New Results

Multifunctional role of GPCR signaling in epithelial tube formation

Vishakha Vishwakarma, Thao Phuong Le, SeYeon Chung
doi: https://doi.org/10.1101/2022.01.06.475238

ABSTRACT

Epithelial tube formation requires Rho1-dependent actomyosin contractility to generate the cellular forces that drive cell shape changes and rearrangement. Rho1 signaling is activated by G protein-coupled receptor (GPCR) signaling at the cell surface. During Drosophila embryonic salivary gland (SG) invagination, the GPCR ligand Folded gastrulation (Fog) activates Rho1 signaling to drive apical constriction. The SG receptor that transduces the Fog signal into Rho1-dependent myosin activation has not been identified. Here, we reveal that the Smog GPCR transduces Fog signal to regulate Rho kinase accumulation and myosin activation in the apicomedial region of cells to control apical constriction during SG invagination. We also report on unexpected Fog-independent roles for Smog in maintaining epithelial integrity and organizing cortical actin. Our data supports a model wherein Smog regulates distinct myosin pools and actin cytoskeleton in a ligand-dependent manner during epithelial tube formation.

INTRODUCTION

The formation of three-dimensional tubes by invagination from flat epithelial sheets is a fundamental process in forming organs, such as lungs and kidneys (Andrew and Ewald, 2010). The Drosophila embryonic salivary gland (SG) is a premier model system to study the mechanisms underlying epithelial tube morphogenesis (Chung et al., 2014; Girdler and Röper, 2014). The SG begins as a two-dimensional plate of cells on the embryo surface. Neither cell division nor cell death occurs once the SG cells are specified; all the morphogenetic changes arise by changes in cell shape and rearrangement.

A major cell shape change during SG invagination is apical constriction, wherein the apical side of epithelial cells shrinks while keeping the nearly constant volume (Lubarsky and Krasnow, 2003; Martin and Goldstein, 2014). During stage 11, SG cells begin to invaginate at the dorsal/posterior region of the placode through apical constriction (Fig. 1A). This leads to forming a narrow invagination pit through which all SG cells eventually internalize. Apical constriction is observed in both invertebrates and vertebrates, driving morphogenetic processes ranging from gastrulation in Drosophila and Xenopus to lens formation in the mouse eye to formation of tubular organs such as the Drosophila SG and chicken lungs (Kim et al., 2013; Myat and Andrew, 2000; Plageman et al., 2011; Sweeton et al., 1991). Defects in apical constriction result in defects in overall tissue shape, suggesting that coordinated apical constriction is required for final tissue architecture (Chung et al., 2017; Guglielmi et al., 2015; Izquierdo et al., 2018).

Figure 1.
Figure 1. Smog transduces Fog signal to activate Rok and myosin in the SG.

(A) Cartoon diagram showing SG placodes (magenta) in the ventral side of a stage 11 Drosophila embryo. Cells that undergo apical constriction anterior to the invagination pit (asterisk) are shown in purple. In those cells, the Fog ligand (yellow) binds to its GPCR (blue) to activate apicomedial myosin. In all stage 11 SG images, anterior is to the left, and dorsal is up. (B, B’) Fluorescent in situ hybridization of smog (magenta) in the invaginating WT SG. Strong smog mRNA signals are shown in apically constricting SG cells near the invagination pit (arrowheads). CrebA (green), SG nuclei. White lines, SG boundary. Arrows, smog signals near the grooves and the ventral midline. (C-D’’) Smog-GFP (green) signals in control (C-C’’) and Fog-overexpressing SGs (D-D’’). Magenta, E-Cad. Control SG cells show puncta of Smog-GFP signals in the apical region near AJs (yellow arrowheads). Considerably higher Smog-GFP signals in the entire apical domain of Fog-overexpressing SG cells (red arrowheads). Insets, magnified images of yellow boxed regions. (C’’, D’’) Z sections across the yellow lines in C and D. (E) Quantification of total Smog-GFP intensity in control and Fog overexpressing SG cells. n= 4 SGs (control); 5 SG (Fog overexpression). *p≤0.05 (Welch’s t-test). (F, G) sqh-GFP (green) and E-Cad (magenta) signals in control (F) and Fog-overexpressing (G) SGs. (F’-G’’’) Magnified images of yellow boxed regions in F and G. Compared to control (F-F’’’), sqh-GFP signals are increased in the apicomedial region of Fog-overexpressing SG cells (G-G’’’). Yellow arrowheads, AJs. Cyan arrowheads, sqh-GFP signals. (H-J) Quantification of apicomedial myosin intensity (H), junctional myosin intensity (I), and the ratio of apicomedial to junctional myosin intensity (J). (K-M’’’) Rok-GFP (green) and E-Cad (magenta) signals in SGs. K’-M’’’, magnified images of yellow boxed regions in K-M. Compared to control (K-K’’’), Fog-overexpressing SG shows overaccumulation of Rok-GFP and wavy cell junctions (L-L’’’). smog knockdown suppresses these phenotypes (M-M’’’). Yellow arrowheads, AJs. Cyan arrowheads, Rok-GFP puncta. (N, O) Quantification of the area of Rok-GFP particles (N) and waviness of AJs (O). Asterisks, invagination pit. n=5 SGs, 75 cells for each genotype in H-J, N, and O. *p≤0.05; **p≤0.01; ***p≤0.001; ****p≤0.0001 (Welch’s t-test).

Contractile actomyosin networks generate the cellular forces driving coordinated cell behaviors during epithelial morphogenesis (Gorfinkiel and Blanchard, 2011; Levayer and Lecuit, 2012; Munjal and Lecuit, 2014). Non-muscle myosin II (hereafter referred to as myosin) uses its motor activity to pull on actin filaments anchored to adherens junctions (AJs) to apply tension on the junction and neighboring cells. Three distinct pools of myosin generate the contractile forces required for SG invagination (Booth et al., 2014; Chung et al., 2017; Röper, 2012). Apicomedial myosin forms in the center of the apical side of the SG cell and drives apical constriction (Booth et al., 2014; Chung et al., 2017; Röper, 2012). Junctional myosin, associated with AJs of SG cells, is involved in cell intercalation (Sanchez-Corrales et al., 2018). A supracellular myosin cable surrounding the entire SG placode generates the tissue-level compressing force (Chung et al., 2017; Röper, 2012). From studies in the past several years by us and others, we are beginning to understand how each myosin pool is created and regulated during SG invagination (Booth et al., 2014; Chung et al., 2017; Le and Chung, 2021; Röper, 2012; Sanchez-Corrales et al., 2018; Sidor et al., 2020).

Localized activation of the RhoA (Rho1 in Drosophila) GTPase is key to creating and/or polarizing contractile myosin structures in epithelial cells (Blanchard et al., 2010; Dawes-Hoang et al., 2005; Martin et al., 2009; Mason et al., 2013). Studies in Drosophila have identified the Folded gastrulation (Fog) pathway as a key signaling pathway regulating Rho1 signaling during epithelial morphogenesis (Manning and Rogers, 2014). During Drosophila gastrulation, two G protein-coupled receptors (GPCRs), Smog (ubiquitously expressed) and Mist (mesoderm-specific), respond to Fog to activate heterotrimeric G proteins, Rho1, and Rho kinase (Rok) to regulate myosin contractility (Costa et al., 1994; Dawes-Hoang et al., 2005; Kerridge et al., 2016; Kölsch et al., 2007; Manning et al., 2013; Mason et al., 2013; Parks and Wieschaus, 1991). In our previous study in the SG, we also revealed a tissue-specific regulation of Fog signaling to activate myosin contractility: the Fork head (Fkh) transcription factor regulates SG upregulation of fog, which promotes accumulation of Rok and myosin specifically in the apicomedial region of SG cells to drive clustered apical constriction (Chung et al., 2017; Fig. 1A). Yet, it is unknown how the Fog signal is sensed and transduced into downstream Rho1 signaling during epithelial tube formation.

