Apical constriction is a widely utilized cell shape change linked to folding, bending and invagination of polarized epithelia. It remains unclear how apical constriction is regulated spatiotemporally during tissue invagination and how this cellular process contributes to tube formation in different developmental contexts. Using Drosophila salivary gland (SG) invagination as a model, we show that regulation of folded gastrulation expression by the Fork head transcription factor is required for apicomedial accumulation of Rho kinase and non-muscle myosin II, which coordinate apical constriction. We demonstrate that neither loss of spatially coordinated apical constriction nor its complete blockage prevent internalization and tube formation, although such manipulations affect the geometry of invagination. When apical constriction is disrupted, compressing force generated by a tissue-level myosin cable contributes to SG invagination. We demonstrate that fully elongated polarized SGs can form outside the embryo, suggesting that tube formation and elongation are intrinsic properties of the SG.https://doi.org/10.7554/eLife.22235.001
Many organs in the human body – like the kidneys, lungs, and salivary glands – are organized as a single layer of cells that surround a hollow tube. There are a number of ways that cells can achieve this particular arrangement. In one mechanism, a small group of cells bud out of a single cell layer to become the end of a new tube or a new branch of an existing tube. Since all the cells are still connected, the first cells bring their neighbouring cells along behind them, rearranging these cells to form the walls of a tube.
In addition to changing position, the cells must change their shape to form a tube. One crucial change in cell shape is called apical constriction, and involves the side of the cell facing the inside of the tube becoming smaller than the other sides. This creates cells with a wedge-like shape that can fit together to form the curved wall of the tube, similar to shaped bricks in an archway. Apical constriction has been widely studied and is controlled by proteins that act like motors moving along protein-based filaments; however the roles of apical constriction in tube formation have not been fully explained.
Using the developing salivary glands of the fruit fly Drosophila melanogaster, Chung et al. confirmed that the motor protein known as myosin II controls apical constriction during tissue invagination. Further examination showed that proteins (called Fork Head and Fog) activate and localize an enzyme (Rho kinase) to control the localized accumulation of myosin II and thereby control apical constriction. Chung et al. then showed that salivary glands could still form tubes if apical constriction was blocked, indicating that it is not an essential part of tissue invagination in this organ. However, blocking apical constriction led the tube to develop unusual shapes at intermediate stages.
More work is now needed to better understand the links between apical constriction, cell rearrangement and tissue invagination. These processes are fundamental for organs to form correctly in many organisms and understanding their control could have wide-ranging impacts. A better understanding of these processes may provide insight into how the tubes can form while keeping all the cells adequately supplied with oxygen and nutrients, and into diseases that result if there are defects in the invagination process.https://doi.org/10.7554/eLife.22235.002
Organs that transport gases and nutrients, as well as those producing and secreting vital hormones and enzymes, are organized as epithelial tubes, many of which arise from already polarized epithelial sheets (Andrew and Ewald, 2010). To enter the third dimension, a flat sheet of polarized epithelial cells bends or invaginates using either of two distinct processes–wrapping or budding (Lubarsky and Krasnow, 2003). During wrapping, the entire epithelial sheet folds in until its edges meet and seal to form an elongated tube, as occurs in vertebrate neural tube formation and in Drosophila gastrulation (Andrew and Ewald, 2010; Massarwa et al., 2014). During budding, a subset of cells extend out of the plane of the epithelium in an orthogonal direction to form a tube; this process is observed during branching morphogenesis of many organs, including the mammalian lungs and kidney, and the primary branches of the Drosophila trachea (Andrew and Ewald, 2010; Lubarsky and Krasnow, 2003).
A limited number of cellular processes are involved in creating three-dimensional structures, which include regulated changes in cell shape, arrangement and position, as well as oriented cell divisions and spatially restricted programmed cell death (Andrew and Ewald, 2010). One cell shape change associated with such tissue remodeling is apical constriction, wherein the nuclei move to a basal position in the cell and the apical domains constrict (Martin and Goldstein, 2014; Sawyer et al., 2010). In polarized epithelial cells that maintain cell-cell adhesion, apical constriction is linked to tissue folding or invagination (Alvarez and Navascués, 1990; Hardin and Keller, 1988; Kam et al., 1991; Lewis, 1947; Sweeton et al., 1991; Wallingford et al., 2013).
Non-muscle myosin II-dependent contractility generates the force that drives this cellular process. Particularly, a pulsatile actomyosin complex in the apical medial region of the cell (hereafter referred to as apicomedial myosin) has been described in tissues that undergo apical constriction (Blanchard et al., 2010; Martin et al., 2009). Studies in early Drosophila embryos have identified the Folded gastrulation (Fog) pathway that regulates apical constriction and apicomedial myosin formation (Manning and Rogers, 2014). During gastrulation, mesodermal cells undergo apical constriction to form the ventral furrow along the anterior/posterior body axis. In those cells, the mesoderm-specific transcription factors Twist and Snail activate G protein-coupled receptor signaling and recruit RhoGEF2 to the apical surface, which, in turn, activates Rho1 (Costa et al., 1994; Kölsch et al., 2007; Manning et al., 2013; Parks and Wieschaus, 1991). GTP-bound Rho1 then activates Rho-associated kinase (Rok), which phosphorylates and activates non-muscle myosin II, which forms an actomyosin complex at the medial apical cortex (Dawes-Hoang et al., 2005). This actomyosin complex causes asynchronous contractions that pull the adherens junctions (AJs) inward. Contractions are maintained between pulses by the actomyosin belt, which serves as a ‘ratchet’ to incrementally reduce apical area (Martin et al., 2009).
Although apical constriction and its associated forces are suggested to drive tissue invagination, the exact role of this cell shape change in tube formation remains controversial (Llimargas and Casanova, 2010). In Drosophila trachea defective for EGF receptor signaling, apical constriction is impaired, but most cells invaginate (Brodu and Casanova, 2006; Nishimura et al., 2007). Similarly, in Drosophila embryos mutant for twist or fog, mesodermal cells with defective apical constriction still invaginate, although the process is both delayed and aberrant (Leptin and Grunewald, 1990; Sweeton et al., 1991). In these mutants, however, apical constriction is not completely blocked; it is simply less extensive and more random (Brodu and Casanova, 2006; Costa et al., 1994; Nishimura et al., 2007; Sweeton et al., 1991), making it difficult to draw any clear conclusions. A recent study, using an optogenetic method to deplete phosphatidylinositol-4,5 bisphosphate (PI(4,5)P2) and actin from the cell cortex, showed that local inhibition of apical constriction is sufficient to cause global arrest of mesoderm invagination during Drosophila gastrulation (Guglielmi et al., 2015). This finding suggests that apical constriction is essential for the invagination by wrapping that occurs during ventral furrow formation. It remains unclear, however, whether apical constriction is also critical for tissue invagination by budding.
