The proteolysis of ZP proteins is essential to control cell membrane structure and integrity of developing tracheal tubes in Drosophila
Abstract
Membrane expansion integrates multiple forces to mediate precise tube growth and network formation. Defects lead to deformations, as found in diseases such as polycystic kidney diseases, aortic aneurysms, stenosis, and tortuosity. We identified a mechanism of sensing and responding to the membrane-driven expansion of tracheal tubes. The apical membrane is anchored to the apical extracellular matrix (aECM) and causes expansion forces that elongate the tracheal tubes. The aECM provides a mechanical tension that balances the resulting expansion forces, with Dumpy being an elastic molecule that modulates the mechanical stress on the matrix during tracheal tube expansion. We show in Drosophila that the zona pellucida (ZP) domain protein Piopio interacts and cooperates with the ZP protein Dumpy at tracheal cells. To resist shear stresses which arise during tube expansion, Piopio undergoes ectodomain shedding by the Matriptase homolog Notopleural, which releases Piopio-Dumpy-mediated linkages between membranes and extracellular matrix. Failure of this process leads to deformations of the apical membrane, tears the apical matrix, and impairs tubular network function. We also show conserved ectodomain shedding of the human TGFβ type III receptor by Notopleural and the human Matriptase, providing novel findings for in-depth analysis of diseases caused by cell and tube shape changes.
Editor's evaluation
Using the Drosophila tracheal system as a model for apical membrane expansion of tubes, the authors convincingly demonstrate that ectodomain shedding of the Zona Pellucida (ZP) domain protein Piopio by the Matriptase homolog Notopleural releases its linkage with the ZP protein Dumpy and thus ensures proper apical membrane function and tube expansion. Given the high degree of conservation of these proteins in other species, the results presented are important for future analysis and will have further implications on tubular development and homoestasis in other systems, including human.
https://doi.org/10.7554/eLife.91079.sa0Introduction
Tube networks are essential for organisms transporting liquids, gases, or cells across bodies. Endothelial and epithelial cells generate such networks with strict hierarchical order and precise tube dimensions as a prerequisite for proper tube network functioning (Ochoa-Espinosa and Affolter, 2012; Potente and Mäkinen, 2017). Defective cell shapes and tube dimensions result in severe syndromes such as chronic obstructive pulmonary diseases (384 million people in 2010) with high global mortality as well as tube dysfunctions like aortic aneurysms (152,000 deaths worldwide) in blood vessel systems and polycystic kidney diseases (1 in 1000 people) (Adeloye et al., 2015; Barnes et al., 2015; Harris and Torres, 2009; Quintana and Taylor, 2019; Zhai et al., 2021). One fundamental question regarding the formation of functional tube systems is how cells balance forces at the cell membranes emerging during organ growth and the simultaneous maintenance of the tubular network integrity. Drosophila melanogaster embryos form a tracheal system, an excellent model for studying molecular mechanisms controlling cell expansion in combination with tube elongation.
The development of tracheal tubes is subject to precise genetic control. Genes that control cell polarity, junction formation at the lateral membranes, cytoskeletal organization, and intracellular trafficking of tube size determinants support apical cell membrane expansion (Behr et al., 2003; Dong et al., 2014; Förster and Luschnig, 2012; Laprise et al., 2010; McSharry and Beitel, 2019; Nelson et al., 2012; Olivares-Castiñeira and Llimargas, 2017; Skouloudaki et al., 2019; Syed et al., 2012; Tonning et al., 2005; Tsarouhas et al., 2007). In contrast, genes controlling the meshwork of chitinous apical extracellular matrix (aECM) formation restrict excessive cell expansion to prevent tube overexpansion (Luschnig et al., 2006; Moussian et al., 2006; Öztürk-Çolak et al., 2016; Petkau et al., 2012; Tiklová et al., 2013; Wang et al., 2006). Subsequently, genetically controlled mechanisms establish tracheal airway clearance, aeration, and tube stabilization (Behr et al., 2007; Behr and Riedel, 2020; Drees et al., 2019; Stümpges and Behr, 2011; Tsarouhas et al., 2007; Tsarouhas et al., 2019).
Previous studies revealed that axial and radial forces affect tracheal tube elongation. The apical membrane grows axially, pulling on the associated aECM until the aECM’s elastic resistance balances the elongation force throughout the tubes (Dong and Hayashi, 2015). Given the association between cell membranes and aECM, ongoing tube expansion and luminal shear stresses inevitably lead to problematic membrane tension. Once forces are out of equilibrium, tracheal tubes show curvy appearance. Similarly, also blood vessels can appear unstable, twist, kink, and buckle. High stresses even lead to vascular damage and aneurysm rupture (Dong et al., 2014; Han et al., 2013). However, it is not known how cells manage to integrate the axial forces to stabilize the cell membrane and aECM.
Zona pellucida (ZP) domain proteins are critical components of apical cell membranes and aECM (Plaza et al., 2010) and assemble into extracellular fibrillar polymers (Jovine et al., 2005; Litscher and Wassarman, 2020). For example, Uromodulin plays a role in chronic kidney diseases and hypertension (Rampoldi et al., 2011). Secreted Uromodulin requires proteolysis at the apical cell membrane for shedding and polymerization within the tube lumen (Brunati et al., 2015). Similarly, the ZP domain protein Piopio (Pio) (Figure 1—figure supplement 1A) is secreted into the tracheal tubes of Drosophila embryos (Grieder et al., 2008; Massarwa et al., 2009). Pio restricts the elongation of autocellular junctions (Jaźwińska et al., 2003). Further, Pio is involved in relocating microtubule organizing center components γ-TuRC (γ-tubulin and Grips, gamma-tubulin ring proteins). This requires Spastin-mediated release from the centrosome and Pio-mediated γ-TuRC anchoring in the apical membrane (Brodu et al., 2010). However, while Pio is a transmembrane protein, it was detected in the tube lumen (Jaźwińska et al., 2003), but the release mechanism remains unknown. A promising candidate for Pio proteolysis is Notopleural (Np), the functional homolog of the human Matriptase. It is a type II single-transmembrane serine protease in tracheal and lung epithelia and is capable of ectodomain shedding (Bugge et al., 2009). Initial in vitro studies prove that Np cleaves the Pio ZP domain (Drees et al., 2019). The elastic luminal matrix is essential for the integrity of the tubular network. During tube elongation, the matrix balances elongation forces in the anterior-posterior direction (Dong et al., 2014). Here, we show that the ZP domain proteins Pio and Dumpy and the protease Np respond to mechanical stresses when tracheal tubes elongate to ensure normal membrane-aECM morphology.
Results
Pio maintains structural cell membrane continuity
The tracheal lumen matrix consists of a viscoelastic material that is coupled to the apical membrane. The precise balance between apical membrane growth and luminal matrix resistance determines tube shape (Dong et al., 2014). Based on these observations, we expect the following scenario: first, apical membrane growth and opposing restriction by the extracellular matrix produce increased tensile stress during tube expansion of stage 16 embryos (Figure 1A). Second, increasing tension impacts membrane-matrix couplings, which provide the proper balance between both. Thus, enhanced tensile stress may lead to either release or remodeling of the membrane-matrix couplings to avoid potential deformations. The Pio protein contains a transmembrane and an extracellular ZP domain, suggesting that it may link tracheal cells to the aECM. Using CRISPR/Cas9, we generated three pio lack of function alleles (Figure 1—figure supplement 1), we analyzed the two independent alleles pio5m and pio17c which showed embryonic lethality and identical tracheal mutant phenotypes. The tracheal phenotypes of pio5m are shown in the supplement (Figure 1—figure supplement 1B–F). In all other figures, we show images of the pio17c allele. The pio17c and pio5m null mutant embryos revealed the dorsal and ventral branch disintegration phenotype known from a previously described pio2R-16 mutation allele which contains an X-ray induced single point mutation that led to an amino acid replacement (V159D) in the ZP domain (Jaźwińska et al., 2003). In addition, late-stage 16 pio17c and pio5m null mutant embryos showed over-elongated tracheal dorsal trunk tubes (see below) in contrast to the pio2R-16 mutation. We compared the dorsal trunk morphology between control and pio mutant embryos by using the septate junction (SJs) marker Megatrachea (Mega) (Behr et al., 2003). The early stage 17 control embryos revealed tight appearance of tracheal cells and adjacent luminal extracellular matrix. In contrast, the corresponding pio mutant embryos showed irregular bulge-like gaps between the Mega-marked cell membrane and apical matrix (Figure 1B). Such gaps were not detectable in late-stage 15 wild-type (wt) or pio mutant embryos (Figure 1—figure supplement 2A). This suggests that the gaps arise in stage 16 pio mutant embryos during tube length expansion.

Pio supports structural continuity of the apical cell membrane.
(A) Model implicates the axial and longitudinal forces (arrows) acting apical on cell membrane and extracellular matrix of stage (st) 16 embryos when tracheal lumen expands in growing tubes. (B) Confocal images of wt and pio mutant stage 17 embryos. Lateral membrane is marked by Megatrachea (Mega; red, arrows) immunostaining, and transmission light visualizes the tracheal cells and lumen. The yellow line indicates the distance between the apical cell surface and the detached luminal apical extracellular matrix (aECM). Scale bars represent 10 µm. (C, C’) Confocal LSM Z-stack (overview) and Airyscan (close up) images of immunostainings are displayed as maximum intensity (3D projection) and orthogonal (ortho) projections (C’) using Uif antibody and WGA. Scale bars indicate 10 µm. Stage 16 control embryo showed straight apical cell membrane and tracheal tubes. All pio null mutant (n=10) embryos revealed curly elongated tracheal tubes and unusual bulge-like apical cell membrane deformations (yellow arrows in overview and close-up images). The Uif (green) and WGA (red) stainings on the right panel show airyscan images of the region, which is marked by the yellow frame in close left. Position of membrane deformations are marked by the green and red lines (XY axes) with the ZEN orthogonal projection. The corresponding orthogonal projections are indicated. Note that membrane bulge-like structures interfere with the tube lumen integrity (ortho-target cross in mutant). Control and pio mutant embryos were fixed and stained together. (D) TEM analysis of late-stage 17 wt embryos reveal SJs and chitin-rich taenidial folds and a cleared tube lumen. The corresponding pio mutant embryos (n=4) showed normal SJs formation but unusual apical cell membrane bulge-like deformations (yellow arrow), reduced chitin (red arrowheads), taenidial folds with disorganized pattern, and unusual extracellular matrix contents within the tube lumen (*). Scale bars represent 500 nm. (E) Quantification of pio mutant bulge-like apical cell membrane deformations sizes in nm of airyscan Z-stacks (mean value 750 nm, n=15) and TEM (mean value 230 nm, n=61) images. Standard deviations are indicated. It is of note that measurements of TEM images do not always capture the three-dimensionality of bulges and may show only parts of them.
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Figure 1—source data 1
Bulge-like gaps between the Mega-marked cell membrane and apical matrix in stage 17 embryos.
- https://cdn.elifesciences.org/articles/91079/elife-91079-fig1-data1-v1.zip
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Figure 1—source data 2
Bulge-like gaps between the Mega-marked cell membrane and apical matrix in stage 17 embryos.
- https://cdn.elifesciences.org/articles/91079/elife-91079-fig1-data2-v1.zip
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Figure 1—source data 3
Uif marked unusual apical cell membrane deformations at the dorsal trunk in stage 16 embryos and quantification.
- https://cdn.elifesciences.org/articles/91079/elife-91079-fig1-data3-v1.zip
To study the role of Pio during tube length expansion, we examined pio mutant stage 16 embryos using the apical cell membrane marker Uif. This revealed unusual apical cell membrane deformations most prominent at the dorsal trunk (Figure 1C). The corresponding control embryos of the same fixations and staining did not show membrane deformations (Figure 1C). Additional analysis of orthogonal projections of confocal Z-stacks revealed straight apical cell membrane of dorsal tracheal trunks in the control embryos while comparable pio mutants contained numerous small membrane deformations along the dorsal trunk (Figure 1C’). Moreover, these membrane deformations may compromise the shape of the dorsal trunk tube lumen in pio mutant. The control dorsal trunk tubes showed straight and uniform lumens in three-dimensional reconstructions (Figure 1C). The corresponding orthogonal projections confirmed the appearance of a ring-like structures of the apical cell membrane (Figure 1C), which was detected with Uninflatable (Uif) (Zhang and Ward, 2009) and with the cell surface marker WGA (wheat germ agglutinin), indicating the tube lumen formation in the control embryos. In contrast, projections of the pio mutant dorsal trunk tubes showed bulge-like deformations that appeared as small protrusions reaching into the tracheal cells in the corresponding orthogonal projections (Figure 1C and C’).
Furthermore, the orthogonal projections of the pio mutant dorsal tubes did not show straight but unusually narrow and collapsed lumen at the site of apical membrane deformations (Figure 1C’; Figure 1—figure supplement 1D). The additional ultrastructural analysis confirmed the appearance of apical cell membrane deformations at the dorsal trunks of stage 17 pio mutant embryos. The TEM study revealed unusual gaps between the membrane and aECM, while control embryos lacked such gaps. These gaps appear like membrane bulges that protrude into the cells (Figure 1D and E; Figure 1—figure supplement 2B). These results indicate that Pio is required to stabilize or maintain structural membrane-matrix formation.
Anti-Pio antibody (kindly provided by Markus Affolter) detects a short stretch within the Pio ZP domain (Jaźwińska et al., 2003). Immunostainings confirmed Pio protein expression in the tracheal system of stage 16 embryos when tubes expand (Figure 2A). In these embryos, tracheal Pio staining is detectable at the membrane and is enriched within the tracheal lumen, consistent with previous findings (Dong et al., 2014; Massarwa et al., 2009). At the membrane of stage 16 control embryos, Pio overlaps at discrete points with the apical cell membrane determinant Crumbs (Crb) and WGA that stains the cell membranes (Figure 2—figure supplement 1A and B). Similarly, Z-stack projections and fluorescence intensity profile analysis in stage 16 control embryos showed overlapping staining of discrete Pio puncta with Uif at the apical cell membrane (Figure 2C–E; Figure 2—figure supplement 2A and B). In crb mutant embryos, the apical membrane is compromised (Laprise et al., 2010), but luminal content is still secreted (Olivares-Castiñeira and Llimargas, 2018; Stümpges and Behr, 2011). These crb mutant embryos lacked Pio staining at the membrane, and instead, Pio concentrated within the tube lumen. Control embryos showed normal Pio distribution at the membrane (Figure 2B).

Pio localization depends on the apical membrane and supports tracheal air-filling.
Shown are stage 16 embryos. Maximum intensity projections of confocal Z-stacks are shown in A, single confocal images in B, and Airyscan microscopy in C–E. Scale bars indicate 10 µm. (A) Pio protein is expressed in tracheal tubes (db, dorsal branches; lb, lateral branches; dt, dorsal trunk) and other ectodermal epithelial organs. Pio accumulates in the tracheal lumen (arrow) of control embryos. In contrast, Pio staining showed unusual accumulation at the apical cell membrane of wurst mutant embryos (arrowheads point to Pio accumulation). It is of note that embryos were stained together. Shown are whole-mount embryos. hg, hindgut. (B) In crb mutant embryos, Pio staining is within the luminal matrix but not at the apical cell membrane (indicated by yellow dashes). Corresponding control embryos showed Pio at the membrane and predominantly within the lumen. The tracheal-specific wurst knockdown shows Pio accumulation at the apical cell membrane similar to the wurst mutant embryo (compare with A). 2A12 detects the apical extracellular matrix (aECM) protein Gasp. WGA stains cell membrane surfaces and chitin predominantly. (C) Imaris 3D projection. In wt Pio (magenta) is detectable in a punctuate pattern at the tracheal apical cell membrane, partially overlapping with Uif (blue) in stage 16 control embryos. Chitin-binding probe (cbp; green) labels chitin. The stage 16 mega mutant embryo (n=5) showed Pio accumulation at the apical cell surface. The tracheal expression of the Chitinase 2 (n=5) showed a disturbed Pio pattern, including accumulation of Pio puncta at the apical cell surface. The lower panel shows close-ups of the apical cell membrane of the framed area in the upper images. The white arrows indicate Pio puncta at the Uif marked apical cell membrane. We chose comparable regions where a gap formed between the cell membrane and chitin matrix to detect the apical Pio puncta. Note that mega mutant embryo (20 Pio puncta) contains twice as much Pio puncta at the Uif stained apical cell membrane as the control (9 Pio puncta); both show tracheal metamers 7–9. (D) ZEN co-localization, which compares histograms of fluorescence intensities between the two channels, airyscan images. The overlapping Pio (red) and Uif (blue) puncta are colored in white. Arrows point to such overlapping Uif and Pio puncta at the apical cell membrane. (E) Images show representative orthogonal projections of dorsal trunks (metamere 6–8) of control, mega mutant, and Cht2 overexpression embryos. Arrows point to Pio puncta (red) at and near the apical cell membrane marked with Uif (blue). Chitin is shown in green. (F) Quantification of the ratio of maximum fluorescence intensity values between membranes and tube lumen. The control embryos showed a ration at 0.35, the Cht2 overexpression at 1.29, and mega mutants a ration of at 0.9. Bars represent ± SD and p-values from t-test are indicated with asterisks (***, p<0.0001).
