Mechanosensitive dynamics of lysosomes along microtubules regulate leader cell emergence during collective cell migration

Domain: physics
Published: 2025-12-21
Editorial outcome: major_revision

Abstract
Collective cell migration during embryonic development, wound healing, and cancer metastasis requires the emergence of leader cells at the migration front. Despite their physiological relevance, the full mechanisms underlying the emergence of leader cells remain elusive. Here we report that leader cells display a peripheral accumulation of lysosomes in diverse model systems for wound healing, including cultured epithelial monolayer, mouse embryonic skin, and Drosophila embryos. This accumulation involves cellular contractile forces driving lysosomal transport along microtubules towards the leading edge. Indeed, we control leader cell emergence by manipulating lysosomal movement on microtubules. We further find that peripheral lysosomes associate with Rac1 molecules at the leading periphery, regulating local Rac1-activity, triggering actin polymerization and promoting lamellipodium formation. Taken together, we demonstrate that beyond their catabolic role, lysosomes act as an intracellular platform that links mechanical and biochemical signals to control the emergence of leader cells.

Introduction
Collective cell migration is the coordinated movement of cells, which is essential for organogenesis, embryo development, wound healing, tissue regeneration, and cancer metastasis 1–13. For successful migration of epithelial cell collectives, a few cells at the migration front undergo phenotypic transformation to form distinct lamellipodial structures and emerge as leader cells 3,14. These cells provide guidance to collective cell migration 13,15,16 and are critical for epithelial wound healing 2,17, branching morphogenesis 18,19, and cancer metastasis 1,11,20,21. Despite this physiological importance, what factors selectively designate only a few cells at the migration front as the leader cells remain largely unknown. Nevertheless, recent works have suggested that both mechanical and biochemical cues are crucial for this process 22,23. Mechanical cues, such as the spontaneous fluctuations in the cellular force field, which is defined by the cell-cell and cell-matrix forces, influence the leader cell formation 22. Prior to the appearance of characteristic lamellipodial structures in leader cells, elevated traction forces and tensile stresses appear behind the would-be leader cells 22. These results suggest a non-cell autonomous regulation of leader cell emergence. In addition to this mechanical regulation, spatial patterns of cellular signaling proteins such as p120-catenin 24, Notch-Dll4 25, and RhoA 23 distinguish the leader cells from other non-leader cells at the migration front and the follower cells behind the leaders. For example, the intrinsic polarized activity of Rac and RhoA GTPases is an integral signature of leader cell emergence 23. Cues for leader cell emergence can also come from the extracellular milieu through the interaction of the migrating cells with the compliant extracellular matrix via focal adhesions 3 and with the soluble signaling via self-generated chemokine gradient 26. Furthermore, a recent study has identified the tumor suppressor protein p53 as a key marker for leader cells 2. Elevated levels of p53 guide leader cell emergence and modulate the p21-CDK pathway to regulate collective migration 2. But a connection between the mechanical cues and the biochemical events leading to leader cell emergence remains elusive.

As the sedentary cells become motile and develop a clear front-to-rear polarity, the intracellular organization undergoes a conspicuous transformation 3. This transformation requires both breaking the existing structures and making new structures. In this respect, previous studies have primarily focused on elucidating the dynamics of the actin cytoskeleton during the emergence of leader cells. However, the role of other intracellular structures and cellular organelles in this process remains unexplored. Relevantly, one may presume that lysosomes, which are crucial for the catabolism and recycling of various biomolecules, might play a critical role in the cellular transformation to a motile state. Indeed, lysosomes can act as a signaling hub regulating cell growth, death, and differentiation 27. They also house various proteins which regulate cell adhesion 28,29 and migration 30. At a single-cell level, lysosomes regulate the dynamics of focal adhesions in migrating fibroblasts 29. During dendritic cell migration, lysosomes accumulate in the direction opposite of migration and locally release calcium to modulate actomyosin dynamics supporting cell propulsion 31. Lysosomes also respond to the mechanical properties of the extracellular matrix by altering their intracellular position 32. Interestingly, a parallel study demonstrates that peripheral lysosomes promote invadopodium-based matrix degradation and invasion of melanoma cells, highlighting the importance of lysosome positioning in carcinogenesis 33. While it seems that lysosomes can integrate mechanical signals and biochemical signals, very little is known about the specific roles of this organelle in collective cell migration.

In this work, we explore the roles of lysosomes in the emergence of leader cells during collective cell migration during epithelial wound healing (or gap closure) and show that the peripheral accumulation of lysosomes facilitates leader cell emergence from the epithelial wound margin. Importantly, previous work has established that peripheral lysosomes contribute to cancer cell invasion primarily through exocytosis and ECM degradation 34. In contrast, our study reveals a mechanosensitive role of lysosomes in collective epithelial migration, demonstrating that their peripheral accumulation is not just a consequence of motility but an active driver of leader cell emergence. We also show that lysosomes regulate Rac1 activation and lamellipodium formation, integrating mechanical cues with biochemical signaling to coordinate epithelial wound healing and morphogenesis. These findings redefine lysosomes as key mechanotransductive organelles that control collective migration beyond their conventional degradative functions.

Results
Lysosomes accumulate at the leading periphery of the leader cells in various wound healing models
The biochemical, molecular, and cellular mechanisms underlying collective cell migration during the wound healing process have been studied employing both in vivo and ex vivo model systems. For instance, stage-15 Drosophila melanogaster embryos have been valuable in vivo models 35, while mouse embryonic skin wounds have been utilized as ex vivo systems 36,37. It is known that the epidermal wound closure in stage-15 Drosophila embryos occurs by a combination of pursed-string contraction of actin cables and lamellipodial crawling 35.

First, to study the localization dynamics of lysosomes in wound healing, we used glass needles (0.5 µm inner diameter and 100 µm outer diameter) mounted on a femtojet microinjection system to create wounds in the stage-15 embryos. Since the wound got closed completely in one-to-two hours, we fixed these embryos 15 min post-wounding to capture an intermediate stage of wound closure (Supplementary Fig. 1A). We then stained these embryos for lysosomes and actin, using an antibody against an established lysosome marker protein, LAMP1, and fluorescence-labeled phalloidin, respectively (Supplementary Fig. 1B) and observed a few cells at the migration front emerging as leader cells having lamellipodial structures (Supplementary Fig. 1B). Interestingly, the images revealed a distinct lysosomal accumulation at the cell periphery in these cells (Supplementary Fig. 1B). Next, we analysed the localization of lysosomes in mouse embryonic skin wounds. We subjected the mouse embryonic skin samples that were dissected out of day 16.5 embryos, to incisions and allowed the wound to heal for 30 min in an air-media interface in a humidified chamber maintained at 37 °C and 5% CO2 (Fig. 1A–C). We then fixed and stained these samples for lysosomes and actin (Fig. 1D, Supplementary Fig. 1C). Here, we also observed the movement of cells into the wound gap of the skin tissue (Fig. 1C) and peripheral lysosome distribution at the wound margin (Fig. 1D, Supplementary Fig. 1C). Although challenging, super-resolution imaging of mouse embryonic skin wounds revealed leader cells within the multilayered tissue. The leader cells with an extended lamellipodia have lysosomes polarized at the cell front, while a non-leader marginal cell did not show any peripheral lysosomal polarization (Fig. 1E). These results suggest that lysosome accumulation at the leading edge of wounds occurs in physiologically relevant model systems and may represent an evolutionarily conserved mechanism in collective cell migration.

However, the complex 3D architecture of these models makes it challenging to track individual leader cells and precisely quantify lysosome dynamics. To investigate the cellular mechanisms underlying lysosomal redistribution and leader cell emergence, we transitioned to a well-established gap closure assay 2,8,12,22,23,38 using cultured epithelial monolayers. This model provided precise single-cell tracking and enabled rigorous quantifications, allowing for a more detailed mechanistic analysis of lysosome positioning during collective migration. We grew Madin-Darby canine kidney (MDCK) epithelial cells to confluence into the wells of a migration chamber (Supplementary Fig. 1D). In this system, after lifting off the confinement, cells migrated as two opposing epithelial sheets. Leader cells emerged from the collectively migrating monolayer (Supplementary Fig. 1E) approximately two hours after confinement lift-off. These cells, positioned at the forefront of the migrating sheet, extended large, actin-rich lamellipodia, which served as a key morphological feature for their identification 25. To further investigate the dynamics of lysosomes in migrating epithelial monolayers, we fixed the migrating epithelial monolayer samples at 0, 15, 30, 60, 120, and 240 min post-confinement lift-off and immunostained them for lysosomes and actin (Supplementary Fig. 2A). We observed that in the cells at the migration front (marginal cells), lysosomes accumulated at the cell periphery, whereas the cells in the follower layer did not show any change in lysosome localization (Supplementary Fig. 2A) indicating that lysosome accumulation at the cell periphery in cells at the wound margin increases in a time-dependent manner. Interestingly, among the marginal cells, we observed further enhanced lysosome accumulation at the leading periphery of the leader cells, which was marked by their extended lamellipodial structures (Fig. 1F, G). We subsequently computed the fraction of peripherally localized lysosomes relative to the total lysosome intensity in a cell and reported it as Peripheral Occupancy Fraction (POF) (Fig. 1H). Our method of measuring POF takes into account the area of the cell, as the fraction is calculated by dividing the integrated densities (mean intensity * area). This quantitative analysis revealed a progressive increase in POF for lysosomes in cells at the margin as the migration progresses with time (Supplementary Fig. 2B). Further, a comparison of POF between the leader and non-leader marginal cells revealed a significantly higher fraction of lysosomes localized to the cell periphery in leader cells than in non-leader cells (Fig. 1I). Also, a closer examination of the distribution of lysosomes in leader and non-leader marginal cells revealed a lower junctional lysosome pool in leader cells as compared to non-leader marginal cells (Fig. 1J). In leader cells, lysosomes predominantly localize to the growing lamellipodia in the direction of migration, with minimal alignment along cell-cell junctions. In contrast, non-leader marginal cells exhibit lysosome enrichment along the actin cytoskeleton, but a significant fraction aligns with cell-cell junctions (Fig. 1G). This junctional occupancy was quantified as the fraction of lysosomal intensity at junctions relative to the total intensity (Fig. 1J). Lysosome accumulation at the cell periphery was measured using the POF analysis for leaders and non-leader marginal cells in mouse embryonic skin wounds (Fig. 1K). In our mouse embryonic skin wound assays, we observe a significantly higher lysosome accumulation at the cell periphery in leader cells compared to non-leader marginal cells. To further quantitatively validate lysosome accumulation to the cell periphery in leader versus non-leader marginal cells in the two-dimensional gap closure assay, we analyzed the images using a custom-written program that measures the mean distance of the weighted center of mass (organelle intensity) of the organelle from the cell edge. We called this parameter Mean Distribution from Edge or MDE. It revealed the distance of the organelle of interest, in our case, lysosomes, from the migrating, gap-exposed edge of the cell. Analysis of lysosomal distribution using MDE revealed that lysosomes are positioned closer to the cell edge in leader cells compared to non-leader marginal cells, indicating a greater peripheral accumulation of lysosomes in leader cells. (Supplementary Fig. 2C, left panel). Next, we quantified the asymmetry of lysosome distribution in collectively migrating MDCK cells at zero- and two-hours post-confinement lift-off. To measure this, we identified the long axis of each cell along the direction of migration (perpendicular to the wound edge) and divided the cell at its midpoint along this axis. We then measured the intensity of the lysosomal marker LAMP1 in the front and rear halves of the cell using FIJI (ImageJ) to assess asymmetrical lysosome distribution. Asymmetry analysis revealed a significant front-rear asymmetry in lysosome distribution in collectively migrating MDCK cells, particularly at two hours post-confinement lift-off, further reinforcing the occurrence of polarized lysosome positioning during migration and leader cell emergence (Supplementary Fig. 2C, right panel). To test the generality of our findings among cultured epithelial monolayer models, we next repeated the measurements in a murine mammary epithelial cell line, EpH4-Ev. Enhanced peripheral localization of lysosomes in leader cells also emerged in this model (Supplementary Fig. 2D, E). Since leader cell emergence and lysosomal accumulation in these cells occurred at a similar timescale, we wondered how these dynamics behaved at the onset of collective migration in epithelial monolayers. Subsequently, we asked whether there might be any correlation between lysosome accumulation at the cell periphery in the marginal cells and collective cell movement. The line plot of two events (Supplementary Fig. 2F) indeed showed a temporal correlation between collective cell movement and POF for lysosomes with different migration time points. As the epithelial cells migrated progressively, we observed an increase in lysosome accumulation starting at T-60 min onwards and peaking at T-120 min post-confinement lift-off.