Studies in early Drosophila embryos have suggested a role of GPCR signaling as a common module for Rho1 activation in different subcellular regions of the cell. In the mesoderm, Smog and Mist function together to activate myosin in the apicomedial region of cells to drive apical constriction (Kerridge et al., 2016). In the ectoderm, where no apical constriction occurs, Smog is required for junctional myosin activation to drive cell intercalation (Kerridge et al., 2016). Recent work further has shown that distinct RhoGEFs activate myosin contractility in the apical and junctional domain of epithelial cells under the control of specific G proteins in early Drosophila embryos (Garcia De Las Bayonas et al., 2019). However, whether all three myosin pools in the SG are regulated by independent mechanisms or by a common module during epithelial tube formation remains unclear.

The contractile force of the actomyosin networks causes an increase of hydrostatic pressure in the cytoplasm (Charras et al., 2005). During epithelial morphogenesis, cells maintain the surface integrity by proper organization of actin at the cell cortex. Loss of organized cortical actin networks leads to the formation of blebs, spherical protrusions of the plasma membrane (Charras, 2008). During Drosophila gastrulation, blebs occasionally arise during apical constriction of mesodermal cells; these blebs correlate with apical F-actin holes (Costa et al., 1994; Jodoin et al., 2015). Disruption of cortical actin by latrunculin B, which sequesters globular actin (G-actin) and prevents filamentous actin (F-actin) assembly, enhances blebbing in epithelial cells in early Drosophila embryos (Kanesaki et al., 2013). Mutations in heterotrimeric G proteins also enhance blebbing (Kanesaki et al., 2013), suggesting a role of G proteins in stabilizing cortical actin. The mechanisms of cortical actin regulation during epithelial tube formation are not well understood.

Here, we reveal multiple roles of Smog GPCR in regulating actomyosin networks during Drosophila SG tube formation. We show that the GPCR Smog transduces Fog signal to regulate myosin contractility in the apicomedial region of SG cells to drive apical constriction during SG invagination. Our study supports a model wherein Smog regulates apicomedial and junctional/supracellular myosin pools in a Fog-dependent and -independent manner, respectively. We further reveal new roles of Smog in regulating the microtubule networks, key apical and junction proteins, and cortical actin organization during SG invagination, suggesting multifunctional roles of GPCR signaling during epithelial tube formation.

RESULTS

Smog transduces Fog signal in the SG

To test the role of Smog in transducing Fog signal during SG invagination, we first examined smog transcripts and Smog protein levels in the developing SG. Consistent with the ubiquitous expression of smog (Kerridge et al., 2016), fluorescent in situ hybridization revealed low levels of smog mRNA expression in the entire embryo, including the SG. Strong smog mRNA signals were observed in SG cells near the invagination pit (Fig. 1B, B’), potentially due to the shrinking apical domain in those cells. We also examined Smog protein localization in the SG using a functional Smog-GFP fusion protein expressed under the control of the sqh promoter (Kerridge et al., 2016). Smog-GFP signals were detected as small punctate structures enriched near AJs in SG cells (Fig. 1C-C’’). Importantly, Smog-GFP signals were dramatically increased in the entire apical domain of Fog-overexpressing SGs (Fig. 1D-D’’; quantification in Fig. 1E), suggesting that Smog is recruited to the apical domain of SG cells by overproduced Fog.

To test whether Smog transduces Fog signal in the SG to regulate myosin contractility, we performed a genetic suppression test. We hypothesized that if Smog is a SG receptor for Fog, knocking down smog in the SG should suppress the gain-of-function effect of Fog. We used sqh-GFP, a GFP-tagged version of myosin regulatory light chain (Royou et al., 2004), and Rok-GFP, a GFP-tagged Rok fusion protein (Abreu-Blanco et al., 2014), as readouts of Fog signaling. sqh-GFP signals were significantly increased in the apicomedial region of Fog-overexpressing SG cells (compare Fig. 1G’, G’’’ to Fig. 1F’, F’’’; quantification in Fig. 1H), suggesting increased myosin levels by Fog overexpression. Notably, sqh-GFP signals at AJs of SG cells were not significantly changed (Fig. 1I), suggesting that Fog promotes Rok accumulation and myosin activation in the apicomedial region of SG cells. Fog-overexpressing SG cells showed highly distorted AJs (Fig. 1G, G’’) compared to control SG cells (Fig 1F, F’’), suggesting increased pulling forces due to increased apicomedial myosin in Fog-overexpressing SG cells. Consistent with our finding that Fog signal promotes accumulation of Rok in the apicomedial region of SG cells (Chung et al., 2017), quantification of areas occupied by Rok-GFP puncta revealed overaccumulated Rok-GFP signals in the apicomedial region of Fog-overexpressing SG cells (compare Fig. 1L’, L’’’ to Fig. 1K’, K’’’; quantification in Fig. 1N). Knockdown of smog in the SG using RNA interference (RNAi) and the SG-specific fkh-Gal4 driver (Henderson and Andrew, 2000) both suppressed the apicomedial accumulation of Rok-GFP signals and reduced the magnitude of AJ distortion (Fig. 1M-M’’’; quantification in 1N, O). These data suggest that Smog transduces Fog signal in the SG to facilitate Rok and myosin accumulation in the apicomedial region of SG cells.

Smog-transfected Drosophila S2 cells contract upon Fog signal

To further test the role of Smog in transducing Fog signal to regulate myosin contractility, we performed an in vitro cell contraction assay using Drosophila S2 cells. S2 cells do not express Fog receptors, and therefore, do not respond to Fog in the normal culture condition (Manning et al., 2013; Fig. 2A-B’). As shown in a previous study (Manning et al., 2013), S2 cells transfected with Mist, the mesoderm-specific receptor for Fog, contracted robustly when cultured in Fog-containing media (Fig. 2C-D’). Since Smog was not detectable in Drosophila S2 cells (modENCODE Cell Line Expression Data; Flybase), we tested if S2 cells transfected with Smog also contract upon Fog treatment. Indeed, Smog-transfected cells showed a significantly higher percentage of contraction upon Fog treatment compared to non-transfected S2 cells (Fig. 2A-B’, E-F’; quantification in Fig. 2G). Thus, Smog can transduce Fog signal to regulate cellular contractility both in the SG and in vitro.

Figure 2.
Figure 2. Smog-transfected S2 cells contract in response to Fog.