The Drosophila salivary gland (SG) is an excellent system to study the role of apical constriction during tissue invagination by budding (Figure 1A–A’’,B,B’,C and C’). The SG begins as a two-dimensional sheet of cells on the embryo surface that internalizes to form an elongated tube (Chung et al., 2014). Since neither cell division nor cell death occurs once the SG has been specified, the entire morphogenetic process must be driven by changes in cell shape and rearrangement. Indeed, apical constriction has been observed in this tissue (Myat and Andrew, 2000a), and an increase in apical myosin has been reported during SG invagination (Escudero et al., 2007; Nikolaidou and Barrett, 2004; Xu et al., 2008). More detailed analyses revealed several distinct myosin structures in the forming SG, including a supracellular myosin cable that surrounds the entire tissue and is thought to be involved in tissue invagination, as well as a web-like myosin structure in the apicomedial region of cells that colocalizes with actin (Röper, 2012). The latter shows pulsatile behavior, suggesting that contractile forces by the apicomedial actomyosin complex may drive apical constriction during SG invagination (Booth et al., 2014).
Here, we elucidate the mechanism by which apical constriction is regulated during SG budding and determine its role in tissue invagination. We show that the spatial and temporal pattern of apical constriction correlates with apicomedial myosin formation during SG invagination. We uncover the molecular pathway through which the FoxA transcription factor Fork head (Fkh) coordinates apical constriction by regulating Fog signaling in the SGs. Through genetic manipulations to completely block apical constriction and detailed quantitative analysis, we show that apical constriction is not required for SG invagination, but is required for proper tissue geometry. SG cells can internalize even without forming apicomedial myosin, suggesting a role for other additional forces. We provide evidence that the compressing force generated by the tissue-level supracellular myosin cable contributes to invagination in SGs with defective apical constriction. By analyzing the externalized SG phenotypes of fog mutants, we also reveal that tube formation can be decoupled from tissue invagination.
To gain a detailed view of temporal and spatial regulation of apical constriction during SG invagination, we analyzed apical area in wild type (WT) SGs of stage 11 embryos. Using labeling with E-Cadherin (E-Cad), an AJ marker, and CrebA, an SG nuclear marker, we segmented apical cell outlines and calculated the apical area of individual cells. By analyzing >60 WT SGs for apical area and depth of invagination, we classified four distinct stages (Figure 1D–G, D'–G'). At early stage 11, all SG cells were on the embryo surface and cells with different apical area were distributed relatively randomly, although cells with small and large apical area tended to be toward the inside and at the periphery of the tissue, respectively (Before invagination; Figure 1D and D’). Shortly after, subsets of cells clustered in specific regions had smaller apical areas, most notably in a middle region close to the posterior edge of the SG where invagination begins (Clustered apical constriction; Figure 1E and E’). The less prominent anterior cluster corresponds to a small indentation that can be observed in lateral views of stage 12 SGs (data not shown). When a few cells in the posterior cluster had internalized to form an invagination pit, many cells near the pit were apically constricted (Beginning of invagination; Figure 1F and F’). At late stage 11, as more cells had invaginated, most cells anterior to the invagination pit were apically constricted (Deep invagination; Figure 1G and G’). At the same time, cells in the periphery of the SG were apically expanded, especially in the region ventral to the invagination pit. Quantification of the percentage and cumulative percentage of cells of different apical area showed a gradual and significant increase in the number of cells with smaller apical area as invagination progressed. Small increases in the number of cells with larger apical area were also observed (Figure 1I and J).
The SGs of fkh null mutants undergo early apoptosis and fail to internalize (Myat and Andrew, 2000a). fkh mutant SGs rescued from cell death by deletion of the pro-apoptotic genes contained within the H99 deficiency also fail to internalize. SGs of H99 deficiency embryos develop completely normally during embryogenesis, indicating that blocking apoptosis does not affect SG morphogenesis (Myat and Andrew, 2000a). Scanning electron microscope (SEM) and confocal images of the late stage 11 fkh H99 SGs did not reveal invagination pits (Figure 1B’’ and C’’), consistent with a complete failure of invagination. Analysis of apical areas of late stage 11 fkh H99 SGs revealed that fkh H99 SG cells had overall smaller apical area than WT cells; both the mean and median values for apical area were lower than those of WT (Figure 1—source data 1). Importantly, cells with the smallest apical areas were not clustered in fkh H99 mutant SGs and were instead randomly distributed (Figure 1H and H’). Moreover, the cumulative percentage of late stage 11 fkh H99 SG cells of different apical areas showed a similar distribution trend to that of WT SG cells before invagination (Figure 1K). Indeed, median scaling normalization of the apical area of fkh H99 late stage 11 SG cells revealed a very similar distribution to the apical area of WT SGs before invagination. Particularly, the percentages of cells with the lower 70% apical area were indistinguishable (Figure 1K). Taken together, our analysis reveals that apical constriction is both temporally and spatially coordinated during SG invagination and does not occur in fkh H99 mutants.
Apical constriction of both vertebrate and invertebrate tissues is associated with contraction of the actomyosin cytoskeleton (Dawes-Hoang et al., 2005; Martin et al., 2009; Nishimura et al., 2007; Roh-Johnson et al., 2012). Specifically, pulsatile accumulation of apicomedial myosin has been linked to apical constriction (Martin et al., 2009; Blanchard et al., 2010; Booth et al., 2014). Therefore, we investigated myosin accumulation in WT and fkh H99 SGs, using spaghetti squash (sqh)-GFP, a functional tagged version of the myosin regulatory light chain (Royou et al., 2004). Consistent with results from (Röper, 2012), several myosin structures were observed, including a supracellular myosin cable that encircles the whole tissue, AJ-associated cortical myosin, as well as an apicomedial myosin web (Figure 2A–C). Importantly, apicomedial myosin formed in a manner that recapitulates the observed temporal and spatial pattern of apical constriction. Before invagination, myosin predominantly localized at the cortical regions in all SG cells, partially colocalizing with E-Cad and often showing higher intensity at vertices (hereafter referred to as junctional myosin; Figure 2A). When invagination began, moderate levels of apicomedial myosin were observed in cells near where invagination initiates (Figure 2B). During deep invagination, apicomedial myosin was quite prominent in the cells anterior to the invagination pit (Figure 2C). Indeed, the measured intensity ratios of junctional to apicomedial myosin decreased over time in the WT SG cells (Figure 2E). In contrast, junctional myosin was predominant throughout stage 11 in fkh H99 SGs, and only dispersed, weak myosin signals were observed in the apical region (Figure 2D). Correspondingly, the ratio of junctional to apicomedial myosin in fkh H99 SGs was comparable to that of WT before invagination (Figure 2E).
