent experiments, with no significant differences (Tukey’s test). C A diagram illustrating how the perinuclear (0–5 μm) and periphery ( > 15 μm) regions of a cell were defined, which were used to quantify the lysosomal distribution. D Quantification of the lysosomal distribution, based on LAMP1 intensity, from the experiments shown in B. Data from three independent experiments are presented as superplots. **P < 0.01, Tukey’s test. E Representative fluorescence images of _KIF1C_-KO#2 U2OS cells transfected with expression plasmid for GFP (mock) or KIF1C-Clover and stained with anti-LAMP1 antibody and DAPI. Scale bars, 10 μm. F Schematic domain structure of wild-type (WT) KIF1C with motor, forkhead-associated (FHA), proline-rich (P-rich), and coiled-coil (CC) 1–4 domains, and its mutants used in this study. Amino acids 794–807 constitute the binding site for Hook3. The lower panel illustrates a model of the complex formed by dynein-dynactin (shown in bright and dark red), Hook3 (green), and KIF1C (yellow) on microtubules, based on Kendrick et al.22. G Quantification of the lysosomal distribution in control and _KIF1C_-KO#2 U2OS cells expressing the indicated KIF1C-Clover constructs. Data from three independent experiments are presented as superplots. *P < 0.05; **P < 0.01; ns, not significant, Tukey’s test, all compared to _KIF1C_-KO#2 cells with mock transfection, except for those indicated by bars.

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KIF1C facilitates the perinuclear positioning of lysosomes independently of its motor activity

To elucidate how KIF1C influences lysosomal distribution, we employed various KIF1C constructs featuring deletions or point mutations as shown in Fig. 1F for rescue experiments in _KIF1C_-KO U2OS cells. Notably, expression of KIF1C constructs lacking the motor domain (∆motor) or exhibiting inactivated motor function (K103A) successfully restored the perinuclear distribution of lysosomes (Fig. 1G). Intriguingly, these motor-deficient constructs led to a more pronounced concentration of lysosomes in the perinuclear region compared to wild-type (WT) KIF1C (Fig. 1G). This suggests that the motor activity of KIF1C is not essential for directing lysosomes to the perinuclear region but rather exerts a suppressive effect on this process. In support of this observation, experiments conducted in wild-type U2OS cells revealed that expressing KIF1C-∆motor or KIF1C-K103A, unlike WT KIF1C, induced significant perinuclear clustering of lysosomes (Fig. S3). This indicates that the constructs of KIF1C lacking motor function exhibit dominant effects over endogenous wild-type KIF1C, leading to enhanced perinuclear clustering of lysosomes.

KIF1C facilitates dynein-driven lysosomal transport via interacting with Hook3

To delineate how KIF1C regulates perinuclear positioning of lysosomes, we utilized KIF1C mutants truncated in the C-terminal region (Fig. 1F), which is known to interact with cargo, adapter, or regulatory proteins20,21,22,26,36. Rescue experiments revealed that a truncation at amino acids (aa) 810 (KIF1C-Δ810–1103) successfully restored normal lysosomal distribution, whereas a truncation at aa 600 (KIF1C-Δ600–1103) did not (Fig. 1G). This indicates that the region between aa 600–809 is crucial for lysosomal positioning. Importantly, this region includes aa 794–807, previously identified as the binding site for Hook3, a dynein-activating adapter capable of scaffolding the interaction between KIF1C and dynein22 (Fig. 1F). The specific deletion of this Hook3-binding site (KIF1C-Δ794–807) also failed to restore lysosomal distribution (Fig. 1G), underscoring the importance of the interaction between KIF1C and Hook3 in regulating lysosomal positioning.

To further investigate the role of Hook3 in KIF1C-mediated lysosomal positioning, we performed Hook3 knockdown in _KIF1C_-KO cells. Our results showed that KIF1C was unable to restore perinuclear lysosome positioning when Hook3 levels were reduced (Fig. 2A–C). In contrast, knockdown of bicaudal D homolog 2 (BICD2), encoding another dynein-activating adapter suggested to interact with KIF1C37, did not impair KIF1C’s ability to restore perinuclear lysosome positioning (Fig. 2A–C). This specificity was further supported by immunoprecipitation assays showing co-precipitation of endogenous Hook3 but not BICD2 with KIF1C-mCherry (Fig. 2D). Moreover, knockdown of dynein heavy chain (DHC) significantly diminished KIF1C’s capacity to restore perinuclear lysosome positioning in _KIF1C_-KO cells (Fig. 2A–C). These findings suggest that KIF1C mediates the perinuclear distribution of lysosomes primarily through its interaction with the dynein-dynactin-Hook3 complex. Notably, DHC knockdown in _KIF1C_-KO cells further decreased perinuclear lysosome distribution and increased their peripheral positioning, while Hook3 knockdown did not produce any additional effect (Fig. 2C). This suggests that while Hook3’s role in lysosomal transport is predominantly dependent on KIF1C, dynein also contributes to retrograde lysosomal transport independently of the KIF1C-Hook3 complex (e.g., through the JIP4-TMEM55B complex).

