Abstract
Fascin is an actin bundling protein that is essential for developmental cell migrations and promotes cancer metastasis. In addition to bundling actin, Fascin has several actin-independent roles. Border cell migration during Drosophila oogenesis provides an excellent model to study Fascin’s various roles during invasive, collective cell migration. Border cell migration requires Fascin. Fascin functions not only within the migrating border cells, but also within the nurse cells, the substrate for this migration. Loss of Fascin results in increased, shorter and mislocalized protrusions during migration. Data supports the model that Fascin promotes the activity of Enabled, an actin elongating factor, to regulate migration. Additionally, loss of Fascin inhibits border cell delamination. These defects are partially due to altered E-cadherin localization in the border cells; this is predicted to be an actin-independent role of Fascin. Overall, Fascin is essential for multiple aspects of this invasive, collective cell migration, and functions in both actin-dependent and -independent manners. These findings have implications beyond Drosophila, as border cell migration has emerged as a model to study mechanisms mediating cancer metastasis.
Introduction
Fascin is an actin-binding protein that bundles or cross-links actin filaments (Hashimoto et al., 2011; Jayo and Parsons, 2010) to promote cell motility and invasion through the formation of filopodia and invadopodia (Adams, 2004; Li et al., 2010; Zanet et al., 2012). While Fascin does promote cell migration in this actin-dependent manner, novel actin-independent roles of Fascin have been discovered (Anilkumar et al., 2003; Jayo et al., 2016; Villari et al., 2015). Fascin directly binds the Linker of the Nucleoskeleton and Cytoskeleton (LINC) complex, which mediates mechanotransduction. Perturbing this interaction impairs nuclear shape deformation essential for single-cell invasive migration (Jayo et al., 2016). Fascin also binds to microtubules and loss of this interaction increases the stability of cellular adhesions causing slower migration (Villari et al., 2015). Additionally, Fascin interacts with Protein Kinase C (PKC), LIM kinases (LIMKs), and, notably, Enabled (Ena; (Anilkumar et al., 2003; Hashimoto et al., 2007; Jayo et al., 2012; Winkelman et al., 2014). Ena is an actin elongation factor, and in vitro Ena processivity is increased on Fascin-bundled actin (Harker et al., 2019; Winkelman et al., 2014). These studies illustrate Fascin has multiple functions within the cell that regulate cell migration.
Fascin is important for both developmental cell migrations and cancer metastasis (Cohan et al., 2001; Hashimoto et al., 2011; Hashimoto et al., 2005). Fascin controls cell migration during development such as, growth cone extension, dendrite formation, and in embryonic fibroblasts (De Arcangelis et al., 2004; Ma et al., 2013). Fascin is also highly upregulated in certain types of cancer, and elevated expression is associated with increased invasiveness, aggressiveness and mortality (Arlt et al., 2019; Hashimoto et al., 2011; Hashimoto et al., 2007). While Fascin has been studied in the contexts of 2D migration and single cell 3D migration, the roles of Fascin in invasive, collective cell migration have yet to be investigated (Adams, 2004; Jayo et al., 2016).
Drosophila oogenesis – specifically border cell migration – is an ideal model to study invasive, collective cell migration. Drosophila oogenesis has 14 developmental stages of the egg chambers or follicles (Spradling, 1993). Each follicle is composed of a single oocyte, 15 germline-derived nurse cells, and a layer of somatic epithelial cells, or follicle cells, surrounding the outside. During Stage 9 (S9) of follicle development, a group of follicle cells at the anterior end are specified to become border cells. This group of 8-10 border cells delaminates from the epithelium and migrate between the nurse cells to reach the nurse cell-oocyte boundary (Montell, 2003; Montell et al., 2012). Delamination is a highly regulated process in which the border cells must maintain cellular adhesions, such as E-cadherin, amongst themselves, but sever adhesions with their neighboring follicle cells and nurse cells (Cai et al., 2014; Montell, 2003). Additionally, border cell migration is very dynamic with protrusions extending and retracting to move the cluster (Fulga and Rorth, 2002; Prasad and Montell, 2007). Upon completing its migration, the border cells produce the micropyle, the structure through which sperm fertilize the egg (Montell et al., 1992; Spradling, 1993). Importantly, the migrating border cells highly express Fascin (Cant et al., 1994). Therefore, we can study the role of Fascin in invasive, collective cell migration in vivo using the simple and genetically tractable model of border cell.