To investigate the dynamics of apicomedial myosin in the SG, we took time-lapse images of sqh-GFP. Apicomedial myosin in the SG was also pulsatile, with an average interval of 131.7s ± 42.8 s between pulses (mean ± s.d., n = 14 cells in 4 SGs, 20 pulses; Figure 2F; Video 1). Pulses were mostly asynchronous between adjacent cells. The apicomedial myosin colocalized with F-actin, indicating formation of an actomyosin complex (Figure 2H; Röper, 2012). Importantly, we observed cell distortions during the peak intensity of pulses where the apicomedial myosin contacts E-Cad (Figure 2F and G'), reminiscent of the transient bending of AJs in constricting mesodermal cells (Martin et al., 2009), suggesting that actomyosin contraction generates a pulling force during SG invagination. Measurements of average myosin intensity in the apical domain of each cell showed a negative correlation with the apical area of cells in the entire SG (Figure 2I, I’ and J). This correlation, combined with the relative increase in apicomedial myosin in cells anterior to the invagination pit, suggests a role for this myosin pool in the clustered apical constriction observed during SG invagination.
Rho kinase (Rok) is a key regulator for myosin phosphorylation and activation (Amano et al., 1996). During ventral furrow formation, Rok is polarized to an apicomedial domain of mesodermal cells, where it promotes assembly of apicomedial myosin (Mason et al., 2013). Therefore, we analyzed Rok localization dynamics using a ubiquitously expressed GFP-tagged Rok transgene (Rok-GFP; Abreu-Blanco et al., 2014). Rok localization recapitulated the temporal and spatial distribution of apicomedial myosin. Rok was initially detected as occasional small punctate structures in the apical domain of all SG cells (Figure 3A), but as invagination proceeded, gradually larger apical puncta were observed in the posterior region of the SG (Figure 3B). During deep invagination, huge globular structures of Rok occupied nearly the entire apicomedial domain (Figure 3C). Live imaging revealed congregating and separating behaviors, suggesting that the large globular structures are dynamic (Figure 3D; Video 2). Importantly, only small punctate signals were detected in the apical region of fkh H99 SG cells throughout all of stage 11 (Figure 3E), indicating that Fkh is required for apicomedial Rok accumulation. In embryos treated with the Rok inhibitor Y-27632, myosin was detected only along the lateral membrane and in the basal region of the cells, suggesting that Rok activity is required for the formation of apicomedial myosin (Figure 3F). Furthermore, the SGs of Y-27632-treated embryos did not invaginate, suggesting a critical role for Rok.
We next asked which effector molecule acts downstream of Fkh to regulate apicomedial accumulation of Rok and myosin in SG cells. A good candidate was Fog, a secreted ligand known to regulate apical constriction of cells in several tissues (Costa et al., 1994; Dawes-Hoang et al., 2005; Nikolaidou and Barrett, 2004). Fluorescent in situ hybridization experiments revealed that fog mRNA is upregulated in the SG prior to and during invagination (Figure 4A; Nikolaidou and Barrett, 2004) and persists until at least stage 14 (data not shown). Importantly, SG expression of fog requires Fkh; only background levels of fog mRNA were observed in fkh H99 SGs (Figure 4A’).
Unlike fkh mutants, fog mutant SGs invaginate. fog mutant SGs formed an invagination pit at a relatively normal position at about the same stage as WT, although the pit was often somewhat larger (Figure 4C). Cell segmentation analyses revealed that the percentage and cumulative percentage of cells of different apical area did not show significant differences between fog mutant and WT SGs (Figure 4D; Figure 4—source data 3). Cells with smaller apical area, however, showed a less coordinated spatial distribution in fog mutants. Unlike in WT SGs, where apically constricted cells were tightly clustered anterior to the invagination pit (Figure 4B and B’), in fog mutant SGs, apically constricted cells were dispersed, with significantly increased dispersion along the dorsal/ventral (D/V) axis of the tissue (Figure 4C, C’, E and F).
Consistent with the uncoordinated apical constriction in fog mutant SGs, analysis of myosin signals revealed less organized web-like structures with lower staining intensity than with age-matched WT samples (Figure 4G; compare to Figure 2C). The ratio of intensity between junctional and apicomedial myosin in fog mutants during deep invagination was more like that of WT glands at earlier stages - between the value measured at the before invagination and at the beginning of invagination stages (Figure 4I; compare to Figure 2E).
We then asked if the apicomedial accumulation of Rok is affected by loss of fog. Although apicomedial Rok has been reported during ventral furrow formation (Mason et al., 2013), its dependence on fog has not been addressed. In fog mutant SGs, we observed Rok in small punctate structures dispersed along the apical domain during deep invagination, rather than in the huge globular structures observed in WT SG cells at this stage (Figure 4H; compare to Figure 3C). These data indicate that reduced Rok accumulation in the apicomedial region of fog mutant SG cells leads to reduced apicomedial myosin.
Since the loss of fog only affected the pattern of apical constriction, we next sought to completely block apical constriction. Overexpression of Crb, which expands the apical domain when overexpressed (Wodarz et al., 1995), or a constitutively-active form of Diaphanous (Dia-CA; Somogyi and Rørth, 2004), a fly formin protein that nucleates and facilitates the elongation of actin filaments (Higgs and Peterson, 2005), from the onset of SG specification throughout development efficiently blocked apical constriction; clustered apical constriction was not observed at any stage of invagination (Figure 5A–H, A’−H’). Moreover, analyses of the percentage and cumulative percentage of cells of different apical area showed that Crb or Dia-CA overexpression not only blocked apical constriction but also caused significant increase in apical area. Before invagination, Crb-overexpressing cells showed apical areas comparable to those of WT cells at early stage 11, and Dia-CA-overexpressing cells showed a small (but significant) increase of apical area (Figure 5I; Figure 5—source data 1). At the beginning of invagination, cells showed a notable increase in apical area, with obvious shifts in the distribution of apical area when compared to WT (Figure 5I; Figure 5—source data 1). Importantly, the percentage of cells in different apical area bins of Crb- or Dia-CA-overexpressing SGs was comparable to that of WT SG cells before invagination, confirming that the cells do not undergo apical constriction (Figure 5I; Figure 5—source data 1). During deep invagination, Dia-CA-overexpressing SG cells still have apical areas comparable to those of WT SG cells before invagination, but overexpression of Crb caused a huge increase in apical area (Figure 5I; Figure 5—source data 1). Importantly, however, neither blocking apical constriction nor increasing apical area arrested SG invagination. SGs overexpressing Crb or Dia-CA formed invagination pits without any delays. Plotting of cells of lower 10%, 20% and 30% of apical area showed that cells with smaller apical area were not as clustered as in WT, with significantly increased dispersion along the anterior/posterior (A/P) axis of the tissue (Figure 5—figure supplement 1A and B). These data indicate that overexpression of Crb or Dia-CA effectively blocks apical constriction and alters the distribution of cells with smaller apical areas.