Fig. 2: Dynein and Hook3 are required for KIF1C-induced perinuclear positioning of lysosomes.

figure 2

A Representative immunoblots showing knockdown efficiency of si_Hook3_ (#1 and #2), si_BICD2_, and si_DHC_ in _KIF1C_-KO#2 U2OS cells. B,C _KIF1C_-KO#2 U2OS cells treated with the indicated siRNAs were further transfected with expression plasmid for KIF1C-WT-Clover and stained with anti-LAMP1 antibody and DAPI. Representative fluorescence images of the cells were shown (B). Scale bars, 10 μm. Lysosomal distribution in cells with (+) and without (−) KIF1C-WT-Clover expression was quantified (C). Data from three independent experiments are presented as superplots. *P < 0.05; **P < 0.01; ns, not significant, Tukey’s test. D Whole-cell lysates (WCL) from U2OS cells expressing mCherry (−) or KIF1C-mCherry (+) were subjected to immunoprecipitation (IP) with anti-RFP nanobody, followed by immunoblotting. The graph displays the relative levels of Hook3 that co-immunoprecipitated (IPed) with KIF1C, normalized to the total Hook3 levels in WCL. The value from cells expressing KIF1C-mCherry in one experiment is set to 1.0. Data are expressed as mean ± SD of three independent experiments. Co-IPed BICD2 was undetectable.

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Additional observations showed that KIF1C accumulates not only at cell protrusions but also in the pericentrosomal region, as reported previously25 (Fig. S4A,B). The pericentrosomal accumulation corresponds to the primary sites where Hook3 and the p150 subunit of dynactin are also observed, as previously reported38,39 (Fig. S4C). Notably, deletion of the Hook3-binding site on KIF1C (KIF1C-Δ794–807) abolished its pericentrosomal accumulation, while not affecting its localization at cell protrusions (Fig. S4A,B). Conversely, deletion of the motor domain (KIF1C-∆motor) or motor inactivation (KIF1C-K103A) of KIF1C prevented its accumulation in cell protrusions but did not affect its pericentrosomal localization (Fig. S4A,B). These observations support the proposed interaction between KIF1C and the dynein-dynactin-Hook3 complex.

KIF1C mediates retrograde transport of lysosomes by linking Hook3 and RUFY3

Supporting our observations, proteomic studies have identified Hook3 in complexes associated with lysosomes and autophagic vesicles, including autolysosomes40,41. However, Hook3 is also localized to the Golgi apparatus and early endosomes42,43, suggesting its ability to couple various cargo vesicles with motor proteins. Based on these observations, we explored the interaction of Hook3 with lysosomes, specifically examining the role of KIF1C in these processes. We particularly focused on the interactions of Hook3 and KIF1C with lysosomal proteins involved in dynein-driven transport, namely RUFY3 and TMEM55B. These proteins are known to associate with dynein through JIP4 (or its neuron-specific paralog JIP3), which functions similarly to Hook3 by activating dynein to mediate perinuclear lysosomal clustering under conditions such as nutrient starvation or cytosol alkalinization16,17,19 (Fig. 3A). Our experiments in control U2OS cells demonstrated that ectopic expression of either RUFY3 or TMEM55B induced perinuclear clustering of lysosomes (Fig. 3B,C). However, in _KIF1C_-KO U2OS cells, only TMEM55B maintained this ability (Fig. 3B,C). These data suggest that while both RUFY3 and TMEM55B can drive lysosomal clustering, only RUFY3 requires KIF1C to perform this function, whereas TMEM55B acts independently of KIF1C. Consistent with these results, co-immunoprecipitation assays have revealed that RUFY3, but not TMEM55B, is associated with KIF1C (Fig. 3D).

Fig. 3: KIF1C is required for RUFY3-mediated retrograde transport of lysosomes.

figure 3

A Schematic model illustrating the current understanding of dynein-driven lysosomal transport along microtubules, mediated by either RUFY3 (blue) or TMEM55B (orange). Both RUFY3 and TMEM55B are capable of recruiting lysosomes to the dynein-dynactin (bright and dark red) through the adapter protein JIP4 (pink)16,17,19. B Representative fluorescence images of control and _KIF1C_-KO#2 U2OS cells expressing GFP-RUFY3 or GFP-TMEM55B and stained with anti-LAMP1 antibody and DAPI. Scale bars, 10 μm. C Quantification of the lysosomal distribution from the experiments shown in A. Data from three independent experiments are presented as superplots. **P < 0.01; ns, not significant, Tukey’s test. D Whole-cell lysates (WCL) from HEK293T cells expressing the indicated proteins were subjected to immunoprecipitation (IP) with anti-GFP nanobody, followed by immunoblotting. The graph displays the relative levels of KIF1C-mCherry that co-immunoprecipitated (IPed) with GFP-RUFY3 or GFP-TMEM55B, normalized to the total KIF1C-mCherry levels in WCL. The value from cells expressing KIF1C-mCherry alone is set to 1.0 in each experiment. Data are expressed as mean ± SD of three independent experiments. *P < 0.05, Student’s _t_-test.

