Cell migration is an essential process driving both development and cancer metastasis. During these processes, cells often migrate as groups or collectives, rather than single cells (Friedl and Gilmour, 2009). Collective cell migration requires that cell-cell adhesions be maintained amongst the cells to support cluster cohesion (De Pascalis and Etienne-Manneville, 2017). Additionally, many collective cell migrations occur in an invasive manner with the group of cells migrating between other cells or through basement membranes (Chang et al., 2019). During invasive migration, the environment puts mechanical forces on the migrating cells, causing them to respond by changing their shape and stiffness, and by modifying properties of their environment, such as extracellular matrix (ECM) composition (Aguilar-Cuenca et al., 2014; Gasparski et al., 2017; Eble and Niland, 2019). Therefore, stiffness has emerged as a critical regulator of collective cell migration.

During invasive, collective cell migration the group or cluster of cells must generate force necessary to invade through the ECM or other cells. Stiffness of the substrate is considered the primary regulator of the migrating cell’s stiffness and ability to migrate (Aguilar-Cuenca et al., 2014). For example, increased substrate stiffness contributes to cancer cell migration and metastasis (Gasparski et al., 2017; Oakes, 2018; Eble and Niland, 2019). Indeed, hard matrices induce migration in breast cancer cells (Ren et al., 2021), and increased substrate stiffness promotes epithelial to mesenchymal transitions (Nieto and Cano, 2012). While the role of substrate stiffness in promoting cell migration is well-established, most of these studies utilized in vitro culture systems. Therefore, it remains poorly understood how migrating cells are regulated in their native environments by the stiffness of their endogenous substrates.

A master regulator of cellular stiffness is Non-Muscle Myosin II (subsequently referred to as Myosin). Myosin is a force generating actin motor (Vicente-Manzanares et al., 2009; Aguilar-Cuenca et al., 2014). It is composed of two copies of three subunits: two heavy chains, two essential light chains, and two regulatory light chains (MRLC; Vicente-Manzanares et al., 2009; Aguilar-Cuenca et al., 2014). Myosin activation is regulated through phosphorylation of its regulatory light chains. This phosphorylation occurs through a number of kinases, including Myosin light chain kinase (MLCK) and Rho-associated kinase (Rok), and dephosphorylation occurs through phosphatases, such as protein phosphatase 1 c (PP1c) and its catalytic subunit, Myosin binding subunit (Mbs Vicente-Manzanares et al., 2009; Aguilar-Cuenca et al., 2014). Myosin generates cortical tension by associating with and acting upon cortical F-actin; this regulates cell stiffness which can influence cell migration (Butcher et al., 2009; Aguilar-Cuenca et al., 2014). Importantly, Myosin regulates stiffness in both cellular substrates and migrating cells during many different cell migrations (Lo et al., 2000; Vicente-Manzanares et al., 2009; Mohan et al., 2015). Additionally, Myosin not only generates mechanical force within a cell but aids in sensing and responding to external forces applied to the cell (Butcher et al., 2009; Vicente-Manzanares et al., 2009; Aguilar-Cuenca et al., 2014).

A recently discovered regulator of Myosin is Fascin. Fascin is an F-actin binding protein that bundles or cross-links actin filaments into fibers (Jayo and Parsons, 2010; Hashimoto et al., 2011). However, recent studies demonstrate that there are many non-canonical roles for Fascin (Lamb and Tootle, 2020). One of these non-canonical functions of Fascin is the regulation of Myosin (Elkhatib et al., 2014). Increasing concentrations of Fascin in an in vitro system decreased Myosin ATP consumption and motor speed along actin filaments (Elkhatib et al., 2014). These data suggest that Fascin limits Myosin activity (Elkhatib et al., 2014). Whether Fascin limits Myosin activity to control substrate stiffness and thereby cell migration remains unknown. Notably, Fascin has well-established roles in promoting cell migration (Lamb and Tootle, 2020). Fascin aids in the formation of cell migratory structures like filopodia (Hashimoto et al., 2011) and invadopodia (Li et al., 2010). Fascin promotes many types of cell migrations in development and disease, including cancer metastasis (Ma and Machesky, 2015). Investigation of Fascin’s role in promoting cell migration has primarily focused on Fascin as an F-actin bundler and it is unknown if Fascin limits Myosin activity to regulate collective cell migration.