To understand how apical constriction is blocked in Crb- and Dia-CA-overexpressing SGs, we asked if the cellular machinery required for apical constriction was affected. Even during deep invagination, Rok signals were detected only as small- to medium-sized puncta (in Crb-overexpressing SG cells; Figure 5J) or as very small puncta (in Dia-CA-overexpressing SG cells; Figure 5L) rather than the huge globular structures detected in WT cells (Figure 3C). However, Crb-overexpressing SGs formed relatively normal pools of apicomedial myosin that are associated with actin and with membrane distortions (Figure 5K), consistent with our findings that in fog mutants even dispersed Rok can still result in some apicomedial myosin formation (Figure 4F and G). These data suggest that in Crb-overexpressing SG cells, the forces of apical expansion overwhelm those of apical constriction despite the actomyosin machinery being in place and functional. Interestingly, in Dia-CA-overexpressing SG cells, most myosin signals were observed in a broad cortical area partially overlapping with actin, but no web-like apicomedial myosin formed (Figure 5M; compare to Figure 2C). A similar increase in cortical myosin was observed in Dia-CA-overexpressing amnioserosa cells, but interestingly, premature apical constriction was observed in those cells (Homem and Peifer, 2008), suggesting possible tissue-specific effects. Moreover, the significant ‘wiggliness’ of the AJs in Dia-CA-overexpressing SGs suggests a decrease of cortical tension (Figure 5E, G, H and L; Blanchard et al., 2010; Choi et al., 2016; Lecuit and Lenne, 2007; Martin et al., 2009). Overall, since SG invagination occurs despite the failure in apical constriction with overexpression of either Crb or Dia-CA, we conclude that apical constriction is not required.
Although invagination pits formed at approximately the right position and at the right time in the SGs with either uncoordinated (in fog mutants) or completely blocked apical constriction (in Crb- or Dia-CA-overexpressing SGs), the pits were abnormal. Specifically, both the fog mutant and Crb-overexpressing SGs frequently formed a pit that was elongated along the D/V axis, rather than the smaller oval-shaped pit of WT SGs (Figure 6A–C, A'–C'). Overexpression of Dia-CA resulted in a huge invagination pit, often larger than half the size of the tissue (Figure 6D and D'). We asked how this aberrant invagination and the continuous increases in apical area affected tube structure. 3D reconstruction of invaginating SGs (late stage 11) revealed a wider lumen in fog mutant and Crb- and Dia-CA-overexpressing SGs compared to WT, indicating that early tube architecture is affected by defective apical constriction and consequent abnormal invagination (Figure 6E–6H). We next analyzed stage 14 embryos, when in WT, all SG cells have fully internalized (Figure 6I). Interestingly, Dia-CA-overexpressing SGs showed a significant delay in cell internalization and migration. All Dia-CA-overexpressing cells eventually internalized, but formed a short gland with increased luminal diameter that remained close to the embryo surface (Figure 6J). Overproduced Crb not only localized to the apical membranes, but also mislocalized to the basal membranes facing outside of the tube; nonetheless, SG cells still formed a single-layered fully internalized tube at stage 14 (Figure 6K). Although Crb overexpression did not delay SG internalization or migration, partially or completely externalized SGs with no clear internal lumen were occasionally observed at stage 15 and 16 (Figure 6L). Since all SGs were internalized at earlier stages, these findings suggest that the Crb-overexpressing SG cells subsequently evaginated from the inside to the outside of the embryo, likely because of continued apical expansion. Similar invagination-followed-by-evagination phenotypes were observed when a myristoylated, membrane-tethered form of Wiskott–Aldrich syndrome protein (Myr-Wasp), an activator of the actin nucleating Arp2/3 complex, was overexpressed in the SGs (Figure 6M; Video 3). Overall, these data suggest that the major role of apical constriction is to ensure the proper tissue geometry during SG invagination.
The supracellular myosin cable that surrounds the entire SG is under tension (Röper, 2012), suggesting that it generates a compression force. Therefore, we asked if this tissue-scale myosin cable is present in the SGs defective for apical constriction and could contribute to invagination. At late stage 11, fog and Crb-overexpressing SGs formed a supracellular myosin cable almost indistinguishable from that of WT (Figure 7A–, A'–C'). However, this cable was defective in both Dia-CA-overexpressing SGs and in fkh H99 mutants, where invagination is either delayed or completely fails, respectively. In Dia-CA-overexpressing SGs, myosin signals were observed in a broad cortical area of individual cells (Figure 5M) and no obvious cable structure surrounding the tissue was observed (Figure 7D and D'). In fkh H99 mutants, myosin cables were observed along the lateral boundaries of the tissue, coinciding with the parasegmental boundaries, but the connecting dorsal cable was not observed (Figure 7E and E'). Similar myosin cables are observed in the lateral boundaries of Scr mutants (Röper, 2012), which do not form SGs (Andrew et al., 1994).
We calculated the circularity of the SG placode as a measure of smoothness and tension (See Materials and methods for details). The circularity of the WT placode is slightly, but significantly increased from early stage 11 to late stage 11, suggesting an increased tension over time (in Figure 7F). At late stage 11, the border of the WT placode where the supracellular cable was observed was very smooth, and circularity of the placode boundary was significantly greater than that of the boundaries shifted outside the placode by one cell (outer boundary) or inside the placode by one cell (inner boundary), suggesting that this big cable is under tension (Figure 7F and G). The circularity of the fog and Crb-overexpressing SG placode was comparable to that of WT, and was significantly greater than that of the outer and inner boundaries (Figure 7F). Consistent with the lack of a supracellular cable, the circularity of the Dia-CA-overexpressing SG boundary was significantly lower than that of WT, and was not significantly different from the circularity of the inner boundary, suggesting disrupted tension at the SG boundary (Figure 7F). The circularity of the fkh H99 mutant SG boundary was between that of early and late stage 11 WT, and was also comparable to the fkh H99 outer boundary, indicating loss of compressive forces on cells at and near the SG boundary (Figure 7F).