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Based on these findings, we hypothesized that RUFY3 may function as an adapter that links lysosomes to the dynein-dynactin-Hook3-KIF1C complex (Fig. 4A). To explore this hypothesis, we tested whether RUFY3 also associates with Hook3, and indeed, RUFY3 was co-immunoprecipitaed with Hook3 (Fig. 4B). However, the C-terminal truncation of Hook3, which removed its KIF1C-binding region (aa 553–718)22, diminished its interaction with RUFY3 (Fig. 4B), indicating that the KIF1C-binding region of Hook3 is critical for the RUFY3-Hook3 interaction. To further confirm the role of KIF1C in the RUFY3-Hook3 interaction, we performed co-immunoprecipitation assays using _KIF1C_-KO cells. Notably, in _KIF1C_-KO U2OS cells, the RUFY3-Hook3 interaction was significantly weaker compared to control U2OS cells (Fig. 4C). Consistently, _KIF1C_-KO cells exhibited significantly reduced colocalization of Hook3 with LAMP1 (Fig. 4D,E). Moreover, while ectopic expression of RUFY3 significantly enhanced Hook3-LAMP1 colocalization in control cells, it failed to induce a similar increase in _KIF1C_-KO cells, resulting in more pronounced differences in Hook3-LAMP1 colocalization between control and _KIF1C_-KO cells (Fig. 4D,E). A previous report described that RUFY3 can be visualized on the lysosomal membrane by permeabilizing the cells with a mild detergent saponin before fixation, which minimizes the fluorescent signal from the cytosolic pool of overexpressed RUFY317. Using this procedure, we observed that RUFY3 colocalizes, at least partially, with either KIF1C or Hook3, and also with both (Fig. S5A,B). Together, these findings suggest that KIF1C may help stabilize the interaction between Hook3 and RUFY3, potentially on lysosomes, thereby facilitating the dynein-driven retrograde transport of lysosomes.

Fig. 4: KIF1C links Hook3 and RUFY3 for retrograde transport of lysosomes.

figure 4

A Schematic model illustrating the proposed mechanism of lysosomal transport driven by a complex consisting of dynein-dynactin (bright and dark red), Hook3 (green), KIF1C (yellow), and RUFY3 (blue) on microtubules. The lower panel shows the schematic domain structure of wild-type (WT) Hook3 and its Δ553–718 mutant, which lacks the KIF1C-binding region. CC, coiled-coil. B,C Whole-cell lysates (WCL) from HEK293T cells (B) and control or _KIF1C_-KO#2 U2OS cells (C) expressing the indicated proteins were subjected to immunoprecipitation (IP) with anti-RFP nanobody, followed by immunoblotting. The graphs display the relative levels of GFP-RUFY3 that co-immunoprecipitated (IPed) with mCherry-Hook3, normalized to the total mCherry-Hook3 levels in WCL. The value from cells expressing GFP-RUFY3 alone is set to 1.0 in each experiment. Data are expressed as mean ± SD of three independent experiments. *P < 0.05; **P < 0.01, Student’s _t_-test. D Representa_t_ive fluorescence images of control and _KIF1C_-KO#2 U2OS cells expressing mock (left panels) or GFP-RUFY3 (right panels) and stained with anti-LAMP1 and anti-Hook3 antibodies. Insets in the right panels show scaled-down images of the GFP channel. Magnified views of the boxed regions, with single and merged channels, are shown to the right of each panel. Scale bars, 10 μm. E Colocalization between Hook3 and LAMP1 from the experiments shown in D was analyzed by measuring the percentage of total Hook3 signal colocalizing on LAMP1 per cell. Data from three independent experiments are presented as superplots. *P < 0.05; **P < 0.01; ns, not significant, Tukey’s test.

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KIF1C is essential for efficient degradation of autophagic and endocytic cargo

The perinuclear localization of lysosomes is crucial for maintaining proper autophagic flux, which encompasses autophagosome formation, their fusion with lysosomes, and the degradation of their contents11,44. Given that the loss of KIF1C led to the dispersion of lysosomes under nutrient-rich conditions, we hypothesized that KIF1C plays a role in regulating basal autophagic flux. To investigate this, we measured the levels of LC3B-II, a lipidated form of LC3B associated with autophagosomes, which correlates with the number of autophagosomes45. In _KIF1C_-KO cells, LC3B-II levels were significantly higher compared to control cells under nutrient-rich conditions (Fig. 5A,B), which may be a consequence of either enhanced autophagosome formation or reduced autophagosome turnover via their fusion with lysosomes and degradation process within the autolysosomes45. To distinguish between these two possibilities, cells were treated with bafilomycin A1, which inhibits lysosomal acidification and blocks degradation process within the autolysosomes. This treatment normalized LC3B-II levels between _KIF1C_-KO and control cells (Fig. 5B), indicating that the observed increase in LC3B-II in _KIF1C_-KO cells results primarily from reduced turnover rather than increased formation.

Fig. 5: KIF1C is required for efficient degradation of autophagic and endocytic cargo.