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Figure 2—source data 1
Quantification of Pio distribution across the dorsal trunks in different stress situations in stage 16 embryos.
- https://cdn.elifesciences.org/articles/91079/elife-91079-fig2-data1-v1.zip
These observations prompted us to address whether Pio misdistribution depends on apical cell membrane organization. Crb genetically interacts with wurst on late airway maturation, including gas-filling (Stümpges and Behr, 2011). The transmembrane protein Wurst (DNJAC22) is a critical component of clathrin-mediated endocytosis (Behr et al., 2007) and controls the internalization of proteins at the apical membrane (Stümpges and Behr, 2011). Stage 16 wurst mutant embryos and tracheal-specific wurst RNAi (interference) knockdown embryos revealed unusually increased Pio accumulation at the apical cell membrane compared to control embryos (Figure 2A and B; Figure 2—figure supplement 1B and C).
These findings suggest Wurst-mediated internalization of Pio and raise the question of how intracellular Pio trafficking may occur. Retromer and ESCRT-mediated endosomal sorting regulate essential proteins of tracheal tube size control (Dong et al., 2014). Pio is not affected by the retrograde transport from endosomes to the trans-Golgi network (Dong et al., 2013). Vps32/Shrub is a subunit of ESCRTIII, which regulates endocytic sorting of membrane-associated proteins leading to lysosomal degradation and is known to be involved in tracheal tube size control in stage 16 embryos (Dong et al., 2014). Loss of shrub leads to the formation of swollen endosomes that accumulate Crb within tracheal cells (Dong et al., 2014). We observed intracellular Crb staining that overlapped with Pio in shrub mutant embryos (Figure 2—figure supplement 1A). Additionally, Crb and Pio overlapped at the apical cell membrane, suggesting that newly synthesized Pio was secreted as it is known for Crb (Dong et al., 2014). These results indicate that Pio localization relies on apical membrane formation, turnover, and intracellular protein trafficking.
To understand the distribution of Pio under different stress situations, we examined mutants that either increase the apical cell surface or cause severe chitin defects. We generated voxel data of 3D Pio staining projections (Imaris, 3D surface rendering) using Airyscan Z-stacks and deconvolution (SVI Huygens). These data identified Pio voxel in punctuate pattern at and next to the apical cell membrane in wt embryos. However, most voxel accumulated around the inner luminal chitin matrix structure (Figure 2C; Figure 2—figure supplement 2A), resembling the confocal pattern of control embryos (Figure 2A and B). The disruption of SJs in mega mutant embryos caused apical cell surface expansion and increased tube length in stage 16 embryos (Behr et al., 2003). These mega mutant embryos showed Pio localization at the chitin matrix but increased staining at and near the apical cell membrane (Figure 2C; Figure 2—figure supplement 2A). The ectopic expression of the chitinase 2 (cht2) in tracheal cells leads to excessive tube dilation due to strongly reduced luminal chitin (Behr et al., 2003; Petkau et al., 2012; Tonning et al., 2005). Also, the tracheal cht2 expression led to increased Pio staining at and near the apical cell membrane (Figure 2C; Figure 2—figure supplement 2A). Additional Z-stack analysis methods support such a shift of Pio localization. The ZEN co-localization tool and orthogonal projections across the tube lumen showed enriched Pio puncta at the membrane of mega mutant and Cht2 overexpressing tracheal cells compared with control (Figure 2D and E).
Further, the fluorescence intensity profile confirmed that fluorescence peaks of Pio antibody staining overlapped with peaks of Uif staining at the apical cell membrane and with the chitin-binding probe (cbp) at the luminal chitin cable (Figure 2—figure supplement 2B–D). The statistical analysis of the fluorescence intensity profiles of Pio, Uif, and cbp confirmed a significant difference between control, mega mutant, and Cht2 overexpression (Figure 2F). Together, these findings provide evidence that Pio staining is distributed in a punctuate pattern at the apical cell membrane in stage 16 embryos, which elongate tracheal tube length. Furthermore, the genetic modifications of the apical membrane and chitin matrix morphology affect the Pio pattern at the cell membrane.
Next, we investigated if Pio affects the tracheal airway function. First, orthogonal projections of confocal Z-stacks of late-stage 16 pio mutant embryos revealed that membrane deformation compromised the shape of the tube lumen, which includes deformations that protrude into cells, which constricted the lumen (Figure 1C and C’; Figure 1—figure supplement 1D). Second, the ultrastructure analysis of stage 17 pio mutant embryos revealed, in addition to the membrane protrusions, reduced chitin of taenidial folds at the aECM of dorsal trunks while control embryos established chitin-loaded taenidial folds at the apical cell surface (Figure 1D; Figure 1—figure supplement 2B). Also, confocal microscopy revealed reduced tracheal chitin staining in stage 16 pio null mutant embryos (see chitin stainings below). Third, pio mutant embryos showed an irregular pattern of taenidial folds in TEM and airyscan analysis, while control embryos showed a regular and narrower spacing of taenidial folds (Figures 1D and 3A; Figure 1—figure supplements 1D and 2B). Fourth, ultrastructure TEM images revealed aECM remnants in the airway lumen of pio mutant stage 17 embryos, while control embryos cleared their airways (Figure 1—figure supplement 2B). Consistently, the in vivo analysis of airways in stage 17 pio mutant embryos revealed lack of tracheal air-filling (Figure 3B). The pan-tracheal expression of Pio in pio mutant embryos rescued the lack of gas-filling (Figure 3B). Thus, TEM images suggest that pio mutant embryos showed impaired tube lumen clearance of aECM, which prevented subsequent airway gas-filling.

Pio is required for taenidial fold morphology and airway gas-filling.
(A) Airyscan images (left) and orthogonal (middle) and 3D projections (right) of control and pio mutant late-stage 16 embryos. cbp (in green and gray) detects chitin at the taenidial folds and within the tracheal lumen of control and pio mutant embryos. The pio mutant embryos show loose taenidial fold patterns with enlarged distances between the ridges. (B) Late-stage 17 wt embryos revealed normal tracheal air-filling (indicated with blue dashes). All pio (n=20) mutant embryos mutant embryos showed tracheal air-filling defects (red dashes indicate liquid filled airways), while almost none of the control embryos showed defects (n=100). Tracheal air-filling defects in embryos were displayed as dark gray bars, and normal air-filling as light gray bars. Error bars indicate the standard deviation. Heterozygous pio mutant embryos (n=10) as well as the btl-Gal4 driven tracheal expression of Pio in pio mutant embryos (n=10) revealed normal tracheal air-filling at the end of stage 17 (indicated with blue dashes). Green signal indicates eYFP expression from the balancer chromosome in the heterozygous pio mutant embryo. (C) The transheterozygous wurst;pio and wurst;pio double mutant stage 17 embryos showed tracheal air-filling defects (red dashes) in bright-field microscopy. Whole-mount embryos are shown left, framed parts of dorsal trunks are shown as close-ups right. Dorsal trunk tubes are indicated with red dashes. Confocal 3D projection of cbp (chitin, green) revealed collapsed and irregular tube lumen morphology in the mutant embryos. Scale bars indicate 10 µm. (D) Quantification of completed air-filling in late-stage 17 embryos. 89% of the transheterozygous (n=27) and all 11 wurst;pio double mutant embryos revealed incomplete air-filling whereas nearly all control embryos managed to complete air-filling. Error bars indicate the standard deviation.
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Figure 3—source data 1
Pattern of taenidial folds in stage 16 embryos.
- https://cdn.elifesciences.org/articles/91079/elife-91079-fig3-data1-v1.zip
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Figure 3—source data 2
Quantification of tracheal air-filling in stage 17 embryos.
- https://cdn.elifesciences.org/articles/91079/elife-91079-fig3-data2-v1.zip
Since Pio localization depends on Wurst, we addressed putative genetic interaction by investigating tracheal air-filling in stage 17 transheterozygous wurst and pio mutant embryos. Heterozygous wurst and pio mutant control embryos showed normal tracheal air-filling (Figure 3B; Behr et al., 2007). In contrast, 88.4% of transheterozygous embryos bearing one copy of the pio and one of the wurst mutant alleles showed air-filling defects, accompanied by lethality before larval hatching (Figure 3C and D), suggesting genetic interaction of pio and wurst mutants. Consistently, all wurst;pio double mutant embryos failed to complete tracheal air-filling and are lethal as late-stage 17 embryos (Figure 3C and D). Moreover, higher resolution images of the not-gas-filled dorsal trunks of wurst;pio transheterozygous and double mutant stage 17 embryos revealed compromised tube lumen morphology accompanied by dilatations in bright-field microscopy and confocal analysis of chitin staining (Figure 3C). The wurst mutant embryos failed to clear luminal matrix content (Behr et al., 2007), including Pio, thus preventing airway clearance at stage 17 (Figure 2—figure supplement 1D). Also, the pio mutant embryos showed tracheal lumen clearance defects of chitin fibers in ultrastructure (TEM) analysis (Figure 1D, Figure 1—figure supplement 2B). In contrast, confocal analysis revealed that well-known chitin matrix proteins, such as Obstructor-A (Obst-A) and Knickkopf (Knk), are removed from the lumen of pio mutants (Figure 3—figure supplement 1A). These results suggest that the Pio function did not affect airway clearance of Obst-A and Knk and therefore did not play a central role in airway clearance like Wurst. Nevertheless, airway clearance defects observed in TEM images in pio null mutant embryos and, in addition, defective tube lumen morphology in wurst;pio transheterozygous mutant embryos explain the occurrence of airway gas-filling defects.
The chitin matrix is required in tracheal tube size control, taenidial fold formation, and tube lumen stability (Behr and Riedel, 2020; Dong et al., 2014; Öztürk-Çolak et al., 2016; Tonning et al., 2005). Essential chitin matrix proteins, such as chitin-binding protein Obst-A, chitin deacetylases Serpentine (Serp), and Vermiform (Verm), as well as chitin-protein Knk restrict tube expansion (Luschnig et al., 2006; Moussian et al., 2006; Petkau et al., 2012; Wang et al., 2006). In the pio mutant stage 16 embryos, luminal Obst-A, Knk, Serp, and Verm antibody staining showed wt-like distribution, but Serp and Verm appeared in reduced levels in the lumen (Figure 3—figure supplement 2). Thus, Pio does not control tracheal chitin-matrix secretion, formation, and organization but may affect their maintenance in stage 16 embryos.
Apical cell membrane growth is another essential cellular mechanism of tube growth (Dong et al., 2014; Laprise et al., 2010). However, the apical cell membrane marker Uif showed wt-like localization in pio mutant trachea (Figure 1C). Further, Crb immunostainings showed normal localization in pio mutant embryos (Figure 4A). Also, actin and tubulin cytoskeleton, which may impact apical cell membrane and tracheal tube formation, did not show gross differences between control and pio mutant stage 16 embryos. The apical cell cortex enriched tubulin and F-actin staining (Figure 4—figure supplement 1). Further, the confocal Mega immunostainings and appearance of SJs in ultrastructure analysis were similar between control and pio mutant embryos (Figure 1B and D; Figure 1—figure supplement 2). Finally, adherens junctions (AJs) revealed a wt-like appearance in pio mutant embryos in ultrastructure images (Figure 1—figure supplement 2B), and also the Armadillo (Arm; Drosophila β-Catenin) immunostainings showed wt-like pattern at the apicolateral membrane of tracheal cells (Figure 4B). These data indicate that Pio is not involved in either apical polarity or lateral membrane formation, which is consistent with previous findings (Brodu et al., 2010). However, we observed an increased distance in the axial direction between the AJs of the dorsal trunk fusion cells. In addition, we determined an enlarged apical cell surface area due to unusual cell elongation in the axial direction (Figure 4B–D). These findings indicate that Pio is required to prevent excessive apical cell growth and membrane deformation.

Apical polarity and AJs localization in pio and Np mutant embryos.
Confocal LSM Z-stacks of tracheal dorsal trunk show single layer (A) and 3D projections (B, C) of stage 16 and 17 embryos. (A) Control and pio mutant late-stage 16 embryos show Crb (red) staining at the apical membrane, the cell surface marker WGA (blue), and cbp (green) in the tracheal apical extracellular matrix (aECM) at the apical cell surface and in the luminal cable-like ECM. A single Crb channel is indicated in gray. (B) Maximum intensity projections of confocal Z-stacks of control and pio mutant late-stage 16 embryos and Np mutant early stage 17 embryo. Upper panels show immunostainings with Armadillo (red) and Wurst (green) at the dorsal tracheal trunk. Yellow dashes mark the tracheal tube. Magnifications of the framed regions in the top panel show Armadillo staining in gray (bottom). Red double arrows indicate tracheal cells. Yellow arrows point to AJs of fusion cells. (C) Yellow double arrows point to magnifications of the Armadillo staining of dorsal trunk fusion cells. The distance of AJs of fusion cells in control, pio, and Np mutant embryos are indicated in representative images. Plots show AJ distances of fusion cells (n>9) in µm and apical cell area in µm2 (n>15). Bars represent ± SD and p-values for AJs distance (pio p=5.8e-5; Np p=0.0022) and cell area (pio p=1.6e-6, Np p=2,5e-10), unpaired t-test. (D) 3D reconstruction (Imaris surface rendering) of confocal Armadillo immunostainings marking the AJs of control, pio, and Np mutant embryos. Yellow arrows point to AJs of fusion cells; white double arrows indicate cell length in the axial direction.
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Figure 4—source data 1
Pattern of adherens junctions in dorsal trunk fusion cells in stage 16 pio mutant embryo and quantifications.
- https://cdn.elifesciences.org/articles/91079/elife-91079-fig4-data1-v1.zip
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Figure 4—source data 2
Pattern of adherens junctions in dorsal trunk fusion cells in stage 16 Np mutant embryos.
- https://cdn.elifesciences.org/articles/91079/elife-91079-fig4-data2-v1.zip
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Figure 4—source data 3
Quantification of pattern of adherens junctions in dorsal trunk fusion cells.
- https://cdn.elifesciences.org/articles/91079/elife-91079-fig4-data3-v1.zip
To understand Pio dynamics in living embryos, we generated CRISPR/Cas9-mediated homology-directed repair piomCherry::pio embryos (Figure 3—figure supplement 1B). The mCherry::Pio expression in stage 16 embryos (Figure 3—figure supplement 1C) resembled Pio protein expression pattern in the tracheal system, digestive organs, epidermis, and spiracles (Figure 2A). Early stage 16 embryo dorsal trunk cells showed apical enrichment of mCherry::Pio (Figure 5A). Airyscan images and fluorescence intensity profiles of Z-stacks revealed a punctuate pattern of mCherry::Pio overlapping with chitin in the chitin cable and at the apical cell membrane surface where taenidial folds form in stage 16 embryos (Figure 5—figure supplement 1A–C). When tube expansions stopped at the end of stage 16, the tracheal mCherry::Pio signal shifted toward the lumen. This mCherry::Pio signal shift is distinct from the chitin matrix pattern (Figure 5A; Figure 5—video 1). Fluorescence recovery after photobleaching (FRAP) experiments demonstrated recovery of mCherry::Pio expression after photobleaching (Figure 5B and B’; Figure 5—video 2). In stage 16 embryos, mCherry::Pio puncta reappeared in tracheal cells within 2 min of bleaching and in the tubular lumen within 6 min. This demonstrates the dynamic mCherry::Pio relocation in tracheal cells and the lumen during tube expansion.

Pio is dynamically localized at the membrane and controls Dumpy secretion.
