Cells that migrate as collectives help establish and organize many tissues and organs in the embryo, yet also promote tumor invasion, dissemination and metastasis (Friedl et al., 2012; Friedl and Gilmour, 2009; Wang et al., 2016; Cheung and Ewald, 2016; Scarpa and Mayor, 2016). A wide variety of cells undergo collective cell migration during development, ranging from neural crest cells in Xenopus, the zebrafish lateral line primordium, and branching mammary glands (Friedl and Gilmour, 2009; Scarpa and Mayor, 2016; Huebner et al., 2016; Shellard and Mayor, 2019), among many other examples. Despite the apparent diversity in collectively migrating cell types, there is remarkable conservation of the cellular and molecular mechanisms that underlie group cell movements. In particular, migrating collectives require fine-tuned organization and cell coordination to move effectively as a unified group. Similar to individually migrating cells, collectively migrating cells display a front-rear polarity, but this polarity is often organized at the group level (Mayor and Etienne-Manneville, 2016). Leader cells at the front extend characteristic protrusions that help collectives navigate tissues. Mechanical cell coupling and biochemical signals then reinforce collective polarity by actively repressing protrusions from follower cells and by maintaining lead cell protrusions that pull the group forward (Mayor and Etienne-Manneville, 2016; Friedl and Mayor, 2017). Importantly, cell-cell adhesions keep collectives together by maintaining strong but flexible connections between cells. Moreover, many cell collectives exhibit a ‘supracellular’ organization of the cytoskeleton at the outer perimeter of the entire cell group that serves to further coordinate multicellular movement (Shellard and Mayor, 2019; Shellard et al., 2018; Hidalgo-Carcedo et al., 2011; Reffay et al., 2014). Despite progress in understanding how single cells become polarized and motile, less is known about the mechanisms that control the global organization, cohesion, and coordination of cells in migrating collectives.

Drosophila border cells are a genetically tractable and relatively simple model well-suited to investigate how cell collectives undergo polarized and cooperative migration within a developing tissue (Montell et al., 2012; Saadin and Starz-Gaiano, 2016). The Drosophila ovary is composed of strings of ovarioles made up of developing egg chambers, the functional unit of the Drosophila ovary. During late oogenesis, four to eight follicle cells are specified at the anterior end of the egg chamber to become migratory border cells. The border cells then surround a specialized pair of follicle cells, the polar cells, and delaminate as a multicellular cluster from the follicular epithelium. Subsequently, the border cell cluster undergoes a stereotyped collective migration, moving between 15 large germline-derived nurse cells to eventually reach the oocyte at the posterior end of the egg chamber (Figure 1A–F). Throughout migration, individual border cells maintain contacts with each other and with the central polar cells so that all cells move as a single cohesive unit (Llense and Martín-Blanco, 2008; Cai et al., 2014). A leader cell at the front extends a migratory protrusion whereas protrusions are suppressed in trailing follower cells (Prasad and Montell, 2007; Bianco et al., 2007; Poukkula et al., 2011). As with other collectives, polarization of the border cell cluster is critical for the ability to move together and in the correct direction, in this case towards the oocyte (Figure 1A–F; Prasad and Montell, 2007; Bianco et al., 2007).

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Polarization of the border cell cluster begins when two receptor tyrosine kinases (RTKs) expressed by border cells, PDGF- and VEGF-receptor related (PVR) and Epidermal Growth Factor Receptor (EGFR), respond to multiple growth factors secreted from the oocyte (Duchek et al., 2001; McDonald et al., 2006). Signaling through PVR/EGFR increases activation of the small GTPase Rac, triggering F-actin polymerization and formation of a major protrusion in the lead border cell (Prasad and Montell, 2007; Poukkula et al., 2011; Duchek et al., 2001; Wang et al., 2010). E-Cadherin-based adhesion to the nurse cell substrate stabilizes this lead cell protrusion via a feedback loop with Rac (Cai et al., 2014). Furthermore, the endocytic protein Rab11 and the actin-binding protein Moesin mediate communication between border cells to restrict Rac activation to the lead cell (Ramel et al., 2013). Mechanical coupling of border cells through E-Cadherin suppresses protrusions in follower cells, both at cluster exterior surfaces but also between border cells and at contacts with polar cells (Montell et al., 2012; Cai et al., 2014). E-Cadherin also maintains border cell attachment to the central polar cells. F-actin and non-muscle myosin II (Myo-II) are enriched at the outer edges of the border cell cluster (Aranjuez et al., 2016; Lucas et al., 2013; Combedazou et al., 2017). Such ‘inside-outside’ polarity contributes to the overall cluster shape, cell-cell organization, and coordinated motility of all border cells (Montell et al., 2012). While progress has been made in understanding the establishment of front-rear polarity, much less is known about how individual border cell behaviors are fine-tuned and adjusted to produce coordinated and cooperative movement of the cluster as an entire unit.