An ideal model to uncover the role of Fascin in regulating Myosin during collective cell migration in a native context is Drosophila border cell migration. Border cell migration occurs during Stage 9 (S9) of oogenesis. During S9, the follicle is composed of an oocyte and 15 germline-derived nurse cells that are surrounded by a layer of somatic epithelial cells called follicle cells (Spradling, 1993). Surrounding the follicle cells is a layer of ECM that envelopes the follicle (Spradling, 1993). Inside the follicle, however, there is limited evidence of any ECM (Medioni and Noselli, 2005). At the beginning of S9, a group of 8–10 follicle cells are specified as border cells and delaminate from the epithelium to start their migration (Montell, 2003). The border cells migrate invasively and collectively between the nurse cells until they reach the nurse cell-oocyte boundary (Figure 1A and B; Montell, 2003). Border cell migration is a cell-on-cell migration in which the nurse cells are the substrate for the migration, there is only a small puncta of ECM on the border cell cluster as it migrates (Medioni and Noselli, 2005) and border cell migration is largely independent of Integrin-based adhesions (Dinkins et al., 2008; Llense and Martín-Blanco, 2008). Importantly, similar to other types of migration, the stiffness of the nurse cell substrate regulates both the stiffness of the border cells and their migration (Aranjuez et al., 2016). Therefore, border cell migration is a powerful model for studying invasive, collective cell migration as the cluster of migrating cells can be visualized in its native context using both fixed and live imaging. Additionally, the factors that regulate border cell migration play conserved roles in other invasive, collective cell migrations, including cancer metastasis (Montell et al., 2012; Stuelten et al., 2018). Indeed, both Fascin and Myosin play roles in promoting cancer metastasis (Hashimoto et al., 2011; Aguilar-Cuenca et al., 2014) and on-time border cell migration (Figure 1C, Edwards and Kiehart, 1996; Lamb et al., 2020). We previously found that Fascin (Drosophila Singed, Sn) is required for both border cell delamination and proper protrusion localization (Lamb et al., 2020). Both loss and activation of Myosin result in similar phenotypes of delayed delamination and mislocalized border cell cluster protrusions (Majumder et al., 2012; Aranjuez et al., 2016; Mishra et al., 2019). These data suggest that the cycling of Myosin between active and inactive forms controls border cell migration. Thus, border cell migration is an ideal system to uncover the relationship of Fascin and Myosin during collective cell migration.

Figure 1

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Here, we demonstrate for the first time that Fascin inhibits Myosin activity in vivo. Loss of Fascin significantly increases the level of active Myosin, reduces Myosin dynamics, and increases nurse cell (aka substrate) stiffness as quantified by atomic force microscopy (AFM) nanoindentation technique. Reducing Myosin in fascin mutant follicles rescues border cell migration delays, indicating that Fascin’s tight regulation of Myosin activity is critical for on-time migration. Further, a phosphomimetic form of Fascin that precludes F-actin bundling is unable to limit Myosin activation, supporting the prior model that Fascin limits Myosin activity by tightly bundling F-actin and preventing Myosin binding to actin filaments (Elkhatib et al., 2014). We used RNAi knockdown and rescue experiments to assess the cell-specific roles of Fascin in regulating Myosin activity and nurse cell stiffness. Based on the literature, we expected that Fascin would primarily function within the nurse cells to control both substrate stiffness and Myosin activity within both the nurse cells and border cells. Surprisingly, we find that knocking down Fascin in the border cells increases the level of active Myosin on both the border cells and the nurse cells, and increases the stiffness of the nurse cells. Similarly, re-expressing Fascin in only the border cells of fascin mutants restores normal Myosin activity levels and stiffness of the nurse cells. These unexpected findings suggest that migrating cells influence the mechanobiology of their substrate to promote their migration. Supporting this, increasing Rok activity in the border cells also results in increased nurse cell stiffness, indicating this migratory cell regulation of substrate stiffness is not a Fascin-specific phenomenom. Together these findings lead to the following model: Fascin acts primarily within the migrating border cells to limit Myosin activation which controls the stiffness of the both the border cells and their substrate, the nurse cells, to promote on-time migration. It is likely that this regulation of Myosin by Fascin and thereby, migrating cells controlling substrate stiffness, is a conserved means of promoting collective cell migration.