(A) Confocal LSM images of dorsal trunks of embryos with endogenous mCherry:: Pio expression stained with anti-mCherry antibody (magenta) and cbp (chitin; green) at indicated embryonic stages. In early stage 16 embryos, mCherry:: Pio enriches apically (arrowheads) and is present in the lumen (arrow). In contrast, at the end of stage 16 mCherry::Pio predominantly localizes within the tracheal lumen (arrows). The luminal mCherry:: Pio staining disappeared during stage 17. (B, B’) Confocal images of a representative fluorescence recovery after photobleaching (FRAP) experiment (n=4) in a live embryo with endogenous expression of Dpy::eYFP and mCherry::Pio and quantification of normalized fluorescence in the bleached area of n=4 embryos (right) are shown. Yellow frames indicate the bleached area. Close-ups (right-most images) show details of the bleached area (below arrows in header). The dashed line indicates apical cell membranes, red arrows indicate luminal mCherry::Pio, and red arrowheads indicate intracellular or membrane-associated mCherry::Pio. A representative movie of an FRAP experiment is presented in Movie S2. Fluorescence intensities refer to the bleached regions of interest (ROIs) as indicated with the frame in corresponding Movie S2 and was measured after correction for embryonic movements. The mCherry::Pio (magenta) reveals recovery of small Pio puncta in the bleached area including the tracheal lumen, while Dpy::eYFP (green) shows no recovery even after 56 min. Scale bars indicate 5 µm in overview panels and 2 µm in bleach close-ups. (C) Confocal LSM images of endogenous expression of Dumpy:eYFP stained with anti-GFP antibody. The wt-like stage 16 control embryos show extracellular Dumpy:eYFP (magenta) in the apical extracellular matrix (aECM) at the cell surface and in the luminal cable (arrow) overlapping with cbp (chitin; green). In contrast, in pio mutant embryos Dumpy::eYFP did not overlap with chitin (cbp, green), but remained intracellularly (arrowhead). Upper rows focus on the dorsal trunk, lower rows show 3D maximum intensity projections of whole tracheal segments. Note the dorsal branch disruption known from hypomorphic pio point mutation allele (Jaźwińska et al., 2003). Single channels are indicated in gray. Db, dorsal branch; Dt, dorsal trunk; Tc, transverse connective. Scale bars indicate 10 µm. (D) Immunoblotting of co-immunoprecipitation (Co-IP) assay of RFP-tagged Dpy constructs and Strep-tagged Pio expressed in Drosophila S2R+ (Schneider) cells reveals binding of Dpy and Pio. Schemata of expressed proteins used in the assay are shown on the right. Strep::Pio is the full-length Pio protein with a Twin-Strep tag inserted C-terminal to the signal peptide (light blue). RFP::DpyZP and RFP::DpyCT both contain the endogenous Dpy signal peptide (light blue) followed by mCherry (RFP) and different length of the C-terminal region of the Dpy isoform A protein as indicated. Transmembrane (TM) domains (yellow), ZP domains (blue), and Furin and Np protease cleavage sites (pcs) in Pio are indicated. Western blots of input cell lysates (left) and anti-Strep IP elutions (right) stained with anti-Strep (top) and anti-RFP (bottom) antibodies are shown. Both RFP::Dpy proteins are only detectable in IP elutions when they were co-expressed with Strep::Pio.
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Figure 5—source data 1
Confocal Z-stack images of dorsal trunk showing mCherry::Pio expression during stages 16 and 17 of embryogenesis.
- https://cdn.elifesciences.org/articles/91079/elife-91079-fig5-data1-v1.zip
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Figure 5—source data 2
Confocal Z-stack images of dorsal trunk showing mCherry::Pio expression during late stages 16.
- https://cdn.elifesciences.org/articles/91079/elife-91079-fig5-data2-v1.zip
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Figure 5—source data 3
Confocal Z-stack images of dorsal trunk showing mCherry::Pio expression during early stage 17.
- https://cdn.elifesciences.org/articles/91079/elife-91079-fig5-data3-v1.zip
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Figure 5—source data 4
Confocal Z-stack images of dorsal trunk showing mCherry::Pio expression during mid stage 17.
- https://cdn.elifesciences.org/articles/91079/elife-91079-fig5-data4-v1.zip
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Figure 5—source data 5
Confocal Z-stack images of dorsal trunk showing Dpy::YFP expression in control and pio mutant embryos.
- https://cdn.elifesciences.org/articles/91079/elife-91079-fig5-data5-v1.zip
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Figure 5—source data 6
Confocal Z-stack images of dorsal trunk showing Dpy::YFP expression in pio mutant embryos.
- https://cdn.elifesciences.org/articles/91079/elife-91079-fig5-data6-v1.zip
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Figure 5—source data 7
Uncropped western blots.
- https://cdn.elifesciences.org/articles/91079/elife-91079-fig5-data7-v1.pdf
Pio binds Dumpy to organize the luminal ZP protein matrix
Dumpy (Dpy) is a giant (3.2 mDa) and stretchable ZP domain protein. In stage 16 embryos Dpy::eYFP (Lye et al., 2014) appears at the tracheal apical cell surface and predominantly within the lumen (Figure 5C; Dong et al., 2014; Jaźwińska et al., 2003). Airyscan images and fluorescence intensity profiles of Z-stacks in stage 16 embryos revealed predominant Dpy::eYFP staining, which overlapped with chitin in the chitin cable. In addition, we observed a punctate Dpy::eYFP pattern at the apical cell surface overlapping with taenidial chitin and mCherry::Pio (Figure 5—figure supplement 1D and E). In contrast, the Dpy::eYFP signal predominantly remained intracellularly in pio mutants (Figure 5C), showing that Dpy secretion depends on Pio. We also performed cell culture experiments to extend our analysis of Dpy secretion. We generated constructs of RFP-tagged Dpy that lacked a portion of EGF and DPY repeats but contained the essential Dpy C-terminal region (ZPD domain, transmembrane domain, cytoplasmic region) (Figure 5D). Only the co-expression of RFP-tagged Dpy with FLAG-tagged Pio resulted in extracellular RFP::Dpy localization in the S2R+ cells (Figure 5—figure supplement 2A). Since extracellular RFP::Dpy is not released from the cells but overlaps with FLAG::Pio at the membrane, it suggests that they co-localize at the cell surface. The S2R+ cells expression products of RFP::Dpy constructs were only pulled down together with Strep::Pio in Strep-IP samples (Figure 5D). These data demonstrate that Pio co-localizes and interacts with Dpy. In contrast to Pio, tracheal Dpy::YFP was immobile in our FRAP experiments (Figure 5B and B’; Figure 5—video 2), supporting findings that the exchange of Dpy is negligible (Dong et al., 2014). Importantly, the current model suggests that the tracheal Dpy matrix stretches very likely by force applied from the cells during tube expansion and thus must be connected to the epithelium (Dong et al., 2014). Our FRAP data suggest that Pio is dynamic at the tracheal ZP matrix, while the static and stretchable Dpy modulates mechanical tension within the matrix, as discussed for Dpy previously (Dong et al., 2014; Wilkin et al., 2000).
Embryos of the lethal dpyolvR allele showed pio2R-16 mutant-like tracheal branch disintegration (Jaźwińska et al., 2003) and additionally twisted and broken dorsal trunks in late-stage 17 embryos (Bökel et al., 2005). Also, previously shown quantification revealed a significantly increased dorsal trunk length in dpyolvR mutant embryos (Dong et al., 2014). These findings suggest that also dpy mutation may impact tube expansion. Chitin staining of dpyolvR mutants showed a sinusoidal appearance of tracheal dorsal trunk tubes and branch disintegration in stage 16 embryos (Figure 5—figure supplement 3A). The higher resolution airyscan images of Uif stainings at the dorsal trunks and three-dimensional projections revealed blister-like apical membrane deformations but less frequent as in the pio mutant embryos (Figure 5—figure supplement 3B). The orthogonal projections of such blisters in dpy mutant dorsal trunks revealed Uif-marked membrane protrusion into tracheal cells similar to pio-mutant embryos (Figure 5—figure supplement 3C). Additionally, we observed enriched puncta of Pio staining in the Uif-marked membrane blisters (Figure 5—figure supplement 3C). The subcellular localization studies in stage 16 dpyolvR mutant embryos revealed a punctuate Pio pattern at the apical cell membrane overlapping with Uif but not within the chitin cable (Figure 5—figure supplement 3D and E). These findings suggest that Dpy is involved in Pio release at the apical cell surface and in apical cell membrane stability during tube length elongation.
Pio release involves the serine protease Notopleural at the apical cell membrane
Furin-like enzymes cleave ZP precursors (Jovine et al., 2005). Pio contains a Furin proteolytic cleavage site (Furin pcs) followed by a C-terminal transmembrane domain. Surprisingly, we detected in stage 16 embryo lysate three mCherry::Pio variants, one correlating with the predicted mass of a full-length protein (115 kDa), a second after furin site cleavage (90 kDa), and a third one correlating with the size after proteolysis within the ZP domain (60 kDa) (Figure 6A). The latter was even the predominant variant in stage 17 (Figure 6A). In contrast, the Np mutant embryo lysates of stages 16 and 17 contained only faint amounts of the 60 kDa mCherry::Pio and an increased level of the small variant was not apparent in stage 17 embryos (Figure 6A). These findings support our recent study suggesting that Np cleaves the Pio ZP domain in vitro and in vivo (Drees et al., 2019). Co-expression of the catalytically inactive NpS990A with mCherry::Pio in Drosophila Schneider cells showed as a prominent signal the 90 kDa mCherry::Pio variant in the cell lysate (Figure 6B), and live imaging revealed mCherry::Pio localization at the cell surface (Figure 5—figure supplement 2B). This was comparable with control cells that expressed mCherry::Pio alone (Figure 6B, Figure 5—figure supplement 2A). In contrast, cells that co-expressed either the functional Np or its homolog, the human Matriptase, revealed as a predominant signal the 60 kDa mCherry::Pio variant in the supernatant fraction (Figure 6B) and no mCherry::Pio localization at the cell surface (Figure 5—figure supplement 2B). These results demonstrate that the enzymatic activity of the serine protease Np is sufficient for ZP domain cleavage, which results in the ectodomain shedding of Pio in Schneider cells.

Np-mediated ZP domain shedding controls Pio dynamics at the apical cell membrane.
(A) Left: Immunoblot of protein lysates from embryos of stages 16 and 17 stained with anti-mCherry antibody show three specific bands in samples from mCherry::Pio expressing embryos. Middle: Schematic presentation of the mCherry::Pio fusion protein with signal peptide (light blue), mCherry (red), ZP domain (blue), and transmembrane domain (yellow). Furin and Np protease cleavage sites (pcs) and expected molecular weights of resulting fragments are indicated. Right: Proportional intensity of the Np-cleaved 60 kDa mCherry::Pio fragment to the whole signal from all three mCherry::Pio fragments normalized to β-tubulin intensity. Data from three biological replicates show that proteolytic processing of mCherry::Pio at the Np cleavage site is highly increased in stage 17 embryos compared to stage 16 embryos in wt genetic background. This difference is not detectable in samples from Np mutant embryos. The endogenous Pio protein has a calculated mass of about 50.82 kDa. (B) Cleavage assay within the Pio ZP domain is mediated by proteolytic activity of Np and the human Matriptase. Immunoblotting of cell lysates and supernatant precipitates from Drosophila S2R+ cells expressing mCherry::Pio alone or together with Np, catalytically inactive NpS990A or human Matriptase with anti-mCherry antibody. Np and human Matriptase cleave mCherry::pio, causing shedding of the mCherry::Pio extracellular domain and a substantial increase of the 60 kDa mCherry::Pio, which correlates in size with cleavage at the ZP domain, in cell culture supernatants. This effect is not observable for catalytically inactive NpS990A. (C) Confocal images of a representative fluorescence recovery after photobleaching (FRAP) experiment in a live Np mutant embryo with endogenous expression of Dpy::eYFP and mCherry::Pio and quantification of normalized fluorescence in the bleached area (yellow frames) of n=4 embryos (right) are shown. Close-ups (right-most images) show details of the bleached area (below arrows in header). The dashed line indicates apical cell membranes, arrowheads indicate intracellular or membrane-associated mCherry::Pio. Arrowhead points mCherry::Pio at the apical cell surface in the untreated area. Representative movie of the FRAP experiments is presented in Figure 6—video 1 (Np mutant), compare with Figure 5—video 2 (wt). The fast recovery of small mCherry::Pio puncta in the tracheal lumen is impeded in Np mutant embryos (compare with wt in Figure 4D). As in wt embryos, Dpy::eYFP (green) shows no recovery even after 56 min. Scale bars indicate 5 µm in overview panels and 2 µm in bleach close-ups. (D) Confocal images of tracheal dorsal trunks of Np mutant embryos with endogenous expression of mCherry::Pio at indicated developmental stages stained with cbp (chitin; green) and anti-mCherry antibody (magenta). Single channels are indicated in gray. Stage 16 Np mutant embryos show intracellular mCherry::Pio at the apical cell surface (arrowhead), which is similar to control embryos (see Figure 4A). In contrast to control embryos, the luminal mCherry::Pio (arrow) is strongly reduced in stage 16 and 17 Np mutant embryos, while the non-luminal mCherry::Pio accumulates in stage 17. The luminal chitin cable is degraded normally in Np mutant embryos but does not condense (Drees et al., 2019) during early stage 17 and instead, remains attached to the tracheal cell surface and fills the whole lumen during degradation (compare with control in Figure 5A). The asterisks mark bulges in Np mutant tubes. Note the two layers of chitin visible at the membrane bulges and the adjacent apical extracellular matrix (aECM) indicating disintegration of the tracheal chitinous aECM (see also Figure 6). Scale bars indicate 10 µm. (E) The tracheal trypsin-like S1A Serine transmembrane protease Filzig (Flz) shows high sequence homology to Np (Drees et al., 2019) and acts in processing the lumen matrix (Rosa et al., 2018). Confocal images of dorsal trunks of flz mutant embryos with endogenous expression of mCherry::Pio at indicated developmental stages stained with cbp (chitin; green) and anti-mCherry antibody (magenta). Single channels are indicated in gray. The flz mutant embryos revealed normal Pio expression, luminal shedding, and clearance from airways (compare with wt in Figure 4A). In contrast to Np mutant embryos, the luminal aECM cable condensed during early stage 17 luminal clearance (arrows) as in wt embryos (compare with Np mutant in D and wt in Figure 4A and A). Scale bars indicate 10 µm.
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Figure 6—source data 1
Confocal Z-stack images of dorsal trunk showing mCherry::Pio expression in Np mutant embryos during stages 16 and 17.
- https://cdn.elifesciences.org/articles/91079/elife-91079-fig6-data1-v1.zip
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Figure 6—source data 2
Confocal Z-stack images of dorsal trunk showing mCherry::Pio expression in Np mutant late stage 16 embryo.
- https://cdn.elifesciences.org/articles/91079/elife-91079-fig6-data2-v1.zip
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Figure 6—source data 3
Confocal Z-stack images of dorsal trunk showing mCherry::Pio expression in Np mutant early stage 17 embryo.
- https://cdn.elifesciences.org/articles/91079/elife-91079-fig6-data3-v1.zip
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Figure 6—source data 4
Confocal Z-stack images of dorsal trunk showing mCherry::Pio expression in Np mutant mid stage 17 embryo.
- https://cdn.elifesciences.org/articles/91079/elife-91079-fig6-data4-v1.zip
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Figure 6—source data 5
Confocal Z-stack images of dorsal trunk showing mCherry::Pio expression in flz mutant embryos during stages 16 and 17.
- https://cdn.elifesciences.org/articles/91079/elife-91079-fig6-data5-v1.zip
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Figure 6—source data 6
Confocal Z-stack images of dorsal trunk showing mCherry::Pio expression in flz mutant stage 17 embryo.
- https://cdn.elifesciences.org/articles/91079/elife-91079-fig6-data6-v1.zip
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Figure 6—source data 7
Uncropped western blots (Figure 6A).
- https://cdn.elifesciences.org/articles/91079/elife-91079-fig6-data7-v1.pdf
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Figure 6—source data 8
Uncropped western blots (Figure 6B).
- https://cdn.elifesciences.org/articles/91079/elife-91079-fig6-data8-v1.pdf
Next, we investigated the consequences of NP-mediated ZP cleavage. FRAP experiments showed only minor intracellular recovery of mCherry::Pio in Np null mutant embryos (Figure 6—video 1). In contrast to the control, extracellular mCherry::Pio is not released into the tube lumen within 56 min after bleaching in Np mutant embryos (Figure 6C, Figure 6—video 1). The luminal mCherry::Pio immobility was comparable to the immobile Dpy::eYFP fraction in control and Np mutant embryos, suggesting that Pio is hampered to diffuse in Np mutant embryos. Second, mCherry::Pio was not released into the tracheal lumen in late-stage 16 Np mutant embryo stainings (Figure 6D, Figure 6—video 2). Third, the tracheal trypsin-like S1A Serine transmembrane protease Filzig had no influence on the mCherry::Pio pattern (Figure 6E, Figure 6—figure supplement 1). The flz mutant (Figure 6—figure supplement 1) and Np mutants stage 17 embryos showed no Dpy clearance from the tracheal lumen (Drees et al., 2019). However, the flz mutation did not affect the release of Pio at the apical cell surface (Figure 6E). Further, the luminal aECM cable condensed during early stage 17 in flz mutant embryos as observed in wt (compare Figure 5A with Figure 6E), indicating that these processes require Np. In summary, our findings prove the inhibition of luminal Pio release in Np mutant embryos. Furthermore, the Np-mediated proteolysis of the Pio ZP domain is specific and plays a central role in Pio dynamics at and near the apical cell membrane during tube expansion.
To further analyze the Np function, we used Uif to examine the tracheal apical cell membrane structure. This revealed unusual bulge-like membrane deformations in late-stage 16 and 17 Np mutant embryos, while control embryos did not show such bulges (Figure 7A and B). Confocal time-lapse series revealed that bulge-like deformations emerged during early stage 16 embryos and grew in size (Figure 7C). Furthermore, the increasing size of the membrane bulges led to the detachment of α-tubulin::GFP marked cells from the aECM at the cell surface (Figure 7C). The control embryos did not show such a separation between cells and adjacent aECM (Figure 1B). However, confocal images of Uif and chitin show residual chitin at the Uif marked membrane and chitin at the detached aECM (Figure 7A). These results indicate the tearing of the tracheal aECM at the apical cell surface in Np mutant embryos.

Np supports structural cell membrane integrity.