In the current study we made the unexpected discovery that Protein phosphatase 1 (Pp1) activity coordinates the collective behavior of individual border cells. Dynamic cycles of protein phosphorylation and dephosphorylation precisely control many signaling, adhesion and cytoskeletal pathways required for cell migration (Larsen et al., 2003). Serine-threonine kinases, such as Par-1, Jun kinase (JNK), and the p21-activated kinase Pak3, as well as phosphorylated substrate proteins such as the Myo-II regulatory light chain (MRLC; Drosophila Spaghetti squash, Sqh) and Moesin regulate different aspects of border cell migration (Llense and Martín-Blanco, 2008; Ramel et al., 2013; Majumder et al., 2012; Felix et al., 2015). In contrast, the serine-threonine phosphatases that counteract these and other kinases and phosphorylation events have not been extensively studied, either in border cells or in other cell collectives. Pp1 is a highly conserved and ubiquitous serine-threonine phosphatase found in all eukaryotic cells (Lin et al., 1999; Verbinnen et al., 2017). Pp1 can directly dephosphorylate substrates in vitro, but specificity for phosphorylated substrates in vivo is generally conferred by a large number of regulatory subunits (also called Pp1-interacting proteins [PIPs]). These regulatory subunits form functional Pp1 complexes through binding to the Pp1 catalytic (Pp1c) subunits and mediate the recruitment of, or increase the affinity for, particular substrates (Verbinnen et al., 2017; Heroes et al., 2013). Thus, despite the potential for pleiotropy, Pp1 complexes have specific and precise cellular functions in vivo, that range from regulation of protein synthesis, cell division and apoptosis to individual cell migration (Ceulemans and Bollen, 2004; Ferreira et al., 2019).

We now show that Pp1 activity controls multiple collective behaviors of border cells, including timely delamination from the epithelium, collective polarization, cohesion, cell-cell coordination, and migration. Remarkably, Pp1-inhibited border cells round up, break off from the main group, and move as single cells or small groups but are generally unable to complete their migration. We determine that Pp1 controls the levels of E-Cadherin and β-Catenin, which are needed to retain border cells within a cohesive cluster. Additionally, Pp1 activity restricts F-actin and Myo-II enrichment to the outer edges of the cluster, maintaining a supracellular cytoskeletal ultrastructure and supporting polarized collective movement. Furthermore, a major Pp1 specific complex for Myo-II activity, myosin phosphatase, coordinates border cell shape and adherence of cells to the cluster. Our work thus identifies Pp1 activity, mediated through distinctive phosphatase complexes such as myosin phosphatase, as a critical molecular regulator of collective cell versus single cell behaviors in a developmentally migrating collective.