figure 5

A Representative immunoblots showing increased basal levels of LC3B-II in _KIF1C_-KO U2OS cells compared to those in control U2OS cells. B Control and _KIF1C_-KO#2 U2OS cells were treated with bafilomycin A1 or DMSO in growth medium for 2 h and analyzed by immunoblotting. The graph shows relative LC3B-II levels normalized to tubulin. The value from control cells treated with DMSO is set to 1.0 in each experiment. Data are expressed as mean ± SD of three independent experiment. **P < 0.01; ns, not significant, Tukey’s test. C Representative fluorescence images of control and _KIF1C_-KO#2 U2OS cells incubated in growth medium or HBSS for 5 h and stained with anti-LAMP1 antibody and DAPI. Scale bars, 10 μm. D Quantification of the lysosomal distribution from the experiments shown in C. Data from three independent experiments are presented as superplots. *P < 0.05; **P < 0.01; ns, not significant, Tukey’s test. E Control and _KIF1C_-KO#2 U2OS cells were incubated in growth medium or HBSS with or without bafilomycin A1 for 2 h and analyzed by immunoblotting. The graph shows relative LC3B-II levels normalized to tubulin. The value from control cells cultured in growth medium is set to 1.0 in each experiment. Data are expressed as mean ± SD of three independent experiment. **P < 0.01; ns, not significant, Tukey’s test.Data are expressed as mean ± SD of three independent experiment. *P < 0.05; **P < 0.01; ns, not significant, Tukey’s test. F Representative fluorescence images of control and _KIF1C_-KO#2 U2OS cells were labeled by DQ-Green-BSA and LysoTracker Red DND-99. Scale bars, 10 μm. G Fluorescence intensity of DQ-green-BSA was quantified. Data from three independent experiments are presented as superplots. *P < 0.05, Student’s _t_-test. H Control and _KIF1C_-KO#2 U2OS cells were incubated in serum-free DMEM containing biotinylated BSA with or without bafilomycin A1 for 1 h and analyzed by immunoblotting. The graph displays the relative levels of biotinylated BSA normalized to tubulin. The value from control cells treated with DMSO is set to 1.0 in each experiment. Data are expressed as mean ± SD of three independent experiment. *P < 0.05; ns, not significant, Tukey’s test.

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We further assessed whether loss of KIF1C affects autophagy under nutrient starvation, a condition known to induce perinuclear lysosome clustering and accelerate autophagy4,11. Under conditions of nutrient deprivation, control cells exhibited perinuclear clustering of lysosomes, whereas in _KIF1C_-KO cells, lysosomes remained dispersed (Fig. 5C,D). Additionally, LC3B-II levels were significantly higher in _KIF1C_-KO cells compared to control cells during starvation, but these levels were normalized following bafilomycin A1 treatment (Fig. 5E), indicating impaired autophagic flux in _KIF1C_-KO cells. These findings suggest that KIF1C is crucial for maintaining efficient autophagic degradation under both basal and stress-induced conditions.

To confirm the role of KIF1C in autophagic degradation, we analyzed the colocalization of GFP-LC3 with LAMP1 under starvation conditions, which assesses the fusion efficiency between autophagosomes and lysosomes. Although _KIF1C_-KO cells exhibited highly dispersed LAMP1-positive compartments, the colocalization with GFP-LC3 did not differ significantly from that in control cells, regardless of the presence or absence of bafilomycin A1 (Fig. S6). This suggests that while KIF1C is not critical for the fusion of autophagosomes with lysosomes, it is necessary for the efficient degradation within autolysosomes.

Beyond their role in autophagy, lysosomes are essential for endocytic degradation processes46. We therefore explored the role of KIF1C in endocytic degradation by assessing the fluorescence intensity of dye-quenched DQ-Green-BSA, which is endocytosed and de-quenched upon proteolytic cleavage in lysosomes. _KIF1C_-KO cells exhibited significantly lower fluorescence intensity compared to control cells (Fig. 5F,G), suggesting impaired endocytic degradation. To further validate this observation, we measured the levels of endocytosed biotin-labeled BSA by immunoblotting analysis. Our results showed that _KIF1C_-KO cells accumulated significantly higher levels of biotin-BSA compared to control cells (Fig. 5H). However, these levels were normalized following bafilomycin A1 treatment (Fig. 5H), indicating a reduced capacity for lysosomal degradation of BSA in _KIF1C_-KO cells. Additionally, we examined the endocytic degradation of platelet-derived growth factor receptor β (PDGFRβ) following ligand stimulation, a process primarily dependent on lysosomal function, as evidenced by its suppression upon bafilomycin A1 treatment (Fig. S7A). Notably, degradation of PGDFRβ was significantly delayed in _KIF1C_-KO cells compared to control cells (Fig. S7B), further suggesting a compromised capacity for endocytic degradation within lysosomes in _KIF1C_-KO cells. To ensure that the observed effects were specific to the degradation process rather than altered endocytic delivery, we assessed the delivery of endocytic cargo using Alexa Fluor 647-dextran and BODIPY-FL-LDL. Our analysis revealed no significant differences in the efficiencies of these cargo delivery to acidic lysosomes between control and _KIF1C_-KO cells (Fig. S8). These results suggest that while KIF1C is not required for the trafficking of endocytic cargo to lysosomes, it plays an essential role in facilitating their subsequent degradation.

Discussion

Our findings underscore the critical role of KIF1C in regulating the retrograde transport of lysosomes for their perinuclear positioning. This function is particularly noteworthy considering its established role in anterograde transport, where it moves diverse cargoes such as secretory vesicles23, α5β1-integrin25, and mRNAs31,32,33 across cells. Intriguingly, KIF1C does not rely primarily on its motor activity for the retrograde transport of lysosomes. Instead, the motor domain appears to serve a negative regulatory role, while the stalk and tail domains are responsible for this process by acting as a regulatory adapter, facilitating dynein-driven lysosomal transport. This distinctive function is likely attributed to the unique structural composition of KIF1C, particularly within the tail domain, which contains an intrinsically disordered region that drives liquid-liquid phase separation47. This structural feature differentiates KIF1C from its kinesin-3 family counterparts, KIF1A and KIF1Bβ, which are known to drive the anterograde transport of lysosomes15. Thus, KIF1C regulates cellular transport processes through both motor-dependent and -independent functions.