(A) Bulge-like tracheal apical membrane deformations appeared in Np mutant embryos as stable structures that grow during late stage 16. Confocal images of dorsal trunks of wt embryos and NpP6/NpC2 embryos stained with cbp (chitin; green) and anti-Uif antibody (magenta) as a marker for apical tracheal membranes. (B) Confocal Z-stack projections of cbp staining of stage 17 Np mutant shows several bulges (arrows) at the dorsal trunk. Quantification of bulges per dorsal trunk (n=16) in Np mutants is shown right. (C) The in vivo time-lapse series of 130 min show bulges arising at Np mutant embryos' dorsal trunk cell membranes. Tracheal cells express Tubulin::GFP in transheterozygous NpP6;NpC2 mutant embryos. Frames 1 and 2 are shown as close-ups (below) of forming bulges. The cell membrane of GFP-expressing cells and parts of the misorganized tracheal cuticle apical extracellular matrix (aECM) (dashed line) separate, as shown in the time-lapse. Yellow arrows point to membrane deformations; red arrowheads point to the tracheal cuticle at the apical cell surface, and red arrows to the luminal aECM cable. Note that chitin is detectable in two layers at the sites of bulges, while Uif is detectable only at the bulges, indicating a disintegration of the tracheal chitinous aECM (A). Scale bars are 10 µm (overviews) and 3.5 µm (details). Note that bulges grew as time progressed (up to 130 min).
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Figure 7—source data 1
Quantification and confocal Z-stack images of bulges at dorsal trunks of Np mutant embryos.
- https://cdn.elifesciences.org/articles/91079/elife-91079-fig7-data1-v1.zip
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Figure 7—source data 2
Quantification of bulges at dorsal trunks of Np mutant embryos.
- https://cdn.elifesciences.org/articles/91079/elife-91079-fig7-data2-v1.zip
Finally, our time-lapse series and the analysis of stage 17 embryos proved that the bulge-like cell membrane structures remained in Np mutant embryos. Moreover, these cell membrane bulges destabilize the normal epithelial barrier. Upon tracheal expression of myr-RFP in Np mutants, we detected accumulation of RFP signal first at the membrane bulge and subsequently in the tube lumen (Figure 7—figure supplement 1). The Np and pio mutant embryos show apical membrane deformations (compare Figures 1A and 7A). Np is located at the membrane (Drees et al., 2019), where it controls Pio release into the tracheal lumen. This suggests that membrane deformations in Np mutant embryos are caused by impaired function of the Pio-mediated ZP matrix due to a lack of Pio shedding.
Any imbalance between membrane and matrix during tube expansion causes tube deformations. Chitin staining was reduced and revealed sinusoidal over-elongated tubes in stage 16 pio mutant embryos (Figure 8A and B; Figure 1—figure supplement 1B and C), proving that pio function prevents tracheal tubes from over-elongation. In Np mutant stage 16 embryos, Pio was not shed into the lumen but remained at the cell, while Dumpy showed normal and immobile localization within the tube lumen (Figure 6C and D; Figure 6—videos 1; 2). This indicates that the Pio-mediated ZP matrix can still restrict tube expansion of the membrane in Np mutants independently from membrane deformations. The Np mutant stage 16 embryos did not show tube overexpansion (Figure 8A and B). Further, Np,pio double mutants did not exacerbate the pio mutant tube length defects suggesting that both act in the same genetic pathway (Figure 8A and B). Our data assumes that Np overexpression may enhance Pio shedding in stage 16 embryos, affecting the Pio-mediated ZP matrix function. Upon breathless (btl)-Gal4-mediated expression of UAS-Np in tracheal cells, we observed a high amount of Pio puncta across the entire tracheal tube lumen, specifically in stage 16 embryos but not in earlier stages (Figure 8—figure supplement 1). Consistently tracheal Np overexpression led to tube overexpansion in stage 16 embryos resembling the pio mutant phenotype (Figure 8A and B). Thus, Np-mediated Pio shedding controls Pio function.

Pio and Np control tube size and their regulatory mechanisms of ZP domain shedding is conserved.
(A) Confocal Z-stack projections of whole-mount stage 16 embryos stained with cbp (chitin) focusing on the tracheal system and close-ups at the right. In contrast to straight branches of control embryos, pio17C null mutant embryos revealed curly elongated tubes indicating excess tube expansion. Note dorsal branch disruption known from a hypomorphic pio point mutation (Jaźwińska et al., 2003). Np mutant embryos show straight wt-like tubes. Embryos that overexpress Np in the tracheal system (btl-Gal4>UAS-Np) show curly elongated tubes and dorsal branch disruption, phenocopying the pio mutant phenotype. Np;pio double mutant embryos do not exacerbate pio mutant tube size defects and show a similar phenotype as pio mutant embryos, respectively. (B) Quantification of normalized dorsal trunk length from 9 stage 16 embryos of each genotype. Anterior-posterior dorsal trunk length was divided by the anterior-posterior length of the embryo. Normalized dorsal trunk length in pio mutant embryos, btl-Gal4>UAS-Np embryos and Np,pio double mutant embryos is significantly increased when compared with wt (p=0.00013, p=0.00007, p=0.00019). Notably, the Np mutant dorsal trunk is relatively straight, while control embryos show slightly convoluted tubes. Also, statistical analysis reveals the tendency of slightly shortened dorsal trunk length in Np mutant. Individual points represent the respective embryos. (C) Human TGFβ type III receptor (TGFβRIII) is a widely expressed ZP domain containing protein. Human TGFβRIII with a cytoplasmic GFP tag and an extracellular Strep tag was expressed in Drosophila S2R+ cells either alone or together with Drosophila Np or human Matriptase. A schema of the tagged TGFβRIII is shown (top). The ZP and transmembrane (yellow) domains, the N-terminal Strep tag (magenta), C-terminal GFP (green) Furin protein cleavage site, (V) and the signal peptide (blue) are indicated. Images display maximum intensity projections of confocal Z-stacks. Shown are S2R+ cells that expressed the Strep::TGFβRIII::GFP construct alone or together with Np or human Matriptase stained with DAPI (blue) anti-GFP (green) and anti-Strep (magenta) antibodies. Single channel panels are indicated in gray. Control cells contain co-localizing GFP and Strep signals. The co-expression of Np or Matriptase reveals strong GFP but faint Strep signals due to extracellular cleavage and shedding of the TGFβRIII ectodomains. Framed boxes in overview images display details in panels on the right side. Scale bars indicate 10 µm.
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Figure 8—source data 1
Quantification of dorsal trunks lengths.
- https://cdn.elifesciences.org/articles/91079/elife-91079-fig8-data1-v1.zip
Since our findings show that Np controls tracheal Pio function by ZP domain cleavage, we addressed whether this is also a putative mechanism of human ZP domain proteins. The type III transforming growth factor-β receptor (TβRIII) acts as a signaling modifier and co-receptor of TGFβ and contains a ZP domain (López-Casillas et al., 1991; Moustakas et al., 1993). TβRIII ectodomain shedding in lung cancer cell models induces epithelial-to-mesenchymal transition and promotes the growth of tumors (Huang et al., 2019). For an in vitro cleavage assay, we co-expressed GFP- and Strep-tagged combined variants of TβRIII together with either human Matriptase or Np in Drosophila cells. The C-terminal GFP tag detects the intracellular part of TβRIII. The N-terminal Strep-tag follows the extracellular ZP domain and recognizes the ectodomain. Cells expressing TGFβRIII without Matriptase or Np revealed strong GFP and Strep co-localization signals. Co-expression of either human Matriptase or Np caused a substantial reduction of the extracellular Strep signal. In contrast, the intracellular GFP signal remained (Figure 8C). This shows that matriptases catalyze the extracellular proteolysis and the extracellular localization of the human TβRIII ZP domain protein. This indicates a new and unexpected conserved mechanism capable of controlling TβRIII function.
Discussion
Tracheal tube lumen expansion requires mechanical stress regulation at apical cell membranes and attached aECM. This involves the proteolytic processing of proteins that set local membrane-matrix linkages. Thus, the membrane microenvironment exhibits critical roles in regulating tube and network functionality.
ZP domain proteins organize protective aECM in the kidney, tectorial inner ear, and ZP (Jovine et al., 2005; Litscher and Wassarman, 2020), as well as in Drosophila epidermis, tendon cells, and appendages (Bökel et al., 2005; Plaza et al., 2010; Ray et al., 2015). Drosophila ZP domain proteins link the aECM to actin and polarity complexes in epithelial cells (Fernandes et al., 2010). Dumpy establishes force-resistant filaments for anchoring tendon cells to the pupal cuticle (Chu and Hayashi, 2021).
We identify that ZP protein-mediated microenvironmental changes increase the flexibility of membrane-matrix association, resulting from the activity of ZP domain proteins (Figures 1 and 7). Shear stress stimulates the activity of membrane-anchored proteases (Kang et al., 2015) and potentially also Np since we did not observe the misdistribution of the tracheal cytoskeleton when blisters arise. The dynamic membrane-matrix association control is based on our findings that loss of Np prevents Pio ectodomain shedding at the apical cell membrane resulting in immobile localization of Pio at the membrane and Dpy localization within the matrix (Figure 6). Direct interaction and overlapping subcellular localization at the cell surface showed that both proteins form a ZP matrix that potentially attaches membrane and ECM (Figure 5; Figure 5—figure supplement 1). Deregulation of Pio shedding blocks ZP matrix rearrangement and release of membrane-matrix linkages under tube expansion and subsequent shear stress. This destabilizes the microenvironment of membranes, causing blister formation at the membrane due to ongoing membrane expansion (Figure 7). Additionally, Pio could be part of a force-sensing signal transduction system destabilizing the membrane and matrix. Our observation that the membrane deformations are maintained in Np mutant embryos supports our postulated Np function to redistribute and deregulate membrane-matrix associations in stage 16 embryos when tracheal tube length expands. In contrast, Np overexpression potentially uncouples the Pio-Dpy ZP matrix membrane linkages resulting very likely in unbalanced forces causing sinusoidal tubes (Figure 8).
The membrane defects observed in both Pio and Np mutants indicate errors in the coupling of the membrane matrix due to the involvement of Pio (Figures 1 and 7). In pio mutants, gaps appear between the deformed membrane and the apical matrix (Figure 1B–D). These changes in apical cell membrane shape are consistent with increased cell and tube elongation in pio mutant embryos because the matrix is uncoupled from the membrane in such mutants (see model Figure 9). In contrast to pio mutants, the large membrane bulges in Np mutants affect the membrane and the apical matrix (Figure 7). Since apical Pio is not cleaved in Np mutants (Figure 6D), the matrix is not uncoupled from the membrane as in pio mutant embryos but is likely more intensely coupled, which leads to tearing of the matrix axially along the membrane bulges (Figures 7 and 9; Figure 7—figure supplement 1), when the tube expands in length. If apical Pio detachment reduces coupling between the matrix and apical membrane, then it is likely that Np mutant embryos may exhibit a reduced tube length phenotype. In Np mutant embryos, average tracheal dorsal trunk length tends to be reduced compared to wt embryos (Figure 8B), suggesting that Pio shedding is critical in controlling tracheal tube lumen length.

Model of apical Pio and Dpy matrix at the apical cell surface and Pio proteolysis and release.
(A) The simplified model shows a Pio-mediated Dpy secretion, the apical ZP domain protein matrix formation, and subsequent Pio ectodomain shedding by Np at wt tracheal cells. In pio null mutant embryos, Dpy is not secreted. The loss of the apical ZP proteins led to membrane matrix disconnection. This results in unstable membrane structures with numerous unusual gaps between membrane and matrix, excess apical cell shape expansion (indicated by the orange double arrow) in the axial direction, and dorsal trunk overexpansion. In contrast, apical ZP domain protein matrix forms in Np mutant embryos, but apical Pio ectodomain shedding is prevented. This led to membrane bulging and rupture (indicated by green arrows) of the apical ECM at bulges and slightly shortened dorsal trunks. (B) Simplified illustration of late embryonic tracheal development. The ZP proteins stabilize the apical membrane-matrix during tube expansion, subsequent taenidial fold formation, and airway clearance. Np cleaves Pio ZP domain at the membrane during tube expansion and sheds luminal Dpy during airway protein clearance. Wurst enables endocytosis at the apical membrane. This altogether enables precise tube length expansion (stage 16), lumen stability and airway gas-filling (stage 17). Basal (stage 16) and high-level (stage 17) endocytosis at the apical cell membrane have been described recently by Tsarouhas et al., 2019.
The btl-Gal4-driven Np expression mimics the endogenous Np from stage 11 onward in all tracheal cells throughout embryogenesis (Drees et al., 2019), suggesting that Np is not expressed at a wrong time point. However, the ratio between Np and Pio is essential. We assume that tracheal Np overexpression increases Pio shedding in stage 16 embryos (Figure 8—figure supplement 1), resulting in a pio loss-of-function-like phenotype. Thus, the tube length overexpansion upon Np overexpression indicates that Pio cleavage is required for tube length control.
Is Pio ectodomain shedding in response to tension? We did not measure tension directly. However, the developmental profile of mechanical tension during tracheal tube length elongation in stage 16 embryos (Dong et al., 2014) is consistent with the profile of Pio shedding. Np cleaves apical Pio during stage 16 when tube length expands (Figures 5 and 6). In contrast, Pio shedding decreases sharply at early stage 17 when tube elongation is completed (Figures 5A and 6A). Our model, therefore, predicts that loss of Pio or increased Pio secretion at stage 16 may reduce the coupling of the membrane matrix so that increased tracheal tube elongation is maintained until the end of stage 16, which is found in pio mutants and upon Np overexpression (Figure 8A and B). Unknown proteases may likely be involved in Pio processing since cleaved mCherry::Pio is also detectable in inactive NpS990A cells. Previously we identified a mutation at the Pio ZP domain (R196A) resistant to NP cleavage in cell culture experiments (Drees et al., 2019). Establishing a corresponding mutant fly line would be essential in determining whether the observed phenotype resembles the phenotype of the Np mutant embryos. In addition, unknown mechanisms, such as distinct membrane connections during development and emerging links to the developing cuticle, may also influence tension at the apical membrane during tube length control.
Indeed, the anti-Pio antibody, which detects all different Pio variants, showed a punctuate Pio pattern overlapping with the apical cell membrane markers Crb and Uif at the dorsal trunk cells of stage 16 embryos (Figure 2; Figure 2—figure supplement 1, Figure 2—figure supplement 2). Additionally, Pio antibody also revealed early tracheal expression from embryonic stage 11 onward, and due to Pio function in narrow dorsal and ventral branches, strong luminal Pio antibody staining is detectable from early stage 14 until stage 17, when airway protein clearance removes luminal contents. In the pio5m and pio17c mutants, Pio stainings were strongly reduced although some puncta were still detectable in the trachea (Figure 1—figure supplement 1G and H). Similarly, Pio antibody staining is intracellular in the trachea of stage 11 pio2R-16 point mutation embryos (Jaźwińska et al., 2003). Interestingly, also dpy mutants showed strongly reduced and intracellular Pio antibody staining (Figure 5—figure supplement 3E).
We generated mCherry::Pio as a tool for in vivo Pio expression and localization pattern analysis during tube lumen length expansion. The mCherry::Pio resembled the Pio antibody expression pattern from early tracheal development onward. However, luminal mCherry::Pio enrichment occurs specifically during stage 16, when tubes expand. The stage 16 embryos showed mCherry::Pio puncta accumulating apically in dorsal trunk cells. Moreover, mCherry::Pio puncta partially overlapped with Dpy::YFP and chitin at the taenidial folds, forming at apical cell membranes. Supported by several observations, such as antibody staining, video monitoring, FRAP experiments, and western blot studies (Figure 5; Figure 8—figure supplement 1; Figure 5—videos 1 and 2; Figure 6—videos 1; 2), these findings indicate that Pio may play a significant role at the apical cell membrane and matrix in dorsal trunk cells of stage 16 embryos.
Furthermore, we show that Np mediates Pio ZP domain cleavage for luminal release of the short Pio variant during ongoing tube length expansion. The luminal cleaved mCherry::Pio is enriched at the end of stage 16 and finally internalized by the subsequent airway clearance process during stage 17 after tube length expansion (Figures 5 and 6). Such rapid luminal Pio internalization is consistent with a sharp pulse of endocytosis rapidly internalizing the luminal contents during stage 17 (Tsarouhas et al., 2007). Wurst is required to mediate the internalization of proteins in the airways (Behr et al., 2007; Stümpges and Behr, 2011). In consistence, during stage 17, luminal Pio antibody staining fades in control embryos but not in Wurst deficient embryos.
Nevertheless, Pio and its endocytosis depend on its interaction with the chitin matrix and the Np-mediated cleavage. In stage 16 wurst and mega mutant embryos, we detect Pio antibody staining at the chitin cable, suggesting that Pio is cleaved and released into the dorsal trunk tube lumen. Also, the Cht2 overexpression did not prevent the luminal release of Pio. However, reduced wurst, mega function, and Cht2 overexpression caused an enrichment of punctuate Pio staining at the apical cell membrane and matrix (Figures 1 and 2). Although the three proteins are involved in different subcellular requirements, they all contribute to the determination of tube size by affecting either the apical cell membrane or the formation of a well-structured apical extracellular chitin matrix, indicating that changes at the apical cell membrane and matrix in stage 16 embryos affect the Pio pattern at the membrane. It also shows that local Pio linkages at the cell membrane and matrix are still cleaved by the Np function for luminal Pio release, which explains why those mutant embryos do not show pio mutant-like membrane deformations and Np-mutant-like bulges. This is in line with our observations that tracheal Pio overexpression cannot cause tube size defects as the Np function is sufficient to organize local Pio linkages at the membrane and matrix. Therefore, it is unlikely that tracheal tube length defects in wurst and mega mutants as well as in Cht2 misexpression embryos are caused by the apical Pio density enrichment.