Finally, to understand how the lysosome dynamics are spatially correlated to actin dynamics within leader cells, we performed live-cell imaging of LifeAct-eGFP-expressing MDCK cells having Dextran-647 (Alexa Fluor 647-labeled Dextran) pre-labeled lysosomes (Fig. 2A, Supplementary video 1). Temporal color-coded map shows lysosome localization and initiation of lamellipodia as coordinated events (Fig. 2B). Live cell imaging of LifeAct-eGFP expressing MDCK cells having Dextran-647 labeled lysosomes, revealed differential lysosome dynamics in leaders and immediate non-leader marginal cells (Fig. 2C). Lysosomes in emerging leader cells showed peripheral accumulation followed by dispersal, faster bi-directional movement and subsequently, the lamellipodial protrusions started emerging (Fig. 2C, leader cell). Whereas in non-leader marginal cells, the lysosomes did not move dynamically as the migration progressed (Fig. 2C, marginal cell). Importantly, the live-cell imaging of these cells clearly showed that lamellipodia extension happens in lysosome-rich zones (Fig. 2D, E, and F-top panel, and Supplementary video 2). In contrast, the actin structure was retracted from lysosome-deficient regions (Fig. 2F, bottom panel, Supplementary video 2). The line plot in Fig. 2G shows the correlation of lamellipodia growth and lysosome accumulation. The increase in lamellipodia area and lysosome enrichment is positively correlated (Fig. 2G, Top-panel), while lamellipodia area decreases when there is little to no change in lysosome accumulation (Fig. 2G, Bottom-panel). Taken together, these results revealed that lysosome accumulation at the periphery of the leader cells is a ubiquitous event during collective cell migration across different model systems and is spatiotemporally correlated with the initiation and extension of lamellipodial structures in the leader cells.

Lysosome transport on microtubule tracks drives its peripheral accumulation
Next, we asked whether the peripheral accumulation of lysosomes in marginal cells, including the leader cells, was a result of increased lysosome biogenesis or of an escalated anterograde trafficking of the organelle. Considering that nuclear localization of Transcription Factor EB (TFEB) is an indicator of enhanced lysosome biogenesis 39,40, we examined TFEB localization. However, we did not observe TFEB accumulation in the nuclei of marginal cells at T-120 min post confinement lift-off (Supplementary Fig. 3A). We further quantified the number of lysosomes in the marginal and follower cells, before and during migration and did not observe any significant change (Supplementary Fig. 3B). These results eliminated the possibility of de novo lysosome biogenesis and pointed towards the polarized transport of lysosomes in leader cells. Previous studies have shown that microtubules are the tracks for bidirectional molecular motor-mediated transport of lysosomes 41. To gain insight into the organization of microtubules during collective epithelial migration, we performed a gap-closure assay with both MDCK and EpH4-Ev monolayers. We allowed the cells to migrate for 0 and 120 min and then fixed and immunostained them for microtubules with anti-Alpha-tubulin and lysosomes with anti-LAMP1. Microtubules show peripheral accumulation in marginal cells in both MDCK and EpH4-Ev cells (Fig. 3A, Supplementary Fig. 3C). Similar to computing the lysosomal POF, we also computed the POF (peripheral occupancy fraction) of microtubules to measure the microtubule accumulation at the cell periphery. We calculated the POF of microtubules by taking the ratio of the intensity of microtubules in the defined ROI to the total intensity of microtubules in the cell of interest. This analysis revealed a significant increase in POF of microtubules at T-2h post confinement lift-off, like that was observed for lysosomes (Fig. 3B). We hypothesized that disrupting microtubules would abolish the dynamic peripheral accumulation of lysosomes. We, therefore, disrupted the microtubules by using nocodazole at a concentration of 10 µM and observed its effect on the collective migration of MDCK cells (Supplementary Fig. 3D, middle panel and supplementary video 3). Nocodazole treatment led to the loss of peripherally localized lysosomes (Fig. 3C, middle panel), and at the same time, we observed a reduced emergence of leader cells upon nocodazole treatment (Fig. 3D, Supplementary Fig. 3D, middle panel). Microtubules are dynamic structures and constantly undergo cycles of assembly and disassembly at the plus end 42. Taxol (Paclitaxel) stabilizes the growing ends of microtubules, altering their dynamics. Treatment with 1 µM Taxol delayed collective migration of MDCK cells (Supplementary Fig. 3D, bottom panel; Supplementary video 3) and disrupted lysosome accumulation at the cell periphery, instead causing their even distribution throughout the cytoplasm of marginal cells (Fig. 3C, right panel). This disruption was accompanied by a significant reduction in leader cell emergence (Fig. 3D). We quantified POF for lysosomes in control, nocodazole-, and taxol-treated cells and found a significant drop in the peripheral accumulation of lysosomes in the drug-treated cells as compared to the control (Fig. 3E). Finally, we observed a temporally congruent polarization pattern for microtubules and lysosomes in MDCK cells, which were allowed to migrate for 0-, 15-, 30-, 60-, 120-, and 240-min post-confinement lift-off (Supplementary Fig. 3E).

We next investigated if cellular organelles such as late endosomes, early endosomes, endoplasmic reticulum (ER), and mitochondria, which also use microtubule tracks for their intracellular transport, show peripheral accumulation. We again allowed MDCK cells to migrate for 0- and 120-min post-confinement lift-off, fixed and immunostained them for late-endosomes (Anti-Rab7), early endosomes (Anti-EEA1), endoplasmic reticulum (Anti-KDEL), and mitochondria (Anti-Tom20) (Supplementary Fig. 4A–E). We found that none of the other organelles exhibited enhanced peripheral localization compared to lysosomes. A comparison of the POF for all organelles (Supplementary Fig. 4F) revealed that lysosomes showed the highest peripheral enrichment at 120-min post-confinement lift-off. These results indicated that lysosomal accumulation at the periphery is a specific, active process rather than a passive consequence of microtubule reorientation during collective cell migration.

However, nocodazole and taxol perturb microtubules, affecting multiple cellular processes and complicating the isolation of their specific role in lysosome relocalization during leader cell emergence. To overcome this, we used a strategy that selectively disrupts lysosome transport without altering microtubule structure. Downstream effectors of small GTPase Arl8b control the lysosome transport on the microtubule track. Among these proteins, lysosome adapter protein SKIP recruits the plus-end-directed kinesin motor to the lysosome 43. We, therefore, hypothesized that transfecting the cells with a kinesin-binding defective mutant of SKIP (SKIP WD->2XA) can specifically disrupt the lysosome dynamics without affecting the microtubule structure. Interestingly, we found reduced peripheral accumulation of lysosomes in cells expressing the mutant SKIP (Fig. 3F, G). The POF of lysosomes SKIP-WT was significantly higher compared to SKIP WD->2XA mutant expressing cells (Fig. 3H). Taken together, these results indicate that the peripheral accumulation of lysosomes in marginal cells is the result of a microtubule-mediated dynamic localization of lysosomes.

Peripheral accumulation of lysosomes is necessary for leader cell emergence
Next, we sought to examine whether the peripheral accumulation of lysosomes is necessary for the emergence of leader cells. We subsequently hypothesized that if that were the case, preventing lysosomes from moving to the cell periphery would reduce the propensity of leader cell emergence. Conversely, forcing lysosomes to the cell periphery would increase this propensity. To test these hypotheses, we resorted to abrogating the interaction between kinesin motor, KLC2 and SKIP using the chemical inhibitor kinesore 44 at a concentration of 12.5 µM. Lysosomes in MDCK cells treated with kinesore were repositioned to the perinuclear region compared to the control cells (Fig. 4A). The effect on lysosome positioning upon kinesore treatment was similar to the previous report 44, thus confirming the inhibitor activity in our system. The POF of lysosomes measured in cells treated with kinesore revealed a significant reduction in the peripheral pool of lysosomes as compared to the control cells (Fig. 4B). The collective movement of the kinesore-treated MDCK cells was delayed, leading to diminished gap closure, and fewer leader cells emerged as compared to the control (Supplementary Fig. 5A, B, Supplementary video 4). Next, we treated mouse embryonic skin with kinesore and subjected these kinesore-treated skins to incision wounds. We then allowed the cells to migrate for 30 min, fixed the samples, and immunostained for lysosomes, actin, and microtubules (Fig. 4C). Here, also lysosome polarization was also lost in kinesore-treated skin wounds as compared to the control. POF of lysosomes measured in kinesore-treated skin wounds revealed a significantly lower accumulation of the organelle at the cell periphery compared to control conditions (Fig. 4D).