Anisotropic Crb distribution at the placode boundary has been suggested to drive supracellular myosin cable formation during SG formation (Röper, 2012). Consistent with this, Crb levels were higher in WT SGs than in surrounding tissues by early stage 11 and further increased by late stage 11, with overt anisotropic localization of Crb in SG cells at the dorsal boundary (Figure 7H,H',I,I'). In these cells, myosin levels were very high at the boundary, where Crb levels were lowest (inset). Importantly, Crb levels were lower in fkh H99 SGs than in WT throughout stage 11 and did not show anisotropy in boundary cells (Figure 7J,J',K,K'). Also, unlike in WT SG cells, myosin levels were never high at the dorsal boundary (Figure 7H’–K’). Taken together, we conclude that the compressing force generated by the tissue-level supracellular myosin cable, which fails to form in fkh mutants where SG levels of Crb are low, contributes to SG invagination.
A range of SG defects was observed in late stage fog mutants, with 34% of SGs either partially or fully externalized (Figure 8A–E, A'–E' and K). Given that loss of fog had relatively minor effects on apical constriction compared to overexpression of Crb or Dia-CA, and that fog mutants also form the circumferential cable surrounding the tissue, these severe tube internalization defects were unexpected. Therefore, we asked if SG expression of Fog in otherwise fog mutant embryos could rescue internalization using the UAS-Gal4 system, which has allowed the rescue of multiple SG mutant phenotypes, including loss of fkh and other transcription factors expressed in the early SG (D. Johnson and D.J. Andrew, unpublished data). The range of defects observed with SG expression of Fog in otherwise fog mutant embryos was similar to that seen with complete loss of fog (Figure 7K), indicating that the Fog requirement for internalization is not SG autonomous. Instead, analysis of early fog mutant embryos suggested that the separation of the SG from the surrounding epithelia causes these defects. In stage 11 fog mutants, deep ingression furrows often formed between the two tissues, and in more than 60% of embryos with such furrows (24/39), some SG cells at the edge were tilted and even perpendicular to the rest of the cells in the SG epithelium, forming a lip-like structure (Figure 8F–J and F'–J'). These cells maintained adhesion to their neighbors and were properly polarized, with their AJs connected to those of neighboring SG cells at the edge. Strong myosin signals at the level of AJs along the edge of the lip (Figure 8H’) suggest that pulling or squeezing forces along the edge of the gland in combination with the pushing forces of the ingression furrow caused the SG cells to slip outside the embryo.
We propose that if the ingression furrow formed at or near the posterior end of the primordia (where invagination always initiated), then the resulting glands would slip outside and form fully externalized tubes (Figure 8J and J'). However, if the ingression furrows formed elsewhere, the resulting glands would either fully or partially internalize, depending on the extent of ingression between the SG primordia and surface epithelium (Figure 8I and I'). Importantly, all fog mutant SGs (whether internalized, externalized, elongated, folded or twisted) showed intact, closed and properly polarized tubes with the apical membrane facing an internal lumen (Figure 8A–E, A'–E'). The completely externalized SGs that elongated along the embryo surface often formed longer and narrower tubes than internalized SGs (Figure 8E and E'), suggesting that, although the SG does not require contact with internal tissues to elongate, both tube diameter and length may be regulated through contact with internal tissues. Altogether, these findings reveal that SG tube formation occurs in a tissue-autonomous manner and can be decoupled from internalization.
Apical constriction often occurs as polarized epithelial precursors bend, fold and invaginate to form different tissues and organs (Martin and Goldstein, 2014). How tissues/cells undergo apical constriction in a temporally and spatially controlled manner and how much this morphogenetic process contributes to tissue invagination in different developmental contexts have been less clear. Using the Drosophila SG as a model allowed us to quantitatively analyze the distribution and magnitude of cell shape changes during invagination and to determine the signaling pathway through which apical constriction is regulated tissue-specifically. Our studies support a model in which apical constriction plays only a minor role in tissues that internalize by budding and that multiple morphogenetic processes contribute to tissue budding.
Fog has been implicated in invagination of several embryonic tissues, including the ventral furrow, the posterior midgut and the SG, and also functions in imaginal disc folding during larval development (Manning and Rogers, 2014; this study). In the ventral furrow, Twist and Snail regulate expression of the Fog ligand and its receptor, respectively (Manning et al., 2013; Seher et al., 2007). Likewise, Caudal, through activation of additional downstream transcription factors, is required for fog expression in the posterior midgut (Costa et al., 1994; Wu and Lengyel, 1998). Here, we demonstrate that Fkh regulates SG expression of fog to coordinate apical constriction of SG cells (Figure 4). Therefore, at least during embryogenesis, the tissue-specificity of Fog signaling is determined by the tissue-specific transcription factor(s) that activate expression of fog (and its receptors). The downstream pathway components are both maternally deposited and widely expressed, and are thus likely to be shared in all cell/tissue types where Fog functions. Indeed, mutations in RhoGEF2, a key component of the Fog signaling pathway in the mesoderm, cause SG invagination defects (Nikolaidou and Barrett, 2004). Due to their ubiquitous expression, localized activation of the downstream components, rather than their localized expression, is likely to determine their site of action. Consistent with this notion, we demonstrated that the robust apicomedial Rok accumulation in the SG cells requires both Fkh and Fog (Figures 3 and 4).
An interesting question is what determines regional specificity for Fog signaling within a tissue. Levels of fog transcripts are relatively uniform across the primordia, yet both Rok accumulation and apicomedial myosin accumulation occur only in a temporally and spatially regulated subset of SG cells. We propose that production and apical release of Fog and/or the apical localization of its transmembrane receptors could be regulated post-translationally. Excellent candidate regulators include Huckebein (Hkb), an SP1-like transcription factor, and its key target Klarsicht (Klar), a putative regulator of dynein ATPase (Mosley-Bishop et al., 1999; Myat and Andrew, 2000b, 2002). The expression of both hkb and klar is limited to the dorsal posterior domain of the SG, where we observe high Rok, high myosin, and apical constriction. Since Klar is required for the delivery of vesicles to the apical SG surface, which is enriched in minus-end microtubules (MT) (Myat and Andrew, 2002), a possible scenario is that during invagination, Klar-dependent apically-targeted vesicles contain Fog ligand and/or its receptor. Consistent with this model, a recent study has demonstrated a requirement for the MT cytoskeleton in the formation of apicomedial myosin (Booth et al., 2014). Moreover, unlike WT, hkb mutant SGs invaginate from the center of the SG placode (Myat and Andrew, 2002).