Using Drosophila border cell migration as a model, we provide the first evidence that Fascin limits Myosin activity in vivo to control tissue stiffness Figure 8. We find that loss of Fascin significantly increases activated Myosin, and this increase contributes to the border cell migration delays observed in fascin mutant follicles during S9. Our data suggest that Fascin’s bundling activity is required to limit Myosin activation, supporting the prior proposed model that Fascin tightly bundles F-actin and precludes Myosin from binding to actin filaments (Elkhatib et al., 2014). The increased Myosin activity in fascin mutants results in substrate stiffening. Using cell-specific knockdown and rescue experiments, we made the suprising finding that Fascin activity in the border cells is necessary and sufficient to regulate Myosin activity and stiffness of the nurse cells. Thus, Fascin activity within the border cells plays a critical role in controlling the balance of forces between the border cells and their substrate, the nurse cells. We also show that this force balance is not specific to Fascin, as directly altering Myosin activity within the border cells phenocopies knockdown of Fascin in these cells. Together our data uncover that in vivo, collectively migrating cells modulate the stiffness of their substrate to control their own migration (Figure 8).

Multiple lines of evidence support the model that Fascin is a critical regulator of cellular and tissue stiffness. Specifically, loss of Fascin results in increased active pMRLC on both the border cell and nurse cell membranes, altered MRLC-GFP dynamics on the border cell cluster, and increased nurse cell stiffness as measured by AFM (Figures 2 and 3). Interestingly, pMRLC staining and MRLC-GFP time-lapse imaging exhibit distinct patterns (Figure 2). In both wild-type and fascin mutant follicles, the MRLC-GFP regions are shorter in length than pMRLC puncta. We suspect this difference is due to the MRLC-GFP time-lapse imaging capturing a small period of just activated Myosin, whereas fixation and pMRLC staining captures a longer period of activity. For instance, when Fascin is lost the MRLC-GFP puncta have a longer lifetime, this longer period of Myosin activity may be captured as an increased number of shorter pMRLC puncta. The short and numerous pMRLC puncta may also suggest that Myosin activity in the border cells is not appropriately spatially regulated in fascin mutants. These data, in conjunction with our cell-specific RNAi and rescue analyses (Figures 6 and 7), reveal that Fascin acts primarily within the border cells to control the stiffness of their substrate, the nurse cells.

Fascin-dependent inhibition of nurse cell Myosin activity and stiffness is essential for on-time border cell migration (Figure 4), raising the question of how Fascin regulates Myosin. Our data supports the previously proposed model that Fascin bundled F-actin prevents Myosin binding to F-actin and thereby, restricts Myosin activity (Elkhatib et al., 2014). Specifically, we find that expression of the phosphomimetic form of Fascin (S52E), which is unable to bundle F-actin, in fascin mutants fails to both inhibit Myosin activation (Figure 5) or fully restore migration (Figure 5—figure supplement 1). It is important to note that phosphorylated Fascin, and likely phosphomimetic Fascin, also exhibits reduced F-actin binding by in vitro assays (Yamakita et al., 1996), raising the possibility that Fascin binding to F-actin, without bundling filaments, is sufficient to inhibit Myosin activation. Further experiments are needed to fully elucidate how Fascin limits Myosin activity.