To migrate collectively, cells need to coordinate and cooperate at the multicellular level. Individual cells within a group must remain together, maintain optimal cell shapes, organize motility of neighboring cells, and polarize. The mechanisms that globally orchestrate single cell behaviors within migrating cell collectives are still unclear. Here we report that Pp1 activity is a critical regulator of key intra- and intercellular mechanisms that together produce collective border cell migration. Loss of Pp1 activity, through overexpression of NiPp1 or Pp1c RNAi, switches border cells from migrating as a cohesive cluster to moving as single cells or in small groups (Figure 8A). A critical aspect of this switch is the redistribution of enriched F-actin and Myo-II to cell contacts between individual border cells, rather than at the cluster periphery, and a concomitant loss of adhesion between cells. We identified one key Pp1 phosphatase complex, myosin phosphatase, that controls collective-level myosin contraction (Figure 8B). Additional phosphatase complexes, through as-yet-unknown regulatory subunits, likely function in border cells to generate collective F-actin organization, maintain cell-cell adhesions, and potentially to restrain overall RhoA activity levels. Our results support a model in which balanced Pp1 activity promotes collective border cell cluster migration, and timely delamination from the epithelium, by coordinating single border cell motility and keeping the cells together (Figure 8A).

Figure 8

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Many collectively migrating cells require a supracellular enrichment of actomyosin at the group perimeter to help organize their movement (Shellard and Mayor, 2019; Shellard et al., 2018; Hidalgo-Carcedo et al., 2011; Reffay et al., 2014). Active Myo-II is required for border cell collective detachment from the epithelium, cluster shape, rotational movement of the cluster, and normal protrusion dynamics (Aranjuez et al., 2016; Combedazou et al., 2017; Majumder et al., 2012; Mishra et al., 2019; Fulga and Rørth, 2002). We show here that Pp1 organizes collective-level Myo-II-contractility during border cell migration. Inhibition of Pp1 shifts the balance of dynamic activated Myo-II from the cluster-level to individual border cells, resulting in rounded, hyper-contractile border cells that dissociate from the cluster. The myosin-specific Pp1 complex, myosin phosphatase, directly dephosphorylates Sqh and inhibits Myo-II activation (Grassie et al., 2011). Depletion of Mbs, the myosin-binding regulatory subunit of myosin phosphatase, causes rounder border cells and fragmentation of the cluster. We previously found that Mbs-deficient border cells have significantly higher levels of phosphorylated Sqh (p-Myo-II) (Majumder et al., 2012). Thus, myosin phosphatase inhibits Myo-II activation to promote coordinated collective contractility of border cells. Myosin phosphatase is a downstream target of Rok, which phosphorylates and inhibits the Mbs subunit (Kimura et al., 1996). Consistent with loss of myosin phosphatase activity, Pp1-inhibition increases phosphorylated active Sqh in individual border cells within the cluster. Thus, myosin phosphatase, downstream of Rok, promotes elevated active Myo-II (p-Sqh/p-Myo-II) and cortical contraction of the entire collective (Figure 8B). Interestingly, expression of constitutively activated RhoA also induces cellular hypercontractility, resulting in amoeboid-like round border cells (Aranjuez et al., 2016; Combedazou et al., 2017; Mishra et al., 2019). RhoA activates Rok, which directly phosphorylates and activates the Myo-II regulatory subunit Sqh (Amano et al., 1996; Matsui et al., 1996). We observe somewhat elevated RhoA activity in the absence of Pp1 activity. Thus, Pp1 may also restrain the overall levels of RhoA activity in border cells through an unknown Pp1 complex, which would further promote the collective actomyosin contraction of border cells (Figure 8B).

Myo-II is activated preferentially at the cluster periphery and not between internal border cell contacts. Mbs and at least one catalytic subunit, Flw, localize uniformly in border cells, both on the cluster perimeter and between cells. Such uniform phosphatase distribution would be expected to dephosphorylate and inactivate Myo-II everywhere, yet phosphorylated Sqh is only absent from internal cluster border cell contacts. Rok phosphorylates and inactivates Mbs in addition to directly activating Myo-II (Kimura et al., 1996). Our previous results indicate that Rok localizes to the cluster perimeter similar to p-Sqh, but there appeared to be overall less Rok between border cells (Aranjuez et al., 2016). Thus, spatially localized Rok could inhibit myosin phosphatase and activate Myo-II preferentially at the outer edges of the cluster (Figure 8A). Other mechanisms likely contribute to collective polarization of Myo-II. For example, during border cell detachment from the epithelium the polarity kinase Par-1 phosphorylates and inactivates Mbs at the cluster rear resulting in increased active Myo-II, whereas the Hippo pathway prevents accumulation of phosphorylated Myo-II between border cells (Lucas et al., 2013; Majumder et al., 2012).