Here we find that Fascin plays a critical role in regulating border cell migration. Using a new quantification method that assesses border cell migration during S9, we find loss of Fascin results in significant delays in border cell migration. Surprisingly, Fascin is necessary in both the germline cells and somatic cells but is only sufficient within the somatic cells to promote border cell migration. Live imaging reveals that loss of Fascin results in border cell clusters with more protrusions emerging from all sides that are shorter in length and duration. These alterations culminate in the fascin-null clusters migrating slower than controls. These defects are due, in part, to Fascin’s role in regulating Ena. Dominant genetic interactions reveal Fascin and Ena work together to regulate border cell migration, and overexpression of Ena suppresses migration defects in fascin-null mutants. Fascin also regulates border cell delamination. In fascin-null mutants, the clusters take longer to delaminate, and display altered localization of E-cadherin. Overall, our data reveal that Fascin regulates multiple aspects of border cell migration, including both protrusion dynamics and delamination. These findings suggest that Fascin regulates invasive, collective cell migration through modulating cellular protrusions by bundling actin and regulating cellular adhesions to control initiation of migration.
Results
Fascin is required for border cell migration
Previously, it was reported that loss of Fascin does not affect border cell migration (Cant et al., 1994). This analysis showed that border cells of fascin-null mutants completed their migration between the nurse cells and reached the nurse cell-oocyte boundary by Stage 10A (S10A) (Cant et al., 1994); we have reproduced these findings (Fig. S1). These findings are surprising as Fascin is highly expressed in the border cells (Cant et al., 1994) and is known to regulate many types of cell migration (Hashimoto et al., 2011; Jayo and Parsons, 2010). Thus, we hypothesized that while border cells reach the nurse cell-oocyte boundary by S10, border cell migration may be delayed during S9 in fascin mutants.
Border cell migration is a highly regulated event; at any point during S9, the distance the border cells have migrated is approximately equal to the distance the outer follicle cells are from the anterior (Fig 1A). Thus, delayed or accelerated migration of the border cells can be quantitatively assessed by comparing their location relative to that of the outer follicle cells. Specifically, by subtracting the distance of the outer follicle cells from the anterior end of the follicle from the distance the border cells have migrated we can calculate the migration index (Fig. 1A). A migration index of ∼0 indicates on-time migration, while a negative value indicates delayed migration and a positive value indicates accelerated migration (Fig. 1A).
To assess border cell migration during S9, we performed immunofluorescent staining for Hts and FasIII, this stain labels both border cells (yellow arrow) and outer follicle cells (yellow line) and enables us to assess border cell migration and quantify migration index (Fig. 1B-G). This stain will be referred to throughout the paper as the border cell migration stain. Using this, we quantified migration index in wild-type and fascin mutant follicles (Fig. 1B-G). Two different null alleles of fascin were used, fascinsn28 and fascinsnX2 (Cant and Cooley, 1996; Cant et al., 1994). S9 follicles that were wild-type or heterozygous for mutations in fascin display on-time border cell migration with the border cell cluster being in line with the outer follicle cells (Fig. 1B-D and G; migration indices of 4.23, −0.48 and −1.35, respectively). Loss of Fascin by both homozygous (fascinsn28/sn28 and fascinsnX2/snX2) and transheterozygous fascin mutations (fascinsn28/snX2 and fascinsnX2/sn28; maternal allele is listed first) results in border cell clusters that are significantly delayed (Fig. 1E-G; migration indices of −21.20 (p<0.0001), −15.27 (p<0.0001), −17.25 (p<0.0001) and −10.85 (p=0.0363), respectively). These data reveal Fascin is required for proper on-time border cell migration during S9 of Drosophila oogenesis.
Fascin is necessary in both the germline and somatic cells for border cell migration
We next sought to identify where Fascin is needed for border cell migration. During S9, while Fascin is most highly expressed in the border cell cluster, the nurse cells and outer follicle cells also express Fascin (see Fig. S2A; (Cant et al., 1994)). Moreover, the nurse cells are the substrate upon which the border cells migrate and changes in nurse cell structure or stiffness perturb border cell migration (Aranjuez et al., 2016; Cai et al., 2016).
We used the UAS/GAL4 system (Rorth, 1998) to express Fascin RNAi constructs to knockdown Fascin in specific cell types and determine the effects on border cell migration. Two different Fascin RNAi lines were used (second chromosome: TRiP.HMJ21813 and third chromosome: TRiP.HMS02450) and yielded similar results; data presented uses the third chromosome line. We knocked down Fascin in all somatic cells (c355 GAL4), in the border cells (c306 GAL4), or in the germline cells (matα GAL4). Knockdown of Fascin was confirmed by immunostaining for Fascin (Fig. S2A-E’). Knockdown of Fascin in all somatic cells (c355 GAL4) causes signification border cell migration delays compared to the GAL4 driver only and RNAi only controls (Fig. 2A, B, E; migration indices of −22.97 compared to 0.76 and −7.14; p<0.0001). Similarly, knockdown of Fascin in only the border cells (c306 GAL4) causes delayed migration, however, this is not significant compared to controls (Fig. 2A, C, E; migration indices of −13.94 compared to −4.92 and −7.14; p=0.051). This mild phenotype is likely due to insufficient knockdown of Fascin during migration, as high levels of Fascin in observed in the border cells at early stages of migration (Fig. S2D-D’) and diminishing levels at the later stages (Fig. S2E-E’). Notably, knockdown of Fascin in the germline cells (matα GAL4) also significantly delays border cell migration compared to controls (Fig. 2A, D, E; migration indices of −27.80 compared to −7.25 and −7.14; p<0.0001). These findings indicate Fascin is necessary in both the somatic and germline for border cell migration.