Nevertheless, oversized tube length due to the misregulation of the apical cell membrane and adjacent chitin matrix may cause changes to local Pio set linkages and the need for Np-mediated cleavage. Strikingly, we observe a lack of Pio release in Np mutants. This shows that Pio density at the membrane versus lumen depends predominantly on Np function. The molecular mechanisms that coordinate the Np-mediated Pio cleavage are unknown and will be necessary for understanding how tubes resist forces that impact cell membranes and matrices. On the other hand, Pio is required for the extracellular secretion of its interaction partner Dpy (Figure 5—figure supplement 2). At the same time, Dpy is needed for Pio localization at the cell membrane and its distribution into the tube lumen (Figure 5—figure supplement 3). Consistently, in vivo, mCherry::Pio, and Dpy::eYFP localization patterns overlap at the apical cell surface and within the tube lumen (Figure 5—figure supplement 1). These observations support our model that Pio and Dpy interact at the cell surface where Np mediates Pio cleavage to support luminal Pio release by the large and stretchable matrix protein Dpy (Figure 9).
Taenidial organization prevents the collapse of the tracheal tube. Therefore, cortical (apical) actin organizes into parallel-running bundles that proceed to the onset of cuticle secretion and correspond precisely to the cuticle’s taenidial folds (Matusek et al., 2006; Öztürk-Çolak et al., 2016). Mutant larvae of the F-actin nucleator formin DAAM show mosaic taenidial fold patterns, indicating a failure of alignment with each other and along the tracheal tubes (Matusek et al., 2006). In contrast, pio mutant dorsal tracheal trunks contained increased ring spacing (Figure 3A). Fusion cells are narrow doughnut-shaped cells where actin accumulates into a spotted pattern. Formins, such as diaphanous, are essential in organizing the actin cytoskeleton. However, we do not observe dorsal trunk tube fusion defects as found in the presence of the activated diaphanous.
On the other hand, ectopic expression of DAAM in fusion cells induces changes in apical actin organization but does not cause any phenotypic effects (Matusek et al., 2006). DAAM is associated with the tyrosine kinase Src42A (Nelson et al., 2012), which orients membrane growth in the axial tube dimension (Förster and Luschnig, 2012). The Src42 overexpression elongates tracheal tubes due to flattened axially elongated dorsal trunk cells and AJ remodeling. Although flattened cells and tube overexpansion are similar in pio mutant embryos, we did not observe a mislocalization of AJ components, as found upon constitutive Src42 activation (Förster and Luschnig, 2012). Instead, we detected an unusual stretched appearance of AJs at the fusion cells of pio mutant dorsal trunks (Figure 4B and C), which to our knowledge, has not been observed before and may play a role in regulating axial taenidial fold spacing and tube elongation.
Self-organizing physical principles govern the regular spacing pattern of the tracheal taenidial folds (Hannezo et al., 2015). The actomyosin cortex and increased actin activity before and turnover at stage 16 drive the regular pattern formation. However, the cell cortex and actomyosin are in frictional contact with a rigid apical ECM. The Src42A mutant embryos contain shortened tube length but increased taenidial fold period pattern due to decreased friction. In contrast, the chitinase synthase mutant kkv1 has tube dilation defects and no regular but an aberrant pearling pattern caused by zero fiction (Hannezo et al., 2015).
In contrast, pio mutant embryos do not contain tube dilation defects or shortened tubes but increased tube length (Figures 1 and 8; Figure 1—figure supplement 1). Furthermore, our cbp and antibody stainings reveal the presence of a luminal chitin cable and a solid aECM structure in pio mutant stage 16 embryos (Figure 8; Figure 1—figure supplement 1; Figure 3—figure supplement 2). In addition, apical actin enrichment in tracheal cells of pio mutant embryos appeared wt-like. Nonetheless, pio mutant embryos show an increased taenidial fold period compared with wt, indicating a decreased friction. Thus, we propose that the lack of Pio reduces friction. Reasons might be subtle defects of actomyosin constriction or chitin matrix, which we have not detected in the pio mutant tracheal cells. Further reasons for lower friction might also be the loss of Pio set local linkages between apical cortex and aECM in stage 16 embryos, which are modified by Np, as proposed in our model (Figure 9).
Heterozygous and homozygous pio mutant embryos generally do not show tubal collapse. However, the loss of Pio and accompanying lack of Dpy secretion in stage 17 pio mutant embryos led to the loss of a Pio/Dpy matrix, impacting the late embryonic maturation and differentiation of a normal chitin matrix at the apical cell surface. TEM images reveal reduced dense chitin matrix material at taenidial folds and misarranged taenidial fold pattern (Figure 1; Figure 2—figure supplement 1), suggesting impaired taenidial function prevents tube lumen from collapsing after tube protein clearance. Wurst knockdown and mutant embryos do not show general tube collapse, but luminal chitin fiber organization is disturbed in stage 17 embryos (Behr et al., 2007). Therefore, transheterozygous wurst;pio mutant embryos may combine both defects and suffer from maturation deficits of the chitin/ZP matrix at the apical cell surface and within the tube lumen, which finally causes a high number of embryos with incomplete gas-filling due to tube collapse. These maturation deficits are even more dramatic in the wurst;pio double mutants, which show no gas-filling.
Our studies on human Matriptase provide evidence for a mechanistic conservation of ZP domain protein as a substrate for ectodomain shedding (Figure 8). The upregulation of Matriptase activity and increased TGFβ receptor density affect human and mouse model idiopathic pulmonary fibrosis cells on pulmonary fibrogenesis (Bardou et al., 2016; Naik et al., 2012). Furthermore, the human Matriptase induces the release of proinflammatory cytokines in endothelial cells, which contribute to atherosclerosis and probably also to abdominal aortic aneurysms (Seitz et al., 2007). The membrane bulges arising in our Drosophila model during tracheal tube elongation upon Np loss of function showed analogy to the appearance of artery aneurysms. Bulges with varying phenotypic expression in different organs can lead to aortic rupture due to fragile artery walls or degeneration of layers in responses to stimuli, such as shear stresses (Kubo et al., 2015). Indeed, aneurysms development is forced by alterations in the ECM (Yoon et al., 1999) and are characterized by extensive ECM fragmentation caused by shedding of membrane-bound proteins (Antalis et al., 2016; Quintana and Taylor, 2019; Yoon et al., 1999; Zhong and Khalil, 2019).
We identified a dynamic control of matrix proteolysis, very likely enabling fast and site-specific uncoupling of membrane-matrix linkages when tubes expand. Such a scenario has not yet been studied in angiogenesis. It may represent a new starting point for genetic studies to decipher the putative roles of ZP domain proteins and Matriptase in clinically relevant syndromes, including the formation of aneurysms caused by membrane deformation and defects in size determination of airways and vessels.
Materials and methods
Reagent type (species) or resource | Designation | Source or reference | Identifiers | Additional information |
---|---|---|---|---|
Antibody | Mouse, anti-Crb, monoclonal | DSHB | Cq4 | 1:10 |
Antibody | Mouse anti-Flag, monoclonal | Merck | F3165 | 1:500 |
Antibody | Mouse anti-Gasp/Obst-C, monoclonal | DSHB | 2A12 | 1:5 |
Antibody | Chicken anti-GFP, polyclonal | Abcam | ab13970 | 1:1000 |
Antibody | Rabbit anti-GFP, polyclonal | Synaptic Systems132003 | 132003 | IF: 1:500 WB: 1:10,000 CIF: 1:1000 |
Antibody | Rabbit anti-Knk, polyclonal | Moussian et al., 2006 | Moussian | 1:50 |
Antibody | Mouse anti-Mega, monoclonal | Jaspers et al., 2012 | Schuh | 1:50 |
Antibody | Rabbit anti-mCherry, polyclonal | Rockland | 600-401P16 | IF: 1:500 WB: 1:10,000 |
Antibody | Rabbit anti-Obst-A, polyclonal | Petkau et al., 2012 | Behr | 1:50 |
Antibody | Rabbit anti-Pio, polyclonal | Jaźwińska et al., 2003 | Affolter | 1:100 |
Antibody | Rabbit anti-Serp, polyclonal | Luschnig et al., 2006 | Luschnig | 1:50 |
Antibody | Rabbit anti-Spalt, polyclonal | Kühnlein et al., 1994 | Schuh | 1:25 |
Antibody | Mouse anti-α-Spectrin, monoclonal | DSHB | 3A9 | 1:10 |
Antibody | Anti-Strep-HRP, mouse monoclonal | IBA | 1-1509-001 | 1:10,000 |
Antibody | Mouse anti-β-Tubulin, monoclonal | DSHB | E7 | 1:100 |
Antibody | Guinea pig ani-Uif, polyclonal | Zhang and Ward, 2009 | Ward | 1:100 |
Antibody | Rabbit anti-Verm, polyclonal | Luschnig et al., 2006 | Luschnig | 1:50 |
Antibody | Donkey anti-goat Alexa488, polyclonal | Dianova | 705-545-003 | 1:400 |
Antibody | Donkey anti-guinea pig Cy3, polyclonal | Dianova | 706-165-148 | 1:400 |
Antibody | Donkey anti-mouse Alexa647, polyclonal | Dianova | 715-605-020 | 1:400 |
Antibody | Donkey anti-mouse Alexa647, polyclonal | Dianova | 715-605-020 | 1:400 |
Antibody | Donkey anti-rabbit Cy3, polyclonal | Dianova | 711-167-003 | 1:400 |
Antibody | Donkey anti-rabbit-AlexaFluor488, polyclonal | Thermo Fisher Scientific | A-11034 | 1:500 |
Antibody | Donkey anti-rabbit-AlexaFluor568, polyclonal | Thermo Fisher Scientific | A-21069 | 1:500 |
Antibody | Goat anti-mouse-HRP, polyclonal | Thermo Fisher Scientific | G-21040 | WB: 1:10,000 |
Antibody | Goat anti-rabbit-HRP, polyclonal | Thermo Fisher Scientific | Thermo Fisher ScientificG-21234 | 1:10,000 |
Reagent | WGA, Alexa Flour 633 | Invitrogen | W21404 | 1:100 |
Reagent | Cbp, Alexa488 | New England Biolabs | 1:200 | |
Reagent | Phalloidin- PromoFluor-488 | PromoKine, VWR | PROMOPK-PF488P-7 | 1:75 |
Genetic reagent | btl-Gal4 | Bloomington Drosophila Stock Center (BDSC) | ||
Genetic reagent | crb2 (crb11A22) | BDSC | Stock ID 3448 | https://flybase.org/reports/FBal0001817.html |
Genetic reagent | dpyolvR/SM5 | BDSC | Stock ID 280 | https://flybase.org/reports/FBal0002971#phenotypic_data_sub |
Genetic reagent | Dumpy::eYFP [CPTI-001769] | Lye et al., 2014 | Sanson | |
Genetic reagent | megaG0012/FM7, act-GFP | Behr et al., 2003 | Schuh; U Schäfer | |
Genetic reagent | shrub4/Cyo | Dong et al., 2014 | Hayashi | |
Genetic reagent | w1118 | BDSC | https://flybase.org/reports/FBal0018186.html | |
Genetic reagent | w*; mCherry::pio/CyO, dfd-eYFP | This manuscript | Drees | Generation as described in the supplement, available from MB |
Genetic reagent | w*; flzC1 | This manuscript | Drees | Generation as described in the supplement, available from MB |
Genetic reagent | w*; flzC1, mCherry::pio/CyO, dfd-eYFP | This manuscript | Drees | Generation as described in the supplement, available from MB |
Genetic reagent | w1118;PBac{681 .P.FSVS-1}flzCPTI001902 | Kyoto Stock Center | Stock ID 115246 | https://flybase.org/reports/FBti0143804 |
Genetic reagent | w*; pio5M/CyO, dfd-eYFP | This manuscript | Drees | Generation as described in the supplement, available from MB |
Genetic reagent | w*; pio17C/CyO, dfd-eYFP | This manuscript | Dress | Generation as described in the supplement, available from MB |
Genetic reagent | w*; NpP6/CyO, dfd-eYFP | Drees et al., 2019 | Drees | Available from MB |
Genetic reagent | w*; NpP6, P{Gal4-btl}/CyO, dfd-eYFP | Drees et al., 2019 | Drees | Available from MB |
Genetic reagent | w*; NpC2, P{UAS-NpS990A}/CyO, dfd-eYFP | Drees et al., 2019 | Drees | Available from MB |
Genetic reagent | w*; NpC2/CyO, dfd-eYFP; P{UASp-GFPS65C-alphaTub84B}3/TM3, Sb1 | Drees et al., 2019 | Drees | Available from MB |
Genetic reagent | w*; NpP6, mCherry::pio/CyO, dfd-eYFP | Drees et al., 2019 | Drees | Available from MB |
Genetic reagent | w*; P{UAS-NpS990A}/P{UAS-NpS990A} | Drees et al., 2019 | Drees | Available from MB |
Genetic reagent | w*; P{UAS-Np}/P{UAS-Np} | Drees et al., 2019 | Drees | Available from MB |
Genetic reagent | wurst162/FM7-actin-GFP | Behr et al., 2007 | Behr | Available from MB |
Genetic reagent | UAS-Cht2 | Tonning et al., 2005 | Uv | |
Genetic reagent | w[1118]; P{w[+mC]=UAS-myr-mRFP}1 | BDSC | Stock ID 7118 | https://flybase.org/reports/FBst0007118.html |
Genetic reagent | UAS-wurst-RNAi | Stümpges and Behr, 2011 | VDRC | |
Cell line (D. melanogaster) | S2R+ cells | Drees et al., 2019 | DGRC | https://flybase.org/reports/FBtc0000150.html#:~:text = S2R%2B%20is%20an%20isolate%20of,to %20the%20original%20S2%20line.&text = S2R%2B%20is%20an%20isolate%20of%20S2 %20that%20has%20receptors%20for%20wg%20signalling. |
Cell line (D. melanogaster) | Kc167 | Drees et al., 2019 | DGRC | https://flybase.org/reports/FBtc0000001.html |
Sequence-based reagent | pio-sgRNA-sense | Eurofins Genomic | CTTCGATTGGGACACCGAGCCACT | |
Sequence-based reagent | pio-sgRNA-antisense | Eurofins Genomics | AAACAGTGGCTCGGTGTCCCAATC | |
Sequence-based reagent | flz-sgRNA-sense | Eurofins Genomics | CTTCGTGGGTTACGCCGG CCTCAA | |
Sequence-based reagent | flz-sgRNA-antisense | Eurofins Genomics | AAACTTGAGGCCGGCGTA ACCCAC | |
Sequence-based reagent | UAS-mCherry::pio-for | Eurofins Genomics | GAATTCATGAAGACAGGCACTCGAATGGACGCTTTCCACACGGCGCTGCACTTAATCACAATCGCAGCTCTGACGACG | |
Sequence-based reagent | UAS-mCherry::pio-rev | Eurofins Genomics | CTCGAGGCCGCCTTTGTAAAGCTCATCC | |
Sequence-based reagent | Pio-5’-HA1-for | Eurofins Genomics | ACTAGTCCGAATTCGCAGG TGATTATCGCCTCTCGGCC ATCAG | |
Sequence-based reagent | Pio-5’-HA1-rev | Eurofins Genomics | AAGCTTCTTTAATTAAAGG GGAAATTTCG | |
Sequence-based reagent | Pio-5’-HA2-for | Eurofins Genomics | ACTAGTGGCAAGCTTACTG GCGATGGATTAGGCC | |
Sequence-based reagent | Pio-5’-HA2-rev | Eurofins Genomics | CACCTGCGATCTTAATCTT GCCAGCGTCTGTC | |
Sequence-based reagent | Pio-3’-HA-for | Eurofins Genomics | TTAAGGAAGAGCACACAG TTGGGCGCTTTGTTAGTCG | |
Sequence-based reagent | Pio-3’-HA-rev | Eurofins Genomics | CGGGGAAGAGCGACGAGA TTGCGCCGGAAAATAAG | |
Sequence-based reagent | UAS-pio-ORF-for | Eurofins Genomics | CTCGAGCCAACGGCAATGAAAGATGCCC | |
Sequence-based reagent | UAS-pio-ORF-rev | Eurofins Genomics | TCTAGATTAGCTGCTGTGCGAGAAG | |
Sequence-based reagent | Dpy-ZP-for | Eurofins Genomics | GCTTTACAAAGGTTACACGGGTAATCCG | |
Sequence-based reagent | Dpy-CT-for | Eurofins Genomics | GCTTTACAAAGGTGGAAATGCCAGGATTG | |
Sequence-based reagent | Dpy-CTZP-rev | Eurofins Genomics | GTGGAGCCGGCCACCATTTATGGAGGTTTC | |
Sequence-based reagent | Dpy-ZP-for | Eurofins Genomics | GGCCACCATTTATGGAGGTTTC | |
Sequence-based reagent | Dpy-ZP-rev | Eurofins Genomics | GGTTCCTTCACAAAGATCCTTTAGGATATGTAATCCGGCG | |
Sequence-based reagent | Strep::TGFBR3::GFP1-for | Eurofins Genomics | CTGAATAGGGAATTGGGAATTCATGACTTCCCATTATG | |
Sequence-based reagent | Strep::TGFBR3::GFP1-rev | Eurofins Genomics | CACCGCTGCCACCTCCTGATCCGCCACCCTTTTCAAACTGCGGATGACTCCATGCACTTTGCACCTCTTCTGGCTCTC | |
Sequence-based reagent | Strep::TGFBR3::GFP2-for | Eurofins Genomics | ATCAGGAGGTGGCAGCGGTGGAAGTGCATGGAGCCATCCCCAATTCGAGAAGGGGAGCGTGGATATTGCCCTG | |
Sequence-based reagent | Strep::TGFBR3::GFP2-rev | Eurofins Genomics | TCACCATACCGCCGCTAGCGGCCGTGCTGCTGCTG | |
Plasmid | pJet1.2 | Thermo Fisher Scientific | ||
Plasmid | pUAST | GAL4/UAS-mediated overexpression; Brand and Perrimon, 1993 | ||
Plasmid | pBFv-U6.2 | Expression of single sgRNA; Kondo and Ueda, 2013 | ||
Plasmid | pBFv-U6.2B | Expression of two sgRNAs; Kondo and Ueda, 2013 | ||
Plasmid | pHD-ScarlessDsRed | Scarless genome editing via HDR | DGRC | |
Plasmid | actin5C-Gal4 | Expression of Gal4 in cultured cells; Usui et al., 1999 | ||
Software, algorithm | Clustal omega algorithm | https://www.ebi.ac.uk/Tools/msa/clustalo/ | ||
Software, algorithm | DNASTAR software suite | Lasergene Software | Lasergene Software | |
Software, algorithm | Flybase | https://www.flybase.org | https://www.flybase.org | BLASTP algorithm |
Software, algorithm | Huygens professional | SVI | 20.10 | |
Software, algorithm | Illustrator | Adobe | CS6 | https://www.adobe.com |
Software, algorithm | Imaris 9.7.2 | Oxford Instruments | Oxford Instruments | https://imaris.oxinst.com/ |
Software, algorithm | NetOGlyc | DTU Health Tech | https://services.healthtech.dtu.dk/ | |
Software, algorithm | Office 365 (Word, Excel) | Microsoft | Microsoft | https://www.microsoft.com |
Software, algorithm | ProP | DTU Health Tech | https://services.healthtech.dtu.dk/ | |
Software, algorithm | SignalP | DTU Health Tech | https://services.healthtech.dtu.dk | |
Software, algorithm | Photoshop CS6 | Adobe | CS6 | https://www.adobe.com |
Software, algorithm | SMART | EMBL | EMBL | |
Software, algorithm | Serial Cloner | Serial basics | 2.6.1 | |
Software, algorithm | TMHMM 2.0 algorithm | DTU Health Tech | https://services.healthtech.dtu.dk/ | |
Software, algorithm | ZEN 2.3 | Zeiss | Zeiss | 2.3, black |
Fly husbandry, gas-filling, and statistics
Request a detailed protocolFor collection of D. melanogaster embryos and larvae, flies of the desired genotype took place at 25°C for collection of embryos for RNAi-mediated knockdown. All used fly strains are listed in the Materials and methods section in the supplement. In all other cases, egg-laying took place at 22°C. The apple-juice agar plates were exchanged with fresh plates at respective points of time to obtain embryos or larvae at certain developmental stages. The mutant alleles were kept with ‘green’ balancers to recognize mutant embryos.