Our data also support a role for Pp1 in controlling F-actin stability, dynamics, and spatial organization. Similar to the pattern of activated Myo-II, cortical F-actin is normally high at the cluster periphery, although low levels are found between border cells (Ramel et al., 2013; Lucas et al., 2013; Wang et al., 2018). Reduced Pp1 activity causes high levels of F-actin to redistribute from the cluster perimeter to surround entire cell cortices of individual border cells. In migrating cells, networks of F-actin produce forces essential for protrusion extension and retraction dynamics that generate forward movement (Ridley, 2011; Caswell and Zech, 2018). Further supporting a role for Pp1 in regulating F-actin, Pp1-inhibited border cells extend fewer protrusions with shorter lifetimes, resulting in altered motility patterns. How Pp1 promotes F-actin organization and dynamics is unknown. One possibility comes from the known function for Rok in regulating F-actin through the downstream effector LIM Kinase (LIMK) (Julian and Olson, 2014). LIMK phosphorylates and inhibits cofilin, an actin severing and depolymerizing factor (Bravo-Cordero et al., 2013). In border cells, cofilin restrains F-actin levels throughout the cluster and increases actin dynamics, resulting in normal cluster morphology and major protrusion formation (Zhang et al., 2011). Although cofilin dephosphorylation, and thus activation, is typically mediated by the dual-specificity phosphatase Slingshot (Bravo-Cordero et al., 2013), Pp1-containing complexes have been shown to dephosphorylate cofilin in a variety of cell types (Huet et al., 2013; Ambach et al., 2000; Zhang et al., 2012; Oleinik et al., 2010). Additionally, RhoA activates formin proteins such as Diaphanous, which nucleate actin to form long filaments (Kühn and Geyer, 2014). There are at least seven formin-related proteins in Drosophila, several of which have domains associated with activation by Rho GTPases. However, which formin, if any, promotes border cell migration and F-actin distribution is unknown. Further work will be needed to determine whether any of these potential targets, or other actin regulatory proteins, control collective level F-actin enrichment via Pp1.

A major consequence of decreased Pp1 activity is fragmentation of the border cell cluster into single border cells and small groups. This raises the question of how Pp1 activity maintains cluster cohesion, which is critical for collective cell movement in vivo. Like many cell collectives, high levels of cadherin-catenin complex proteins are detected between all border cells (Niewiadomska et al., 1999). The cadherin-catenin complex is required for border cells to adhere to the central polar cells as well as to provide migratory traction of the entire cluster upon the nurse cells (Cai et al., 2014; Niewiadomska et al., 1999). We found that Pp1 maintains E-Cadherin and β-Catenin levels between border cells. Indeed, other mutants that disrupt the levels and localization of adhesion proteins in border cells often also disrupt cluster shape and cohesion. For example, loss of JNK signaling causes border cell clusters to dramatically elongate, with downregulation of adhesion resulting in incomplete separation of border cells (Llense and Martín-Blanco, 2008; Melani et al., 2008). Raskol, a putative Ras guanine nucleotide activating protein (GAP), maintains E-cadherin at BC-BC contacts and cohesion of the cluster (Raza et al., 2019). However, while loss of Raskol causes a significant number of border cells to fully dissociate from the cluster (~35%) (Raza et al., 2019), similar to what we observe with knockdown of the cadherin-catenin complex, this is less than what we observe upon inhibition of Pp1 activity (~90%). Thus, while cluster fragmentation caused by Pp1 inhibition is at least partly due to deficient cadherin-catenin adhesion, other targets likely contribute.