Our findings revealed the essential role of the dynein-activating adapter Hook3 in KIF1C-mediated lysosomal positioning. It has been reported that Hook3 alleviates the autoinhibition of KIF1C by interacting within its stalk domain21. Furthermore, Hook3 can serve as a scaffold for linking dynein and KIF1C22, enabling bidirectional motility on microtubules with each motor responsible for transport in different directions along microtubules22. Our results suggest that KIF1C binding to Hook3, likely linked with the dynein-dynactin complex, may bring this complex to RUFY3 on lysosomes, promoting their retrograde transport. Thus, we propose that Hook3 activates both dynein and KIF1C motors within the dynein-dynactin-Hook3-KIF1C complex, where dynein primarily drives lysosomal movement towards the perinuclear region, and KIF1C may allow this complex to interact stably with lysosome-associated RUFY3 while acting as a brake that slows the dynein motor so that lysosomes do not approach too close to the nucleus (Fig. 6i). Consequently, dysfunction in the KIF1C motor results in pronounced perinuclear clustering of lysosomes, attributable to unregulated, dynein-mediated lysosomal transport (Fig. 6ii), while KIF1C KO induces lysosomal dispersion throughout the cell, likely due to a compromised interaction between the dynein-dynactin-Hook3 complex and lysosomes (Fig. 6iii). Since our co-immunoprecipitation experiments involving KIF1C, RUFY3, and Hook3 relied on overexpressed proteins, which may lead to false-positive results, further studies using endogenous proteins will be necessary to strengthen the conclusions of this study.

Fig. 6: Model for the role of KIF1C in lysosomal positioning.

figure 6

(i) In wild-type cells, KIF1C (yellow) interacts with the dynein-dynactin complex (bright and dark red) via Hook3 (green), facilitating the recruitment of this complex to RUFY3 (blue) on lysosomes. This interaction promotes the retrograde transport of lysosomes towards the perinuclear region, optimizing the degradation of cellular cargo in autophagic and endocytic pathways. (ii) Dysfunction in the KIF1C motor, which may include mutations associated with hereditary spastic paraplegia (HSP), leads to significant perinuclear clustering of lysosomes. This phenomenon is primarily due to the aberrant activation of dynein-mediated lysosomal transport, which alters the normal lysosomal distribution and potentially impairs their functional capacity. (iii) KIF1C knockout results in the dispersion of lysosomes throughout the cell, due to a compromised interaction between the dynein-dynactin-Hook3 complex and lysosomes. This disruption in lysosomal transport interferes with normal lysosomal degradation processes in autophagic and endocytic pathways.

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Previous studies have reported that RUFY3 can interact with dynein either directly or indirectly through JIP3/416,17. Additionally, other proteins like TMEM55B and DENND6A have also been shown to link lysosomes to dynein18,19. These findings, coupled with our own, raise the question of why there are multiple mechanisms for dynein-driven lysosomal motility. One possible explanation is that multiple adapters are required to recruit enough dynein motors to overcome the force exerted by a kinesin, equivalent to the strength of eight dynein-dynactin complexes48. Alternatively, different adapters might be required in various physiological conditions, such as during nutrient starvation or under oxidative stress, to ensure lysosomal accumulation in the perinuclear region, a process that is critical for efficient cellular response11,19. Our results suggest that KIF1C, through its interaction with Hook3, may enhance the recruitment of dynein to the lysosomal transport system and fine-tune its activity, thus regulating lysosomal positioning within the cell. Exploring how KIF1C influences the formation of other adapter-dynein complexes is essential to fully understand its specific role in regulating dynein-driven lysosomal transport.

This study further elucidated the role of KIF1C in autophagic and endocytic pathways. During conditions of nutrient starvation, lysosomes typically cluster around the nucleus, facilitating their fusion with mature autophagosomes and enhancing autophagic efficiency11. However, our observations indicated that the loss of KIF1C does not significantly affect the fusion between autophagosomes and lysosomes. Instead, the disrupted autophagic flux in _KIF1C_-KO cells was primarily attributed to an impairment in the degradation of autolysosomal contents, not in the initial fusion process. Additionally, the observed reduction in endocytic degradation capacity in _KIF1C_-KO cells indicated a broader role of KIF1C in lysosomal function. Currently, it is unclear how KIF1C regulates the lysosomal function. It has been reported that the balance between minus-end and plus-end motility of lysosomes is crucial for lysosome tubulation, which serves as a platform for lysosome reformation, a process essential for replenishing the functional lysosome pool49,50. Therefore, the absence of KIF1C may disrupt this balance, impeding lysosome reformation and consequently impairing lysosomal degradative capacity.