Somatic expression of Fascin rescues border cell migration
We next asked in what cell types is Fascin sufficient for normal border cell migration. The UAS/GAL4 system was used to express GFP-Fascin in specific cell types of fascin mutant follicles to determine where restoring expression rescues border cell migration. We expressed GFP-Fascin in the somatic cells (c355 GAL4), the germline cells (oskar GAL4), or in both the germline and somatic cells (actin5C GAL4) (see Table S1 for all statistical comparisons). Expression of GFP-Fascin in the somatic cells of fascin mutant follicles restores border cell migration (Fig. 3A, B, G; migration indices −0.35 compared to −20.19; p=0.0004). Conversely, expression of GFP-Fascin in the germline cells of fascin mutant follicles fails to rescue border cell migration (Fig. 3C, D, G; migration indices of −20.97 compared to −26.49; p=0.49). Finally, expression of Fascin in both the somatic and germline cells of fascin mutant follicles restores border cell migration (Fig. 3E-G; migration indices of 10.99 compared to −26.43; p<0.0001). Thus, Fascin is necessary but not sufficient in the germline cells, whereas Fascin is both necessary and sufficient within the somatic cells to promote proper border cell migration.
Fascin regulates protrusion dynamics in the migrating border cell cluster
To determine how loss of Fascin causes delayed border cell migration we utilized live imaging. We visualized border cell migration with membrane localized GFP expressed under the control of the slbo promoter (slbo>mCD8-GFP), which specifically labels the border cells and allows us to analyze cluster protrusions.
During migration, the border cell cluster typically forms one or two large protrusions that extend and retract from the leading end of the cluster as it migrates (Bianco et al., 2007; Prasad and Montell, 2007). In agreement with this, control clusters (fascinsn28/+) typically have one or two main protrusions extending and retracting from the front of the cluster (Fig. 4A-A”, Movie 1). Conversely, in fascin-null mutants (fascinsn28/sn28) the clusters extend many protrusions from their front, sides, and back (Fig. 4B-B”, Movie 2). Clusters in control follicles have just one protrusion in 64% of the frames analyzed versus 34% of the frames in fascin-null follicles (Fig. 4C). Furthermore, the clusters in fascin-null follicles have a higher percentage of frames with 3-4 protrusions (19%) compared to those of controls (1%) (Fig. 4C; p<0.0001, Pearson’s chi-squared test). Moreover, we assessed the localization of the protrusions on the cluster: front (0° to 45° and 0° to 315°), sides (45° to 135° and 225° to 315°), or back (135° to 225°) of the cluster (Sawant et al., 2018). The fascin-null clusters have a significantly altered protrusion localization with 43% of the protrusions emerging from either the side or back of the cluster compared to 17% for the control clusters (Fig. 4D; p<0.0001, Pearson’s chi-squared test).
In addition to quantifying protrusions per frame, we measured the protrusion length and binned them based on their directionality in the same manner as described above. The protrusions that emerge from the front of the cluster are typically longest in length (Bianco et al., 2007; Prasad and Montell, 2007). Protrusions extending from the front of the cluster were significantly longer in control clusters compared to fascin-null clusters (Fig. 4E; 9.3μm compared to 7.5μm, respectively; p=0.045). Additionally, in control clusters, the protrusions extending from the front are significantly longer than the protrusions extending from the sides (Fig. 4E; front=9.3μm, sides=6.6μm; p=0.047). Conversely, fascin-null clusters extend protrusions of similar lengths from all sides of the cluster (Fig. 4E; front=7.5μm, sides=6.8μm, and back=7.2μm). Additionally, protrusion duration is significantly shorter in the fascin-null clusters, with the average duration being 20min compared to 43.4min for controls (Fig. 4F, p<0.0001).
Lastly, we quantified the migration speed of clusters during mid-migration. Loss of Fascin results in significantly slower migration (0.24μm/min) compared to controls (0.51μm/min; Fig. 4G; p=0.0019). Overall, these data indicate the loss of Fascin impairs protrusion formation and regulation within the cluster, and these impairments cause slower migration speeds.
Fascin regulates Ena to promote border cell migration
We next wanted to determine how Fascin regulates protrusions during border cell migration. Recent findings demonstrate that Fascin cooperates with the actin elongation factor Ena to promote actin polymerization and filament formation in vitro by enhancing Ena processivity (Harker et al., 2019; Winkelman et al., 2014). Additionally, loss of Ena causes border cell migration defects (Gates et al., 2009). Based on these data, we hypothesize Fascin regulates border cell cluster protrusions by promoting Ena activity.