Our results indicate that E-Cadherin, β-Catenin, and α-Catenin maintain adhesion of border cells to each other in addition to known roles in keeping border cells attached to the polar cells (Cai et al., 2014). Knockdown of the cadherin-catenin complex members in both border cells and polar cells causes border cells to significantly dissociate from the cluster. The requirement in border cells for cadherin-catenin in cluster cohesion may have been masked in prior studies due to the inability of strong loss-of-function cadherin-catenin mutant border cells to move at all (Cai et al., 2014; Niewiadomska et al., 1999; Sarpal et al., 2012; Desai et al., 2013). While RNAi for E-Cadherin, β-Catenin, and α-Catenin each strongly knock down the respective protein levels, it may be that a small amount of each protein is still present. Such remaining cadherin-catenin proteins may provide just enough traction for border cells to partially migrate upon the nurse cells. We speculate that movement of cadherin-catenin-deficient border cells within the confining tissue would provide mechanical stresses that break the cluster apart at weakened border cell-border cell contacts. Indeed, a mutant α-Catenin protein that lacks part of the C-terminal F-actin-binding domain was shown to partially rescue the migration defects caused by loss of α-Catenin; however, these rescued border cell clusters split into several parts along the migration path (Desai et al., 2013). Further supporting this idea, Pp1-inhibited border cells fall apart during their effort to migrate between the nurse cells.

How do Pp1 phosphatase complexes molecularly promote cluster cohesion? Given the effects of Pp1 on E-Cadherin and β-Catenin at internal border cell contacts, and the requirement for cadherin-catenin complex proteins in maintaining cluster integrity, Pp1 could directly regulate cadherin-catenin protein stability and/or adhesive strength. In mammalian and Drosophila cells, phosphorylation of a conserved stretch of serine residues in the E-Cadherin C-terminal tail region regulates E-Cadherin protein stability, binding of E-Cadherin to β-Catenin, and cell-cell junction formation and turnover (Stappert and Kemler, 1994; McEwen et al., 2014; Chen et al., 2017). Serine-phosphorylation of α-Catenin is also required for adhesion between epithelial cells and possibly for efficient border cell migration (Escobar et al., 2015). More work will be needed to determine whether a to-be-identified Pp1-containing phosphatase complex directly dephosphorylates E-Cadherin and/or α-Catenin, as the roles for phosphatases in cadherin-catenin junctional stability are still poorly understood.

Alternatively, or in addition, Pp1-dependent restriction of collective actomyosin contraction to the cluster periphery could allow internal cluster cell-cell junctions to be maintained. Pp1-inhibition greatly alters actomyosin distribution, causing individual border cells to contract and round up. The forces transmitted by high cell contractility alone could weaken adherens junctions, causing the border cells to break apart during migration (Figure 8A). Myosin phosphatase-depleted border cells, which have elevated phosphorylated Sqh (Majumder et al., 2012), and thus active Myo-II, are round, highly contractile, and fall off the cluster. In support of this idea, overexpression of a phosphorylation mutant form of Sqh (Sqh^E20E21), which mimics activated Myo-II, causes border cells to have a similar round shape and separate from the cluster (Mishra et al., 2019). Thus, collective-level active actomyosin contraction contributes to keeping border cells adhered to the cluster. Myo-II and cadherin-catenin complexes have dynamic and quite complex interactions that influence stability of cell-cell junctions, and which may depend on cellular context (Mège and Ishiyama, 2017; Yap et al., 2018). In border cells, the cadherin-catenin complex promotes enrichment of actomyosin to the cluster periphery, whereas Myo-II does not greatly influence cadherin-catenin levels within the cluster (this study) (Mishra et al., 2019). However, Pp1 is required for the proper distribution (or stability) of cadherin-catenin at cell contacts between border cells and prevents the enrichment of actomyosin in individual border cells. Moreover, NiPp1 expression disrupts cluster cohesion to a greater extent than knockdown of either myosin phosphatase or cadherin-catenin complex members alone. This suggests that cadherin-catenin stability and optimal collective-wide actomyosin activity both contribute to cluster cohesion through distinct Pp1 phosphatase complexes, although this possibility remains to be formally tested (Figure 8B).