(A-F’) S2 cells cultured in the absence (A, C, E) or presence (B, D, F) of Fog. (A-B’) non-transfected S2 cells. (C-D’) Mist-transfected S2 cells. (E-F’) Smog-transfected S2 cells. A-F, phase contrast; A’-F’, merged images of phase contrast and V5 signals for transfected cells (green). White arrowhead in E’, Smog-transfected S2 cell is not contracting in the absence of Fog. Magenta arrowheads, contracting cells. (G) Quantification of percentage of contracting cells. n=∼100 cells per condition from three independent experiments. ****p≤0.0001 (Two-way ANOVA with Tukey’s multiple comparison post hoc analysis).

Smog is required for apical constriction in the SG and epithelial morphogenesis during embryogenesis

To test the possibility that Smog functions as a Fog receptor in the SG, we knocked down smog in the SG (using fkh-Gal4) using RNAi and compared apical areas of SG cells to those in fog mutants. Two independent short hairpin RNAi lines, TRiP.HMC03192 (a stronger line; Fig. 3B, B’; also used for genetic suppression assay in Fig. 1J-J’’’) and TRiP.GL01473 (a weaker line; Fig. S1C, C’), were used. Cell segmentation analysis revealed that SG cells with smaller apical areas showed a less coordinated spatial distribution in smog RNAi SGs compared to control (Fig. 3A-B’; Fig. S1B-C’), similar to the previously reported fog mutant phenotype (Chung et al., 2017; Fig. S1A, A’). Quantification of percentage and cumulative percentage of cells of different apical areas also revealed that SGs with smog knockdown with either RNA line show larger apical areas compared to control (Fig. 3G; Fig. S1D), suggesting defective apical constriction upon smog knockdown.

Figure 3.
Figure 3. smog knockdown and loss result in apical constriction defects in the SG and disrupted epithelial morphology.

(A-F) Confocal images of SGs in control (A), with zygotic knockdown of smog by using TRiP.HMC03192 (B), with M/Z knockdown of smog by using TRiP.GL01473 (C), WT (D), smog null [mild] (E), and smog null [severe] (F). (A’-F’) Cell segmentation based on apical areas of A-F. Red and white lines, SG boundary. Asterisks, invagination pit. (G, H) Quantification of percentage (left) and cumulative percentage (right) of cells with different apical areas. Mann-Whitney U test (for percentage of cells) and Kolmogorov-Smirnov test (for cumulative percentage of cells). n=8 SGs (control, 902 cells; smog RNAi (Z), 823 cells; smog RNAi (M/Z), 881 cells; WT, 892 cells; smog null [mild], 859 cells); 6 SGs (smog null [severe], 672 cells). ***p<0.001; ****p<0.0001. (I-L) 3D reconstruction of confocal z-sections of SGs using Imaris for WT (I), M/Z knockdown of smog (J), smog null [mild] (K), and smog null [severe] (L). Green, E-Cad. Magenta, CrebA. Asterisks, invagination pit. Arrows in J and L, exaggerated grooves. Arrowheads in J and L, enlarged SG cells. (M, N) Tile scan confocal images of stage 11 embryos in WT (M) and M/Z knockdown of smog (N). Green, E-Cad. Magenta, CrebA. Arrows, ventral midline. Arrowheads, SG.

Since smog has a strong maternal contribution (Flybase Developmental RNA-Seq), we also knocked down smog both maternally (using matα-Gal4; Häcker and Perrimon, 1998) and zygotically in the SG (using fkh-Gal4) and observed slightly more severe apical constriction defects in SG cells (hereafter referred to as smog M/Z knockdown; Fig. 3C, C’; quantification in 3G). Maternal knockdown of smog using the stronger RNAi line (TRiP.HMC03192) resulted in severe defects in egg-laying. Therefore, all M/Z knockdown experiments were performed using the weaker line (TRiP.GL01473). Consistent with the ubiquitous expression of smog and its role in epithelial morphogenesis in the early Drosophila embryo (Kerridge et al., 2016), smog M/Z knockdown also resulted in a slight disruption of the overall embryonic morphology, including defects in the head region, a wavy embryo surface, and an irregular ventral midline (Fig. 3M, N). Unlike the circular SG placode of WT, smog M/Z knockdown also resulted in a SG placode that was elongated along the dorsal/ventral axis (compare Fig. 3A, A’ to 3C, C’). 3D reconstruction of confocal images revealed disrupted epithelial morphology in smog M/Z knockdown, with wide and wavy grooves, enlarged epithelial cells, and an elongated SG placode (Fig. 3I, J).

smog zygotic null mutants (Kerridge et al., 2016) had a range of defects similar to those observed with smog knockdown. 73.8% of embryos (59/80) showed relatively normal embryonic morphology with mild apical constriction defects in SG cells, like smog zygotic knockdown (classified as ‘smog null [mild]’; Fig. 3D-E’, K; quantification in 3H); 26.2% of embryos (21/80) showed a disrupted embryonic morphology with SGs elongated along the dorsal/ventral axis (Fig. 3F, F’, L) and more severe apical constriction defects (Fig. 3H), like smog M/Z knockdown (classified as ‘smog null [severe]’). All embryos at late stages (stages 15-17) smog null mutants showed relatively normal embryonic morphology (Fig. S2A, B), suggesting that severely defective embryos with smog loss do not survive/develop past stage 14. Notably, these embryos have relatively normal internalized SGs (Fig. S2F, G), except for rare cases of crooked SG morphology (Fig. S2D), confirming our previous finding that apical constriction is not required for SG internalization (Chung et al., 2017). Altogether, these data suggest that Smog is required for coordinated apical constriction during SG invagination and proper epithelial morphogenesis during embryogenesis.

Smog regulates apicomedial and junctional myosin in a Fog-dependent and - independent manner in the SG

We next examined myosin levels and distribution in the SGs in smog knockdown or smog mutants using sqh-GFP. Apicomedial myosin, which generates the pulling force for apical constriction during SG invagination (Booth et al., 2014), is the only myosin pool defective in the SGs of fog mutants (Chung et al., 2017). Consistent with the idea that Smog transduces Fog signal in the SG, both smog knockdown (either zygotic or M/Z knockdown) and smog loss resulted in a significant reduction in the intensity of apicomedial myosin in SG cells near the invagination pit, compared to control (Fig. 4 A-D’’; quantification in 4I). Moreover, compared to clear web-like structures of apicomedial myosin in control SG cells (Fig. 4A-A’’), SG cells in smog knockdown or smog null mutants showed reduced areas of sqh-GFP puncta, suggesting dispersed myosin along the entire apical surface in SG cells (Fig. 4K). Interestingly, the intensity of junctional myosin was also significantly reduced in SG cells with smog knockdown or smog loss, compared to control (Fig. 4A-D’’, quantification in 4J). This phenotype is distinct from what was observed in fog mutants, where junctional myosin was unaffected in SG cells (Chung et al., 2017). We conclude that Smog is required for maintaining normal levels of both apicomedial and junctional myosin in the SG in a Fog-dependent and - independent manner, respectively.

Figure 4.
Figure 4. smog regulates all three pools of myosin in the SG in a Fog-dependent and -independent manner.