Our discovery that Fascin limits Myosin activity in vivo is unlikely to be restricted to Drosophila. Indeed, both Fascin and Myosin play critical roles during cancer metastasis (Hashimoto et al., 2011; Aguilar-Cuenca et al., 2014; Ma and Machesky, 2015). Increased Myosin activation and consequently, increased stiffness are a common phenotype observed in cancer cells and their substrate (Tse et al., 2012; Aguilar-Cuenca et al., 2014; van Helvert and Friedl, 2016; Ren et al., 2021). Increased substrate stiffness promotes migration in a wide range of cancers, suggesting increased Myosin activity can lead to increased cancer metastasis (Aguilar-Cuenca et al., 2014; Emon et al., 2018; Mierke, 2020; Ren et al., 2021). Additionally, Fascin is highly expressed in many types of cancers, notably carcinomas (Hashimoto et al., 2011; Ma and Machesky, 2015). High Fascin expression in these cancers is correlated with increased migration (Grothey et al., 2000; Hashimoto et al., 2007), invasion (Adams et al., 1999; Minn et al., 2005), and metastasis (Li et al., 2014; Alburquerque-González et al., 2020). However, according to our model, increased Fascin would reduce Myosin activity. Our finding that Fascin-dependent bundling is required to limit Myosin activity and substrate stiffness suggests that phosphorylated Fascin may promote cancer metastasis by allowing high Myosin activation and potentially other bundling-independent functions. Supporting this idea, expression of a S39 phosphomimetic form of Fascin, which cannot bundle F-actin, promotes human colon carcinoma migration (Hashimoto et al., 2007), suggesting phosphorylated Fascin could promote cancer metastasis by allowing increased Myosin activation and cell stiffness.

Our finding that phosphomimetic Fascin only partially rescues the migration delay in fascin mutants suggests that non-bundling roles of Fascin also contribute to border cell migration. Indeed, Fascin has many functions besides F-actin bundling, such regulating microtubules (Villari et al., 2015) and acting within the nucleus (Groen et al., 2015). Additionally, S52 phosphorylated Fascin functions as an adaptor for the Linker of the Nucleoskeleton and Cytoskeleton (LINC) Complex (Groen et al., 2015; Jayo et al., 2016). This LINC Complex role of Fascin is required for nuclear shape changes necessary for mammalian single-cell invasive migration (Jayo et al., 2016), raising the idea that Fascin may be similarly required for the invasion of the border cells between the nurse cells. Further experiments are needed to understand how the different functions of Fascin are coordinated to promote migration.

Our results suggest that increased stiffness in the border cell cluster affects the stiffness of its substrate, the nurse cells (Figures 6 and 7). This non-autonomous function of the border cells in altering the stiffness of the nurse cells was unexpected, as previous data suggested the nurse cells exert force on the border cells and the border cells respond to this force (Aranjuez et al., 2016). This balance of forces is necessary to promote the migration of the cluster through the tightly packed nurse cells (Aranjuez et al., 2016). Specifically, a previous study observed that overexpression of a Rho GEF in the nurse cells, which both increased Myosin activation and caused the nurse cells to change their shape and become more circular, ultimately impairs border cell migration (Aranjuez et al., 2016). As we do not observe any obvious changes in nurse cell shape when Fascin is lost or knocked down in the nurse cells, it may be that loss of Fascin does not cause a severe enough change in nurse cell Myosin activity and cell stiffness to cause the border cell cluster to respond. Instead, our data suggest that the border cells play a larger role in this balance of forces by exerting force on the nurse cells to control nurse cell stiffness. This interaction could potentially allow the border cell cluster to stiffen the nurse cells as the cluster migrates. Interestingly, in the context of cancer cell migration, a stiffer substrate often promotes cell migration (Parekh and Weaver, 2016; Oakes, 2018; Ren et al., 2021). Further, there is growing evidence that one means of directing migration is a gradient of substrate stiffness, such that cells move from softer to stiffer substrates; this is termed durotaxis (Sunyer and Trepat, 2020; Shellard and Mayor, 2021). Indeed, durotaxis has emerged as a property of collectively migrating cells. Specifically, it has been suggested that clusters of migrating cells are better able to sense differences in stiffness and respond more effectively (Martinez et al., 2016; Sunyer et al., 2016). Therefore, it is tempting to speculate that the border cells exert force on the nurse cells to stiffen them to aid in migration.