In recent years, mutations in KIF1C have been found in patients with complex forms of hereditary spastic paraplegia (HSP) or cerebellar dysfunction, underscoring its critical role in neuronal function37,51,52,53. Notably, most of these mutations are located within the motor domain, although the exact mechanisms by which they contribute to disease pathology remain elusive. In neurons, soma-derived degradative lysosomes are continuously delivered to distal axons to maintain local degradation capacity in axons54. It is hypothesized that mutations associated with HSP in KIF1C impair its motor activity. This impairment could facilitate the activation of retrograde lysosomal transport via the dynein-dynactin-Hook3 pathway, as depicted in Fig. 6ii, thereby preventing their delivery to distal axons. Such a scenario would likely lead to the accumulation of autophagosomes and pathological cargoes within distal axons. In fact, lysosomal and autophagic pathways have been linked to various forms of HSP and other neurodegenerative diseases55,56.

In conclusion, our study reveals an unconventional role of KIF1C in activating dynein-driven lysosomal transport, independent of its traditional motor functions. This study establishes KIF1C as a unique player in cellular transport, performing dual roles: it facilitates anterograde transport of various cargoes as a conventional motor protein, and it uniquely supports retrograde lysosomal transport in a motor-independent manner. This latter role is critical for efficient degradation during autophagy and endocytosis. Understanding the regulatory mechanisms governing the dual functions of KIF1C is essential for deepening our insights into its role in cellular dynamics and its implications in neurodegenerative disorders.

Methods

Plasmids, siRNAs, and preparation of anti-GFP and -RFP nanobodies

Expression plasmids for human KIF1C-Clover and KIF1C-mCherry were constructed as described previously30. The cDNAs encoding KIF1C lacking amino acids (aa) 1–349 (∆motor), aa 810–1103 (∆810–1103), aa 600–1103 (∆600–1103), and aa 794–807 (Δ794–807) were amplified by PCR and subcloned into pClover-N1 vector. A lysine-to-alanine substitution at aa 103 (K103A) within the motor domain was introduced into KIF1C-Clover by site-directed mutagenesis. The cDNA encoding human TMEM55B was isolated from PC9 cells and subcloned into pEGFP-C1 vector. Expression plasmid for human RUFY3 in the pEGFP-C1 vector was kindly provided by S. Arai (Fukushima Medical University, Fukushima, Japan). Expression plasmid for GFP-LC3 was kindly provided by J. Lippincott–Schwartz (Janelia Research Campus, Ashburn, VA, USA). Expression plasmids for LAMP1-mScarlet (#98827) and the Hook3 cDNA (#198525) were purchased from Addgene. The cDNAs encoding full-length Hook3 and its truncated version lacking aa 553–718 (Δ553–718) were subcloned into pmCherry-C1 vector. The sequences of the respective siRNAs used were as follows: si_DHC_; 5′-(GAGCATGTCTTTACAGATCTT)-3′; and si_BICD2_; 5′-(GGAGCUGUCACACUACAUGTT)-3′. si_Hook3_#1(SASI_Hs01_0015178), si-Hook3#2(SASI_Hs01_0015179), and MISSION® siRNA Universal Negative Control #1 (siCtrl) were purchased from Sigma-Aldrich (St. Louis, MO, USA). E.coli BL21(DE3) cells harboring pGEX6P1-GFP-Nanobody and pGEX6P1-mCherry-Nanobody plasmids (gifts from K. Nakayama at Kyoto University, Kyoto, Japan)57 were cultured in the presence of 0.1 mM IPTG for 4 h at 30 °C to induce the expression of GST-tagged anti-GFP and -RFP nanobodies, respectively. The resulting GST-fusion proteins were purified using glutathione–Sepharose 4B beads (GE Healthcare. Chicago, IL, USA).

Cell culture and transfection

U2OS (ATCC, Manassas, VA), HeLa-S3 (Japanese Collection of Research Bioresources Cell Bank, Osaka, Japan), and HEK293T cells (RIKEN BRC, Tsukuba, Japan) were cultured in DMEM (Nacalai Tesque, Kyoto, Japan) containing 10% (v/v) FBS (growth medium). Transfections were performed using Lipofectamine 3000 (Thermo Fisher Scientific, Waltham, MA, USA) or PEI-Max (Polyscience, Inc., Warrington, PA, USA) for plasmids, and RNAiMAX (Thermo Fisher Scientific) for siRNAs, following the manufacturer’s instructions.

Generation of _KIF1C_-KO cells

_KIF1C_-deficient U2OS and HeLa-S3 cells were produced using the Alt-R CRISPR-Cas9 system (Integrated DNA Technologies [IDT], Coralville, IA, USA) following the manufacturer’s protocol. Briefly, U2OS and HeLa-S3 cells were transfected with an RNP complex composed of Cas9 nuclease V3 (IDT) and the following predesigned gRNAs against the KIF1C gene (IDT): KIF1C#1 (Design ID: Hs.Cas9.KIF1C.1.AA), KIF1C #2 (Design ID: Hs.Cas9.KIF1C.1.AD) or negative control guide RNA#1 (IDT). After transfection, monoclonal U2OS and HeLa-S3 cells were obtained through limiting dilution, and successful knockout was confirmed by immunoblotting.