Although the chemical inhibitor kinesore specifically abolishes lysosome transport on microtubule tracks by disrupting the interaction between KLC2 and SKIP, the effect is not limited to the cells at the migration margin. Hence, to precisely modulate lysosome distribution in cells at the migration margin and monitor its implications, we switched to a method that allowed us to manipulate lysosome localization in individual cells. To this end, we used the Reversible Association of Motor Protein (RAMP) system 45. In this system, the interaction between streptavidin bound to motor protein and streptavidin-binding protein (SBP) fused to an organelle-localizing protein drives the preferential localization of the target organelle. To this end, we generated stable cell lines expressing streptavidin-bound to motor protein, mCh-Kif5b*-Strep or Strep-Kifc1*-mCh (Supplementary Fig. 5C, D, mCh: mCherry). mCh-Kif5b*-Strep generates plus-end-directed trafficking, accumulating the target organelle at the cell periphery (Fig. 4E, F, respectively). In contrast, Strep-Kifc1*-mCh generates minus-end-directed trafficking, accumulating the target organelle near the cell nucleus (Fig. 4G, H, respectively). Further, we transiently transfected these cells with LAMP-1-SBP-GFP, targeting lysosomes, and seeded them in the migration chamber. By the interaction between Streptavidin and SBP, lysosomes were either localized to the cell periphery in mch-Kif5b*-Strep-expressing cells or the perinuclear region in Strep-Kifc1*-mCh-expressing cells (Fig. 4F, H). To validate the preferential positioning bias of lysosomes using the RAMP system, we quantified the POF of lysosomes for LAMP1-SBP-GFP positive-cells in either mch-Kif5b*-Strep-expressing or Strep-Kifc1*-mCh-expressing MDCK cells (Supplementary Fig. 5E). Using time-lapse imaging, we then followed the trajectories of LAMP1-SBP-GFP-expressing cells at the migration front (Fig. 4I, J, Supplementary videos 5, 6). Interestingly, LAMP1-SBP-GFP-expressing cells in mCh-Kif5b*-Strep background, which had peripherally localized lysosomes (Fig. 4E, F, I), showed a significantly increased propensity for leader cell formation than LAMP1-SBP-GFP-expressing cells in Strep-Kifc1*-mCh background, which had lysosomes mostly localized to the perinuclear region (Fig. 4G, H, J). Next, we calculated the probability of leader emergence in the two scenarios by dividing the number of co-transfected cells at the margin with lamellipodia (identified as leader cells) by the total number of co-transfected cells at the migrating edge. The probability of cells with peripheral lysosomes becoming leaders was 55% ± 3% (mean ± SD), while the probability of cells with perinuclear lysosomes was 7% ± 2% (Fig. 4K). Relevantly, the baseline probability was 20% ± 5% in mCh-Kif5b*-Strep and 16% ± 2% in Strep-Kifc1*-mCh motor-only controls (Fig. 4K). We also examined whether the positional bias of mitochondria and the endoplasmic reticulum (ER) toward the cell periphery or center influences leader cell emergence during collective epithelial migration. Stable MDCK cell lines expressing mCh-Kif5b*-Strep or Strep-Kifc1*-mCh were transfected with Tom20-GFP-SBP (for mitochondria) or VAPA-GFP-SBP (for ER), and leader cell emergence was quantified (Fig. 4K). Peripheral or perinuclear repositioning of mitochondria and ER had no significant effect on leader emergence compared to Kif-only controls (Fig. 4K), indicating that lysosomal positioning plays a unique role in this process. Moreover, we found that cells co-expressing mCh-Kif5B*-strep and LAMP1-SBP-GFP possessed prominent lamellipodial structures and mature focal adhesions oriented perpendicular to the migrating front (Fig. 4L, top panel). In contrast, cells co-expressing Strep-KifC1*-mCh and LAMP1-SBP-GFP possessed peripheral actin cables and less prominent focal adhesions oriented parallel to the migrating front (Fig. 4L, bottom panel). These results suggested that mCh-Kif5B*-strep expression and subsequent peripheral localization of lysosomes promoted leader-like cell morphology, while mCh-Kifc1*-Strep expression and subsequent perinuclear localization of lysosomes promoted follower-like cell morphology 22,23. We also compared the movement of cells transiently expressing either mCh-Kif5B*-strep+LAMP1-SBP-GFP or Strep-Kifc1*-mCh+LAMP1-SBP-GFP and observed no significant difference in cell motility between the two sets (Supplementary Fig. 5F). To further strengthen our findings, we monitored live collective cell migration in cells globally expressing LAMP1-SBP-GFP along with either Strep-Kifc1*-mCh or mCh-Kif5b*-Strep (Fig. 4M, Supplementary Fig. 5G, H). We observed the emergence of more pronounced leader cells in the LAMP1-SBP-GFP and mCh-Kif5b*-Strep condition, where lysosomes were globally biased toward the cell periphery, compared to the LAMP1-SBP-GFP and Strep-Kifc1*-mCh condition, which localized lysosomes toward the cell center (Supplementary Fig. 5G, H). The effect of globally manipulating lysosome distribution on leader cell emergence was quantified as the fraction of leader formation, as shown in Fig. 4M. These analyses together showed that while overall migration distance remains unchanged, cells with peripherally positioned lysosomes are more likely to form lamellipodia and emerge as leaders, whereas those with perinuclear lysosomes migrate within the cohort without extending lamellipodia. Leader emergence is thus a systems-level event 22,23, allowing compensation by other cells, but global disruption of lysosome transport, such as with kinesore, impairs this compensatory mechanism, leading to slower wound closure.

Finally, we obtained similar results using EpH4-Ev cells, where cells transiently transfected with mCh-Kif5B*-strep and LAMP1-SBP-GFP or Strep-KifC1*-mCh and LAMP1-SBP-GFP showed lamellipodium or actin cables, respectively (Supplementary Fig. 5I). Noticeably, abrogating lysosomal association with kinesin motor using the kinesin binding defective mutant of SKIP also showed a similar phenotype of reduced lamellipodial formation and hence leader emergence when compared to cells expressing wild type SKIP (Figs. 3F, G, 4N). In summary, we were able to regulate the propensity of leader cell emergence at the migrating front by altering lysosome localization. Collectively, these results suggested that peripheral localization of lysosomes is both necessary and sufficient for leader cell emergence.

Actomyosin contractility-generated cellular forces drive peripheral lysosome localization
In a cohesively migrating epithelium, leader cells emerge through an active mechanical process, where actomyosin contractility-generated cellular forces play critical roles 4,6,9,46. We, therefore, asked whether the peripheral accumulation of lysosomes in leader cells might be driven by the actomyosin contractility of a specific cell within the monolayer (Supplementary Fig. 7A). This experiment involved blue light-mediated photo-stimulation of cryptochrome 2 (CRY2)-bound catalytic domain of a RhoA-activator, ARHGEF11. CIBN, one of the binding partners of CRY2, was engineered to localize either at the plasma membrane or mitochondria. Upon photo-stimulation, the CRY2-CIBN association triggered plasma membrane (Supplementary Fig. 7A–C, Supplementary video 8) or mitochondrial localization (Supplementary Fig. 7E, F, Supplementary video 9) of the RhoA-activator. When localized to the plasma membrane, RhoA-activator increased the actomyosin contractility 49. We used the mitochondrial localization of the RhoA-activator as the negative control for this experiment.

Here, building on the leader-follower interactions observed during epithelial wound healing, we hypothesized that a highly contractile, photo-stimulated cell would generate pulling forces on its neighboring cells, similar to the tension exerted by contractile follower cells on leader cells in a migrating tissue 22,47. This idea stems from the notion that mechanical cues, such as the forces generated by actomyosin contractility, can influence the behavior of neighboring cells 22,47. We reasoned that in response to this mechanical tension, lysosomes in the adjacent cells would polarize toward the cell periphery, moving away from the contractile cell. This observation would suggest that lysosome repositioning is a mechanosensitive process, regulated by tension-mediated signaling rather than being a process exclusive to the leading edge of migrating cells (Supplementary Fig. 7A–C). Indeed, upon photo-stimulation, lysosomes in the cells adjacent to the excited cell moved toward the cell periphery, away from the stimulated cell (Supplementary Fig. 7C). However, in the negative control, where RhoA-activator localized to mitochondria, lysosomes in the cells adjacent to the stimulated cell did not undergo any such changes in localization (Supplementary Fig. 7E–G). One limitation of this experiment is that Opto-RhoA GEF activation induces a brief (~2 min) and localized contraction in a single cell, which distributes pulling forces across multiple neighbors. As a result, only subtle changes in lysosome positioning are observed, indicating that while mechanical forces can influence lysosome localization, this assay likely underestimates the magnitude of the effect seen during collective migration (Supplementary Fig. 7D, G). Consistent with this interpretation, the modest but reproducible outward shift of lysosomes in neighbors was observed only under membrane-targeted RhoA activation and not under the mitochondrial control, reinforcing that the assay does capture contractility-dependent polarization, albeit at a reduced magnitude. Thus, the opto-RhoA GEF results should be viewed as supportive, lower-bound evidence for force-driven lysosome repositioning rather than a full recapitulation of the robust polarization seen during collective migration. Nevertheless, together with the contractility inhibitor and cell doublet experiments, these results are consistent with the idea that forces transmitted through cell-cell interfaces can contribute to lysosome polarization. Lysosomes may therefore participate in coupling intercellular tension among follower cells to leader cell emergence.

Finally, we hypothesized that if the mechanical forces regulated peripheral lysosome localization during the emergence of leader cells, then any cue that spatially biases leader cell emergence should also spatially bias the peripheral localization of lysosomes. One such cue is wound geometry. Having highly positive-curved beaks at the migration front leads to the emergence of leader cells at the tip of the beak, due to strong force concentration at those points 38,50,51. Would such beaks show enhanced peripheral localization of lysosomes? To answer this question, we used a micropatterning approach to fabricate silicone stencils made of polydimethylsiloxane (PDMS). These stencils would create wounds with a single beak to allow one or two cells to have the geometrical cue to become the leader cells (Fig. 5H, Supplementary Fig. 7H). We then seeded cells in these micropatterned chambers, labeled lysosomes using Dextran-647, lifted off the confinement, and performed live-cell imaging. The cells seeded in these micropatterned chambers moved collectively and covered the gap (Supplementary Fig. 7I, Supplementary video 10). Upon lifting the stencil, we indeed observed enhanced lysosome accumulation at the tip of these beaks as compared to their sides (Fig. 5I, J, Supplementary video 11). At the same time, lamellipodial structures emerged out of the beak region while actin fibers ran along both sides (Fig. 5I). These results suggested that the spatial concentration of mechanical force in the epithelial monolayer correlates with the enhanced peripheral localization of lysosomes, supporting the premise of mechanosensitive localization of lysosomes during leader cell emergence. It also strengthened the correlation (Fig. 2E) between peripheral localization of lysosomes and lamellipodia formation through the modification of the actin cytoskeleton.