Our study implicates Pp1 as a major regulator of collective cohesion and migration in border cells. Pp1 catalytic subunits and their regulatory subunits are conserved across eukaryotes (Lin et al., 1999; Verbinnen et al., 2017; Heroes et al., 2013; Ferreira et al., 2019). The roles of specific Pp1 complexes in collective cell migration during development and in cancer have not been well studied. Intriguingly, Mypt1 (Mbs homolog) promotes polarized mesodermal migration during zebrafish gastrulation (Weiser et al., 2009). Similar to what we observe in Mbs-depleted border cells, inhibition of zebrafish Mypt1 switched cells from an elongated mesenchymal mode of migration to a hyper-contractile amoeboid mode of migration. Another Pp1 phosphatase complex containing the Phactr4 (phosphatase and actin regulator 4) regulatory subunit promotes the chain-like collective migration of enteric neural crest cells, which colonize the gut and form the enteric nervous system during development (Zhang et al., 2012). Phactr4, through Pp1, specifically controls the directed migration and shape of enteric neural crest cells through integrin, Rok, and cofilin. Given the conservation of these and other phosphatase complexes, our study highlights the importance of balanced Pp1 phosphatase activity in the organization and coordination of migrating cell collectives.

All data generated or analysed during this study are included in the manuscript and supporting files.

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Author details

1. Yujun Chen

   Division of Biology, Kansas State University, Manhattan, United States

   Contribution

   Conceptualization, Formal analysis, Funding acquisition, Validation, Investigation, Visualization, Methodology

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-3190-6713

2. Nirupama Kotian

   Division of Biology, Kansas State University, Manhattan, United States

   Contribution

   Formal analysis, Validation, Investigation, Visualization, Methodology

   Competing interests

   No competing interests declared

3. George Aranjuez

   Lerner Research Institute, Cleveland Clinic, Cleveland, United States

   Present address

   Division of Immunity and Pathogenesis, Burnett School of Biomedical, Sciences, University of Central Florida College of Medicine, Orlando, United States

   Contribution

   Formal analysis, Validation, Investigation, Visualization, Methodology

   Competing interests

   No competing interests declared

4. Lin Chen

   LBCMCP, Centre de Biologie Intégrative (CBI), Université de Toulouse, CNRS, UPS, Toulouse, France

   Contribution

   Formal analysis, Validation, Investigation, Visualization, Methodology

   Competing interests

   No competing interests declared

5. C Luke Messer

   Division of Biology, Kansas State University, Manhattan, United States

   Contribution

   Formal analysis, Validation, Investigation, Visualization, Methodology

   Competing interests

   No competing interests declared

6. Ashley Burtscher

   Lerner Research Institute, Cleveland Clinic, Cleveland, United States

   Contribution

   Formal analysis, Validation, Investigation, Visualization, Methodology

   Competing interests

   No competing interests declared

7. Ketki Sawant

   Division of Biology, Kansas State University, Manhattan, United States

   Contribution

   Formal analysis, Validation, Investigation, Visualization, Methodology

   Competing interests

   No competing interests declared

8. Damien Ramel

   LBCMCP, Centre de Biologie Intégrative (CBI), Université de Toulouse, CNRS, UPS, Toulouse, France

   Present address

   Institute of Metabolic and Cardiovascular Diseases (I2MC), Université de Toulouse, Institut National de la Santé et de la Recherche Médicale (INSERM) UMR1048, Toulouse, France

   Contribution

   Conceptualization, Formal analysis, Validation, Investigation, Visualization, Methodology

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-9089-1497

9. Xiaobo Wang

   LBCMCP, Centre de Biologie Intégrative (CBI), Université de Toulouse, CNRS, UPS, Toulouse, France

   Contribution

   Conceptualization, Resources, Formal analysis, Supervision, Funding acquisition

   Competing interests

   No competing interests declared

10. Jocelyn A McDonald

    Division of Biology, Kansas State University, Manhattan, United States

    Contribution

    Conceptualization, Formal analysis, Supervision, Funding acquisition, Validation, Investigation, Visualization, Methodology

    For correspondence

    jmcdona@ksu.edu

    Competing interests

    No competing interests declared

    "This ORCID iD identifies the author of this article:" 0000-0002-7494-1466

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