To assess if Fascin regulates Ena during border cell migration we used dominant genetic interactions studies. Reduced levels of Fascin (fascin-/+) or Ena (ena-/+) alone should be sufficient to maintain normal border cell migration. If Fascin and Ena function together to mediate border cell migration, then reduced levels of both (fascin-/+; ena-/+) will exhibit delayed border cell migration. Partial loss of Fascin (data not shown) or two different ena alleles ena210/+ (Fig. 5A) and ena23/+ (data not shown) exhibit on-time border cell migration (Fig. 5C; migration indices of 1.29, −0.85, and −0.86, respectively). However, double heterozygotes of fascin with either ena allele (fascin-/+; ena-/+) causes significant border cell migration delays (Fig. 5B-C; migration indices of −15.58 (p=0.0015) and −20.25 (p=0.0002)). While the dominant genetic interaction results support our hypothesis that Fascin regulates Ena to control border cell migration, if our hypothesis is correct then overexpression of Ena is predicted to suppress the border cell migration delay observed in fascin-null follicles. Indeed, expression of RFP-tagged Ena in fascin mutant follicles restores on-time migration (Fig. 5D-F; migration indices of −5.26 compared to −31.63; p=0.0002). These findings support the model that the defects observed in the protrusion dynamics of fascin mutant clusters are due, at least in part, to Fascin’s role in regulating Ena to promote actin elongation.
Fascin regulates the delamination of the border cells
In addition to regulating protrusions during migration, Fascin also contributes to border cell delamination. Delamination is the process by which the border cell cluster detaches from the surrounding follicle cells to begin its migration. Live-imaging of follicles during delamination revealed fascin-null follicles spend more time detaching from the follicular epithelium (Fig. 6B-B” and Movie 4 compared to 6A-A” and Movie 3). We quantified this change in delamination time by measuring the amount of time elapsed from cluster formation to when the cluster is fully delaminated during early S9. The fascin-null clusters take significantly longer to delaminate (320min) compared to control clusters (147min, Fig. 6C; p<0.0001). Additionally, 3 fascin-null clusters failed to delaminate during the course of imaging (Fig 6C, indicated by x’s). These results suggest that Fascin promotes border cell delamination.
One process that is essential for delamination is disassembly of cell-cell adhesions between border cells and neighboring follicle and nurse cells (Cai et al., 2014; De Graeve et al., 2012; Niewiadomska et al., 1999). One adhesion molecule that must be regulated is E-cadherin (Cai et al., 2014; De Graeve et al., 2012). Both increasing or decreasing E-cadherin levels in the nurse cells impairs border cell migration (Cai et al., 2014). We were unable to assess dominant genetic interactions between e-cadherin and fascin mutants because heterozygosity for mutations in e-cadherin resulted in border cell migration delays (data not shown). Therefore, we assessed E-cadherin by immunofluorescence. As initial differences in E-cadherin between wild-type and fascin-null delaminating clusters were subtle, samples were stained in the same tube for further analyses. Delaminating border cell clusters in fascin-null follicles retain intense E-cadherin localization at all cell-cell boundaries (Fig. 7C-D compared to A-B), with stronger E-cadherin intensity at the periphery of the cluster (border cell-nurse cell boundary) compared to control clusters as observed by both intensity labeling (Fig. 7D compared to B, yellow arrowheads) and line-scan analysis (Fig. 7F compared to E). Additionally, unlike fascin-null clusters (Fig. 7C-D, F), E-cadherin intensity in control clusters is much lower at the cluster periphery (border cell-nurse cell boundary) than the border cell-polar cell boundaries (Fig. 7A-B, E). These results suggest that Fascin is required for reducing E-cadherin at the border cell cluster boundary, which is necessary for delamination.
Discussion
Here we provide evidence that Fascin regulates invasive, collective cell migration through multiple functions. Specifically, loss of Fascin results in delays in border cell migration during S9 of Drosophila oogenesis (Fig. 1). Fascin functions not only within the border cells, but also in the nurse cells, the substrate on which the border cells migrate, to mediate migration (Figs. 2-3). While Fascin’s role within the nurse cells remains unknown, it may involve both actin-dependent and -independent functions (see further discussion below). Within the border cells, Fascin regulates cluster protrusions (Fig. 4). This regulation is likely achieved through the actin bundling function of Fascin and regulation of the actin elongation activity of Ena (Fig. 5). Additionally, loss of Fascin impairs border cell delamination (Fig. 6), which may be the result of altered E-cadherin localization in the delaminating cluster (Fig. 7). This defect, as discussed below, is likely due to an actin-independent function of Fascin. Ultimately, our findings suggest that Fascin’s roles in invasive, collective cell migration can be attributed to both its actin-dependent and actin-independent functions.