Rac1 GTPase hitchhikes on lysosomal membranes
We then asked whether the peripheral localization of lysosomes promotes lamellipodia formation by locally modifying the actin cytoskeleton. In migrating epithelial cells, actin dynamics depend on the localization and activation of small Rho GTPases, including Rho, Rac1, and Cdc42 3,10,23,52. These proteins switch between the GTP-bound active state and the GDP-bound inactive state. To determine whether any Rho family GTPases associate with lysosomes, we performed a screen by transiently transfecting MDCK cells with constitutively active (CA, GTP-locked) and dominant-negative (DN, GDP-locked) forms of RhoA, Cdc42, and Rac1. After allowing migration for two hours post-confinement lift-off, we immunostained lysosomes using anti-LAMP1 and analyzed their association (Supplementary Fig. 8A–G). Interestingly, we observed that Rac1-GTPase associates with lysosomes (Supplementary Fig. 8E, G). Other Rho family GTPases did not show any appreciable colocalization with lysosomes (Supplementary Fig. 8A–D). Also, the wild type (Rac1-WT) is majorly present on the plasma membrane, and a few punctate structures were observed, while Rac1-CA (Q61L) was primarily plasma membrane-localized, whereas Rac1-DN (T17N) was distributed as punctae, some of which were on lysosomes in both MDCK and EpH4-Ev epithelial cells (Supplementary Fig. 8E–I). Mander’s coefficient analysis between Rac1-WT, Rac1-CA, and Rac1-DN with the lysosomal membrane marker LAMP1 confirmed this localization pattern in both MDCK and EpH4-Ev cells (Supplementary Fig. 8H, I). Next, we examined the localization of endogenous Rac1 with lysosomes in collectively migrating MDCK cells. Endogenous Rac1 was majorly localized to the plasma membrane and a few punctate structures, some of which were LAMP1 positive (Fig. 6A, B). Co-localization of Rac1 and lysosome was recapitulated in our ex vivo mouse embryonic skin wound model system (Fig. 6C, D). To further validate our findings, we performed organelle fractionation from MDCK cells grown to 90% confluency using an OptiPrep density gradient and assessed Rac1 association across different organelle fractions (Fig. 6E). In the 15%–30% gradient, lysosomal signals were detected in fractions 1 and 3–7, with fraction 4 showing a strong lysosomal presence along with a faint mitochondrial contribution. Notably, endogenous Rac1 was predominantly found in the lysosome-enriched fractions 1 and 3, which were exclusively lysosomal, as well as in a few other fractions (Fig. 6E). Taken together, our experiments and data strongly point to the association of Rac1 with lysosomes in collective epithelial cells. To understand the dynamics of Rac1 and lysosome association, we performed live imaging of Dextran-647 labeled lysosomes and GFP-tagged Rac1 wild type overexpressed in MDCK cells. Time-lapse imaging of a leader-like cell, having protrusive lamellipodial structures, revealed a subpopulation of lysosomes positive for Rac1 (Fig. 6F, supplementary video 12), thus supporting our previous findings.

These findings led us to investigate whether lysosomes actively shuttle Rac1 to the plasma membrane or vice versa. To examine the dynamic association between lysosomes and Rac1, we employed a photoconversion strategy. We expressed mEos-Rac1-WT in MDCK cells with lysosomes labeled using Dextran-647. Upon 405 nm illumination, mEos fluorescence shifted from green to red, enabling real-time tracking of Rac1 dynamics. In this experiment, migrating mEos-Rac1-expressing cells were photoconverted near the growing lamellipodial protrusions, and the redistribution of photoconverted Rac1 was monitored (Fig. 6G, H). Live imaging and kymograph analysis revealed punctate structures forming from photoconverted mEos-Rac1. Notably, these puncta frequently merged with nearby lysosomes, showing significant colocalization (Fig. 6H, Supplementary video 13), suggesting an active interaction between Rac1 and lysosomes during migration. We also tracked the photoconverted Rac1-DN (T17N) and Rac1-CA (Q61L) proteins for their redistribution within the cell (Supplementary Fig. 9A–C). As depicted by kymographs, live-tracking revealed puncta formation events for photoconverted mEos-Rac1-DN. Interestingly, these punctate structures merged with lysosomes positioned nearby, showing colocalization between the two (Supplementary Fig. 9A, C, Supplementary video 14). In contrast, photoconverted mEos-Rac1-CA diffused throughout the plasma membrane, and puncta formation was negligible in this case (Supplementary Fig. 9B, C, Supplementary video 15). These results indicate that Rac1 hitchhikes on peripheral lysosomes, which may play an essential role in branched actin formation and lamellipodia growth in an emerging leader cell.

Next, we investigated the influence of lysosome localization on Rac1 activity. Interestingly, imaging actin in control and kinesore-treated mouse embryonic skin wounds already indicated a potential link between lysosome positioning and Rac1 activity. In control samples, actin was distributed in a dynamic network at the leading edge, whereas in kinesore-treated wounds, we observed the formation of a prominent actin bundle (Fig. 4C), a hallmark of low or absent Rac1 activity 23,52. This structural change correlated with a clear loss of lysosome polarization at the leading edge in kinesore-treated samples. However, to probe the Rac1 more directly, we measured the Rac1 activity with a FRET-based Rac1 sensor 51, which was expressed in ~80-85% of cells due to high transfection efficiency (Supplementary Fig. 9D). We measured Rac1 activity in the presence or absence of kinesore, which induces perinuclear lysosome accumulation. We also used NSC23766 (Rac1 inhibitor)-treated cells as a negative control. As expected, NSC-treated cells showed no significant FRET signal compared to controls, confirming the functionality of the Rac1 activity sensor (Fig. 6J). Next, we analysed the mean FRET index in control and kinesore-treated cells, yielding three key findings. First, at T-0 (quiescent state), Rac1 activity levels were similar between untreated control and kinesore-treated cells, with no significant difference (Fig. 6I, K). Second, there was no significant change in Rac1 activity within kinesore-treated cells between T-0 and T-2h. Third, at T-2h, Rac1 activity was significantly lower in kinesore-treated cells compared to controls at the same migration time point (Fig. 6L). Together, these results indicate that peripherally localized lysosomes enhance Rac1 activity, linking force-induced lysosome accumulation at the cell periphery to lamellipodium formation and leader cell emergence during collective migration (Fig. 7). Interestingly, in migrating MDCK monolayers, Rac1+ lysosomes were frequently observed near early endosomes (Supplementary Fig. 9E–G), suggesting potential crosstalk between these compartments during leader cell emergence.

Methods
Cell culture
Madin-Darby canine kidney (MDCK) and EpH4 (ATCC; CRL-3063) epithelial cells were used in this study. Tetracycline-resistant wild-type MDCK (MDCK-WT) cell line was a gift from Yasuyuki Fujita. MDCK cells were cultured in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with GlutaMax (Gibco) with 5% fetal bovine serum (tetracycline-free FBS, Takara Bio) and 10 U mL−1 penicillin and 10 μg mL−1 streptomycin (Pen-Strep, Invitrogen) in an incubator maintained at 37 °C and 5% CO2, unless mentioned otherwise. EpH4-Ev cells (ATCC; CRL-3063) were cultured in Dulbecco’s modified Eagle’s medium supplemented with GlutaMax (Gibco) with 10% fetal bovine serum (tetracycline-free FBS, Takara Bio) and 1.2 μg ml−1 puromycin (Gibco) in an incubator maintained at 37 °C and 5% CO2.

MDCK cells stably expressing mCh-Kif5b*-Strep, Strep-KifC1*-mCh (mCh: mCherry), cells were transfected with the respective plasmids using Lipofectamine 2000 (Invitrogen) following the instructions provided by the manufacturer. 16–18 h-post-transfection, cells were seeded via serial dilution in a 96-well plate, such that each well had a single cell. Forty-eight hours post-cell seeding, the cells were subjected to selection media (DMEM-GlutaMax plus 5% FBS) containing 400 μg mL−1 of Geneticin (Invitrogen). The growth of single-cell-derived colonies was monitored over two weeks, following fluorescence confirmation. Colonies with a homogeneous expression of the fluorescence-tagged protein of interest were expanded and further maintained in 100 μg mL−1 Genetic containing media. All transient transfections were done using Lipofectamine 2000 following the instructions provided by the manufacturer. To obtain LAMP1-SBP-GFP and mCh-Kif5b*-Strep or Strep-KifC1*-mCh double-positive cells, LAMP1-SBP-GFP was nucleofected using the SE-Cell Line 4D-Nucleofector™ X Kit (V4XC-1024, LONZA), followed by selection for GFP-positive cells. mCh-Kif5b*-Strep or Strep-KifC1*-mCh was then expressed in the LAMP1-SBP-GFP–expressing cells, and the cells were sorted by FACS for GFP- and mCh-double positive populations. The resulting double-positive cells were seeded into iBidi migration chambers to form a monolayer, followed by live imaging to monitor their collective motion.

Cell migration assays were performed using adhesive biocompatible silicone culture inserts from iBidi. 5 × 104 cells suspended in an 80 μL cell culture medium were seeded in a culture insert stuck to glass-bottom dishes and incubated at 37 °C in a 5% CO2 humidified chamber. After 18 h of cell seeding, the cells formed a monolayer, and the culture insert was lifted to allow the cells to migrate. All migration experiments were carried out at 37 °C and in a 5% CO2 humidified chamber (standalone or stage-top incubator).

Drosophila melanogaster embryo microinjection wounding assay
For fixation and immunostaining, a previously described protocol was followed 59. Briefly, Stage 15 Drosophila melanogaster embryos were collected on an apple juice plate. These embryos were then aligned and stuck to a double-sided tape on a glass slide. They were left at room temperature for 5 min, followed by halocarbon oil application and subsequent wounding using glass needles mounted on an Eppendorf Femtojet system. At 15 min post-wounding, halocarbon oil was drained from the slide and flushed with heptane to release the embryos from the double-sided tape. Next, 0.8 mL 4% Formaldehyde (in PBS) was added to a fresh tube, and embryos were transferred with heptane to this tube, which was then placed on rotation in an orbital shaker for 25 min. The aqueous solution was then removed from the bottom of the tube, and subsequently, heptane was removed too. To devitellinize the embryos, heptane was rapidly added, followed by adding double to triple the volume of 90% ethanol and mixing it vigorously by vortexing. We then proceeded with embryos that settled down in the tube. The embryos were given 5 quick washes with 1X PBS and then blocked in PBS + 0.3% Triton-X100 + 0.5% BSA (PBTB) for 30 min at room temperature. A primary antibody against LAMP (Abcam) was prepared in PBTB. Embryos were incubated overnight at 4 °C, followed by three washes done in PBS + 0.3% Triton X-100 (PBT). Secondary antibody was prepared in PBTB and incubated for 2 h at room temperature, followed by 5 washes with PBT. DAPI and Phalloidin staining were done along with the secondary antibody. The embryos were mounted in Fluoromount (Sigma) on a glass slide and imaged using confocal microscopy.

Mouse embryonic skin wounding
C57/6J strain mice, 8–12-week-old adult females, were used in the study for all experiments involving mice. No statistical methods were used to predetermine sample size. The results are independent of the sex of the animals. The development and physiology of the embryonic epidermal tissue is not known to be different in different sexes; therefore, sex of the animal was not considered a parameter in the study. The protocol was approved by the Institutional Animal Ethics Committee, TIFR Hyderabad, on behalf of the Committee for the Purpose of Control and Supervision of experiments on Animals, India.

For mouse embryonic wounding assays, E14.5 - E16.5 mouse embryos were dissected out. The dorsal skin from these embryos was then excised. The skin was placed on a filter paper (Whatman filter, 8 μm pore size) with the epidermal side up. Using a scalpel/fine scissor, incision wounds were made on the epidermis. The filter paper, along with the tissue, was kept floating in a 6-well plate containing 2 mL of DMEM with 5% fetal bovine serum (FBS) and 1% Pen Strep. After the desired incubation period, the tissue was fixed in 4% paraformaldehyde (PFA) overnight at 4 °C. The tissue was washed in PBS and transferred to 90% ethanol in PBST (0.2% Triton X-100 in 1X PBS) overnight at −20 °C. The tissue was then rehydrated using graded ethanol (75%, 50% and 25% in PBST). After two more washes in PBST, the tissue was blocked using 10% normal goat serum (NGS) in PBST for 2 h. Primary antibodies were added and incubated overnight at 4 °C. This was followed by subsequent PBST washes, after which secondary antibodies were added and incubated for 2 h at room temperature. After washing, the tissue was mounted on a slide using Fluoroshield (Sigma Aldrich) mounting media. The slide was cured overnight and imaged using an upright confocal (Leica Stellaris). For actomyosin contractility inhibition in mouse embryonic skin wounds, 50 μM of Blebbistatin (Sigma) and 30 μM Y27632 (Sigma) were used. SKIP-Kinesin1 interaction inhibitor kinesore (Torcis) was used at a final concentration of 50 μM in the ex vivo mouse embryonic skin wounds. The concentration of inhibitors used for this model system is higher compared to their use for cell monolayers because of the extremely tight epithelial in the tissue, leading to no effect of these compounds at a lower concentration. 3-D renders were created using Imaris (Version 9.2.1).