A key remaining question is whether border cells regulates substrate stiffness in a spatial and/or temporal manner. Specifically, do the border cells push on the nurse cells as they are migrating, driving a wave of local Myosin activation and stiffening of the nurse cell substrate? In other systems, such local stiffening appears to be cell-type and context specific, and can occur at the front or back of the migrating cells (Doyle et al., 2021). Based on both our fixed- and live-imaging of Myosin activity, we believe the stiffening of the nurse cell substrate occurs on all sides of the border cell cluster. This phenotype is consistent in wild-type and fascin-null follicles with fascin-null follicles displaying an overall increase in pMRLC. However, our current means of assessing Myosin activity and cellular stiffness lack the resolution necessary to determine how Myosin activation and changes in stiffness propagate through the nurse cells. Future studies using tools that allow Myosin activity in the border cells to be distinguished from that in its nurse cell substrate, in conjunction with higher resolution, rapid, non-photobleaching microscopy approaches, and cell-specific means of assessing individual nurse cell stiffness, such as laser ablation recoil velocity assessments, are need to uncover the detailed dynamics of how the balance of forces between the border cells and their substrate, the nurse cells, drives collective cell migration.

It remains unclear how Myosin activity at the cell cortex is connected to cellular adhesions in the border cells. Prior work indicates that the border cells migrate directly on the nurse cells, as there is little to no ECM present (Medioni and Noselli, 2005), and thus, Integrin-based ECM adhesions are not critical for border cell migration (Dinkins et al., 2008; Llense and Martín-Blanco, 2008; Montell et al., 2012). Instead, border cells may utilize E-Cadherin based adhesions (Niewiadomska et al., 1999; Cai et al., 2014) or perhaps uncharacterized means to drive migration. Thus, additional work is needed to determine the cellular connections mediating Myosin-dependent force transmission between the border cells and the nurse cells.

While it is clear that the balance of forces between the border cells and the nurse cells is critical for border cell migration, the mechanisms by which force imbalances impair migration remain poorly understood. We speculate that the increased Myosin activity in fascin mutants delays migration by impacting delamination and protrusion dynamics. We previously found that Fascin is required for on-time delamination of the border cell cluster from the follicular epithelium and for restricting the number and location of protrusions to the leading edge of the border cell cluster (Lamb et al., 2020). Similarly, both loss and constitutive activation of Myosin within the border cells delays delamination and causes excessive and misdirected protrusions (Majumder et al., 2012; Aranjuez et al., 2016; Mishra et al., 2019). These data suggest that it is not only the level of Myosin activity but is ability to cycle between active and inactive states that contributes to these two aspects of border cell migration. Based on our data, we suspect altering Myosin activity in the border cells ultimately changes the stiffness of the nurse cells. Too little activation of Myosin would result in a soft substrate and too much would result in a stiff substrate. Such changes in substrate stiffness could alter the polarization of the cluster, resulting in mislocalized and increased protrusions which not only delay migration but impair delamination. Supporting this idea, Myosin regulates active Rac polarization within the border cells (Mishra et al., 2019). Rac activation is highest in the leading cell of the border cell cluster and is require to generate forward directed protrusions (Fulga and Rørth, 2002; Bianco et al., 2007; Mishra et al., 2019). Increased Myosin activation in the border cell cluster disrupts this polarization, resulting in mislocalized protrusions (Mishra et al., 2019). This loss of polarization could function cell-autonomously, but, based on our data, it may also increase nurse cell stiffness. Such increased substrate stiffness could impair delamination and cause mislocalized protrusions by physically altering the topography of the nurse cells, which has recently been shown to be critical for border cell migration and forward directed protrusions (Dai et al., 2020). Additionally, increased substrate stiffness could disrupt durotactic signaling or alter the diffusion of the ligands directing migration. Thus, Fascin’s role in limiting Myosin activation likely contributes to the delayed delamination and aberrant mislocalized protrusions observed during border cell migration in fascin-null follicles.