Multiple cell types within the follicle require Fascin for border cell migration. We discovered that Fascin is necessary in both the somatic and germline cells, and sufficient within the somatic cells for border cell migration (Figs. 2-3). The roles of Fascin within the border cells are discussed below, here we speculate on the roles of Fascin within the nurse cells, the substrate on which the border cells migrate. We hypothesize that loss of Fascin alters the stiffness of the nurse cells which impairs border cell migration. Increasing nurse cell stiffness by enhancing non-muscle myosin II contractility impairs border cell migration (Aranjuez et al., 2016; Cai et al., 2016). Interestingly, in vitro Fascin inhibits non-muscle myosin II (Elkhatib et al., 2014). Therefore, loss of Fascin in the nurse cells may increase non-muscle myosin II contractility resulting in stiffer nurse cells and delayed border cell migration. Another means by which Fascin may alter the nurse cell stiffness is by controlling nurse cell-nurse cell adhesion. Indeed, we find that E-cadherin levels are higher on all cell membranes, including the nurse cells during S9 (Fig. 7). Such increased adhesion may impede border cell migration. Finally, Fascin may regulate the structure of the cortical actin in the nurse cells to control stiffness, as loss of Fascin results in cortical actin breakdown during mid-oogenesis (Groen et al., 2012). Further studies are needed to understand how Fascin functions within the germline to modulate border cell migration.
One way by which Fascin functions within the border cells to regulate migration is through controlling protrusion formation and dynamics. Loss of Fascin results in shorter and more protrusions (Fig. 4B-E). Consistent with this finding, in both Drosophila and cancer cells loss of Fascin results in shorter protrusions during single cell migration (Alam et al., 2012; Zanet et al., 2012). Additionally, Fascin is regulated by and interacts with PKC (Adams et al., 1999) and disruption of this interaction increases cellular protrusions (Anilkumar et al., 2003). Moreover, atypical PKC zeta regulates border cell migration (Wang et al., 2018), but it is unclear if other forms of PKC also do this. Further exploration of PKC regulation of Fascin in border cell protrusion formation and migration is warranted. Protrusion duration is also shorter in fascin-null clusters (Fig. 4F). This observation is consistent with the finding that Fascin contributes to protrusion persistence by stabilizing actin bundles (Bear et al., 2000). Fascin may stabilize protrusions by regulating the actin elongation factor Ena. Previous in vitro studies uncovered that Ena has increased processivity on actin bundled specifically by Fascin (Harker et al., 2019; Winkelman et al., 2014). These findings suggest that Fascin may regulate the activity of Ena to promote proper protrusion formation and dynamics required for border cell migration. Supporting this idea, dominant genetic interaction studies indicate that Fascin and Ena work within the same pathway to regulate border cell migration (Fig. 5A-C). Additionally, overexpression of Ena restores border cell migration in fascin mutants (Fig. 5D-F). These findings indicate that loss of Fascin results in decreased Ena activity and supports the model that Fascin acts to increase Ena processivity to promote stable protrusions necessary for mediating border cell migration. Further studies are needed uncover how this interaction influences protrusions during migration in vivo.
Another role of Fascin during border cell migration is regulating the delamination of the cluster. Loss of Fascin results in significantly longer delamination times (Fig. 6). Contributing to this delamination defect is the retention of high levels of E-cadherin on the membranes of the cluster, particularly at the border cell-nurse cell boundaries (Fig. 7). Proper levels of E-cadherin between the nurse cells and border cells are necessary for migration, as knockdown or overexpression of E-cadherin in the border cells or nurse cells results in impaired border cell migration (Cai et al., 2014; Niewiadomska et al., 1999). Therefore, persistence of E-cadherin along this boundary may impair border cell delamination. Other cell adhesion proteins are also required for border cell migration and delamination, including integrins (Dinkins et al., 2008; Villari et al., 2015). Integrins are highly dynamic during cell migration, and either increased or decreased stability of these adhesions impedes migration (Delon and Brown, 2007). Interestingly, Fascin has been proposed to promote integrin-based adhesion dynamics through its interaction with microtubules (Villari et al., 2015). Disruption of Fascin binding to microtubules leads to increased integrin adhesion stability resulting in decreased cell migration (Anilkumar et al., 2003; Villari et al., 2015). This data leads us to speculate that Fascin may control integrin dynamics during border cell delamination and migration through interaction with microtubules. Future studies are needed to test the role of Fascin in regulating cell adhesion dynamics required for border cell delamination and migration.
Fascin may also regulate border cell migration by mediating mechanotransduction. A key mediator of mechanotransduction is the LINC complex. The LINC Complex interacts with the cytoskeletal filaments within the cytoplasm and extends into the nucleus where it interacts with the nuclear lamina (Lombardi et al., 2011; Lombardi and Lammerding, 2011). This structure allows transmission force from the outside of the cell to the nucleus and plays a critical function in regulating nuclear shape and position during invasive cell migration (Alam et al., 2015; Harada et al., 2014; Lombardi et al., 2011; Lombardi and Lammerding, 2011). Fascin binds directly to the cytoplasmic part of the LINC complex in both the Drosophila ovary and mammalian cultured cells, and disruption of this interaction impairs nuclear deformation required for mammalian single cell invasive migrations (Jayo et al., 2016). As border cell migration is highly invasive, we hypothesize that disrupting the Fascin-LINC complex interaction will lead to defects in border cell nuclear deformation and impair migration.