(A-D’’) Confocal images of SGs with sqh-GFP (green) and E-Cad (magenta) signals. Compared to control (A-A’’), sqh-GFP signals are dispersed in SGs with zygotic (B-B’’) and M/Z (C-C’’) knockdown of smog and in smog null [mild] embryos (D-D’’). (A’-D’’) Higher magnification of the yellow boxed region in A-D. (E-H’’) Confocal images of SGs with Rok-GFP (green) and E-Cad (magenta) signals. Compared to control (E-E’’), Rok-GFP signals are dispersed in SGs with zygotic (F-F’’) and M/Z (G-G’’) knockdown of smog and in smog null [mild] embryos (H-H’’). (E’-H”) Magnified images of the yellow boxed region in E-H. (I-L) Quantification of apicomedial myosin intensity (I), junctional myosin intensity (J), area of apicomedial myosin particles (K), and area of Rok-GFP particles (L). *p≤0.05; **p≤0.01; ***p≤0.001; ****p≤0.0001 (Welch’s t-test). n=5 SGs, 100 cells (I-K) and 5 SGs, 75 cells (L) for each genotype. (M) Control SG with high myosin intensity at the placode boundary (arrowheads in insets in M, M’’). (N) smog null [severe] SGs have low myosin intensity. Arrowheads in insets in N, N’’, discontinuous myosin at the SG boundary. (O) Analysis of the circularity of the SG placode boundary as a measure of smoothness and tension. An example of a WT SG stained for E-Cad (magenta) and sqh-GFP (green) with the boundaries measured. Red line, SG boundary; gray line, cell boundary one cell row outside the SG boundary; blue line, cell boundary one cell row inside the SG boundary. (P) Quantification of circularity of the SG, outer, and inner boundary in control and smog null [severe] SGs. Asterisks, invagination pit. *p≤0.05; **p≤0.01; ***p≤0.001 (Welch’s t-test). n=5 SGs for each genotype.

We also tested Rok distribution in SG cells in smog RNAi and smog null mutants, using Rok-GFP. Consistent with our previous study (Chung et al., 2017), Rok-GFP formed large punctate structures in the apicomedial region in control SG cells (Fig. 4E-E’’). However, Rok-GFP failed to accumulate and was dispersed along the entire apical domain in SG cells in smog knockdown and smog null mutants (Fig. 4F-H’’; quantification in 4L), similar to the dispersed Rok-GFP distribution observed in fog mutant SG cells (Chung et al., 2017). These data suggest that Smog is required for Rok accumulation in the apical domain of SG cells.

smog loss leads to defects in the tissue-level myosin cable

Reduced myosin levels and the elongated SG placode in smog null [severe] embryos led us to test the third pool of myosin in the SG, the supracellular myosin cable surrounding the SG placode. Consistent with the idea that this myosin cable is under tension (Chung et al., 2017; Röper, 2012), WT SGs showed a relatively smooth tissue boundary (Fig. 4M-M’’). However, SGs in smog null [severe] embryos showed an irregular boundary and a less clear myosin cable (Fig. 4N-N’’). We calculated the circularity of the SG placode as a measure of smoothness and tension of the SG boundary. The circularity of the WT placode was significantly higher than the circularity of one cell row inside (inner boundary) or outside of SG placode (outer boundary), suggesting a higher tension of the myosin cable in WT (Fig. 4O, P). Consistent with the irregular SG boundary in smog null [severe], the circularity of the SG placode in smog null embryos was significantly lower than that of WT and was not statistically significant from the inner or outer boundaries, suggesting a lack of tension at SG boundary in smog null mutants (Fig. 4P). Overall, our data suggest that the loss of smog affects all myosin pools in the SG, including the myosin at the SG boundary, leading to reduced tension at the SG boundary and defects in generating the compressing force.

Smog is required for proper localization of apical and junctional components and microtubule networks in epithelial cells

Since we observed a range of defects in whole embryo morphology in smog mutants, we explored the potential cellular basis. In stage 11 smog null [severe] embryos, we occasionally observed large areas in the epidermis and the SG where E-Cad (Fig. 5B, B’) and Crb (Fig. 5D, D’) signals were absent or significantly reduced. Similar defects were observed in late-stage (stage 14) embryos. Crb levels were significantly reduced in large areas; in the most severe cases, Crb was dispersed in the entire epidermis (Fig. S3B, B’). These data suggest that Smog is required for maintaining epithelial polarity and junctional integrity by regulating E-Cad and Crb levels or distribution. To better understand the basis of the reduced E-Cad and Crb in smog mutants, we examined smog null mutant SG cells at higher magnification. Even SGs that do not contain large patches where E-Cad/Crb are missing showed discontinuous E-Cad/Crb signals, resulting in small gaps in signal along the SG cell boundary (Fig. 5F, G, I, J). Such gaps were observed in both smog null [mild] and smog null [severe] embryos. Although the number and the length of gaps in E-Cad signals in smog null [mild] were comparable to those of WT, they were significantly higher in smog null [severe] compared to WT (Fig. 5F, G; quantification in 5K, L). The number and the length of gaps in Crb signals in SG cells were significantly higher in both mild and severe smog null mutant embryos compared to WT (Fig. 5I, J; quantification in 5M, N). These data suggest a role of Smog in regulating Crb and E-Cad levels and continuity during epithelial morphogenesis, including the SG.

Figure 5.
Figure 5. Smog is required for maintaining epithelial integrity and the microtubule networks.

(A-D) Confocal images of stage 11 WT (A, C) and smog null [severe] (B, D) embryos (A, B) Embryos stained for E-Cad (green) and CrebA (magenta). (A’, B’) Magnified view of the white boxed region in A and B. Compared to uniform and continuous E-Cad signals in WT embryonic epithelial cells (A’), E-Cad signals are absent in regions of epithelial cells in smog mutants (arrowheads in B’). (C, D) Embryos stained for Crb (green) and CrebA (magenta). (C’, D’) magnified view of the white box region in C and D. In contrast to continuous WT Crb signals (C’), Crb is almost absent in regions of epithelial cells in smog mutants (arrowheads in D’). (E-J) E-Cad (E-G) and Crb (H-J) signals in WT (E, H), smog null [mild] (F, I), and smog null [severe] (G, J) embryos. Insets, magnified view of yellow boxed region in the SG. Arrowheads, gaps in E-Cad or Crb signals. (K-N) Quantification of the number of gaps in E-Cad (K) and Crb (M) signals and the ratio of gap length to total junctional length for E-Cad (L) and Crb (N). n=5 SGs, 50 cells per genotype. **p≤0.01; ***p≤0.001 (Welch’s t-test). (O-R) Compared to WT (O, Q), acetylated tubulin signals are almost absent in the SG in smog mutants (P), and tyrosinated tubulin signals are also reduced in smog mutants (R). Red dotted lines, SG boundary. (S) Quantification of the ratio of tyrosinated tubulin intensity in SG cells to the intensity in cells outside the SG placode. n=4 SGs (control); 5 SGs (smog null). *p≤0.05 (Welch’s t-test). Asterisks, invagination pit.

Anisotropic localization of Crb at the SG boundary has been suggested to drive the formation of supracellular myosin cable in SG (Röper, 2012). Consistent with this, in control SGs, Crb levels were higher in the junctions that did not contribute to the myosin cable, whereas myosin was highly enriched at the junction forming the cable at the SG boundary (Fig. S3C-E’’’). However, in several smog null [severe] SGs, the overall Crb intensity was quite low, and Crb did not show an anisotropic localization at the SG boundary (Fig. S3F-H’’). This data suggests that smog loss results in the reduction of both myosin and Crb, failing to form the supracellular myosin cable at the SG boundary.