We used w1118 as control (referred to as wt). Rescue experiments: w*; pio17C, btl-Gal4/Cyo, dfd-eYFP were mated with w*; pio5M /Cyo, dfd-eYFP; UAS-Pio to receive the rescue in the progenoty w*; pio17C, btl-Gal4/pio5M; UAS-pio / +.
For gas-filling assay, we transferred stage 17 embryos and freshly hatched larvae onto agar plates and studied those by bright-field microscopy. Significance was tested using t-tests in Excel 2019; asterisks indicate p-values (*p<0.05, **p<0.01, ***p<0.001); error bars indicate the standard deviation. Relevant guidelines (e.g. ARRIVE) were followed in this study.
Embryo dechorionation, fixation, and immunostainings
Request a detailed protocolIn general, we analyzed for control minimum n>20 embryos, and for pio or other mutants n>10 embryos. Embryos were washed from the apple-juice agar plates into close-meshed nets, incubated for 3 min in a bleach solution (2.5% sodium hypochlorite) for dechorionation.
For subsequent fixation, embryos were incubated at 250 rpm for 20 min in 1 ml 10% (vol/vol) formaldehyde solution (50 mM EGTA, pH 7.0), 2 ml HEPES solution, and 6 ml heptane. The fixative was removed and 8 ml methanol added and incubated at 500 rpm for 3 min to detach the vitelline membrane. Finally, embryos were washed with methanol and stored at –20°C.
All used antibodies are listed in the Materials and methods section in the supplement. For antibody staining, embryos were 5 min washed three times with BBT followed by blocking for 30 min. Subsequently, pre-absorbed primary antibodies diluted in blocking solution were applied to the embryos and incubated overnight at 4°C. Primary antibody was incubated overnight washed off six times with BBT. After blocking for 30 min pre-absorbed secondary antibodies diluted in blocking solution were added to the embryos and incubated for 2 hr. If required, Alexa488-conjugated cbp was added to the secondary antibody dilutions at a 1:200 dilution for staining of chitin. Finally, embryos were washed six times with PBT for 5 min and mounted either in Prolong mounting medium (Thermo Fisher Scientific) or in phenol-free Kaiser’s glycerol-gelantine (Carl Roth).
Light microscopy, image acquisition, and image analysis
Request a detailed protocolFor bright-field and dark-field light microscopy, we used a Zeiss Axiophot 2e upright microscope (A-PLAN 10×/0.25 and 25×/0.8 LCI Plan Neofluar Objectices and Zeiss Glycerin) and images were acquired with an AxioCam (HRc/mRc) and the acquisition software (Zeiss Axiovision 12). For handling confocal analysis, we used ZEN software (2.3, SP1 black) and Zeiss LSM780-Airyscan (Zeiss, Jena) microscopes. Overview imaging were taken with 25×/0.8 LD-PLAN Neofluar and magnifications with Plan-Apochromat 40×/1.4 Oil DIC M27 and with 63×/1.3 PLAN Neofluar M27 objectives and Zeiss oil/water/glycerol medium. For imaging standard ZEN confocal microscopy (Pinhole Airy1) settings and image processing (maximum intensity projection and orthogonal section) and orthogonal projections were used.
ZEN black 2.3 fluorescence intensity profile was used with the arrow toolset across the tube lumen to measure and compare the fluorescence pixel intensities of individual channels. For quantification, we determined the fluorescent intensities profile across the tube to identify values at apical membrane and tube lumen at a minimum 10 different positions of DTs (metameres 5–6) of two distinct embryos for each genetic background. The maximum values of membranes versus tube lumen were set into ratio and compared between control, mega mutant, and Cht2 overexpression. Statistical analysis was calculated with Excel 2019. To quantify protein co-localization and overlap in confocal and airyscan images, we used ZEN black 2.3 co-localization tool, set two channels for comparison, and considered quadrant 3, which represents a pixel with high-intensity levels in both channels, as co-localized pixels. Those pixels were set to white for better visualization in the images.
The ‘express’ deconvolution of SVI Huygens pro was used with standard settings. Images were transferred to ZEN for further analyses. For Airyscan acquisition, standard ZEN settings and optimal Z-stack distances were used. For 3D visualization (HP Z4 graphics workstation), deconvolved confocal and Airyscan Z-stacks were processed in Imaris (version 9.7.2) to convert voxel-based data into surface objects. We cropped Images with Adobe CS6 Photoshop and designed figures with CS6 Illustrator. For equipment details, see https://bioimaging.uni-leipzig.de/equipment.html.
CRISPR/Cas9-mediated mutagenesis and genome editing
Request a detailed protocolUsing CRISPR/Cas9-mediated mutagenesis, we generated three independent pio lack of function alleles [pio17C, pio5M, and pio11C] and one flz lack of function allele (flzC1) that carry frameshift mutations in the ORF. The pio mutations led to truncated Pio proteins containing only short N-terminal stretches but lacking all critical for Pio function. These mutations in pio17C, pio5M, and pio11C alleles and flzC1 allele caused embryonic lethality. The in vivo analysis of airways in pio17C, pio5M, and pio11C homozygous and pio17C/pio5M transheterozygous mutant stage 17 embryos revealed lack of tracheal air-filling. The sequences of sgRNAs are listed in the Materials and methods section in the supplement.
The CRISPR Optimal Target Finder tool was used to identify specific sgRNA targets in the D. melanogaster genome. DNA oligos corresponding to the chosen sgRNA target sequences were annealed and ligated into the pBFv-U6.2 vector via BbsI restriction sites (Kondo and Ueda, 2013). For simultaneously targeting two sgRNA recognition sites, one sgRNA target encoding annealed DNA oligo was ligated into the pBFv-U6.2 vector and the other one into the pBFv-U6.2B vector. Subsequently, the U6.2 promotor and sgRNA target sequence of the pBFv-U6.2 vector were transferred to the sgRNA target sequence containing pBFv-U6.2B vector via EcoRI/NotI endonuclease restriction sites. Vectors for sgRNA expression were injected into nos-Cas9-3A embryos by BestGene. Adult flies that developed from the injected embryos were then crossed with balancer chromosome strains. Single flies of the resulting F1 generation were again crossed with balancer chromosome strains and the resulting F2 stock was analyzed regarding lethality of the putatively mutated chromosome. The regions of the genomic sgRNA target sites from the obtained lethal stocks were amplified by PCR followed by purification of the amplicon and sequencing.
The CRISPR/Cas9 system was used for targeted insertion of transgenes by homology-directed repair. Sequences of homology arms (HAs) were amplified by PCR from genomic DNA of nos-Cas9-3A flies. HAs were then purified and cloned into the pHD-ScarlessDsRed donor vector for editing of pio. A mCherry encoding sequence was inserted into the pio ORF at the 3' end of the 5' HA in the pHD-ScarlessDsRed donor vector. The donor vector together with the vector for sgRNA expression were injected into nos-Cas9-3A embryos by BestGene. Adult flies that developed from the injected embryos were crossed with w* flies and resulting F1 flies were selected for the presence of 3xP3-DsRed marker. Stocks were then established by crosses with balancer chromosome strains. Correct insertion of the transgenes was verified by amplification of the targeted genomic regions by PCR and subsequent purification of the amplicons and sequencing. The 3xP3-DsRed marker gene that was inserted in the pio locus was removed by crosses with the tub-PBac strain and subsequent selection of F2 flies that lacked the tub-PBac balancer chromosome and the 3xP3-DsRed marker gene.
Cell culture-based experiments
Request a detailed protocolD. melanogaster S2R+ cells (DGRC) and Kc167 cells (DGRC) were kept in flasks containing Schneider’s Drosophila medium (Thermo Fisher Scientific) supplemented with 1% penicillin/streptomycin (Thermo Fisher Scientific) and 10% FBS (Sigma-Aldrich) at 25°C. Handling of cells was performed in the sterile environment of a clean bench. The Drosophila S2R+ and Kc167 cells were ordered from the Drosophila Genomics Resource Center (DGRC) where extensive authentication of the cell lines is carried out. Negative mycoplasma status was tested by PCR. Frequent observation of cell shape, growth rate, and ability to adhere to surfaces was carried out to ensure that key parameters of the cell line remained constant during passaging of the cells for ongoing experiments.
Confluent cells were detached, diluted 1:6, and transferred either to 10 cm diameter Petri dishes, or to six-well plates (Greiner Bio One), or to glass bottom micro-well dishes (MatTek; 6 ml per dish), approximately 24 hr before transfection. The UAS/Gal4 system was used for protein overexpression in cultured cells. Cells in six-well plates and glass bottom micro-well dishes were transfected with 500 ng actin5C-Gal4 vector and 500 ng pUAST-responder vector (1 µg total DNA), while cells in 10 cm diameter Petri dishes were transfected with twice the amount of each vector (2 µg total DNA). The Effectene transfection reagent (QIAGEN) was used for cell transfection according to suppliers’ guidelines. For transfections of cells in six-well plates and glass bottom micro-well dishes, vector DNA was mixed with 190 µl Effectene EC buffer (QIAGEN) followed by adding 8 µl Effectene Enhancer (QIAGEN), vortexing and incubation for 5 min. Subsequently, 20 µl Effectene were added and the mix was vortexed for 10 s followed by incubation for 15 min. The transfection mix was then carefully trickled onto the cells with a pipette. Incubation steps were performed at room temperature. For transfections of cells in 10 cm diameter Petri dishes with 2 µg DNA, volumes of the transfection mix components were doubled.
Cells in glass bottom micro-well dishes were incubated at 25°C for approximately 48 hr after transfection followed by imaging with a confocal LSM.
For immunostainings cells were incubated at 25°C for 48 hr after transfection and then washed with PBS twice and fixed for 15 min by adding formaldehyde solution. The cells were then washed twice with PBS and incubated in PBTX for 150 s for permeabilization. Subsequently, cells were washed three times with PBS followed by 30 min blocking. Primary antibodies diluted in blocking solution were added to the cells and incubated for 2 hr. Then cells were washed three times with PBS followed by incubation with secondary antibodies diluted in blocking solution for 1 hr. Finally, the cells were washed twice with PBS followed by mounting in Prolong mounting medium with DAPI (Thermo Fisher Scientific). Images were acquired with a confocal LSM. All incubation and washing steps were performed at room temperature.
For pull-down assays Strep-tagged Pio was co-expressed in S2R+ cells together with either of two independent RFP-tagged Dpy fragments for pull-down assays. One RFP-tagged Dpy contains the Dpy C-terminal region, and the second exclusively the Dpy-ZP domain. Expression products of both RFP::Dpy constructs were only pulled down together with Strep::Pio in the Strep-IP samples. For western detection see below. All used oligos to generate constructs are listed in the Materials and methods section in the supplement.
Protein purification and western blotting
Request a detailed protocolWe dechorionated embryo collection, squashed them with a needle and pulsed with ultrasound for 30 s, added 25 ml 4× SDS sample buffer, heated for 9 min at 96°C, centrifuged at 11,000 rpm for 20 min. Lysate was stored in a fresh cup at −80/–20°C. Schneider’s cells were manually detached, centrifuged at 900 rpm, PBS was replaced by 35 µl 4× SDS sample buffer and heated 9 min at 96°C, and stored at –80°C. For supernatant samples, we centrifuged at 15,000 rpm at 4°C for 20 min, washed with acetone and resuspended in 100 µl 1× SDS sample buffer, heating it at 96°C for 9 min. We used 4–20% gradient Mini-Protean TGX Precast Protein Gels (Bio-Rad) together with PageRuler prestained protein ladder (Thermo Scientific), MiniProtean chamber (Bio-Rad), PowerPac Basic power supply (Bio-Rad) for 40 min at 170 V. The gels were then equilibrated in transfer buffer and packed into a western blotting sandwich. Sandwich blotting was performed with a PVDF transfer membrane with 0,2 µm pore size (Thermo Fisher Scientific) for embryos lysates or in other cases, Amersham Hybond-ECL membrane with 0.2 µm pore size (GE Healthcare). Protein transfer to the membrane was performed on ice in a MiniProtean chamber (Bio-Rad) that was filled with transfer buffer at 300 mA for 90 min.
Fluorescence recovery after photobleaching
Request a detailed protocolThe stage 16 embryos were dechorionated, transferred, and fixed on the coverslip of small Petri dishes (ibidi, Germany; https://ibidi.com/dishes/185-glass-bottom-dish-35-mm.html) and mounted in PBS. We used LSM780 confocal for FRAP experiments. To define the region of interest (ROI) we used the Zeiss Zen software. The bleaching was performed with 405 nm full laser power (50 mW) at the ROI for 20 s. A Z-stack covering the whole depth of the tracheal tubes in the ROI were taken at each imaging step. The confocal images were taken every 2 min until 60 min after bleaching. To correct for movements of the embryos, images that are presented in figures and supplemental movies were manually overlayed to center them in the same focal plane and to correct for movements in the X and Y axis. Fluorescence intensity in the bleached ROIs was measured after correction for embryonic movements using Fiji.