The different functions of Fascin must by tightly regulated to ensure they are employed properly to mediate migration. One of the ways that Fascin is regulated is through phosphorylation (Adams et al., 1999; Anilkumar et al., 2003; Zanet et al., 2012). PKC phosphorylates Fascin in response to integrin activation (Anilkumar et al., 2003). Following this phosphorylation Fascin cannot bundle actin and binds to PKC to control integrin dynamics (Anilkumar et al., 2003). Future studies are needed to determine if this interaction controls the balance between Fascin bundling actin and promoting integrin dynamics during border cell migration. Additionally, previous work in our lab demonstrated that Fascin is regulated by prostaglandins (PGs) (Groen et al., 2012). PGs are lipid signaling molecules that regulate a wide variety of biological processes, including cytoskeletal dynamics (Bulin et al., 2005; Peppelenbosch et al., 1993; Tamma et al., 2003; Tootle, 2013). We previously showed that PGs regulate actin bundling during Drosophila oogenesis through Fascin (Groen et al., 2012). Exactly how PGs regulate Fascin has yet to be determined, however we believe this may occur through regulating Fascin phosphorylation (Groen and Tootle, unpublished data) and localization (Groen, 2015; Jayo et al., 2016). Indeed, loss of PGs prevents Fascin’s localization to the nuclear periphery where it interacts with the LINC Complex (Jayo et al., 2016). Other work in our lab has found that PGs also regulate border cell migration (Fox, Mellentine, and Tootle, manuscript in preparation). As described above, Fascin regulates Ena during border cell migration, and interestingly, PGs regulate Ena localization and activity in the nurse cells (Spracklen et al., 2014); these findings suggest that PGs may regulate the interaction between Fascin and Ena. Thus, PGs may regulate multiple functions of Fascin to control border cell migration. Altogether, many regulatory mechanisms control how Fascin functions during cell migration, and future studies are needed to define the means of regulating Fascin during border cell migration.
Border cell migration has emerged as an excellent model to study cancer metastasis in vivo. Border cell migration recapitulates the collective cell migration often seen in cancer metastasis and enables us to study essential aspects of this migration, such as cluster adhesion or polarization (Friedl and Gilmour, 2009; Montell et al., 2012). Fascin’s role in promoting cancer metastasis is well documented in several types of carcinomas (Gross, 2013; Hashimoto et al., 2011). Fascin is not typically expressed in adult epithelial tissue, however elevated expression of Fascin in epithelial cancers has been correlated with increased aggressiveness, mortality, and notably, metastasis (Arlt et al., 2019; Hashimoto et al., 2011; Yoder et al., 2005). In fact, knockdown of Fascin decreases metastasis in a xenograft tumor model of colon cancer (Hashimoto et al., 2007). Here we identified Fascin as a new regulator of border cell migration and find that Fascin influences both protrusion and adhesion dynamics to control this collective invasive migration. Thus, border cell migration is a simplified, in vivo, and genetic tractable system to define the actin bundling-dependent and -independent roles of Fascin in regulating invasive, collective cell migration.
Materials and methods
Fly stocks
Fly stocks were maintained on cornmeal/agar/yeast food at 21°C, except where noted. Before immunofluorescence and live imaging, flies were fed wet yeast paste daily for 2-4 days. Unless otherwise noted, yw was used as the wild-type control. The following stocks were obtained from the Bloomington Stock Center (Bloomington, IN): snX2, ena210, ena23, matα GAL4 (third chromosome), c355 GAL4, c306 GAL4, actin5C GAL4, and UASp-RNAi-Fascin (TRiP.HMS02450 and TRiP.HMJ21813). The sn28 line was a generous gift form Jennifer Zanet (Université de Toulouse, Toulouse, France; (Zanet et al., 2012), the oskar GAL4 line (second chromosome) was a generous gift from Anne Ephrussi (European Molecular Biology Laboratory, Heidelber, Germany; (Telley et al., 2012), the UASp-GFP-Fascin wild-type transgenic fly line was a generous gift from Francois Payre (Université de Toulouse, Toulouse, France; (Zanet et al., 2009), the UASp-RFP-Ena wild-type transgenic fly line was a generous gift from Mark Peifer (University of North Carolina, Chapel Hill, NC, unpublished) and the slbo>mCD8-GFP transgenic fly line was a generous gift from Xiaobo Wang (French National Centre for Scientific Research, Toulouse, France). Expression of UASp-RNAi-Fascin was achieved by crossing to mata GAL4, c355 GAL4, and c306 GAL4, maintaining crosses at 25°C and progeny at 29°C. The sn28, c355 GAL4 flies were generated by recombining sn28 and c355 GAL4 onto the same chromosome. Briefly, sn28, c355 GAL4 males were identified by selecting for the singed phenotype (marker for sn28) and w+ eyes (marker for c355 GAL4). Recombination was verified by crossing sn28, c355 GAL4/FM7 flies to sn28; UASp-GFP-Fascin and assessing both GFP expression and singed phenotype. A similar recombination scheme was performed to generate sn28, c306 GAL4/FM7 flies. Expression of UASp-GFP-Fascin was achieved by crossing to oskar GAL4, c355 GAL4, and actin5C GAL4, crosses were maintained at 25°C and progeny at 29°C. Expression of UASp-RFP-Ena was achieved by crossing to c355 GAL4, crosses were maintained at 25°C and progeny at 29°C.