Antibodies and reagents

Following antibodies were obtained commercially: anti-KIF1C (A301-027A, BETHYL, Montgomery, TX, USA), anti-Hook3 (15457-1-AP, Proteintech, Rosemont, IL, USA), anti-γ-tubulin (ab11316, Abcam, Cambridge, MA, USA), anti-horseradish peroxidase-conjugated anti-α-tubulin (PM054-7, MBL), anti-GFP (598, MBL), anti-RFP (165-3, MBL), anti-EEA1 (M176-3, MBL), anti-GM130 (PM061, MBL), anti-Tom20 (sc-17764, Santa Cruz, Dallas, TX, USA), anti-LAMP1 (sc-20011, Santa Cruz), anti-DHC (DYNC1H1) (12345-1-AP, Proteintech), anti-BICD2 (ab117818, Abcam), anti-LC3B (L7543, Sigma-Aldrich), anti-PDGFRβ (13449-1-AP, Proteintech). Bafilomycin A1 was purchased from LKT Laboratories (Minneapolis, MN, USA). LysoTracker Red DND-99 (L7528), Alexa Fluor 647-dextran (D22914), DQ-Green BSA (D12050), and BODIPY-FL LDL (L3483) were purchased from Thermo Fisher Scientific. Biotinylated BSA (A8549) was purchased from Sigma-Aldrich. PDGF-BB was purchased from FUJIFILM Wako Pure Chemical Corporation (Osaka, Japan).

Immunoprecipitation and immunoblotting

Cells were lysed in ice-cold lysis buffer (50 mM Tris-HCl, pH7.4, 0.5% (v/v) Nonidet P-40, 150 mM NaCl, 5 mM EDTA, 50 mM NaF, 1 mM Na3VO4, 1 mM _p_-phenylmethylsulfonyl fluoride, 10 μg/ml leupeptin, and 10 μg/ml aprotinin). For immunoprecipitation, the GST-tagged anti-GFP or -RFP nanobody beads were mixed with cell lysates. The resultant samples were subjected to SDS-PAGE and transferred to polyvinylidene difluoride membrane filters (Immobilon-P; Merck, Darmstadt, Germany). The membranes were immunoblotted with the respective antibodies, and the bound antibodies were visualized with horseradish peroxidase-conjugated secondary antibodies using chemiluminescence reagent (Immobilon ECL Ultra Western HRP Substrate; Merck).

Immunofluorescence, imaging, and quantification

Cells untransfected or transfected with expression plasmids and/or siRNAs were seeded on coverslips (18 mm ø) and cultured. Cells were fixed with 4% (w/v) paraformaldehyde, permeabilized with 0.2% (v/v) TritonX-100, and then blocked with 5% (w/v) BSA. Blocked cells were stained with primary antibodies overnight at 4 °C and then Alexa Fluor-conjugated secondary antibodies and DAPI for 1 h at room temperature. To detect colocalization of mCherry-RUFY3 with KIF1C-Clover and Hook3 (shown in Fig. S5), free cytosolic proteins were removed by permeabilizing cells with 50 μg/ml digitonin in Hepes buffer (20 mM HEPES, pH7.4, 110 mM potassium acetate, 2 mM magnesium acetate, 5 mM sodium acetate, 0.5 mM EGTA) for 5 min at 4 °C before fixation with 4% (w/v) paraformaldehyde. Fluorescence images were obtained using A1 confocal microscope (Nikon, Tokyo, Japan) with 60x/1.45 and 100x/1.49 oil immersion objective lenses and Andor Technology-iXonEM EMCCD camera and processed using Fiji software (National Institutes of Health, Bethesda, MD, USA). Dotted lines in microscopy images represent the cell periphery.

To assess lysosomal distribution, serial optical z sections through the cells, stained with anti-LAMP1 antibody and DAPI, were collected and stacked. Lysosomal distribution was quantified from the stacked images by using Fiji. A single cell was selected along the periphery using a freehand tool, and neighboring cells were cleared. A line outlined the nucleus was incremented by 5 μm till the cell periphery as shown in Fig. 1C. LAMP1 intensities in perinuclear (0–5 μm) and periphery ( > 15 μm) regions were normalized by the whole-cell LAMP1 intensity. Distribution of early endosomes and mitochondria, based on EEA1 and Tom20 intensities, respectively, was quantified using the same methodology. Number of Golgi fragments was quantified by using the particle analysis tool of Fiji. Colocalization was analyzed by quantifying the percentage of total intensities from one fluorescence channel that colocalized with another per cell using Fiji. Pearson’s correlation coefficient was measured using the Coloc2 plugin in Fiji.

Endocytic degradation

Activity of endocytic degradation within lysosomal compartments was quantitatively assessed using a fluorescence assay with dye-quenched DQ-Green, which fluoresces upon degradation in lysosomes58. Cells were incubated with 10 μg/mL DQ-Green BSA and 1 μM LysoTracker Red DND-99 in growth medium for 1 h, fixed with 4% paraformaldehyde, and examined via confocal microscopy. Colocalization of DQ-Green-BSA and LysoTracker Red fluorescence was confirmed, and fluorescence intensity of DQ-Green-BSA was quantified using Fiji software. To assess the endocytic degradation of BSA by immunoblotting, cells were incubated in serum-free DMEM containing 10 μg/mL biotinylated BSA, with or without 100 nM bafilomycin A1, for 1 h. Following incubation, cell lysates were collected and analyzed by Western blotting using horseradish peroxidase-conjugated streptavidin. To assess the endocytic degradation of PDGFRβ following ligand stimulation, cells were serum-starved for 24 h and then treated with 20 ng/mL PDGF-BB in the presence of 50 μg/mL cycloheximide, with or without 100 nM bafilomycin A1. After incubation, cell lysates were collected and analyzed by immunoblotting.