Electron microscopy
Request a detailed protocolStage 17 Drosophila embryos were dechorionated, transferred to a 150 μm specimen planchette (Engineering Office M. Wohlwend GmbH), and frozen with a Leica EM HBM 100 high-pressure freezer (Leica Microsystems). Vitrified samples were embedded with an Automatic Freeze Substitution Unit (EM AFS; Leica Microsystems) at −90°C in a solution containing anhydrous acetone, 0.1% tannic acid, and 0.5% glutaraldehyde for 72 hr and in anhydrous acetone, 2% OsO4, and 0.5% glutaraldehyde for additional 8 hr. Samples were then incubated at −20°C for 18 hr followed by warm-up to 4°C. After subsequent washing with anhydrous acetone, embedding in Agar 100 (Epon 812 equivalent) was performed at room temperature. Counterstaining of ultrathin sections was done with 1% uranylacetate in methanol.
Alternatively, the dechorionated embryos were fixed by immersion using 2% glutaraldehyde in 0.1 M cacodylate buffer at pH 7.4 overnight at 4°C. The cuticle of the embryos was opened by a cut to allow penetration of fixative. Postfixation was performed using 1% osmium tetroxide. After pre-embedding staining with 1% uranyl acetate, tissue samples were dehydrated and embedded in Agar 100. Ultrathin sections were evaluated using a Talos L120C transmission electron microscope (Thermo Fisher Scientific, MA, USA).
Data availability
All data confocal raw data used for this this study are either included in the manuscript as supporting files or available from the research data repository of the saxon universities, Open Access Repository and Archive OpARA, https://doi.org//10.25532/OPARA-240.
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Open Access Repository and Archive OpARAproteolysis of ZP protain Pio at the apical cell membrane.https://doi.org/10.25532/OPARA-240
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Decision letter
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Elisabeth KnustReviewing Editor; Max-Planck Institute of Molecular Cell Biology and Genetics, Germany
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Claude DesplanSenior Editor; New York University, United States
In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.
[Editors' note: this paper was reviewed by Review Commons.]
https://doi.org/10.7554/eLife.91079.sa1Author response
1. General Statements
We thank the three reviewers for carefully reading our manuscript and for all considerations, ideas, suggestions, and comments. These were all very helpful for us to strengthen the scientific statements of our manuscript. Please, note that all changes are marked in red in the manuscript and supplement. Below you will find, point by point, our responses to all questions and comments.
2. Point-by-point description of the revisions
Reviewer #1
Overall, this is an exciting work. There are, however, several open questions that the authors could address to facilitate understanding of their work. These points are:
1. On page 5, lines 113ff, the authors mention the membrane bulges that they analyse in figure 1. They show these deformations by light (confocal) and electron microscopy. However, the bulges seen by confocal microscopy seem to be bigger that those seen by electron microscopy. The authors could quantify the sizes of the bulges for clarification.
We quantified the size of the membrane bulges. At the confocal we measured in average 750nm as mean value of identified bulges (n=12) with 650nm as minimal and 890nm as maximal sizes. At the TEM we measure ~243nm as mean value (n=61), with a range between 62nm as minimum and 442 as maximum value. These measurements are shown as Figure 1E.
Please note that measurements of TEM images do not always capture the three-dimensionality of bulges and may show only parts of them. In addition, ultrastructure is more sensitive and can easily detect small membrane changes that we cannot observe with confocal and airsycan microscopy. In contrast, even with our high-quality objective (63x Zeiss Plan Neofluar, Glycerin, 1.3 NA), standard confocal analysis is limited at ~200nm on the XY axis (airyscan ~110nm) and ~450nm on Z-axis. Therefore, TEM analysis detects smaller bulges than confocal analysis, and consequently, this method detected a large range of bulge lengths between 63nm and 441nm.
In contrast, the airyscan method detected a range of bulge length between 0.65 and 0.83 µm. However, confocal and TEM analyses provide evidence of membrane bulges in pio mutant embryos. Please note that we extended our studies and now show membrane bulges in two different pio mutant alleles (17C and 5M) with airsycan microscopy.
2. The subject of the manuscript is rather complicated; presentation of data from Figure 1C and D on lines 113ff and 169ff is confusing.
We apologize and thank the reviewer for careful reading. We revised both paragraphs (lines 108 – 123 and lines 166 – 174) and are confident that the descriptions are now much more understandable. All changes are marked in red.
3. The quality of the sub-images of Figure 2E differs. Especially, the phenotype of the wurst, pio transheterozygous embryo is not well visible.
We apologize for it. We repeated the experiment with wurst;pio transheterozygotes, and generated wurst;pio double mutant embryos to improve the quality. The gas filling assay is shown in Figure 3. With brightfield microscopy in overview images (10x air objective) and closeups of the dorsal trunks (25x Glycerin objectives). Both show the gas-filling defects of dorsal trunk tubes. In a subsequent confocal analysis of chitin stainings in late-stage 17 embryos, we found that tracheal tube lumens are collapsed in the transheterozygotes and double mutant embryos.
4. Lines 246ff: the protein size are given for the mCherry:chimeric proteins; an estimate of the native Pio portions should be given.
The endogenous Pio protein has a calculated mass of about 50.82 kDa. We state it now in the according legend of Figure 6.
5. In Figure 6A, the appearance of chitin in the wildtype tube is different compared to the Np mutant situation, more filamentous. Can the authors comment on that?
The author is correct. The chitin cable formation in Np mutant embryos is normal but lacks the condensation process, and, therefore, fiber structure of the chitin matrix differs from control embryos in late stage 16 and stage 17 embryos (see Drees et al., PLOS Genetics, 2019).
6. In the Discussion section, I would appreciate if the timing of events was discussed or even shown in a model. The central question is: how are the functions of Pio and Np coordinated in time? As I understand, Np should not cleave Pio before morphogenesis is completed. Is there any example in the literature for how such an interaction could be controlled? The overexpression of Np shows that either the ratio between Np and Pio is important, or the btl promoter expresses Np at the "wrong" time point.
We thank the reviewer for this interesting comment.
Of course, we did not measure forces, but it has been published that axial forces appear at the apical cell membrane during stage 16 tube expansion. Our data show that Np cleaves Pio ZP domain and subsequent release increase during stage 16. The cleaved and released Pio enriches in the lumen during stage 16, from where cleaved Pio is internalized during stage 17 with the help of Wurst-mediated endocytosis. This is supported by several in vivo studies, video microscopy, antibody stainings and biochemical data, such as the interaction of Pio and Dumpy as well as the identification of different Pio products with and without Np cleavage. Moreover, we found membrane bulges that increase in size during stage 16 and identified a subsequent tear-off of the chitin matrix in Np mutant embryos. Thus, we propose that Np is required to cleave Pio-Dumpy linkages at the membrane-matrix when tubes elongate and postulated forces appear at the cell membrane during tube elongation in stage 16 embryos.
We stated this in the discussion as follows:
“The membrane defects observed in both Pio and Np mutants indicate errors in the coupling of the membrane matrix due to the involvement of Pio (Figures 1,7). …, the large membrane bulges in Np mutants affect the membrane and the apical matrix (Figure 7). Since apical Pio is not cleaved in Np mutants (Figure 7D), the matrix is not uncoupled from the membrane as in pio mutant embryos but is likely more intensely coupled, which leads to tearing of the matrix axially along the membrane bulges (Figures 7, 9), when the tube expands in length.”
How could Np be regulated at the membrane? Np is a zymogen that very likely undergoes ectodomain shedding for activation, similar to what has been described for matriptases. Additionally, human matriptase requires transient interaction of the stem region with its cognate inhibitor HAI-2, which Drosophila lacks (see Drees et al., PLOS Gen, 2019). Thus, the regulation of Np activation is not known.
Further, we observed that Dumpy is not degraded in Np mutant embryos during stage 17.
Nevertheless, in a previous publication, we showed that btl-G4 driven Np expression rescues Np mutant phenotypes in a time-specific manner. We used the btl-G4 driver line for these rescue experiments to express Np in tracheal cells. This restored tracheal Dumpy degradation in Np mutant embryos. Thus, btlG-G4 driven Np overexpression is able to rescue Np mutant tracheal phenotypes in a time-specific manner, although Gal4 is expressed from early tracheal development onwards. Further, btl-Gal4 driven Np expression mimics the endogenous Np, which is expressed from stage 11 onwards in all tracheal cells throughout embryogenesis (see Drees et al., PLOS Gen, 2019).
Based on these experiments, we conclude that the btl-G4-driven Np overexpression can cleave Pio ZP domain in stage 16 embryos at the correct time.
However, the ratio of Np expression and Pio is essential in the way that btl-Gal4 driven Gal4 Np overexpression may cause cleavage of a higher number of Pio proteins and the release of critical Pio-Dumpy linkages at the cell membrane and matrix. Thus, increased Pio shedding into the lumen reduces Pio linkages at the membrane, resulting in a pio mutant like tracheal overexpansion in btl-Gal4 driven Gal4 Np overexpression.
Finally, we were able to prove the reviewer’s question in a new experiment. We used btl-Gal4 driven UAS-Np embryos for Pio antibody staining. This revealed Pio enrichment at the tracheal chitin cable in stage 14 and 15 embryos. In contrast, stage 16 embryos showed numerous Pio puncta appearing across the entire tube lumen, indicating that Np mediates Pio shedding specifically in stage 16 embryos and not before. This Np-controlled Pio releases modifies tube length control.
Therefore, we stated this in the manuscript as follows:
Results:
“Our data assumes that Np overexpression may enhance Pio shedding in stage 16 embryos, affecting the Pio-mediated ZP matrix function. Upon breathless (btl)-Gal4-mediated expression of UAS-Np in tracheal cells, we observed a high amount of Pio puncta across the entire tracheal tube lumen, specifically in stage 16 embryos but not in earlier stages (Figure S13). Consistently tracheal Np overexpression led to tube overexpansion in stage 16 embryos resembling the pio mutant phenotype (Figure 8A,B). Thus, Np-mediated Pio shedding controls Pio function.”
Discussion:
“The btl-Gal4-driven Np expression mimics the endogenous Np from stage 11 onwards in all tracheal cells throughout embryogenesis (Drees et al., 2019), suggesting that Np is not expressed at a wrong time point. However, the ratio between Np and Pio is essential. We assume that Np overexpression increases Pio shedding, resulting in a pio loss-of-function phenotype. Thus, the tube length overexpansion upon Np overexpression indicates that Pio cleavage is required for tube length control.
Our observation that the membrane deformations are maintained in Np mutant embryos supports our postulated Np function to redistribute and deregulate membrane-matrix associations in stage 16 embryos when tracheal tube length expands. In contrast, Np overexpression potentially uncouples the Pio-Dpy ZP matrix membrane linkages resulting very likely in unbalanced forces causing sinusoidal tubes.”
7. Also for the discussion: We have two situations where Pio amounts/density are enhanced at the apical plasma membrane. The wurst experiments on lines 136ff show that Pio amount and density depends on endocytosis; is the wurst phenotype (Figure 2), at least partially, due to over-presentation of Pio? Likewise, in Figure 2C, there is more Pio in Cht2 overexpressing tracheae (but there is overall more Pio in these tracheae) – is actually endocytosis reduced in chitin-less luminal matrices? First: does the Pio signal at the apical plasma membrane correspond to membrane-Pio or free-Pio? Second, as in the case of wurst: would more Pio on the membrane (density) affect tracheal dimensions in Cht2 over expressing tracheae? Or are the consequences of Pio accumulation in the apical plasma membrane different in Cht2 and wurst backgrounds? Maybe cleavage of Pio and its endocytosis are dependent on its interaction with the chitin matrix. These questions connect to the question immediately above: how are the functions of the different players coordinated in space and time? We need a discussion on this issue.
We thank the reviewer for this very important idea to discuss the functions of the different players in a coordinated space and time and apologize that we haven’t done before. As this is an important point, we tried to figure out all questions raised by the reviewer and discussed it in several new paragraphs in the discussion:
"Indeed, the anti-Pio antibody, which can detect all different Pio variants, showed a punctuate Pio pattern overlapping with the apical cell membrane marker Uif at the dorsal trunk cells of stage 16 embryos. Additionally, Pio antibody also revealed early tracheal expression from embryonic stage 11 onwards, and due to Pio function in narrow dorsal and ventral branches, strong luminal Pio staining is detectable from early stage 14 until stage 17, when airway protein clearance removes luminal contents.
[…]
Heterozygous and homozygous pio mutant embryos generally do not show tubal collapse. However, the loss of Pio and accompanying lack of Dpy secretion in stage 17 pio mutant embryos led to the loss of a Pio/Dpy matrix, impacting the late embryonic maturation and differentiation of a normal chitin matrix at the apical cell surface. TEM images reveal reduced dense chitin matrix material at taenidial folds and misarranged taenidial fold pattern (Figures 1; S2), suggesting impaired taenidial function prevents tube lumen from collapsing after tube protein clearance. Wurst knockdown and mutant embryos do not show general tube collapse, but luminal chitin fiber organization is disturbed in stage 17 embryos (Behr et al., 2007). Therefore, transheterozygous wurst;pio mutant embryos may combine both defects and suffer from maturation deficits of the chitin/ZP matrix at the apical cell surface and within the tube lumen, which finally causes a high number of embryos with incomplete gas filling due to tube collapse. These maturation deficits are even more dramatic in the wurst;pio double mutants, which show no gas filling.”
8. The sentence on line 242ff should be rephrased: "dynamic" and "elastic" are not opposites.
We thank the reviewer for careful reading. We revised the sentence as follows:
“Our FRAP data suggest that Pio is the dynamic part of the tracheal ZP-matrix, while the static Dpy modulates mechanical tension within the matrix”.
9. A central question to me is the amounts and the density of factors in different genetic backgrounds as mentioned above. Is there any mechanism adjusting the amounts or the density of the players according to the size of the apical plasma membrane or the tracheal lumen? Pio seemingly responds to these changes.
We would like to know the molecular mechanisms that control the density of players at the apical membrane. This question is important and could be the starting point for novel scientific investigations. Mechanisms of protein trafficking, such as exocytosis, recycling and endocytosis regulate delivery and internalization of proteins at the apical cell membrane. Furthermore, protein junctions at the lateral membrane may recognize and therefore may respond to low and high mechanical stresses between cells that appear during tube length expansion. However, we did not observe any hint for misregulation of Pio expression levels in the different mutants which affect endocytosis, SJs and luminal ECM. But we observed a shift of Pio levels between apical cell membrane/matrix and lumen in wurst, mega mutants and Cht2 overexpression. This shift is analyzed with diverse ZEN tools and quantified (Figure 2D-F; Figure S4B). As discussed in the new paragraph, this shift is very likely caused by changes at the apical cell membrane and chitin matrix which impact Pio shedding. Moreover, we observe the lack of Pio release in Np mutants. This shows that Pio density at the membrane versus lumen depends predominantly on Npmediated cleavage. As discussed above, how Np is activated at the apical cell membrane to cleave Pio is not known.
10. The connection of Pio and taenidia is mentioned in the Results section (page 7) but not discussed.
We appreciate the careful reading and comments of the reviewer very much. We included the connection of Pio and taenidial in the Discussion section as follows:
“Taenidial organization prevents the collapse of the tracheal tube. Therefore, cortical (apical) actin organizes into parallel-running bundles that proceed to the onset of cuticle secretion and correspond precisely to the cuticle's taenidial folds (Matusek et al., 2006; Öztürk-Çolak et al., 2016). Mutant larvae of the F-actin nucleator formin DAAM show mosaic taenidial fold patterns, indicating a failure of alignment with each other and along the tracheal tubes (Matusek et al., 2006). In contrast, pio mutant dorsal tracheal trunks contained increased ring spacing (Figure 3A). Fusion cells are narrow doughnut-shaped cells where actin accumulates into a spotted pattern. Formins, such as Diaphanous, are essential in organizing the actin cytoskeleton. However, we do not observe dorsal trunk tube fusion defects as found in the presence of the activated diaphanous.
[…]
Heterozygous and homozygous pio mutant embryos generally do not show tubal collapse.
However, the loss of Pio and accompanying lack of Dpy secretion in stage 17 pio mutant embryos led to the loss of a Pio/Dpy matrix, impacting the late embryonic maturation and differentiation of a normal chitin matrix at the apical cell surface. TEM images reveal reduced dense chitin matrix material at taenidial folds and misarranged taenidial fold pattern (Figures 1; S2), suggesting impaired taenidial function prevents tube lumen from collapsing after tube protein clearance. Wurst knockdown and mutant embryos do not show general tube collapse, but luminal chitin fiber organization is disturbed in stage 17 embryos (Behr et al., 2007). Therefore, transheterozygous wurst;pio mutant embryos may combine both defects and suffer from maturation deficits of the chitin/ZP matrix at the apical cell surface and within the tube lumen, which finally causes a high number of embryos with incomplete gas filling due to tube collapse. These maturation deficits are even more dramatic in the wurst;pio double mutants, which show no gas filling.”
11. Dp remains cytoplasmic in pio mutant background – is the pio mutant phenotype due to defects by lack of Pio AND Dp function? What is the tracheal phenotype of dp mutants?
It has been discussed that dumpyolvr and pio mutants show similar phenotypes in early tracheal development (Jazwinska, 2003) and it has been discussed that dumpyolvr mutant embryos compromise tube size in combination with shrub mutants. The additional quantifications of the dumpyolvr mutant showed significantly increased tube length (Dong 2014). We used dumpyolvr mutant [In(2L)dpyolvr], an X-ray induced mutation of the dumpy gene locus (Wilkin 2000).
dumpyolvr mutant resemble pio null mutant tracheal phenotypes including detached dorsal and ventral branches and oversized tracheal dorsal trunk with curly appearance in late embryos. We included chitin and Uif staining’s of stage 16 dumpy mutant embryos (Figure S10).