Microtubules are required for forming and maintaining apical myosin structures (Booth et al., 2014; Ko et al., 2019) and for transporting several key apical and junctional proteins in Drosophila tubular organs, including E-Cad and Crb (Le and Chung, 2021; le Droguen et al., 2015). We recently showed that microtubule-dependent intracellular trafficking has an important role in regulating apical constriction and apicomedial myosin during SG invagination (Le and Chung, 2021). To examine the microtubule networks in the SG of wild-type and smog mutant embryos, we stained with antibodies against acetylated-α-tubulin, a marker of stable, long-lived microtubules, and tyrosinated-α-tubulin, a marker of dynamic, short-lived microtubules (Westermann and Weber, 2003). We observed a reduction of both acetylated-α-tubulin and tyrosinated-α-tubulin signals in the apical region of the SG in smog mutants, compared to wild type (compare Fig. 5O, Q to Fig. 5P, R; quantification in 5S), suggesting disrupted microtubule networks in the SG in smog mutants. This finding suggests that loss of smog could affect apical pools of myosin, Crb and E-Cad through effects on microtubule abundance, distribution and/or polarity.

Smog is required for cortical actin organization in the SG

We observed numerous blebs in the apical membrane of SG cells in smog null [severe] mutants, which were most prominent near the invagination pit (compare Fig. 6A-A’’’ to Fig. 6B-B’’’). Such blebs were not observed in SGs in WT, smog null [mild], or fog mutant embryos (Fig. S4A-C’’). Unlike dispersed Rok-GFP signals in smog knockdown or smog null [mild] embryos (Fig. 4F-H’’), Rok-GFP was enriched in many of these blebs (Fig. 6B-B’’’), consistent with Rok’s recruitment to retracting blebs (Aoki et al., 2016). In smog null [severe] embryos, blebs were also occasionally observed in cells outside the SG placode (Fig. S4D-E’’). During Drosophila gastrulation, bleb formation correlates with cortical F-actin holes (Jodoin et al., 2015), and disruption of cortical actin enhances blebbing (Kanesaki et al., 2013). We therefore tested if Smog affects the cortical actin network during apical constriction in the SG. Indeed, phalloidin staining revealed that whereas strong F-actin signals were observed in the apical domain of SG cells near the invagination pit in WT, F-actin was significantly reduced in SGs in smog mutants (Fig. 6C-C’, 6D-D’).

Figure 6.
Figure 6. Smog is required for maintaining cortical actin networks during SG invagination.

(A-B’’’) Rok-GFP (green) and E-Cad (magenta) in SGs in control (A-A’’’) and smog null [severe] (B-B’’’) embryos. (A’-B’’) Higher magnification of the white boxed regions in A and B. (A’’’, B’’’) x-z sections across the yellow lines in A and B. Red arrowheads, normal Rok-GFP signals accumulated in the apicomedial region of control SG cells. White arrowheads, blebs in SG cells in smog mutants enriched with Rok-GFP signals. (C, D) SGs stained for Dia (green) and phalloidin (magenta) in WT (C) and smog null mutant (D) embryos. Compared to high intensity of Dia (red arrowheads in C, C’’) and phalloidin (cyan arrowheads in D, D’’) in the WT SG, Dia and phalloidin signals are reduced in SGs in smog null mutants (red and cyan arrowheads in D-D’’). (E-G) Quantification of Dia intensity in apicomedial (E) and junctional (F) regions and the ratio of apicomedial to junctional Dia intensity (G). (H-I’’) Rok-GFP (green) and E-Cad (magenta) in chic RNAi in the wild-type (H-H’’) and smog mutant background (I-I’’). (H’-I’’) Higher magnification of the white boxed regions in H, I. In chic RNAi SGs, Rok-GFP signals accumulate in the apicomedial region of SG cells (red arrowheads in H’ and H’’). smog null [mild] SG cells with chic knockdown show membrane blebs with enriched Rok-GFP signals (white arrowheads in I’ and I’’). Asterisks, invagination pit. *p≤0.05; **p≤0.01 (Welch’s t-test). n=5 SGs, 50 cells for each genotype.

Apical F-actin formation in Drosophila tubular organs requires the formin family actin nucleator Diaphanous (Dia) (Massarwa et al., 2009; Rousso et al., 2013). Since the apical localization and activity of Dia are critical for restricting F-actin formation to the correct membrane domain, we tested the localization and levels of Dia in the SG in smog mutants. Similar to reduced F-actin, we observed reduced Dia signals in the apical domain of SG cells, most prominently near the invagination pit in smog mutants, compared to WT (Fig. 6C-C’’, D-D’’). Quantification of the Dia intensity revealed that Dia signals are reduced in the apicomedial but not in the junctional region of SG cells (Fig. 6E-G). Our data suggest that Smog is required for the cortical actin networks during SG invagination, at least in part via modulating Dia levels.

We next tested if the reduction in actin levels in smog mutants is due to defects in actin turnover. To test this, we modulated the level of actin regulators in the SG in smog mutants and assayed the blebbing phenotype. Using RNAi, we knocked down chickadee (chic), which encodes Drosophila profilin, a protein that increases F-actin by promoting actin polymerization (Cooley et al., 1992), in the smog mutant background. If there is a shift from F-actin to monomeric G-actin in smog mutants, we expect enhancement of the disorganized actin phenotype and bleb formation with a reduced chic level. Blebs were not observed in smog null [mild] embryos or in SGs knocked down for chic in the otherwise wild-type background (Fig. 6H-H’’; Fig. S4B-B’’). However, when chic was knocked down in the SG in the smog mutant background, we observed enhanced blebbing even in mildly defective SG cells (Fig. 6I-I’’). Since profilin promotes formin-mediated actin filament assembly (Romero et al., 2004; Zweifel and Courtemanche, 2020), our data suggests that reduced profilin by chic knockdown aggravated defects in cortical actin organization with reduced Dia in smog mutants. Overall, our data suggest that Smog is required for cortical actin organization as well as myosin activation during epithelial morphogenesis.

DISCUSSION

Smog regulates different pools of myosin in a Fog-dependent and -independent manner during SG invagination

Fog signaling triggers epithelial cell shape changes driving tissue folding and invagination during development of Drosophila and other insects (Benton et al., 2019; Manning and Rogers, 2014). During Drosophila embryogenesis, fog is upregulated in multiple tissues undergoing apical constriction and tissue invagination, such as ventral furrow (mesoderm), the posterior midgut (endoderm), and the SG (ectodermal derivative) (Nikolaidou and Barrett, 2004). During Drosophila gastrulation, the ubiquitously expressed GPCR Smog and the mesoderm-specific GPCR Mist respond to Fog signal to regulate myosin contractility during mesoderm invagination (Kerridge et al., 2016; Manning et al., 2013). Here, we provide evidence that Smog functions as a SG receptor for Fog to activate myosin during SG invagination. Knockdown of smog by RNAi suppresses the gain-of-function effect of Fog in the SG (Fig. 1). Also, Smog-transfected S2 cells contract upon Fog signal (Fig. 2), suggesting that Smog responds to Fog to regulate myosin contractility both in vivo and in vitro. Our data show that Fog overexpression leads to Smog recruitment to the apicomedial region of SG cells (Fig. 1). Consistent with our data, Fog overexpression induces oligomerization of Smog in early Drosophila embryos (Jha et al., 2018).