Immunofluorescence
Cells were fixed in 4% formaldehyde diluted in 1X PBS for 15 min at room temperature. For most antibody staining, cells were permeabilized with 0.25% Trition X-100 in PBS for 10 min at room temperature, followed by three quick washes with 1X PBS to remove the detergent. Cells were treated with blocking/staining solution (0.1% Triton X-100 in 1X PBS + 2% BSA) for 1 h at room temperature, washed thrice with 1X PBS, and further incubated with primary antibody prepared in blocking solution for 2 h at room temperature. Post-primary antibody incubation, cells were washed thrice with 1X PBS and incubated with secondary antibody, DAPI, and Phalloidin dilutions prepared in the blocking/staining solution for 1 h at room temperature. Finally, the samples were given 2 quick and 5 min wash with 1X PBS before proceeding for confocal microscopy.

For staining LAMP1 (Abcam), cells were incubated with a blocking solution (0.2% Saponin + 5% BSA in 1X PBS) for 30 min at room temperature, followed by three washes with 1X PBS. Cells were incubated with the primary antibody in staining solution (PBS + 0.2% Saponin) for 2 h at room temperature, washed thrice with 1X PBS and further incubated for 1 h with secondary antibodies, DAPI (4′,6-diamidino-2-phenylindole, 1 μg mL−1 in PBS) and Phalloidin made in staining solution. Cells were washed thrice with 1X PBS and imaged using confocal microscopy.

Lysosome labeling for live cell imaging
Lysosomes of MDCK cells for live-cell imaging experiments were labeled with Alexa-fluor conjugated 10,000 MW Dextran (Dextran-647 from Invitrogen). Briefly, cells were incubated with Dextran-647 at a final concentration of 0.25 mg ml−1 in complete media for 6 h at 37 °C in a 5% CO2 humidified chamber. Media for cells was then replaced with complete media and incubated for another 12 h to completely chase Dextran-647 to lysosomes.

Confocal microscopy
Fluorescence images were acquired using a 60X oil objective (Plan Apo N 60x Oil, N.A = 1.42, Olympus) and 100X oil objective (UPlanSApo, 100X/1.40 oil) mounted on an Olympus IX83 inverted microscope equipped with a scanning laser confocal head (Olympus FV3000). Time-lapse images of live samples were taken in the live-cell chamber provided with the microscope setup.

Photoconversion studies were done using MDCK cells that had been transiently transfected with mEos-Rac1 WT, DN or mEos-Rac1 CA. Transiently transfected cells were seeded in culture inserts and proceeded for lysosome labeling using Dextran-647 as described above. Cells were allowed to migrate for 60 min to allow for lamellipodia formation in migrating cells. Stimulation was done on a point region of interest juxtaposed to the site of growing lamellipodia using a 405 nm laser at 2% intensity, looped over 25 times with a scan speed of 1000 μs/pixel. This was followed by LSM imaging of the green and red channels, and Rac1 dynamics were tracked for 5 min.

The photoactivation experiment was carried out using a 60X oil objective (Plan Apo N 60x Oil, NA=1.42, Olympus) mounted on an Olympus IX83 inverted microscope equipped with a scanning laser confocal head (Olympus FV3000), supported with a live-cell imaging setup. 20 mM HEPES (Gibco) was used to maintain CO2 levels. MDCK cells expressing optoGEF-RhoA and CIBN-GFP-CAAX or optoGEF-RhoA and Mito-CIBN-GFP (control) 49, stained with lysosomal marker Dextran-647, were used for the experiment. Pulse activation (20 iterations) was performed in the selected rectangular ROIs (whole cell or cell-cell junction) using a 488 nm laser with 2.0% laser intensity at a 4.0 μsec/pixel scan rate. This step was immediately followed by time-lapse (5 min, 10-s intervals) LSM imaging in 561 nm and 647 nm channels, to visualize RhoA activation and monitor lysosomal movement, respectively. Endogenous Rac1 and lysosome localization images were acquired using a 100X oil objective (N.A.=1.45) Zeiss Airy-Scan microscope (LSM900) on SR mode, and fluorescence images of leader cells in mouse embryonic skin wounds were acquired using a Nikon inverted research microscope, Eclipse Ti2-E with Yokogawa CSU-W1 SoRa unit.

Förster resonance energy transfer (FRET)-based Rac1-activity measurements
FRET experiments for the Rac1 activity sensor were carried out in the live-cell confocal setup (Olympus FV3000). MDCK cells were plated in a 24-well plate and transiently transfected with Rac1 sensor plasmids 52. After 6 h, cells were trypsinized and seeded in migration chambers and left overnight to settle and form confluent monolayers in the chamber. Cells were simultaneously labeled for lysosomes using Dextran-647. Control and AR treatment were given to cells as described in the Global perturbation of lysosome positioning using Ringer’s solution method section. The images were acquired for control and AR-treated cells just after lifting off the confinement (T-0) and post 2 h of migration (T-2 h). The exposure times for donor, acceptor, and FRET channels were kept constant. Images of Dextran-647-labeled lysosomes were also acquired. Each field yielded three 800×800-pixel images representing donor, acceptor, and FRET channels. Images were analysed using custom software written in MATLAB (Mathworks), described previously 60.

Inhibition studies
For all inhibition studies, cells were pre-treated with the desired concentration of the inhibitor in opti-MEM for 30 min at 37 °C in a 5% CO2 humidified incubator, before lifting the confinement/culture insert. During migration, cells were kept in complete media containing the inhibitor at a given concentration for the required time. Actomyosin contractility was altered using Blebbistatin (Sigma), a myosin inhibitor and Y27632 (Sigma), a ROCK inhibitor, at concentrations of 5 μM and 10 μM, respectively, for MDCK cells. For microtubule disruption in MDCK cells, Nocodazole (Sigma) was used at 10 μM, and Taxol/Paclitaxel (Sigma) was used at 1 μM. For SKIP-Kinesin-1 interaction inhibition, Kinesore (cis) was used at 12.5 μM as recommended in the previous study 44. Kinesore inhibition in MDCK cells was performed in OptiMEM for the completed duration of the treatment, and the control with these experiments were also kept in OptiMEM to maintain similar conditions. Cells were fixed and immunostained for the desired antibodies post-inhibitor treatment. For inhibitor studies in ex vivo mouse embryonic skin wounds, 50 μM of Blebbistatin (Sigma), 30 μM Y27632 (Sigma) were used, and SKIP-Kinesin1 interaction inhibitor kinesore (cis) was used at a final concentration of 50 μM. The concentration of inhibitors used for this model system is higher compared to their use for cell monolayers because of the extremely tight epithelial tissue, leading to no effect of these compounds at a lower concentration. Ex vivo mouse embryonic skin wounds were incubated with inhibitor-containing media for 30 min, fixed and immunostained.

Lysosome enrichment using density gradient separation
To perform lysosome enrichment, subcellular fractionation was carried out using the Lysosome Enrichment Kit (Thermo Scientific), using the manufacturer’s protocol. Briefly, the MDCK cells were grown to 90% confluency, followed by collecting cells on ice by cell scraping. The cell pellet was washed in ice-cold PBS, resuspended in Buffer-A, provided in the kit and homogenized with a dounce homogenizer on ice (~25 strokes). To confirm cell lysis, microscopic examination of the homogenate was done by adding 0.5% trypan blue dye. The homogenate was mixed with an equal volume of Buffer-B and subjected to centrifugation at 500 × g for 10 min at 4 °C, and post-nuclear supernatant (PNS) was diluted in OptiPrep gradient buffer to a final concentration of 15% OptiPrep. The sample was then carefully overlayered on the top of a discontinuous density gradient (17, 20, 23, 27, and 30%). The gradient was subjected to ultracentrifugation at 145,000 × g in a swinging bucket rotor (Beckman Colter) for 2 h at 4 °C. After the spin, ten fractions of 800 µl each were collected from top to bottom. The fractions were spun again at 18,000 × g for 20 min at 4 °C, and the resulting pellet was suspended in 4X SDS-sample buffer, boiled for 10 min, and analyzed by SDS-PAGE and immunoblotting.

Antibodies and plasmids
Source and dilution information for all primary and secondary antibodies used in immunofluorescence staining are given in Supplementary Table 1. Details of plasmids used in this study are listed in Supplementary Table 2 with their source.

Micropatterning
PDMS stencils with pillars of defined shapes were created using soft photolithography. Briefly, 150 um thick SU-8 2075 (Kayaku advanced materials, Y111074) was coated on a silicon wafer, followed by soft baking at 65 °C for 5 min and 95 °C for 20 min. SU-8 was exposed in the desired pattern using the MicroWriter ML 3 Pro (Durham Optics Magneto Ltd), followed by post-exposure baking at 65 °C for 7 min and 95 °C for 15 min. The unexposed photoresist was removed by immersing the wafers in the SU-8 developer (Kayaku advanced materials, Y020100) for 8–10 min. Patterned SU-8 wafers were then hard-baked at 170 °C for 45 min. PDMS (Sylgard 184, Dow Corning) was mixed in a ratio of 1:10, degassed, and poured over the patterned wafers. After curing at 85 °C for 2 h, stencils with pillars were peeled off. The stencils were then treated with 2% Pluronic acid for 2 h followed by 15 min of 1X PBS wash. PDMS stencil and a petri dish were plasma cleaned for 10 s, and the stencil was then placed upon the dish. Cells were seeded around the pillars, and after confluence, the stencils were removed, allowing the cells to migrate into the gaps.

Image analysis
All image analysis for this study was performed using Fiji. All images acquired as Z-stacks were converted to a single Z-projected ‘Sum of slices’ image before proceeding with analysis. Peripheral occupancy Fraction (POF) was measured for cells at the margin of a migrating collective in our two-dimensional cell culture and mouse embryonic skin wound models. A leader cell was selected as the cell that is in the front of a migrating monolayer of cells, having lamellipodial extensions. The ROI box with a fixed width of 2.5 μM from the cell edge facing the gap was drawn over the cell of interest. The box was duplicated for the lysosome channel, and lysosome marker intensity was measured. For each cell, the total lysosomal intensity was also calculated by manually marking the cell of interest. For calculating the POF, the lysosomal intensity in the ROI was divided by the total intensity of lysosomes in that cell. 20 marginal and ~10 leader cells per experiment were quantified from three independent experiments. For calculating the junctional occupancy fraction for lysosomes, we measured the lysosomal marker intensity in the cell-cell junctions of the leader cells and non-leader marginal cells by drawing an ROI of 2.5 μM from the junction, followed by dividing it by the total lysosomal intensity in the cell. For measuring the POF for different organelles, we followed the same approach.