Endocytic trafficking of dextran and LDL

For dextran trafficking, cells were incubated with 1 μM LysoTracker Red DND-99 in growth medium for 1 h at 37 °C. Cells were further incubated with 0.2 mg/ml Alexa Fluor 647-dextran for 30 and 120 min at 37 °C. For LDL trafficking, cells were loaded with 7.5 μg/ml BODIPY-FL LDL together with 1 μM LysoTracker Red DND-99 in serum-free DMEM for 1 h at 37 °C. Cells were then washed and incubated in growth medium for 0, 30 and 120 min at 37 °C. At the end of the incubation period, cells were washed with PBS and fixed. Confocal images were collected and analyzed for colocalization of dextran or LDL with LysoTracker Red DND-99.

Autophagy assay

Autophagic flux was assessed by monitoring the levels of LC3B-II in cells treated with 100 nM bafilomycin A1 or DMSO in growth medium. For starvation treatment, cells were gently washed three times with PBS and cultured in Hank’s Balanced Salt Solution (HBSS; 14025-092, Thermo Fisher Scientific). Cells were lysed and assessed by immunoblotting as described above. To quantify the fusion between autophagosomes and lysosomes, cells were transfected with GFP-LC3 plasmid, cultured for 48 h, and incubated with 1 μM LysoTracker Red DND-99 for 1 h. Cells were then starved in HBSS containing 100 nM bafilomycin A1 for 2 h, followed by fixation. Confocal images were collected and analyzed for colocalization of GFP-LC3 with LysoTracker Red DND-99.

Statistics and reproducibility

All statistical analyzes were performed using Prism 9.0 (GraphPad Software, Boston, MA, USA), and p values were calculated using two-tailed Student’s _t_-test for two-group comparisons and one-way ANOVA followed by the Tukey’s test for multiple data set comparisons. All statistical analyzes were conducted with a significance level of α = 0.05 (p < 0.05). All experiments were conducted at least three times. Superplots were generated as described by Lord et al.59, displaying individual data points (small dots) and the corresponding mean values (large circles) for each experiment, with each experiment’s data color-coded. Mean values between conditions were statistically compared using the tests specified in the figure legends. The bars in the superplots represent the mean ± SD derived from the means of each replicate.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

Data reported in this study are available in the article and the supplementary material. The source data file underlying all graphs can be found in Supplementary Data. Uncropped Western blot images can be found in Supplementary Fig. S9. All other data are available from the corresponding author on reasonable request.

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Acknowledgements

We thank Drs. S. Arai (Fukushima Medical Univ.) for pEGFP-RUFY3 plasmid, J. Lippincott–Schwartz (Janelia Research Campus) for pEGFP-LC3 plasmid, and K. Nakayama (Kyoto Univ.) for pGEX6P1-GFP-Nanobody and pGEX6P1-mCherry-Nanobody plasmids. This work was supported in part by a grant from JST Moonshot R&D [JPMJMS2022 (Y.M.); JPMJMS2025-14 (Y.O.)], JST CREST [JPMJCR20E2 (Y.O.)], the Japan Agency for Medical Research and Development (AMED) [18gm5010001s0901 (Y.M.)], and MEXT/JSPS KAKENHI [23K14599 (T.S.); 23K06678 (M.N.); 19H05794 & 19H05795 (Y.O.)] from MEXT.

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Authors and Affiliations

  1. Department of Biochemistry, Fukushima Medical University School of Medicine, Fukushima, Japan
    Takeshi Saji & Michiru Nishita
  2. Division of Cell Physiology, Department of Physiology and Cell Biology, Graduate School of Medicine, Kobe University, Kobe, Japan
    Mitsuharu Endo & Yasuhiro Minami
  3. Department of Cell Biology, Graduate School of Medicine, The University of Tokyo, Tokyo, Japan
    Yasushi Okada
  4. Laboratory for Cell Polarity Regulation, RIKEN Center for Biosystems Dynamics Research (BDR), Osaka, Japan
    Yasushi Okada
  5. Department of Physics, Graduate School of Science, The University of Tokyo, Tokyo, Japan
    Yasushi Okada
  6. Universal Biology Institute (UBI) and International Research Center for Neurointelligence (WPI-IRCN), The University of Tokyo, Tokyo, Japan
    Yasushi Okada

Authors

  1. Takeshi Saji
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  2. Mitsuharu Endo
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  3. Yasushi Okada
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  4. Yasuhiro Minami
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  5. Michiru Nishita
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Contributions

T. Saji performed data collection and analysis; T. Saji and M. Nishita conceived the study and wrote the manuscript. Y. Okada developed experimental tools and provided critical feedback on the data interpretation and the manuscript. M. Endo and Y. Minami provided critical suggestions and helped to write the manuscript.

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Correspondence toMichiru Nishita.

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Communications Biology thanks Frederic Darios and the other, anonymous, reviewer(s) for their contribution to the peer review of this work. Primary Handling Editor: Manuel Breuer. A peer review file is available.

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Saji, T., Endo, M., Okada, Y. et al. KIF1C facilitates retrograde transport of lysosomes through Hook3 and dynein.Commun Biol 7, 1305 (2024). https://doi.org/10.1038/s42003-024-07023-6

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