Our study also dissects the roles of Smog in myosin activation during SG morphogenesis and reveals Fog-independent roles of Smog. Whereas fog mutants exhibit a decrease in only the apicomedial myosin pool in SG cells (Chung et al., 2017), smog loss and SG-specific knockdown of smog result in a significant reduction of both apicomedial and junctional myosin (Fig. 4). We propose that Smog regulates junctional myosin during epithelial morphogenesis in response to an unknown, ubiquitously expressed ligand and regulates apicomedial myosin in response to Fog in tissues with high Fog signals, such as the mesoderm (Kerridge et al., 2016; Manning et al., 2013) and the SG (this study). Consistent with this model, during Drosophila gastrulation, Smog activates different myosin pools in tissues with different Fog levels. In the mesoderm, where Fog levels are high, Smog and Mist transduce Fog signal to activate myosin in the apicomedial region and drive apical constriction. In the ectoderm, where

Fog levels are very low, Smog is required for junctional myosin activation to drive cell intercalation. SG cells undergo both apical constriction and cell intercalation, and our data shows that Smog regulates both pools of myosin in the SG in a Fog-dependent and -independent manner. It is possible that in tissues with high Fog signals, an additional tissue-specific GPCR(s) function with Smog in response to Fog to fine-tune the Fog signaling for proper apical constriction, like Mist does in the mesoderm. It will be interesting to determine whether additional SG-specific GPCRs function with Smog during SG invagination.

The downstream components that transduce signals from the Smog GPCR to regulate myosin in distinct regions of SG cells await discovery. Good candidates include two Rho guanine nucleotide exchange factors (RhoGEFs), RhoGEF2 and Dp114RhoGEF, which have been shown to activate Rho1 signaling at the apicomedial and junctional domain of epithelial cells, respectively (Garcia De Las Bayonas et al., 2019). This process requires upstream activation by heterotrimeric G proteins Gα12/13/Cta and Gβ13F/Gγ1 at the apicomedial and junctional domains, respectively (Garcia De Las Bayonas et al., 2019). Thus, distinct ligand-receptor binding with Smog may activate distinct G proteins and/or RhoGEFs in the apicomedial and junctional domains.

Smog regulates cortical actin organization during SG invagination independent of Fog

We also discover that Smog regulates cortical F-actin organization during SG invagination (Fig. 6). smog loss significantly reduces levels of F-actin and the formin protein Dia (Fig. 6). Rho1 regulates actin polymerization via Dia (Goode and Eck, 2007), and apical F-actin formation in the Drosophila SG requires apical localization and activity of Dia (Massarwa et al., 2009; Rousso et al., 2013). Our data suggest that Smog activates Rho1 signaling to regulate both cortical F-actin (via Dia) and myosin (via Rok) (Fig. 7). Since mutations in fog do not enhance blebbing during SG invagination (Fig. S4), Smog’s role in regulating cortical actin is independent of Fog. Supporting our data, during Drosophila gastrulation, fog mutants show normal GFP-Moesin signals in the ventral furrow, suggesting normal cortical actin organization upon fog loss (Kanesaki et al., 2013). Gα12/13/Cta is a key component downstream of Fog that recruits RhoGEF2 to the apical membrane to regulate myosin contractility (Barrett et al., 1997; Dawes-Hoang et al., 2005). Interestingly, mutations in Gαi (G-iα65A) and Gβ13F/Gγ1, but not Gα12/13/Cta, disrupt cortical actin organization in early Drosophila embryos (Fox and Peifer, 2007; Kanesaki et al., 2013). Therefore, Smog may signal through Gαi and/or Gβ13F/Gγ1 to organize cortical actin, independent of Fog and Gα12/13/Cta.

Figure 7.
Figure 7. A proposed model for roles of Smog during epithelial tube formation.

During SG invagination, Smog responds to Fog signal to promote Rok accumulation and apicomedial myosin formation to control apical constriction (red arrows). Independent of Fog (and responding to an as-yet-unknown ligand X), Smog regulates epithelial integrity by regulating junctional and the supracellular myosin pools, the microtubule networks, and key apical/junctional components (grey arrows). Smog is also required for cortical actin organization during SG invagination, through regulating Dia levels (grey arrows). Different ligands may recruit specific G proteins and RhoGEFs to activate distinct downstream effectors.

The link between heteromeric G proteins and Rho1 signaling occurs through activation of specific RhoGEFs (Vázquez-Victorio et al., 2016). Actin polymerization resulting from Fog-independent Smog activation may imply the recruitment of distinct proteins modulating the activation of Rho1-mediated pathways. For example, in the absence of Fog, a specific RhoGEF(s) downstream of Gαi or Gβ13F/Gγ1 may link Smog and Rho1 signaling to regulate cortical actin organization; upon high Fog signals, a different Rho1 modulator(s) downstream of Gα12/13/Cta may be recruited to facilitate apicomedial myosin formation. Further work is needed to identify the link between Smog and Rho1 activation in the presence or absence of Fog during SG invagination. It will be also interesting to determine whether Fog-independent roles of Smog–regulating cortical actin and junctional myosin–involve common or distinct Rho1 modulators.

Smog regulates epithelial integrity during development

smog null mutant embryos show a range of defects, allowing us to dissect the roles of Smog in epithelial morphogenesis. Our data reveal that Smog is required for epithelial integrity by affecting levels and localization of Crb and E-Cad (Fig. 5). Since the decrease in the level of Crb and E-Cad in smog mutants correlates with significantly reduced myosin levels, disrupted epithelial integrity may be due to disrupted actomyosin networks. Myosin II is critical for cells to concentrate E-cadherin at cell-cell contacts (Shewan et al., 2005). In early Drosophila embryos, the absence of contractile myosin leads to the disassembly of junctions (Weng and Wieschaus, 2016). A possible scenario is that Smog is required to recruit key Rho1 regulators, such as RhoGEF2, Dp114RhoGEF, or other SG-upregulated RhoGEFs, to AJs during SG invagination to promote junctional myosin assembly. Since Dia helps coordinate adhesion and contractility of actomyosin (Homem and Peifer, 2008) and formin-mediated actin polymerization at AJs stabilizes E-Cad and maintains epithelial integrity (Rao and Zaidel-Bar, 2016), reduced Dia levels in smog mutants may also contribute to loss of epithelial integrity.

Reduced and discontinuous signals of apical and junctional components in smog mutant embryos can also be due, in part, to compromised intracellular trafficking. Disorganized microtubule networks in SGs in smog mutants (Fig. 5) support this idea. Our recent study showed that microtubule- and Rab11-dependent intracellular trafficking regulates apical myosin pools and apical constriction during SG invagination, via apical enrichment of Fog and the continuous distribution of Crb and E-Cad along junctions (Le and Chung, 2021). The mechanism of how Smog affects microtubule organization remains to be revealed. Overall, our findings suggest the multifaceted roles of Smog during epithelial tube formation in regulating distinct myosin pools and cortical actin in different subcellular domains in a ligand-dependent manner.

MATERIALS AND METHODS

Fly stocks and genetics

The fly stocks used in this study are listed in Table S1. All the crosses were performed at 25°C.