Immunofluorescence
Whole-mount Drosophila ovary samples were dissected into Grace’s insect media and fixed for 10 minutes at room temperature in 4% paraformaldehyde in Grace’s insect media (Lonza, Walkersville, MD or Thermo Fischer Scientific, Waltham, MA). Briefly, samples were blocked using Triton antibody wash (1X phosphate-buffered saline, 0.1% Triton X-100, and 0.1% bovine serum albumin) six times for 10 minutes each. Primary antibodies were diluted with Triton antibody wash and incubated overnight at 4°C. The following primary antibodies were obtained from the Developmental Studies Hybridoma Bank (DSHB) developed under the auspices of the National Institute of Child Health and Human Development and maintained by the Department of Biology, University of Iowa (Iowa City, IA): mouse anti-Hts 1:50 (1B1, Lipshitz, HD; (Zaccai and Lipshitz, 1996), mouse anti-FasIII 1:50 (7G10, Goodman, C; (Patel et al., 1987); mouse anti-Fascin 1:20 (sn7c, Cooley, L; (Cant et al., 1994), rat anti-DCAD2 1:20 (Umemura, T; (Oda et al., 1994). Additionally, the following primary antibody was used: rabbit anti-GFP 1:2000 (pre-absorbed on yw ovaries at 1:20 and used at 1:100; Torrey Pines Biolabs, Inc., Secaucus, NJ) and rabbit anti-dsRed 1:300 (Clontech, Mountain View, CA). After 6 washes in Triton antibody wash (10 minutes each), secondary antibodies were incubated overnight at 4°C or for ∼4 hours at room temperature. The following secondary antibodies were used at 1:500: AlexaFluor (AF)488::goat anti-mouse, AF568::goat anti-mouse, AF488::goat anti-rabbit, AF568::goat anti-rabbit (Thermo Fischer Scientific) and AF647::goat anti-mouse and AF488::goat anti-rat (Jackson ImmunoResearch Laboratories, Inc., West Grove, PA). AF647-, rhodamine, or AF568-conjugated phalloidin (Thermo Fischer Scientific) was included with primary and secondary antibodies at a concentration of 1:250. After 6 washes in Triton antibody wash (10 minutes each), 4’,6-diamidino-2-phenylindole (5 mg/ml) staining was performed at a concentration of 1:5000 in 1X PBS for 10 minutes at room temperature. Ovaries were mounted in 1 mg/ml phenylenediamine in 50% glycerol, pH 9 (Platt and Michael, 1983). All experiments were performed a minimum of three independent times.
Image acquisition and processing
Microscope images of fixed Drosophila follicles were obtained using LAS AS SPE Core software on a Leica TCS SPE mounted on a Leica DM2500 using an ACS APO 20x/0.60 IMM CORR -/D objective (Leica Microsystems, Buffalo Grove, IL) or using Zen software on a Zeiss 700 LSM mounted on an Axio Observer.Z1 using a Plan-Apochromat 20x/0.8 working distance (WD) = 0.55 M27 or a EC-Plan-Neo-Fluar 40x/1.3 oil objective (Carl Zeiss Microscopy, Thornwood, NY). Maximum projections (two to four confocal slices), merged images, rotations, and cropping were performed using ImageJ software (Abramoff et al., 2004).
Quantification of migration index
Quantification of the migration index of border cell migration during S9 was performed on confocal image stacks of follicles stained with anti-Hts and anti-FasIII. Measurements of migration distances were obtained from maximum projections of 2-4 confocal slices of deidentified 20x confocal images using ImageJ software (Abramoff et al., 2004). Briefly, a line segment was drawn from the anterior end of the follicle to the front or posterior of the border cell cluster and the distance measured, this was defined as the distance of border cell migration. Additionally, a line segment was drawn from the anterior end of the follicle to the anterior end of the main-body follicle cells and the distance measured, this was defined as the distance of the follicle cells. The migration index was calculated in Excel (Microsoft, Redmond, WA) by subtracting the follicle cell distance from the border cell distance. Data was compiled, graphs generated, and statistical analysis performed using Prism (GraphPad Software, La Jolla, CA).
Line scan analysis of E-cadherin
Line scan analysis was performed on maximum projections of 2 confocal slices of a 40x confocal image using ImageJ software (Abramoff et al., 2004). Briefly, a line segment was drawn across a delaminating border cell cluster and the plot profile function was used to generate a fluorescent intensity plot for E-cadherin. Raw data was graphed in Prism (GraphPad Software). The cell boundaries were defined as the peaks in fluorescent intensity.