During ventral furrow formation, where apical constriction plays a major role in tissue invagination (Guglielmi et al., 2015), myosin is absent from the cortex and accumulates in only the apicomedial region. Since AJ-associated cortical myosin has been linked to junctional remodeling and cell rearrangement in the intercalating ectodermal cells of the Drosophila embryo (Bertet et al., 2004; Blankenship et al., 2006; Irvine and Wieschaus, 1994; Rauzi et al., 2008; Zallen and Wieschaus, 2004), one might expect that cell rearrangement does not take place during ventral mesoderm formation. Indeed, ventral furrow cells invaginate without changing their position relative to each other; cells constrict predominantly in the ventral/lateral direction, remaining longer along the A/P axis to form a long, narrow furrow along the axis (Martin et al., 2010; Sweeton et al., 1991).
We show that apical constriction is not required for SG invagination per se, but is needed to acquire proper geometry of the invagination pit and the tube that forms from it (Figures 5 and 6). In the SG, where cells sequentially invaginate through a relatively small invagination pit to make a tube, combinatorial forces that drive different morphological changes are expected to contribute to tissue invagination (Figure 8L). Correspondingly, in tracheal invagination, at least three independent processes have been implicated: EGF-dependent myosin accumulation (Brodu and Casanova, 2006; Nishimura et al., 2007), an oriented final cell division (Kondo and Hayashi, 2013), and FGF-mediated migratory forces. Only when all three processes are disrupted, do tracheal cells fail to internalize (Kondo and Hayashi, 2013). However, since no cell divisions occur once the SG has been specified (Chung et al., 2014), oriented cell divisions cannot contribute. Similarly, our finding that the non-autonomous functions of fog can lead to fully formed external SGs (Figure 8) reduces the likelihood that signaling between the SG and neighboring tissues is critical.
We show at least one additional force contributes to tissue invagination when apical constriction is blocked: the supracellular myosin cable that is under tension and sequentially closes in as SG cells internalize. This circumferential myosin cable forms normally in both fog mutant SGs and in Crb-overexpressing SGs, where invagination occurs relatively normally, but fails to form in fkh mutant SGs and in Dia-overexpressing SGs, where invagination either fails or is delayed (Figure 7). These findings support the idea that tension generated by the myosin cable contributes to and facilitates invagination (Figure 7). This closing process in the SG is reminiscent of the ‘purse-strings’ observed during wound healing in both flies and vertebrates (Martin and Parkhurst, 2004) and also during Drosophila dorsal closure (Franke et al., 2005). It is notable that multiple forces have been suggested to contribute to dorsal closure; dorsal closure fails only when both the forces of apical constriction of amnioserosa cells and of purse strings from surrounding epidermal cells are ablated (Kiehart et al., 2000), although this model of a combinatorial force-component system has been challenged by recent studies showing that the epidermal actin cable tension may not contribute to dorsal closure (Pasakarnis et al., 2016).
The difference in invagination defects in Dia-CA-overexpressing SGs (delayed) and fkh H99 SGs (completely failed), where both apical constriction and the circumferential myosin cable are absent, indicate that additional other forces contribute to SG invagination. We propose that cell rearrangement linked to junctional myosin may also be important. A strong junctional myosin pool is observed throughout SG invagination, with high levels at cell vertices (Figure 2; Röper, 2012). We have direct and indirect evidence for cell rearrangement in early WT SGs from live imaging and measurements of cell topology (S. Chung, S. Kim and D.J. Andrew, unpublished data), suggesting that this process is linked to tissue invagination. Since myosin localizes in a broader cortical region in Dia-CA-overexpressing SGs (Figure 5), it is possible that cell rearrangement of Dia-CA-overexpressing cells is somewhat affected, explaining the delayed invagination and abnormal tissue morphology observed early (Figure 6D and H). Nonetheless, cell rearrangement must still occur in the Dia-CA overexpressing glands since they form elongated tubes with an approximately WT arrangement of cells at later stages (Figure 6J). The complete failure of invagination in fkh H99 SGs raises the possibility of failed cell rearrangement. Interestingly, changes in cell topology occur independently of fkh function. WT and fkh H99 mutant SGs show a similar distribution of n-sided cells during early stages of invagination (S. Chung and D.J. Andrew, unpublished data). Consistent with the idea that cortical enrichment of myosin has a role in these topological changes, junctional myosin is intact in fkh H99 SGs (Figure 2). In any case, more direct measurements of cell rearrangement in both WT and fkh mutants will be necessary to learn how directed cell rearrangement contributes to SG internalization. Overall, our findings suggest that multiple forces are required for SG invagination and that the system is remarkably robust. Importantly, our finding that polarized and fully elongated tubes can form outside the embryo indicates that SG morphogenesis is largely a tissue-autonomous process.
fkh6 H99 recombinants (RRID:DGGR_130500; RRID:BDSC_1576; Myat and Andrew, 2000a) were used for the analysis of fkh mutant SGs. UAS-Crb (RRID:DGGR_108289; Wodarz et al., 1995), UAS-HA-Dia-CA (RRID:BDSC_27616; Somogyi and Rørth, 2004) and UAS-Myr-Wasp-GFP (C-terminal GFP tag added to construct described in FBal0243602) driven by fkh-Gal4 (FBtp0013253/FBti0027904; Henderson and Andrew, 2000) were used. fkh-Gal4 driving UAS-Tmem-GFP (Ftp0011013 [aka UAS-GFP-CAAX]; Kakihara et al., 2008) was used as a control. Two null alleles for fog (fogS4 and fogRA67; RRID:DGGR_106665; RRID:BDSC_6218) were analyzed; both resulted in indistinguishable phenotypes. UAS-fog (FBtp0021363; Dawes-Hoang et al., 2005) driven by fkh-Gal4 in fogRA67 embryos at 25°C was used for SG-specific rescue experiment of fog. sqhAX3; sqh-GFP (RRID:BDSC_57144; Royou et al., 2004) and sqh-Rok-GFP (RRID:BDSC_52289; Abreu-Blanco et al., 2014) were used to analyze myosin and Rok accumulation, respectively.