Immunofluorescence and confocal microscopy

Drosophila embryos were collected on grape juice agar plates supplemented with yeast paste at 25 °C. Embryos were dechorionated with 50% bleach. For most samples, embryos were fixed in 1:1 heptane: formaldehyde for 40 minutes and devitellinized with 80% EtOH. For Rok-GFP, Sqh-GFP, and phalloidin staining, embryos were hand-devitellinized. Embryos were then stained with primary and secondary antibodies in PBSTB (1X PBS with 0.1% Triton X-100 and 0.2% bovine serum albumin) and PBTr (1X PBS with 0.1% Triton X-100) for EtOH devitalization and hand-devitalization, respectively. Antibodies used in our experiments are listed in Table S2. Embryos were mounted in Aqua-Poly/Mount (Polysciences, Inc) and imaged with a Leica TCS SP8 confocal microscope using 63x, NA 1.4 and 40x, NA 1.3 objectives. Images were acquired as z-stacks (each 0.3 µm apart) that span the apical and junctional domains of cells in the SG placode.

Fluorescent in situ hybridization

Embryos were fixed using formaldehyde/heptane for 30 min followed by devitellinization with methanol. SP6 polymerase-synthesized digoxigenin (DIG)-labeled antisense probe was prepared using the following primers: smog-F, 5’-ACAGAGCCCACCTGTGTAGG-3’; smog-R, 5’-TCGCTGATCGAAAATGATCTC-3’. Fluorescent in situ hybridization was performed using standard methods as described in (Knirr et al., 1999). Briefly, the embryos were pre-hybridized for 1 hour at 56°C post-fixation. Following this step, the embryos were hybridized with the probe overnight at 56°C. The next day, embryos were stained for DIG using the anti-DIG primary antibody and biotin-conjugated secondary antibody. The embryos were then incubated with AB solution (PK-6100, VECTASTAIN Elite ABC kit). Tyramide signal amplification reaction was performed using Cy3 fluorescent dye (diluted 1:50 using amplification diluent) (Perkin Elmer Life Sciences, Inc., NEL753001kt). The embryos were co-stained with CrebA and Crb antibodies for visualizing SG nuclei and apical cell boundaries, respectively.

Total Smog-GFP intensity

Maximum intensity projections of two z-sections that span the apical and the junctional region of SG cells were used. Intensity means of Smog-GFP signals of SG cells in the entire SG placode were measured using Imaris software. The mean intensity of Smog-GFP was normalized by the median deviation. The integrated density of Smog-GFP was calculated by multiplying the apical area of the cell with the mean intensity of each cell. Total Smog-GFP signals within the whole placode were calculated as the sum of the integrated density of all cells. For background correction, mean gray values of Smog-GFP in ten cells outside of the SG placode were measured. The average value of mean gray values of Smog-GFP in these ten cells was used to subtract the background of the cells inside the placode from the same embryo. Four and five SGs were used to quantify control and Fog overexpression, respectively. P values were calculated using Welch’s t-test in the GraphPad Prism software.

Cell segmentation and apical area quantification

SGs that were invaginated within the range of 4.8-9.9 µm depth were used for quantification. Two or three z-sections of the apical domain were used to generate a maximum intensity projection (Leica LasX software). SG cells were marked using CrebA and segmented along E-Cad signals using the Imaris program. Apical areas of segmented cells were calculated using Imaris, and cells were color-coded based on their apical domain size. Frequency distribution was performed using GraphPad Prism. Apical areas from eight SGs were quantified in control, smog knockdown (Z), and smog null [mild] embryos. Six SGs were used for quantification for smog knockdown (M/Z) and smog null [severe] embryos. Statistical significance was determined using Mann-Whitney U test (percentage of cells) and Kolmogorov-Smirnov test (cumulative percentage of cells).

Quantification of myosin and Dia intensity in the apicomedial and junctional domains

Maximum intensity projection of two z-sections spanning apical and junctional regions of SG cells with Sqh-GFP and E-Cad signals was used. The images were then exported to ImageJ and converted to RGB stack image type. Twenty cells near the invagination pit in each of five SGs were used for quantification. To calculate the apicomedial and junctional myosin intensity, regions were drawn manually along the inner and outer boundary of the E-Cad signal of each cell, and mean gray values were measured for apicomedial and junctional myosin. For background subtraction, the average mean gray value of sqh-GFP signals for ten cells outside the SG placode was subtracted from the mean gray value of apicomedial and junctional myosin for each SG cell. SuperPlots (Lord et al., 2020) were used to address the variability of datasets in each SG. Each data point in the graph represents one SG consisting of twenty cells used for quantification. Statistical significance was determined using Welch’s t-test. Dia intensity in the apicomedial and junctional domain of SG cells was quantified using the same strategy, with ten cells near the invagination pit in each of five SGs.

Quantification of areas of Rok-GFP and apicomedial myosin particles

A single z-section of the confocal image with the strongest Rok-GFP signals in the apicomedial region was used. Rok-GFP signals were converted to black and white using the threshold tool in Adobe Photoshop. Junctional Rok-GFP signals were removed manually based on E-Cad signals. Areas of Rok-GFP puncta were determined using the Analyze Particles tool in ImageJ. Rok-GFP puncta with area ≥0.02μm2 was measured. Fifteen cells in the dorsal posterior region near the invagination pit of the SG placode were used for quantification. The same strategy was used for Sqh-GFP signals to quantify the area of apicomedial myosin. SuperPlots show mean values of data points of five SGs. Statistical significance was determined using Welch’s t-test.

Quantification of the waviness of junctions

Using a single z-section with the highest E-Cad signals and ImageJ software, the shortest (L0) and the actual distance (L) between vertices were measured in SG cells. The ratio of these distances (L/L0) was used as the waviness of junctions. 10-15 cells near the invagination pit in each of five SGs were used for quantification. Statistical significance was determined using Welch’s t-test.

Quantification of number and length of gaps of Crb and E-Cad

The number and the length of gaps for Crb and E-Cad signals were measured using confocal images and ImageJ. A single z-section with the highest Crb or E-Cad signals was chosen for quantification. Gaps with a length ≥0.2 µm were used for quantification. All junctions from ten cells in five SGs were quantified. If there were multiple gaps on a junction, we added the length of all the gaps on that junction to calculate the ratio of the length of gaps to junctional length. Statistical significance was determined using Welch’s t-test.

Circularity of the SG boundary

Using confocal images and ImageJ, cell boundaries were manually drawn along E-Cad signals at the SG placode boundary and one cell row outside and inside the placode. For most samples, strong myosin cable signals at the dorsal, anterior, and posterior boundaries of the placode and CrebA signals in SG cells were used to determine the SG boundary. In smog null [severe], where myosin signals were significantly reduced in all cells, CrebA signals were used to determine the boundary. The ventral midline was used as the ventral boundary of the SG in all cases. The perimeter and the area of the SG corresponding to these boundaries were measured using ImageJ, and circularity was calculated using the formula, C = 4π area/perimeter2. For a perfect circle, the value of circularity should be 1. Five SGs were used for quantification, and statistical significance was determined using Welch’s t-test.