To measure Mean Distribution from Edge (MDE), we used a custom-written MATLAB code to quantify the spatial distribution of lysosomes relative to the cell edge in migrating MDCK cells. Briefly, Z-projected sum-of-slices images were first processed to generate a separate cell boundary image from the actin cytoskeleton channel, which was then fed into MATLAB along with the lysosomal marker (LAMP1) image. The code detects the cell edge, allowing for precise spatial mapping of lysosome distribution. To compute MDE, the algorithm determines the center of mass (CoM) of lysosome fluorescence intensity within each cell, representing the weighted centroid of lysosome distribution, where pixel intensities contribute proportionally to their respective positions. The Euclidean distance between the CoM of lysosomal intensity and the detected cell edge is then calculated. A lower MDE value indicates greater lysosome enrichment at the periphery, while a higher MDE value reflects a more centrally distributed lysosome population. For each experiment, 20 marginal cells and approximately 10 leader cells were analyzed, with data collected from three independent experiments. This approach provides an unbiased, quantitative measure of lysosome positioning relative to the migration front, allowing for precise assessment of lysosome polarization dynamics.

Asymmetry of lysosome distribution in collectively migrating MDCK cells was measured at T-0 and T-2 h post-confinement lift-off. Specifically, we first identified the long axis of each cell along the direction of migration (perpendicular to the wound edge) and divided the cell at its midpoint along this axis, creating two distinct regions: the front half facing the migration direction, and the rear half positioned away from the leading edge. To quantify lysosome distribution, we measured the fluorescence intensity of the lysosomal marker LAMP1 in each half using ImageJ, followed by whole-cell LAMP1 intensity measurements. To compute the occupancy fraction for lysosomes at the front and rear ends of the cells, the intensity in each ROI was divided by the total LAMP1 intensity per cell. N = 3, with 15–20 cells per experiment analyzed for each time point. Statistical significance was determined using a one-way ANOVA test in GraphPad Prism.

To measure POF for experiments involving OptoGEF-RhoA, cells adjoining the photoactivated cell were marked manually using the DIC channel to mark cell boundaries. The image was duplicated for the lysosome channel. Lysosome intensity was determined using Fiji. This was followed by marking the cell periphery of the same cell by using the Edit → Selection → Enlarge tool of Fiji to reduce the outline by 2.5 μm. Using the Edit → Clear tool, the lysosomes in the perinuclear region were omitted, and the intensity of remaining lysosomes in the cell periphery was determined. To get the POF for cells, the intensity of lysosomes was divided by the total cell intensity of lysosomes for each cell. POF was determined for pre-activation and post-activation states. Change in POF was compared between pre- and post-activation for the same cell, and 60 cells were analysed per treatment from three independent experiments.

To measure leader emergence in experiments using the RAMP plasmid system, we quantified the total number of cells at the wound margin expressing both the motor (mCh-Kif5b*-Strep or Strep-KifC1*-mCh) and the cargo protein of interest (LAMP-, Tom20-, or VAPA-SBP-GFP), as well as the number of these transiently transfected cells that formed leader cells. The fraction of leader emergence was calculated by dividing the number of leader-forming cells by the total number of transfected cells at the wound margin. The method was used for measuring leader emergence fraction in cells expressing the KLC 2 motor with SKIP-WT or SKIP WD->2XA mutant. For inhibitor treatments, the leader emergence was measured by dividing the number of leaders by the total number of cells facing the wound gap. Colocalization analysis of different forms of small GTPase Rac1 with lysosomal marker LAMP1 was performed using the JACoP tool of Fiji. For measuring Mander’s coefficient, cytoplasmic expression was analysed by marking the inside of the cell boundary using a freehand tool in Fiji. The cytoplasmic Rac1 expression includes its vesicular component, and its co-localization with lysosomes was measured.

Tracking of cells during collective migration was done using the Manual Tracking tool of Fiji. The displacement in X and Y was determined from the manual tracking approach for tracking cells with biased lysosome positioning (Cell lines expressing mch-Kif5b*-Strep, Strep-KifC1*-mCh co-transfected with LAMP1-SBP-GFP plasmids). To measure lysosome speed from time-lapse videos, the TrackMate tool of Fiji was used with the following parameters: Vesicle diameter-1 μm; Detector-DoG; Initial thresholding-none; Tracker-Simple LAP tracker; Linking max distance-2 μm; Gap-closing max distance-2 μm, Gap-closing max frame gap-2; Filters-none. The gap closure was measured by marking the gap area manually in Fiji at T-0, 4, 8, 12 and 16 h.