For most samples, dechorionated embryos were fixed in 1:1 heptane:formaldehyde for 40 min and devitellinized with 80% EtOH. For phalloidin staining, embryos were hand-devitellinized. Antibodies used include: mouse α-Crb (RRID:AB_528181; DSHB; 1:10), rabbit α-CrebA (RRID:AB_10805295; Fox et al., 2010; 1:3000), chicken α-GFP (RRID:AB_300798; Abcam; 1:1000), rat α-E-Cad (AB_528120; DSHB; 1:10), rabbit α-Dia (Afshar et al., 2000; 1: 5000), mouse α-HA (RRID:AB_514506; Roche; 1:50), guinea pig α-Sage (RRID:AB_2632603; Fox et al., 2013); 1:100), rabbit α-Fkh (RRID:AB_2632955; Fox et al., 2013; 1:2000), rabbit α-β-Gal (AB_788167; Novus; 1:1000). Alexa fluor 488, 555, 647-labelled secondary antibodies were used at 1:200 (Molecular probes: RRID:AB_142924; RRID:AB_141822; RRID:AB_142754; RRID:AB_141693; RRID:AB_141373; RRID:AB_141778; RRID:AB_143165; RRID:AB_162543). Alexa fluor-546-labeled phalloidin was used for F-actin labelling (Molecular Probes: RRID_AB2632953; 1:250). Fluorescent in situ hybridization with an RNA probe specific for fog and antibody staining with anti-Sage was performed as described (Knirr et al., 1999). For the in situ analysis, a PCR-labelled product of the fog ORF was made from cDNA generated by reverse transcription of total RNA using the following primers: fog-5’: ATGTCTCCGCCCAATTGTCT and fog-3’: GATGACTGAAAAGCGGCGGC. All images were taken with a Zeiss LSM 780 confocal microscope.
Dechorinated embryos were fixed in 25% glutaraldehyde in cacodylate buffer and hand-devitellinized. Embryos were post-fixed in 1% OsO4 and dehydrated through an EtOH series. They were dried using Hexamethyldislazane, coated with gold palladium, and examined and photographed in a LEO/Zeiss Field-emission SEM.
E-Cad/CrebA stained SGs were imaged with a Zeiss LSM 780 confocal microscope. Two or three focal planes (0.37 µm apart) of apical domain were used to generate a maximum intensity projection (Zeiss Zen software; Germany). Cell segmentation was performed along the E-Cad signals and the apical area for each cell was calculated using the Bitplane Imaris program. During ‘Before invagination’ and ‘Clustered apical constriction’, all cells were at the surface in the same focal plane. During ‘Beginning of invagination’, a few cells in the posterior/dorsal region of the placode were found in a basal position relative to neighboring cells (at 0.37–1.11 µm deeper focal planes). ‘Deep invagination’ included samples with cells that had internalized >2 µm.
Frequency distribution of cells with a different apical area (bin width = 4) was performed using a GraphPad Prism program. For median scaling normalization of fkh H99 SG cells (Figure 1K), the apical area of each cell was multiplied by 1.2.
The x and y coordinates for each cell, which correspond to the position along the A/P and D/V axis of the tissue, respectively, were obtained using Imaris. The position of SG cells relative to the center of the invagination pit was calculated and plotted. Standard deviations for the positions along the A/P or D/V axis were compared to show dispersion of cells.
Cells in the posterior/dorsal region of the placode were chosen for quantification. For all quantifications, maximum intensity projections that span the apical and the junctional region of SG cells were used (Zeiss Zen software) and measurements were performed with Fiji software. Regions were drawn manually along the inner or outer boundary of E-Cad signals of each cell to calculate the apical and junctional intensity. Integrated density for myosin signals was measured after background correction and mean values from three individual measurements were used.
Cell segmentation for six WT SGs at deep invagination was performed as described above. Intensity mean was measured for the entire myosin signals for each segmented cell and re-scaled for 0 to 100 (a.u.) for plotting. Correlation (Pearson) and P values were calculated using the GraphPad Prism software.
sqhAX3; sqh-GFP or sqh-GFP-Rok embryos were dechorinated and mounted ventral side up on a glass slide coated with heptane glue (double-sided tape soaked in heptane). Spacer coverslips (No. 1.5) were attached to prevent embryos from being squeezed. Halocarbon oil mixture 27:700 (1:1) was added on top of the embryos. A No. 1 coverslip was placed on top and sealed with nail polish. Images were taken every 5s with a 3i Marianas/Yokogawa Spinning Disk Confocal microscope to capture apicomedial myosin pulsing and every 20 s with a Zeiss LSM 780 confocal microscope to capture apicomedial Rok signals.
sqhAX3; sqh-GFP embryos were collected and aged until stage 9. Dechorinated embryos were immersed for five minutes in a 1:10 dilution of Citrasolv in water (Citra-Solv, Danbury, CT) to render embryos permeable (Rand et al., 2012). Embryos were rinsed with water and PBS several times and were incubated in Y-27632 (500 µM in PBS) for three hours. Embryos were fixed as described above and immuno-stained.
3D reconstruction of SGs was performed with the confocal stacks using the Imaris program.
Cell boundaries were manually traced in ImageJ to quantify the length and area of the boundary corresponding either to the SG placode boundary or the boundary shifted one cell layer outside or inside the placode boundary. The ventral midline of the embryo was used as the ventral boundary of the traced circle in all cases. For fog mutants, only circularity of the SG boundary and inner boundary was measured, since the cells in the neighboring epithelia were often out of the focal plane due to the twisted phenotype of the embryos. Circularity was calculated based on the fact that for a perfect circle the circularity C = 1 = 4 π area/perimeter2 (Lawrence et al., 1999). Circularity values were plotted in Prism and statistical significance in differences in the circularity of outer and inner boundaries compared to the circularity of the cable was analyzed using a two-tailed t test.
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Hugo J BellenReviewing Editor; Baylor College of Medicine, United States
In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.
[Editors’ note: this article was originally rejected after discussions between the reviewers, but the authors were invited to resubmit after an appeal against the decision.]
Thank you for choosing to send your work, "Uncoupling apical constriction from tissue invagination", for consideration at eLife. Your initial submission has been assessed by a Senior Editor in consultation with a member of the Board of Reviewing Editors (Hugo Bellen), and two reviewers. Although the work is of interest, we regret to inform you that the findings at this stage are too preliminary for further consideration at eLife.
As you will notice one reviewer is very positive, while one reviewer is positive, but feels that additional data are required to warrant publication in eLife. I am ambivalent. Can you let me know if you can address the comment: "Given the extensive descriptions in the Roper paper, however, and the limited treatment of the cable in the present paper, a more careful mechanical treatment, including quantification of the cable and cell shapes, couple with biophysical modeling would be required to justify publication in eLife". Please let me know if you can and are willing to address this issue, and if so, how. I do not feel that modeling is crucial but the first portion of the comment seems important to me.