Tyrosinated tubulin intensity quantification

Three z sections spanning the most apical region of SG cells were used for quantification. The mean intensity of tyrosinated tubulin signals in the whole SG placode was measured using ImageJ. The area of the SG was drawn manually based on the E-Cad and CrebA signals to label the cell boundary and SG nuclei, respectively. For background correction, the mean intensity of ten cells outside the SG of the same embryo was used. The ratio of the mean intensity of tyrosinated tubulin signals inside the placode to the mean intensity of control cells outside the placode was calculated and plotted. Four and five SGs were used for control and smog mutant, respectively. t-test with Welch’s correction was used to calculate the p value.

Cell contractility assay

cDNA clones for smog (RE70685) and mist (RE13854) were obtained from Drosophila Genomics Resource Center (DGRC). Isolated cDNAs were cloned into the pMT-V5-HisB vector (Addgene) using the conventional restriction digestion and ligation method. Primers with restriction sites used to isolate the open reading frame are stated below (restriction sites underlined).

  • smog CDS-EcoRI-5’, 5’-CCGGAATTCATGGAACTGTGCATAGCAAC-3’

  • smog CDS-NotI-3’, 5’-ATTTGCGGCCGCATTGGTCGTGATTGTATCTTTGG-3’

  • mist CDS-EcoRI-5’, 5’-CCGGAATTCATGGACAGGAGTCGGAGTAGC-3’

  • mist CDS-NotI-3’, 5’-ATTTGCGGCCGCAGCAAATGGTCTCCATTTTG-3’

S2 and S2-Mt-Fog-myc cells (Manning et al., 2013) were obtained from DGRC. Shields and Sang M3 insect media (HiMedia) was used with 10% Fetal Bovine Serum (Corning) and 1:1000 Antibiotic-Antimycotic (Gibco). Cells were cultivated at 25°C either in 25-cm2 flask in 5 ml media or 75-cm2 flask in 15 ml media. To prepare for the Fog media, S2-Mt-Fog-myc cells were grown undisturbed for 4-8 days to attain approximately 100% confluency. 50 µl of 100 mM CuSO4 was added to nearly 100% confluent cells in 75-cm2 flask to induce metallothionein promoter. The Fog-containing media was collected by centrifugation and further concentrated using 3,000 MW concentrators. The presence of Fog was confirmed by Western blot using the Myc-antibody (Invitrogen; RRID:AB_2533008). Non-transfected S2 cells were used as a control. Cell contractility assay was performed as described in (Manning et al., 2013). Smog- or Mist-transfected S2 cells were induced with CuSO4. Cells were then transferred to Concanavalin A-coated coverslips and fixed using 10% paraformaldehyde in PBTr for 15 minutes at room temperature (RT). After being blocked in 5% NGS in PBTr for 20 minutes at RT, cells were stained with the V5 antibody (Invitrogen; RRID:AB_2556564) to differentiate between transfected and non-transfected cells. Cell contractility of transfected cells was monitored using a phase-contrast filter in a Leica DM2500 microscope using the 40X, 0.8 NA objective. Three independent experiments were performed. Two-way ANOVA was performed to calculate statistical significance.

Fly strains used (Table S1)

Table

Antibodies Used (Table S2)

Table

Figure Legends

Supplementary Figure S1.
Supplementary Figure S1. smog knockdown using a weaker RNAi line also shows apical constriction defects in SG cells.

(A, A’) A representative confocal image (A) of the stage 11 SG in fog mutants and the corresponding heat map of apical areas (A’). The embryo surface is often uneven in fog mutant embryos due to additional folding and grooves, resulting in some cells outside the SG being out of focus and not shown in the image. (B-C’) Confocal images of SGs in control (B) and zygotic knockdown of smog by using TRiP.GL01473 (C) and corresponding heat maps of apical areas (B’ and C’). Red and white lines, SG boundary. Asterisks, invagination pit. (D) Quantification of percentage and cumulative percentage of cells with different apical areas. Mann-Whitney U test (for percentage of cells) and Kolmogorov-Smirnov test (for cumulative percentage of cells). n=8 SGs (control, 902 cells; smog RNAi(Z), 856 cells). **p≤0.01.

Supplementary Figure S2.
Supplementary Figure S2. SGs in smog null mutants have relatively normal morphology at stage 14 and later during embryogenesis.

(A, B) Confocal images of stage 15 WT (A) and smog null embryos stained for E-Cad (green) and CrebA (magenta). (C-G) SGs stained for E-Cad in WT (C, E), smog M/Z knockdown (F), and smog null mutant (D, G) embryos. All embryos at stage 14 and later have relatively normal morphology, including the SG, except for a rare case of the crooked SG in smog null mutants (D).

Supplementary Figure S3.
Supplementary Figure S3. smog loss leads to the reduction of Crb levels in the whole embryo and loss of Crb anisotropy at the SG boundary.

(A, B) Confocal images of stage 14 WT (A) and smog null [severe] mutant (B) embryos stained for Crb (green) and CrebA (magenta). (A’, B’) Magnified view of the white boxed region in A and B. Compared to clear Crb signals at the cell boundary in WT (A’), Crb levels are reduced and diffused in some smog null mutant embryos with a severely defective morphology (B’). (C-H’’’) control (C-E’’’) and smog null [severe] embryos (F-H’’’) showing sqh-mCh (green) and Crb (magenta) signals. (D, G) Cells at the SG boundary. (E, H) SG cells near the invagination pit (asterisk). (D’-H’’’) Higher magnification of the boxed regions. In contrast to high myosin (red arrowheads in D’’’) and low Crb (cyan arrowheads in D’’) levels in the SG boundary in control embryos, the overall Crb levels are reduced in smog null embryos, showing loss of anisotropic localization of Crb at the SG boundary (G-H’’’).

Supplementary Figure S4.
Supplementary Figure S4. smog loss leads to bleb formation in SG cells as well as outside the SG placode.

(A-C’’) Confocal images for Rok-GFP (green) and E-Cad (magenta) signals in stage 11 SGs in control (A-A’’), smog null [mild] (B-B’’), and fog mutant (C-C’’) embryos. Blebs are not observed in these embryos. Due to the uneven embryo surface in fog mutants, apical/junctional and more basal regions are shown for SG cells near and far from the invagination pit, respectively, in the single z section. (D-E’’) Confocal images for Rok-GFP signals in control (D-D’’) and smog null [severe] (E-E’’) embryos. Blebs are observed both in non-SG cells outside the SG placode (red arrowheads in E’) and SG cells (cyan arrowheads in E’’). Yellow arrowheads in D’’, normal Rok accumulation in the apicomedial region of SG cells. Asterisk, invagination pit.

Acknowledgments

We thank the members of the Chung laboratory for comments and suggestions. We thank A. Martin, T. Lecuit and the Bloomington stock center for fly stocks, and D. J. Andrew and the Developmental Studies Hybridoma Bank for antibodies. We thank Flybase for the gene information. We are grateful to D. J. Andrew, C. D. Hanlon, J. Kim, and J. Matthew for their helpful comments on the manuscript. This work is supported by start-up fund from Louisiana State University and the grant from the Board of Regents Research Competitiveness Subprogram GR-00005224 to S.C.

Footnotes

  • A few minor edits have been made, including correcting labeling and typos.

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Posted January 10, 2022.
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Multifunctional role of GPCR signaling in epithelial tube formation
Vishakha Vishwakarma, Thao Phuong Le, SeYeon Chung
bioRxiv 2022.01.06.475238; doi: https://doi.org/10.1101/2022.01.06.475238
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