Live imaging
Whole ovaries were dissected from flies fed wet yeast past for 2-3 days and maintained at 25°C until the last 16-24 hours when they were moved to 29°C. Genotypes used for live imaging were sn28/FM7; slbo>mCD8-GFP and sn28/sn28; slbo>mCD8-GFP. Ovaries were dissected in Stage 9 (S9) medium (Prasad et al. 2007): Schneider’s medium (Life Technologies), 0.6x penicillin/streptomycin (Life Technologies). 0.2 mg/ml insulin (Sigma-Aldrich, St. Louis, MO), and 15% fetal bovine serum (Atlanta Biologicals, Flowery Branch, GA). S9 follicles were hand dissected and embedded in 1.25% low-melt agarose (IBI Scientific, Peosta, IA) made with S9 media on a coverslip-bottom dish (MatTek, Ashland, MA). Just prior to live imaging, fresh S9 media was added to coverslip-bottom dish. Live imaging was performed with Zen software on a Zeiss 700 LSM mounted on an Axio Observer.Z1 using a Plan-Apochromat 20x/0.8 working distance (WD) = 0.55 M27 (Carl Zeiss Microscopy, Thornwood, NY). Images were acquired every 5-5.5 mins for at least 3 hours. Maximum projections (two to five confocal slices), merge images, rotations, and cropping were performed using ImageJ software (Abramoff et al., 2004). To aid in visualization live imaging videos and stills were brightened by 50% in Photoshop (Adobe, San Jose, CA).
Quantification of live imaging
Quantification of live imaging videos was based on analyses done in Sawant et al. (Sawant et al., 2018). Analyses were performed in ImageJ (Abramoff et al., 2004) using maximum projection of 2-5 confocal slices time-lapse videos of border cell migration. Parameters quantified include number of protrusions per frame, protrusion length, protrusion duration, and migration speed. For number of protrusions per frame, the number of protrusions emerging from the front (0° to 45° and 0° to 315°), sides (45° to 135° and 225° to 315°), and back (135° to 225°) of the cluster was counted per frame for an hour of migration. For protrusion length, a protrusion was defined as an extension longer than 4 μm from the cluster body. The length of the protrusions was measured and binned into groups based on the direction emerging from cluster: front (0° to 45° and 0° to 315°), sides (45° to 135° and 225° to 315°), and back (135° to 225°). Protrusion duration was measured by quantifying the amount of time elapsed between the protrusion beginning to extend and the protrusion fully retracting. Migration speed was calculated during mid-migration by measuring cluster displacement dividing by time elapsed. For delaminating clusters, delamination time was defined as the amount of time elapsed from early S9 to when the border cell cluster completely detached from the epithelium. Data was compiled, graphs generated, and statistical analysis performed using Prism (GraphPad Software).
Movie 1. Control border cell migration. Video of S9 control follicle (fascinsn28/+; slbo>mCD8-GFP/+). Time listed in minutes. Images were acquired every 5.5 mins with a 20x objective. Anterior is to the right. Scale bar = 50μm. The control cluster displays single front-oriented protrusions that extend and retract throughout the migration.
Movie 2. fascin-null border cell migration. Video of S9 fascin-null follicle (fascinsn28/sn28; slbo>mCD8-GFP/+). Time listed in minutes. Images were acquired every 5 mins with a 20x objective. Anterior is to the right. Scale bar = 50μm. The fascin-null cluster displays aberrant protrusion extensions with many protrusions extending at the same time and from the sides and back of the cluster.
Movie 3. Control border cell delamination. Video of early S9 control follicle (fascinsn28/+; slbo>mCD8-GFP/+). Time listed in minutes. Images were acquired every 5 mins with a 20x objective. Anterior is to the right. Scale bar = 50μm. The control cluster delaminates considerably faster (104min) than the fascin-null follicle (Movie 4).
Movie 4. fascin-null border cell delamination. Video of early S9 fascin-null follicle (fascinsn28/sn28; slbo>mCD8-GFP/+). Time listed in minutes. Images were acquired every 5 mins with a 20x objective. Anterior is to the right. Scale bar = 50μm. The fascin-null cluster delaminates significantly slower (390min) than the control follicle (Movie 3).
Acknowledgements
We thank the Westside Fly Group and Dunnwald lab for helpful discussions and the Tootle lab for helpful discussions and careful review of the manuscript. We thank Xiaobo Wang for the slbo>mCD8-GFP fly stock. Stocks obtained from the Bloomington Drosophila Stock Center (NIH P40OD018537) were used in this study. Information Technology Services – Research Services provided data storage support. This project is supported by National Institutes of Health R01GM116885. M.C.L. is partially supported by the University of Iowa Summer Graduate Fellowship and has previously been supported by the Anatomy and Cell Biology Department Graduate Fellowship.