The critical role of apical constriction in cell rearrangements from gastrulation onward has been recognized for decades but the mechanisms underlying it have remained more elusive. The last six years have seen a revolution in our thinking about the mechanisms by which apical constriction is driven, based in large part on studies of the Drosophila invaginating mesoderm. This has revealed a novel and unexpected role for an apical myosin network created in response to cell-cell signaling and regulated by apical Rok. However, the generality of this model, in particular in invaginating tubular organs, has remained unexamined. Here Chung et al. present a stunning examination of this, using the extremely tractable Drosophila salivary gland as a model. In a series of experiments combining sophisticated genetic tools, beautiful imaging and extensive quantitative analysis, they lay out a pathway leading from the transcription factor Fkh through the cell signaling molecule Fog to Rok and myosin. They also find some exceptionally surprising things along the way-most particularly that the system is remarkably robust, such that eliminating apical construction does not block organ invagination. I think these results will be of very broad interested to both cell and developmental biologists.
I had some questions and suggestions that I think would strengthen the story.
Figure 1A. I found the dual 3D images here very informative, as they give insight into how apical constriction combines with other cell rearrangements. Could the authors add one of an intervening stage as the pit begins to form? Also, would it be possible to create similar images of the fog, crb- and Dia-CA expressing embryos?
Figure 1D-G. I found it intriguing that there seem to be two clusters of invaginating cells, anterior and posterior, only the posterior of which goes into the pit. This might be worth noting.
Subsection “Blocking apical constriction does not arrest SG invagination”, last paragraph. The authors state that "the significant 'wiggliness' of the AJs suggests a decrease of cortical tension." I think this is unlikely-in cultured mammalian cells, we and others have found that decreasing cortical tension (e.g., by Rok or myosin inhibition) leads to smoothly rounded cell borders – really wiggly cell borders are likely to reflect actin filaments oriented perpendicular to the membrane.
Subsection “Blocking apical constriction does not arrest SG invagination”. When during the process of convergent elongation of the organ primordium does Fog expression go off?
Subsection “Blocking apical constriction does not arrest SG invagination”. In interpreting their Dia-CA experiments the authors might take a look at Homem and Peifer (Development. 2008 Mar;135(6):1005-18) in which it was observed that Dia-CA increased cortical myosin in the amnioserosa and altered apical constriction.
The only set of data I found was weak was the attempt to rescue Fog by SG specific Fog re-expression. Given the likelihood that levels and timing were not precisely replicated, I think the speculation about non-autonomous functions was not well supported and I would suggest removing this data.
Discussion. The authors mention a number of times the incredibly provocative fact that they may have identified a GPCR related to Mist expressed in the SG. This would be very interesting, but the degree of discussion of this unpublished data in the manuscript seemed unwarranted unless they wanted to add this data.
Subsection “Additional morphogenetic processes take place during SG invagination”, last paragraph. The authors should add the data showing that the tissue level circumferential myosin cable does form in fog mutants – this is a key part of their model. More generally, I think a more careful description of the progression of junctional myosin in wildtype and mutants would be beneficial – it appeared, for example, that it was AP planar polarized before the process began.
The final circuitry diagram in Figure 7L is nice but would be better if supplemented with a diagram showing the cell shape changes themselves and how the authors think apical constriction and tissue level constriction work together.
The manuscript describes the cell biology behind the invagination of salivary gland primordia in the Drosophila embryo. The changes in cell morphology that occur during this process has been published previously, but the present paper incorporates new features (e. g., the pulsatile apical medial myosin networks) that allow comparison to other well characterized morphogenetic movements in Drosophila, especially the ventral furrow and posterior midgut. In many respects (e. g., the involvement of RoK and Folded gastrulation) the SC pathway shows a regulation of apical constriction and myosin activity very similar to these other processes.
Mutations in Fork head disrupt the invagination process, although they allow apparent initial specification of the primordia, given the CrebA marker expression. Some of this has been previously shown, but the authors extend their earlier phenotypic characterizations by blocking the associated apoptosis using the H99 deficiency. Under these conditions, the pulsing medial myosin population observed in the SG of wild type embryos is not observed, nor does Rok-GFP accumulate in the large globular structures observed in the posterior SG of wildtype There is also no Fog expression in mutants, and pattern of apical constriction is randomized. Conclusions based on these phenotypes are difficult, however, given the uncertainty about how normal or unnormal cells blocked in apoptosis are, and where exactly Fork head intervenes in the hierarchy between programming and cell behavior. Fork head's role is more extensive than just apical constriction, given that the mutants apparently fail to form complete actin cables surrounding the primordial (poorly documented, but see below).
To inhibit all apical constriction, the authors use Crb- or Dia-CA-overexpression. These manipulations are associated with increased apical area (i.e., decreased apical constriction), but surprisingly, even in the most extreme cases, do not block SG invagination. The invaginations are abnormally shaped, however, elongated along the DV axis. The authors attribute these defects to the failure of apical constriction, but overexpression of Crb and Dia may have complex cumulative effects not related to their earlier effects on apical constriction. The most interesting result here, however, as emphasized in the paper's title, is that even in these extreme circumstances, invagination still occurs. The paper would be much stronger if the authors had developed experiments that actually address in a quantitative rigorous way the underlying mechanism that accomplished this invagination.
One potential mechanism has in fact been proposed earlier by Roper (Developmental Cell 2012) and involves a supracellular myosin cable surrounding the SG primordium. Contraction of this cable could act as a "purse string" that would drive internalization of the primordium. By blocking apical constriction, the present paper provides additional arguments for the importance of this cable. Given the extensive descriptions in the Roper paper, however, and the limited treatment of the cable in the present paper, a more careful mechanical treatment, including quantification of the cable and cell shapes, couple with biophysical modeling would be required to justify publication in eLife. In the Discussion, the authors mention another mechanism that might also contribute to invagination (myosin accumulation at cell vertices) but these possibilities need to be developed in greater experimental and theoretical detail.
Overall, I think this paper is a very solid contribution to understanding salivary gland morphogenesis, but does not contain sufficiently new unexpected experimental observations or general concepts. This cannot be improved by simple adding a few additional experiments and in its present form is more suitable to a more specialized journal.https://doi.org/10.7554/eLife.22235.021
- Deborah J Andrew
The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.
We thank Bloomington, E Chen, A Martin and S Parkhurst for fly stocks, Flybase for information on gene structure, and C Hanlon, R Loganathan, C O’Kane and P Paul for their helpful comments on the manuscript.
- Hugo J Bellen, Reviewing Editor, Baylor College of Medicine, United States
- Received: October 9, 2016
- Accepted: February 7, 2017
- Version of Record published: March 6, 2017 (version 1)
© 2017, Chung et al.
This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.