Statistical analysis
Statistical analyses were carried out in GraphPad Prism 9. Statistical significance was calculated by Unpaired t-test with Welch’s correction or One-way ANOVA as mentioned in the corresponding figure. Scatter-bar plots were displayed as mean ± s.e.m. P-values greater than 0.05 were considered to be statistically not significant. No statistical methods were used to predetermine sample size. Quantification was done using data from at least three independent biological replicates. For analysis involving live-imaging experiments, data were collected from three independent experiments. All the experiments with representative images were repeated at least thrice. Data distribution was assumed to be normal, but this was not formally tested. No statistical method was used to predetermine sample size, but our sample sizes are consistent with the standard of the field and are appropriate for the types of analysis performed. Samples were not allocated using randomization, as this study involved observational analysis of epithelial cells under defined experimental conditions. All groups were subjected to identical culture, treatment and imaging protocols to control for potential covariates, ensuring comparability across conditions. Data collection and analysis were not performed blind to the conditions of the experiments. No data points were excluded from the analyses.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "In this revised manuscript, Marwaha and colleagues investigate the role of localized lysosomes in promoting the emergence of leader cells within migrating collectives. Using cell culture and in vivo wound healing models in Drosophila and mouse, the authors find that lysosomes localize to the periphery of leader cells via microtubules and actomyosin contraction. Re-localizing lysosomes away from the periphery prevents leader cell formation, whereas forcing lysosomes to the periphery enhances leader cells. Rac1 GTPase associates with lysosomes and loss of peripheral lysosomes resulted in lower Rac1 activity. The authors propose that lysosomes at the cell periphery increase Rac1 activation, leading to formation of lamellipodia and thus leader cell emergence.\nIn the revised manuscript, the authors added more controls, better quantifications, new drug treatments, and additional in vivo mouse skin explant assays. In response to my comments, the authors added better labels to the figures, added quantifications with more consistent data presentation, and added additional experiments to manipulate lysosome positioning and measure leader cell emergence (e.g., Kinesore treatments). Overall, the manuscript is improved and the general findings should be of high interest, especially the function of localized lysosomes in collective cell migration.\nHowever, a few major results remain less convincing and need to be addressed (see below).\nMajor points:\n1) The authors have added more experiments with their mouse skin wounding explant model. While I appreciate the inclusion of an in vivo model, I find the images very difficult to interpret. For example, while there are many lysosomes labeled in the tissue, it is very hard to be convinced that there is an enrichment at the periphery of leading cells (Fig. 1C,D; Supplemental Fig. 1C). The added labels certainly help with the fly embryo (Supplemental Fig. 1B), but even in this model it is unclear how many cells are “emerging leader cells” and how frequently this occurs in the wounding model. The images for the mouse skin in Fig. 4B and Fig. 5C are better but lysosome positions (and numbers of leader cells) are not quantified. The cell culture models throughout the manuscript are more convincing.\n2) A major conclusion of the manuscript is that “mechanical forces transmitted through the cell-cell interface polarized lysosomes away from the interface” (p. 9, lines 48-49). However, the results with Opto-RhoAGEF (Fig. 4D-H; Videos 8 and 9) are not convincing. The effects appear to be small. The videos are extremely difficult to interpret. The cell expressing opto-RhoAGEF/CIBN-CAAX in Fig. 4F appears to have quite a bit of membrane localization already at the “pre-activation” stage. In the video and in Fig. 4G, post-activation, this cell looks the same and not contracted. A major concern is that the cell is already activated prior to the experimental exposure to blue light, thus impacting interpretation of the results of the experiment. It may be that the timescales of activation are insufficient to see a strong effect on cell contraction, and thus on lysosome positioning. Either the conclusion needs to be toned down or a more convincing set of experiments needs to be done.\nMinor points:\n1) Time stamps are still missing in places, for example Fig. 4F and 4I, Fig. 6F and H, Supplementary Fig. 7H.\n2) The experiments in Supplemental Figure 9E-G are only discussed in the Discussion (p. 13, lines 30-34). It would be helpful to have that data in the Results.", "reviewer_2": "This revised manuscript investigates the importance of lysosome positioning for the emergence of leader cells in models of collective cell migration. The authors argue that peripheral lysosome positioning is required for leader cell specification. The topic is intriguing and the manuscript is improved from the original submission, but some shortcomings remain. Given that the manuscript has now been transferred to Nature Communications, it is probably suitable for publication with only modest modifications.\nSpecific issues\n1. The new data with mouse skin wound closure is really hard to interpret. The authors highlight cells with more peripheral lysosomes, but this is not always very clear and there are other nearby cells in the images that do not seem to fit the authors’ model. I accept that analysis of subcellular localisation in tightly packed 3D cells is challenging, but the results presented are currently rather equivocal. The analysis is further hindered because it is not clear which cells are leader cells in the authors’ model, and which cells are at the wound edge, but not leading. Thus, I consider the level of evidence presented weak. The authors should seek to improve the analysis and quantification of these data.\n2. The authors have not addressed the question (point 2) about how many cells need to be able to regulate lysosome positioning for collective migration to proceed efficiently. I understand the argument about the interchangeable nature of leader cells making it hard to score the overall effect of some manipulations on cell migration; however, there must be a minimal proportion of leader cells that are required. The authors have not sought to determine this, but perhaps this is not essential for publication in Nature Comms.\n3. The other more technical issues have largely been addressed.", "reviewer_3": "I have read the authors’ response carefully. The observation that lysosomes are more peripherally localized in emerging leader cells and during collective migration remains interesting. However, the fundamental issue of causality is still unresolved. The central question—whether peripheral lysosome relocalization is a driver of this process or merely a consequence—has not been convincingly addressed.\nThe authors rely heavily on pharmacological agents such as nocodazole, blebbistatin, and now kinesore to explore this relationship. While these compounds may offer some insight, they are inherently limited by their lack of specificity and broad cellular effects. The authors have included new data using kinesore, but even the group that identified this compound cautions that off-target effects are likely at the concentrations used (25–50 μM). In their own words: “we make no claim regarding the specificity of kinesore outside of this limited functional space...” (see https://doi.org/10.1073/pnas.1715115115). This significantly undermines the interpretability of the phenotypes observed.\nFigure 4 is intended to address causality—and while the RAMP system is indeed elegant, it is, in my view, insufficient on its own to definitively resolve the issue. Furthermore, it remains unclear whether changes in lysosome positioning result in functional consequences for migration. While this may be challenging to assess in vivo, several in vitro systems could be employed to investigate this relationship more directly, such as the assay shown in Supplementary Figure 4A. However, Supplementary Figure 5A shows no effect on migration, making it difficult to determine whether changes in leader cell emergence have any meaningful impact on the migratory process.\nThe authors state that “global disruption of lysosome transport, such as with kinesore, impairs this compensatory mechanism leading to slower wound closure,” but as discussed above, kinesore is not a sufficiently specific tool to demonstrate this convincingly, as it may also disrupt broader mechanotransductive processes required for leader cell emergence and migration, independent of lysosome positioning.\nFor example, would blocking peripheral lysosome localization across cells—using non-pharmacological methods—affect migration, as the authors state? Conversely, would forcing lysosomes to the periphery have a measurable effect when applied broadly across cells?\nWithout complementary and specific approaches to manipulate lysosome positioning—while clearly demonstrating functional effects on migration—the causal link remains uncertain."}, "author_response": "Response to Reviewers\nRebuttal to Reviewer Comments\nReviewer #1 (Remarks to the Author):\nIn this revised manuscript, Marwaha and colleagues investigate the role of localized lysosomes\nin promoting the emergence of leader cells within migrating collectives. Using cell culture and\nin vivo wound healing models in Drosophila and mouse, the authors find that lysosomes\nlocalize to the periphery of leader cells via microtubules and actomyosin contraction. Re-\nlocalizing lysosomes away from the periphery prevents leader cell formation, whereas forcing\nlysosomes to the periphery enhances leader cells. Rac1 GTPase associates with lysosomes and\nloss of peripheral lysosomes resulted in lower Rac1 activity. The authors propose that\nlysosomes at the cell periphery increase Rac1 activation, leading to formation of lamellipodia\nand thus leader cell emergence.\nIn the revised manuscript, the authors added more controls, better quantifications, new\ndrug treatments, and additional in vivo mouse skin explant assays. In response to my\ncomments, the authors added better labels to the figures, added quantifications with more\nconsistent data presentation, and added additional experiments to manipulate lysosome\npositioning and measure leader cell emergence (e.g., Kinesore treatments). Overall, the\nmanuscript is improved and the general findings should be of high interest, especially the\nfunction of localized lysosomes in collective cell migration. However, a few major results\nremain less convincing and need to be addressed (see below).\nResponse: We thank the reviewer for their valuable comments. Below, we summarize the new\nresults and analyses incorporated to address these concerns.\n1. Super-resolution imaging of leader cells in ex vivo mouse embryonic skin (MES)\nwound assays (Fig. 1E).\n2. POF quantification of lysosome distribution in leader vs. non-leader cells (Fig. 1K).\n3. Drug treatment analysis with Blebbistatin, Y27632, and Kinesore (Figs. 4D and 5D).\nMajor points:\n1) The authors have added more experiments with their mouse skin wounding explant model.\nWhile I appreciate the inclusion of an in vivo model, I find the images very difficult to interpret.\nFor example, while there are many lysosomes labeled in the tissue, it is very hard to be\nconvinced that there is an enrichment at the periphery of leading cells (Fig. 1C,D; Supplemental\nFig. 1C). The added labels certainly help with the fly embryo (Supplemental Fig. 1B), but even\nin this model it is unclear how many cells are “emerging leader cells” and how frequently this\noccurs in the wounding model. The images for the mouse skin in Fig. 4B and Fig. 5C are better\nbut lysosome positions (and numbers of leader cells) are not quantified. The cell culture models\nthroughout the manuscript are more convincing.\nResponse: We appreciate the reviewer’s constructive comments. In the previous version of the\nmanuscript, confocal imaging of MES wound assays revealed that cells at the migration front\ndisplay coordinated actin-microtubule polarization, and peripheral lysosomal enrichment,\nwhich were consistent with our 2D monolayer observations. To address the concern raised by\nthe reviewer, we have now performed super-resolution microscopy (Super Resolution via\nOptical Re-assignment: SoRa) by a confocal field scanning spinning disk system on our ex vivo\nMES samples and successfully identified leader cells, characterized by extended lamellipodia\n(Fig. 1E). The identified leader cells display peripheral lysosomal accumulation which is\nconsistent with our observation in our two-dimensional cell monolayer gap closure assays. To\nfurther support our findings, we quantified the Peripheral Occupancy Fraction (POF) of\nlysosomes in both leader and non-leader marginal cells (Fig. 1K). The POF analysis revealed\n1\nsignificantly greater peripheral lysosome accumulation in leader cells, as shown in the dot-\ncolumn graph (Fig. 1K). We have now also included POF analysis performed on MES wounds\ntreated with chemical inhibitors such as Blebbistatin, Y-27632, and Kinesore to measure\nlysosome accumulation (Figs. 4C-D and 5C-D). Our super-resolution imaging and lysosomal\nPOF measurements in MES wound assays align with findings from our two-dimensional gap\nclosure models. Together, they strengthen the evidence that peripheral lysosome distribution is\nessential for leader cell emergence across different systems.\n2) A major conclusion of the manuscript is that “mechanical forces transmitted through the\ncell-cell interface polarized lysosomes away from the interface” (p. 9, lines 48-49). However,\nthe results with Opto-RhoAGEF (Fig. 4D-H; Videos 8 and 9) are not convincing. The effects\nappear to be small. The videos are extremely difficult to interpret. The cell expressing opto-\nRhoAGEF/CIBN-CAAX in Fig. 4F appears to have quite a bit of membrane localization\nalready at the “pre-activation” stage. In the video and in Fig. 4G, post-activation, this cell looks\nthe same and not contracted. A major concern is that the cell is already activated prior to the\nexperimental exposure to blue light, thus impacting interpretation of the results of the\nexperiment. It may be that the timescales of activation are insufficient to see a strong effect on\ncell contraction, and thus on lysosome positioning. Either the conclusion needs to be toned\ndown or a more convincing set of experiments needs to be done.\nResponse: We thank the reviewer for this insightful comment. We have now reperformed\nexperiments with additional precautions and replaced the Opto-RhoA GEF images (Figs. 5E–\nG; Supplementary Fig. 7E; Supplementary videos 8–9) to remove potential pre-activated cells\nand improve clarity. Because RhoA GEF activation lasts only ~2 minutes, and contraction\noccurs in a single cell distributing force among several neighbors, the resulting changes in\nlysosome positioning are subtle. Nevertheless, together with the contractility inhibitor and cell\ndublet experiments, these results demonstrate that mechanical forces transmitted through cell-\ncell interfaces drive lysosome polarization away from the junctions. Thus, lysosomes likely\nserve as the mechanosensitive link coupling intercellular tension among follower cells to leader\ncell emergence.\nFollowing reviewer’s suggestion, we have now revised the text (Page 10, Lines 4-9) to\nexplicitly clarify these limitations and appropriately tone down our interpretation of this\nexperiment.\nMinor points:\n1) Time stamps are still missing in places, for example Fig. 4F and 4I, Fig. 6F and H,\nSupplementary Fig. 7H.\nResponse: We apologize for missing the time stamps on some of our representative time-lapse\nimages. We have now added time stamps to Figs. 4F, 4I, 6F, 6H, and Supplementary Fig. 7H.\n2) The experiments in Supplemental Figure 9E-G are only discussed in the Discussion (p. 13,\nlines 30-34). It would be helpful to have that data in the Results.\nResponse: As suggested by the reviewer, we have now added the results discussing\nSupplementary Figure 9E-G in the results section (Page 12, Lines 12-14).\nThis revised manuscript investigates the importance of lysosome positioning for the emergence\nof leader cells in models of collective cell migration. The authors argue that peripheral\nlysosome positioning is required for leader cell specification. The topic is intriguing and the\nmanuscript is improved from the original submission, but some shortcomings remain. Given\nthat the manuscript has now been transferred to Nature Communications, it is probably suitable\nfor publication with only modest modifications.