The mechanical communication between migrating cells and their substrate is a growing area of research. The overarching premise in the field has been that substrate stiffness regulates the mechanical properties of the migrating cells and thereby, alters their ability to migrate. For example, in a model of breast cancer cell migration, high substrate stiffness promotes migration (Ren et al., 2021). Additionally during zebrafish development, the underlying mesoderm must stiffen to induce the epithelial to mesenchymal transition (EMT) and migration of the neural crest cells (Barriga et al., 2018). Together these studies highlight the current paradigm that substrate stiffness is the driving force that regulates the migrating cells to control their migration. However, the roles of the migrating cells in controlling their substrate stiffness are less understood. Numerous studies support that migrating cells can degrade surrounding ECM, creating paths for easier migration (for example: Wolf et al., 2007); such changes likely decrease substrate stiffness in the local environment. Migrating cells also pull on their local environment, applying a strain on the environment and aligning ECM fibers, which ultimately causes a local increase in substrate stiffness (Hall et al., 2016; van Helvert and Friedl, 2016). Notably, collectively migrating cells exert 4 x more force on their environment than single cells (van Helvert and Friedl, 2016). These studies indicate migrating cells can, at least locally, influence the stiffness of their substrate. This increase in substrate stiffness promotes cell migration, increasing migratory cell force generation in a process termed mechanoreciprocity (van Helvert et al., 2018). Whether migrating cells control substrate and underlying tissue stiffness in native, physiological contexts, and whether local stiffness changes are propagated through the tissue remain poorly understood. Supporting that this may occur, cancer cells drive stromal changes, including increased fibrosis which stiffens the tissue (van Helvert et al., 2018; Chandler et al., 2019; Piersma et al., 2020). These changes have been proposed as potential mechanism by which cancer cells drive their own invasion and make environments supportive of metastatic colonization (Cox and Erler, 2014). Our finding that Fascin activity in the migrating border cells controls substrate stiffness to promote migration positions Drosophila border cell migration as a robust system to uncover the mechanisms controlling this means of force balance.

Here, we propose that migrating cells modulate their own stiffness to regulate substrate stiffness. Our findings suggest that during collective cell migrations, such as those during development and cancer metastasis, the migrating cells apply force to induce the stiffening of their substrate, this results in a reciprocal mechanical communication between the migrating cells and their substrate which drives migration. Further, we demonstrate that Fascin, an F-actin bundling protein, limits the activity of Myosin in the migrating cells to regulate substrate stiffness. Overall, our findings expand our understanding of the mechanical relationship between migrating cells and their substrate, shifting the paradigm in the field from the substrate controlling migrating cell stiffness and thereby, migration, to the migrating cells playing a key role in altering their environment and substrate stiffness to promote their own migration.

All data generated or analysed during this study are included in the manuscript as Figure-specific source data files.

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Author details

1. Maureen C Lamb

   Anatomy and Cell Biology, University of Iowa Carver College of Medicine, Iowa City, United States

   Contribution

   Conceptualization, Formal analysis, Investigation, Methodology, Visualization, Writing - original draft

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-4522-1910

2. Chathuri P Kaluarachchi

   Department of Chemistry, University of Iowa, Iowa City, United States

   Contribution

   Formal analysis, Investigation, Writing - review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-2538-3952

3. Thiranjeewa I Lansakara

   Department of Chemistry, University of Iowa, Iowa City, United States

   Contribution

   Formal analysis, Investigation, Methodology

   Competing interests

   No competing interests declared

4. Samuel Q Mellentine

   Anatomy and Cell Biology, University of Iowa Carver College of Medicine, Iowa City, United States

   Contribution

   Formal analysis, Investigation

   Competing interests

   No competing interests declared

5. Yiling Lan

   Department of Chemistry, University of Iowa, Iowa City, United States

   Contribution

   Formal analysis

   Competing interests

   No competing interests declared

6. Alexei V Tivanski

   Department of Chemistry, University of Iowa, Iowa City, United States

   Contribution

   Supervision, Writing - review and editing

   Competing interests

   No competing interests declared

7. Tina L Tootle

   Anatomy and Cell Biology, University of Iowa Carver College of Medicine, Iowa City, United States

   Contribution

   Conceptualization, Funding acquisition, Supervision, Writing - review and editing

   For correspondence

   tina-tootle@uiowa.edu

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-1515-9538

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