\nResponse: We thank the reviewer for their valuable comments. Below, we summarize the new\nresults and analyses incorporated to address these concerns.\n1. Super-resolution imaging of leader cells in ex vivo mouse embryonic skin (MES)\nwound assays (Fig. 1E).\n2. POF quantification of lysosome distribution in leader vs. non-leader cells (Fig. 1K).\n3. Drug treatment analysis with Blebbistatin, Y27632, and Kinesore (Figs. 4D and 5D).\nSpecific issues\n1. The new data with mouse skin wound closure is really hard to interpret. The authors highlight\ncells with more peripheral lysosomes, but this is not always very clear and there are other\nnearby cells in the images that do not seem to fit the authors’ model. I accept that analysis of\nsubcellular localisation in tightly packed 3D cells is challenging, but the results presented are\ncurrently rather equivocal. The analysis is further hindered because it is not clear which cells\nare leader cells in the authors’ model, and which cells are at the wound edge, but not leading.\nThus, I consider the level of evidence presented weak. The authors should seek to improve the\nanalysis and quantification of these data.\nResponse: We appreciate the reviewer’s constructive comments. In response, we have\nstrengthened our analysis of lysosome localization in mouse embryonic skin (MES) wound\nassays. Using super-resolution microscopy (Super Resolution via Optical Re-assignment:\nSoRa) by a confocal field scanning spinning disk system, we were able to clearly identify leader\ncells, characterized by prominent lamellipodia at the wound edge (Fig. 1E). These leader cells\nexhibit pronounced peripheral lysosomal enrichment, consistent with our two-dimensional\nmonolayer observations. To substantiate this, we quantified the Peripheral Occupancy Fraction\n(POF) of lysosomes in leader and non-leader marginal cells, revealing significantly higher\nperipheral accumulation in leader cells (Fig. 1K). We further extended this analysis to MES\nwounds treated with contractility modulators (Blebbistatin, Y-27632, and Kinesore), which\nyielded corresponding changes in lysosome distribution (Figs. 4C–D and 5C–D). In summary,\nsuper-resolution imaging and POF analysis in MES wound assays corroborate our 2D findings,\nsupporting that peripheral lysosome distribution contributes to leader cell emergence across\nsystems.\n2. The authors have not addressed the question (point 2) about how many cells need to be able\nto regulate lysosome positioning for collective migration to proceed efficiently. I understand\nthe argument about the interchangeable nature of leader cells making it hard to score the overall\neffect of some manipulations on cell migration; however, there must be a minimal proportion\nof leader cells that are required. The authors have not sought to determine this, but perhaps this\nis not essential for publication in Nature Comms.\nResponse: We thank the reviewer for this insightful comment. Related to lysosome\nmanipulation experiments, quantifying the number of “engineered” leader cells required for\n3\nefficient collective migration is inherently difficult, as leader cells continuously emerge,\nretract, and are replaced during wound closure. This dynamic interchangeability enables the\ntissue to maintain coordinated migration even when only a subset of cells can properly regulate\nlysosome positioning, often masking population-level effects of such manipulations. In our\nepithelial monolayer migration assays, we typically observe 3-4 leader cells within a 640 µm\nwound segment, consistent with previous estimates (e.g., Vishwakarma et al., Nat. Commun.\n2018 - https://doi.org/10.1038/s41467-018-05927-6), suggesting that only a small fraction of\ncells acquiring leader-like properties is sufficient to sustain coordinated migration, with\nlysosomal polarization serving as a key hallmark of this state. Nevertheless, we have now\ndiscussed this point in the revised manuscript (Page. 13, Line 24-28) as one of the limitations\nof the study.\n3. The other more technical issues have largely been addressed.\nResponse: We thank the reviewer for their assessment and feedback on our manuscript.\nReviewer #3 (Remarks to the Author):\nI have read the authors’ response carefully. The observation that lysosomes are more\nperipherally localized in emerging leader cells and during collective migration remains\ninteresting. However, the fundamental issue of causality is still unresolved. The central\nquestion—whether peripheral lysosome relocalization is a driver of this process or merely a\nconsequence—has not been convincingly addressed.\nThe authors rely heavily on pharmacological agents such as nocodazole, blebbistatin, and now\nkinesore to explore this relationship. While these compounds may offer some insight, they are\ninherently limited by their lack of specificity and broad cellular effects. The authors have\nincluded new data using kinesore, but even the group that identified this compound cautions\nthat off-target effects are likely at the concentrations used (25–50 μM). In their own words:\n“we make no claim regarding the specificity of kinesore outside of this limited functional\nspace...” (see https://doi.org/10.1073/pnas.1715115115). This significantly undermines the\ninterpretability of the phenotypes observed.\nFigure 4 is intended to address causality—and while the RAMP system is indeed elegant, it is,\nin my view, insufficient on its own to definitively resolve the issue. Furthermore, it remains\nunclear whether changes in lysosome positioning result in functional consequences for\nmigration. While this may be challenging to assess in vivo, several in vitro systems could be\nemployed to investigate this relationship more directly, such as the assay shown in\nSupplementary Figure 4A. However, Supplementary Figure 5A shows no effect on migration,\nmaking it difficult to determine whether changes in leader cell emergence have any meaningful\nimpact on the migratory process.\nThe authors state that “global disruption of lysosome transport, such as with kinesore, impairs\nthis compensatory mechanism leading to slower wound closure,” but as discussed above,\nkinesore is not a sufficiently specific tool to demonstrate this convincingly, as it may also\ndisrupt broader mechanotransductive processes required for leader cell emergence and\nmigration, independent of lysosome positioning.\nFor example, would blocking peripheral lysosome localization across cells—using non-\npharmacological methods—affect migration, as the authors state? Conversely, would forcing\nlysosomes to the periphery have a measurable effect when applied broadly across cells?\nclearly demonstrating functional effects on migration—the causal link remains uncertain.\nResponse: We thank the reviewer for their assessment and comments on our manuscript. Here\nwe provide a point wise response of the results and analysis performed to address the issues\nraised by the reviewer.\n1) Specificity of Kinesore at a concentration of 25 µM: In the study by Randall et al.\n(https://doi.org/10.1073/pnas.1715115115), complete abrogation of KLC2 (kinesin motor) and\nSKIP (kinesin-binding lysosomal adaptor) interactions was demonstrated using biochemical\nassays at 25 µM of Kinesore. Following that, in our previous version of the manuscript draft,\nwe also used 25 µM Kinesore to achieve complete inhibition of the KLC2-SKIP interaction,\nwhich resulted in perinuclear lysosome localization and delayed migration of MDCK\nmonolayers. In response to the reviewer’s comment, we have now tested Kinesore at a lower\nconcentration of 12.5 µM, which has previously been reported to suppress the KLC2-SKIP\ninteraction by up to 50% (Randall et al. PNAS, 2017). Our new data show that even cells\ntreated with 12.5 µM Kinesore display both a clear delay in collective cell migration\n(Supplementary Figs. 5A–B) and a marked reduction in peripheral lysosome localization, as\nquantified by our POF analysis (Figs. 4A–B). These results demonstrate that even partial\ninhibition of the KLC2-SKIP interaction is sufficient to perturb lysosome positioning and\nimpair coordinated migration, thereby reinforcing the mechanistic link between kinesin-driven\nlysosome transport and leader cell behavior.\nWe fully acknowledge that pharmacological inhibitors may exert broad cellular effects.\nHowever, our conclusions are strengthened by the convergence of evidence from three\nindependent approaches:\n1. Kinesore-mediated disruption of KLC2-SKIP binding,\n2. The RAMP system to artificially bias lysosome positioning (Figs. 4E-L), and\n3. The SKIP WD-A mutant that uncouples lysosomes from kinesin motors (Figs. 3F-H\nand Fig. 4M).\nEach of these manipulations, despite their distinct modes of action, consistently produced\nconcordant phenotypes in lysosome localization and leader cell emergence. This\nmethodological triangulation provides robust support for the conclusion that lysosome\npositioning plays a causal and conserved role in regulating collective migration.\n2) We appreciate the reviewer’s comments and would like to clarify the distinction between\nSupplementary Figs. 4A and 5A. Supplementary Fig. 4A does not present a migration assay,\nas suggested by the reviewer; it instead depicts the spatial organization of organelles during\nleader cell emergence. In contrast, Supplementary Fig. 5A specifically shows the results of\nmigration assays, with time-stamped images comparing control and Kinesore-treated MDCK\nmonolayers. Cells treated with 25 µM Kinesore exhibit delayed leader emergence and\nincomplete wound closure even after 16 hours, whereas control monolayers close the gap\nefficiently - contrary to the interpretation that no migration defect was observe. To further\nvalidate this, we repeated the assay using 12.5 µM Kinesore and again observed a significant\ndelay in migration, accompanied by perinuclear lysosome localization (Figs. 4A–B;\nSupplementary Figs. 5A–B).\n3) The reviewer raises an interesting point regarding the outcome of migration when lysosome\nlocalization is biased toward either the perinuclear or peripheral regions across cells in a non-\npharmacological manner. To address this, we transfected LAMP1-SBP-GFP stable cells with\neither mCh-Kif5B*-Strep (peripheral lysosomes) or Strep-KifC1*-mCh (perinuclear\nlysosomes) and monitored collective cell migration in live imaging. Consequently, we\nobserved higher migration length and higher number of leader cells for LAMP1-SBP-GFP+\nmCh-Kif5b*-Strep double positive cells, where all cells have peripherally polarized lysosomes.\nHowever, LAMP1-SBP-GFP+ Strep-KifC1*-mCh double positive cells with perinuclear\nlysosomes, show delayed migration and lesser number of leader cells.\nRebuttal Figure 1. Double positive MDCK cells (LAMP1-SBP-GFP with Strep-KifC1*-mCh\nor mCh-Kif5b*-Strep) were seeded in an iBidi migration chamber (Left side - LAMP1-SBP-\nGFP+ Strep-KifC1*-mCh; Right side - LAMP1-SBP-GFP+ mCh-Kif5b*-Strep). The cells\nwere allowed to migrate for 24 hrs post confinement lift-off. Yellow arrowheads mark the\nleader cells."}, {"round": 2, "reviewer_comments": {"reviewer_1": "Marwaha and colleagues find that localized lysosomes promote the emergence of leader cells within migrating collectives during wound healing. In the revised manuscript, the authors performed better imaging of mouse embryonic skin (MES) wound assays, quantification of lysosomes in the MES wounds, and reperformed opto-RhoA GEF to prevent pre-activation within the cells. However, I am still less convinced by the opto-RhoA GEF experiments (Fig. 5E-I, Supplementary video 8). Especially in the video, it is hard to see that there is any additional change in lysosome localization. I appreciate that the authors are now more cautious with their conclusions from this set of experiments. Even so, these experiments are not as convincing as those using the myosin and Rho kinase inhibitors (Blebbistatin and Y-27632; Fig. 5A-C). Indeed, the experiments with contact inhibition of locomotion (CIL) of MDCK cells shown in Supplemental Fig. 7A-C are more convincing, showing that pulling forces polarize lysosomes towards the cell front. I would suggest either putting the CIL experiments into the main figure or further clarifying the results of the opto-RhoA GEF experiments. Otherwise, I am satisfied with the improvements in the manuscript due to the new experiments.", "reviewer_2": "I think the authors have made satisfactory attempts to address the comments raised, including those of the other reviewers.", "reviewer_3": "I have read the authors’ comments. Rebuttal Figure 1 presents convincing data that do not rely on pharmacological agents and appear to directly affect leader-cell emergence and collective migration in cells with altered distributions of perinuclear versus peripheral lysosomes. This type of experiment provides a much cleaner approach to testing the effects of lysosome positioning, in contrast to the extensive pharmacological data presented by the authors, and the methodology aligns well with other studies that manipulate lysosome positioning through genetic means (for example, PMID: 31130364, 21394080). In my opinion, the manuscript would benefit from the inclusion of these additional data—provided that they have been statistically validated, independently replicated, and shown to correlate with changes in lysosome positioning."}, "author_response": "Rebuttal to Reviewer Comments\nReviewer #1 (Remarks to the Author):\nMarwaha and colleagues find that localized lysosomes promote the emergence of leader cells\nwithin migrating collectives during wound healing. In the revised manuscript, the authors\nperformed better imaging of mouse embryonic skin (MES) wound assays, quantification of\nlysosomes in the MES wounds, and reperformed opto-RhoA GEF to prevent pre-activation\nwithin the cells. However, I am still less convinced by the opto-RhoA GEF experiments (Fig.\n5E-I, Supplementary video 8). Especially in the video, it is hard to see that there is any\nadditional change in lysosome localization. I appreciate that the authors are now more cautious\nwith their conclusions from this set of experiments. Even so, these experiments are not as\nconvincing as those using the myosin and Rho kinase inhibitors (Blebbistatin and Y-27632;\nFig. 5A-C). Indeed, the experiments with contact inhibition of locomotion (CIL) of MDCK\ncells shown in Supplemental Fig. 7A-C are more convincing, showing that pulling forces\npolarize lysosomes towards the cell front. I would suggest either putting the CIL experiments\ninto the main figure or further clarifying the results of the opto-RhoA GEF experiments.\nOtherwise, I am satisfied with the improvements in the manuscript due to the new experiments.\nResponse: We thank the reviewer for these valuable comments. We have further clarified the\nresults of the opto-RhoA GEF experiments. Please see Lines 15-20, Page No. 10.\nIn addition, we have moved CIL experiments to Main figure 5, and the OptoGEF experiments\nto Supplementary figure 7.\nReviewer #2 (Remarks to the Author):\nI think the authors have made satisfactory attempts to address the comments raised, including\nthose of the other reviewers.\nResponse: We thank the reviewer for the valuable comments throughout the review process.\nReviewer #3 (Remarks to the Author):\nI have read the authors’ comments. Rebuttal Figure 1 presents convincing data that do not rely\non pharmacological agents and appear to directly affect leader-cell emergence and collective\nmigration in cells with altered distributions of perinuclear versus peripheral lysosomes. This\ntype of experiment provides a much cleaner approach to testing the effects of lysosome\npositioning, in contrast to the extensive pharmacological data presented by the authors, and the\nmethodology aligns well with other studies that manipulate lysosome positioning through\ngenetic means (for example, PMID: 31130364, 21394080). In my opinion, the manuscript\nwould benefit from the inclusion of these additional data - provided that they have been\nstatistically validated, independently replicated, and shown to correlate with changes in\nlysosome positioning.\nResponse: We thank the reviewer for this suggestion. We confirm that the Rebuttal Figure 1\nfrom the previous rebuttal letter has now been incorporated into the main manuscript, with the\nimages now included in Supplementary Figure 5 and the corresponding quantification with\nstatistics (from 3 independent experiments) presented in Figure 4M. Please also see Lines 17-\n25, Page No. 8 for relevant discussions of this experiment."}]

Decision letter