This data suggest that Pio mutant phenotype is due to a lack of Pio and Dumpy, which would support our model, of Pio and Dumpy protein interaction in the extracellular space of the tube lumen.
In wt embryos Pio is predominantly in the luminal chitin cable, in contrast in dumpy mutant embryos most Pio is predominantly not at the luminal chitin cable. Less luminal Pio staining in dumpy mutant embryos but Pio accumulation apically shows that Dumpy is required for luminal Pio release in stage 16 embryos. This supports our model that Pio and Dumpy interaction may link membrane and matrix and that this link reacts on mechanical stress during tube expansion by Np-mediated cleavage of Pio and its accompanied luminal release due to linked Dumpy.
12. Lines 374ff: the reduced dorsal trunk in Np mutants is not significant; the respective statement should be formulated carefully. If we believe the statistics (no significance), this would mean that attachment of the apical plasma membrane to the luminal chitin via Pio is needed to restrict axial extension; release of Pio is needed for differentiation (taenidia formation, luminal clearance) beyond morphogenesis.
We agree with the reviewer that the reduction of the dorsal trunks in Np mutant is statistically not significant. However, the mean value is clearly below that of WT. Therefore, we revised our statement as follow: “In Np mutant embryos, tracheal dorsal trunk length shows the tends to be reduced compared to wt embryos.” Further, the btlG4-driven UAS-Np overexpression of Np suggests strong Pio release from the apical membrane and therefore resembles the pio mutant tube length overexpansion (Figure 8A,B; Figure S13). Thus, our current observations indicate that Np-mediated Pio release at the cell membrane enables precise tube length elongation. We thank the referee for discussing that Pio is needed for taenidial fold formation which would fit to our findings in pio null mutant embryos. Pio mutant embryos show the appearance of taenidial folds in stage 16 embryos (airyscan) and stage 17 embryos (TEM images). However, TEM images also show chitin matrix reduction in pio mutant stage 17 embryos. Further, costainings of Pio with Crb and Uif, as well as co-stainings of mCherry::Pio with Dpy-GFP and cbp confirms that the Pio localize at the apical cell membrane where taenidial folds form in late stage 16 embryos. Thus, our observations suggest that Pio and Dumpy are required at the apical membrane and matrix to stabilize taenidial folds and tube lumen during 17. This also includes the Np-mediated Pio release at the apical cell membrane. As requested by the referee we summarized Pio function during late tracheal development in our simplified model (see Figure 9).
However, it is of note that Np-mediated Pio release increases at late stage 16 (Figure 5A, 6D; Figure S13) but is strongly reduced in stage 17 embryos. In contrast, thin taenidial fold are formed at late stage 16 and becomes thicker and form at fusion points during stage 17 and reach their most mature form when the intraluminal chitin cable is cleared (Öztürk-Colak et al., eLife, 2016). Thus, the pattern of Pio release and taenidial fold differentiation do not fully match. Moreover, in preliminary experiments we observe Pio antibody staining in stage 17 embryos at the apical cell membrane of dorsal trunks (data not shown). Furthermore, lumen clearance of Obst-A, Knk, Sepr and Verm are not affected in pio mutant embryos, but unknown luminal ECM contents remained (Figure 1D). Therefore, we will follow this very interesting idea in future experiments.
Nonetheless, we state in the results that Pio shedding is essential:
“Our data assumes that Np overexpression may enhance Pio shedding in stage 16 embryos, affecting the Pio-mediated ZP matrix function. Upon breathless (btl)-Gal4-mediated expression of UAS-Np in tracheal cells, we observed a high amount of Pio puncta across the entire tracheal tube lumen, specifically in stage 16 embryos but not in earlier stages (Figure S13). Consistently tracheal Np overexpression led to tube overexpansion in stage 16 embryos resembling the pio mutant phenotype (Figure 8A,B). Thus, Np-mediated Pio shedding controls Pio function.”
13. Why don't we see the apical Pio signal in Figure 4B?
The red arrowhead points to apical mCherry::Pio punctuate staining in the Figure 5B (before 4B) in the close up of the “bleached area” before bleaching and 56min post bleaching. However, in vivo bleaching experiments do not allow additional antibody stainings to detect precisely the apical cell membrane. Further, the Dpy::eYFP marks the tube lumen and the apical cell surface. The latter showed adjacent mCherry::Pio punctuate staining. However, due to bleaching Dpy signal was not detectable in the area.
14. The Strep signals in the merges in Figure 7C are not well visible.
We are not sure which Strep signal the reviewer is referring to in Figure 7C, which is now Figure 8C.
The top panel shows the Strep signal (right panel) overlapping with GFP in cells that do not express Np or human matriptase. Thus, the TGFB3 ZP domain is not cleaved, and the intracellular GFP and also the extracellular Strep signals are maintained and overlap. In contrast, when Np or human matriptase is added, the TGFB3 ZP domain is cleaved and only the intercellular GFP signal is retained, whereas the extracellular Strep signal is released from the cell surface. This explains why the Strep signal is barely detectable in the middle and lower panels of Figure 8C.
Reviewer #1 (Significance (Required)):
This work brings together several factors (Pio, Dp, Np, Wst etc) already known to be needed for tracheal morphogenesis and differentiation in the embryo of D. melanogaster. Having worked myself with some of these factors, however, I recognize that the interaction between these factors is novel and very exciting. The experiments strongly indicate a new mechanism of cellECM connection that seems to be conserved to some extent (as they provide preliminary data on an example from humans). By integrating the functions of different factors, the work provides ample opportunity for future projects to elucidate this mechanism in detail. Therefore, I expect that it will have a significant impact not only on the field of developmental cell biology but also, due to the conserved proteins involved (ZP proteins, Matriptase), on the field of cell biology of human diseases.
Reviewer #2
The figures are clear, and the questions well addressed. However, I find that some of the claims are not completely backed by the data presented and have some suggestions that will hopefully make some points clearer.
Major comments
1. In the abstract and at the end of the introduction the authors claim that they show that Pio, Dpy and Np support the balancing of mechanical stresses during tracheal tube elongation. However, this is not shown in this manuscript, where tension or mechanical stress were not measured and it is therefore speculative.
As requested by the reviewer, we deleted “support balancing of” at the final sentence of the Introduction. Please, note that we did not use the term balancing of mechanical stresses at the abstract.
However, we revised the abstract.
It has been shown previously that forces and mechanical tension rise when apical membrane expands and elastic extracellular matrix, which is anchored to the membrane balances theses forces (Dong et al., 2014). Furthermore, its has been shown that the gigantic and elastic Dumpy protein modulates mechanical tension (Wilkin et al., 2000). Thus, these previous publications state that mechanical tension rise at the apical cell membrane and matrix when tubes expand during stage 16 and that Dpy is part of that molecular process, which we included in the abstract as essential background information.
“The apical membrane is anchored to the apical extracellular matrix (aECM) and causes expansion forces that elongate the tracheal tubes. The aECM provides a mechanical tension that balances the resulting expansion forces, with Dumpy being an elastic molecule that modulates the mechanical stress on the matrix during tracheal tube expansion.”
Nonetheless, our results show that Np-mediated Pio cleavage increases during stage 16 as response to tube length expansion which is accompanied by forces as postulated by others (see above). We further observe that the membrane bulges and chitin matrix tear off, when Pio cleavage does not occur in Np mutant embryos. Our data further show that Pio and Dumpy interact and that Pio release is prevented in Dpy mutant embryos. Altogether this suggests that the Np-mediated Pio cleavage responds to tube expansion and requires Dpy for luminal Pio release.
We therefore claim in the final sentence of the introduction that “…ZP domain proteins Pio and, Dumpy, as well as the protease Np respond to mechanical stresses when tracheal tubes elongate”. The according changes are marked in red.
2. The authors state that all pio CRISPR/Cas9 generated mutants display identical tracheal phenotypes, however these data are not shown. Tracheal phenotypes, in particular DT phenotypes, of all mutants generated should be shown in supplementary materials.
As requested by the reviewer, we included the data in the supplement. The pio5M and pio11R alleles showed embryonic lethality and a 100% gas filling defect resembling the pio17C allele. Additionally, we extended the tracheal analysis with the pio5M allele and identified tube size defects, irregular pattern of taenidial folds and apical membrane deformation, altogether resembling the pio17C allele. These new data are shown in the supplement Figure S1.
We clarify this in the Results section as follows:
“The tracheal phenotypes of pio5m are shown in the supplement (Figure S1B-F). In all other Figures, we show images of the pio17c allele. “
3. At stage 16, pio null mutants display DT overelongation phenotypes (Figure 1). The authors should quantify this phenotype.
As requested by the reviewer, we quantified the DT overelongation phenotypes for pio5M (Figure S1). The quantification of pio17C was shown already in Figure 6B, now Figure 8B.
4. The authors analyse Pio distribution under tubular stress, using mega mutants and Chitinase overexpression. Pio localization changes in these genetic backgrounds and this is shown in Figure 2 only in a qualitative manner. The authors should measure Pio localization at the lumen and at the membrane and provide quantitative data.
As requested by the referee, we measured Pio localization recognized by the anti-Pio antibody at the lumen and at the membrane to provide quantitative data. These are shown in Figure 2E. All images were taken with a Zeiss Airyscan. For statistical analysis we used the profile tool of the Zeiss ZEN 2.3 black software. This tool allows the measurement and comparison of fluorescence pixel intensities of individual channels. We determined the fluorescent intensities profile across the tube to identify values at apical membrane and tube lumen at minimum 10 different position of DTs (metameres 5 to 6) of two distinct embryos for each genetic background. The maximum values of membranes versus tube lumen were set into ratio and compared between control, mega mutant and Cht2 overexpression. The control embryos showed a ration below 0.4, the Cht2 overexpression a ratio of 1.2 and mega mutants a ratio of about ~0.9. These quantitative data confirm the statement that Pio localization increases at and near the apical cell membrane with respect to the lumen in mega mutants and in Cht2 overexpression embryos.
5. Surprisingly and interestingly, wurst;pio transheterozygotes display very strong tracheal defects. The authors say they observe gas filling defects; however it is not clear from figure 2E if this indeed the case. From the panel in the figure, it looks like these embryos suffer from strong tracheal morphogenetic defects. It would be necessary to have a better analysis of these embryos. What is the penetrance of this phenotype. If this is 100% penetrant, one would expect it to be lethal. Therefore, double mutant balanced stocks are not viable? Having analyzed the phenotypes and confirmed which morphogenetic defects the transheterozygote embryos present, how does this genetic interaction fit with the model presented?
We are thankful to the reviewer for this interesting point of view suggesting that the wurst;pio embryos display tracheal morphogenetic defects. First, our data show that only 11.6% of the wurst;pio transheterozygous embryos completed gas filling and survived until adulthood. In contrast, 88.4% of transheterozygous wurst;pio mutant embryos did not complete gas filling which is now presented in Figure 3B. The corresponding quantifications is presented in Figure 3D. Importantly, the 88.4% wurst;pio transheterozygous embryos which show gas filling defects do not hatch as larvae and die.
As requested, we performed a better morphogenetic analysis, which is presented in Figure 3C.
Analysis of the gas filling defects with light microscopy were repeated with a better objective (Zeiss Apochromat 25x Gly; 0.8 NA). Indeed, this analysis revealed a strongly compromised tube lumen morphology with irregular tube lumen pattern as if tubes twist and bend. This tube lumen deformation was further confirmed with the confocal analysis of chitin staining (cbp). The tube lumen of stage 17 transheterozygous wurst;pio mutant embryos showed irregular lumen pattern with unusual twists and even partially collapsed tubes.
Furthermore, as asked by the referee, we generated the wurst,pio double mutation. All wurst,pio double mutant embryos lacked gas filling. In a more in-depth analysis of the tube lumen with a high-performance objective we could not identify any normal tube lumen in stage 17 embryos. Instead the double mutant embryos revealed completely collapsed tracheal tubes. This was confirmed by the chitin staining and confocal analysis. All new data are presented in the supplement.
As shown in our manuscript and in previous publications, neither pio nor wurst mutant embryos affect cell polarity or gross organization of the actin and tubulin cytoskeleton. However, we found that wurst mutant embryos showed irregular apical membrane expansion at tube lumen (Behr et al., 2007; legend Figure 4), irregular chitin fiber organization and to some extend collapsed tube lumen. In pio mutant embryos we found deformed apical membrane of DTs, irregular pattern of taenidial folds and to some extend collapsed tube lumen. Thus, the apical membrane is their common target of both proteins in late embryonic development, suggesting that pio functions provide stability and wurst functions the internalization of proteins at the apical membrane.
We discussed it as follows:
“Nevertheless, Pio and its endocytosis depend on its interaction with the chitin matrix and the Np-mediated cleavage. In stage 16 wurst and mega mutant embryos, we detect Pio antibody staining at the chitin cable, suggesting that Pio is cleaved and released into the dorsal trunk tube lumen. Also, the Cht2 overexpression did not prevent the luminal release of Pio. However, reduced wurst, mega function, and Cht2 overexpression caused an enrichment of punctuate Pio staining at the apical cell membrane and matrix (Figures 1,2). Although the three proteins are involved in different subcellular requirements, they all contribute to the determination of tube size by affecting either the apical cell membrane or the formation of a well-structured apical extracellular chitin matrix, indicating that changes at the apical cell membrane and matrix in stage 16 embryos affect the Pio pattern at the membrane. It also shows that local Pio linkages at the cell membrane and matrix are still cleaved by the Np function for luminal Pio release, which explains why those mutant embryos do not show pio mutant-like membrane deformations and Np-mutant-like bulges. This is in line with our observations that tracheal Pio overexpression cannot cause tube size defects as the Np function is sufficient to organize local Pio linkages at the membrane and matrix. Therefore, it is unlikely that tracheal tube length defects in wurst and mega mutants as well as in Cht2 misexpression embryos are caused by the apical Pio density enrichment.”
“Heterozygous and homozygous pio mutant embryos generally do not show tubal collapse. However, the loss of Pio and accompanying lack of Dpy secretion in stage 17 pio mutant embryos led to the loss of a Pio/Dpy matrix, impacting the late embryonic maturation and differentiation of a normal chitin matrix at the apical cell surface. TEM images reveal reduced dense chitin matrix material at taenidial folds and misarranged taenidial fold pattern (Figures 1; S2), suggesting impaired taenidial function prevents tube lumen from collapsing after tube protein clearance. Wurst knockdown and mutant embryos do not show general tube collapse, but luminal chitin fiber organization is disturbed in stage 17 embryos (Behr et al., 2007). Therefore, transheterozygous wurst;pio mutant embryos may combine both defects and suffer from maturation deficits of the chitin/ZP matrix at the apical cell surface and within the tube lumen, which finally causes a high number of embryos with incomplete gas filling due to tube collapse. These maturation deficits are even more dramatic in the wurst;pio double mutants, which show no gas filling.”
6. mCherry::Pio Dpy::eYFP time lapse analysis and FRAP experiments is very interesting. However, it is not clear to which degree bleaching occurs in the tracheal lumen. The authors claim that recovery is very fast and can be seen from minute 2, however, frame-by-frame analysis of Movie S2 does not show a clear different between luminal Pio from minute 0 to minute 2. Rough comparison with the luminal area surrounding the bleached area, does not show a clear difference in luminal Pio before and after photobleaching. To claim fast recovery of luminal Pio after photobleaching, the authors should quantify luminal Pio, before and after bleaching.
We agree with the reviewer and deleted “fast”. The Video2 shows intracellular mCherry::Pio recovery within 2min after photobleaching. The Video 2 shows extracellular (luminal) recovery within 6min after photobleaching, when first large mCherry::Pio puncta appear at the apical surface of the bleached area. Nonetheless, mCherry::Pio puncta appear in the lumen indicating recovery, whereas Dpy::eYFP did not.
We state this in the Results section as follows:
“In stage 16 embryos mCherry::Pio puncta reappeared in tracheal cells within 2 minutes of bleaching and in the tubular lumen within 6 minutes.”
In addition, in figure 4D, the normalized mCherry::Pio fluorescence in the graph what does it refer to? Intracellular Pio?
Figure 4D, now 5D, shows Western Blot signals. We guess that you refer to Figure 4B which is Figure 5B.
We are sorry for confusion and named it now Figure 5B’.
We stated in the Material section:
“The bleaching was performed with 405nm full laser power (50mW) at the ROI for 20 seconds. A Z-stack covering the whole depth of the tracheal tubes in the ROI were taken at each imaging step. “Fluorescence intensity in the bleached ROIs was measured after correction for embryonic movements using Fiji.”
Thus, to clarify this point, we added to the legends:
“Fluorescence intensities refer to the bleached ROIs as indicated with the frame in corresponding Movie S2 and was measured after correction for embryonic movements.”
7. When mCherry::Pio Dpy::eYFP time lapse analysis and FRAP experiments was done in an Np mutant background, the authors describe lack of Pio recovery within the lumen (Movie S3). However, when comparing control and Np mutant background embryos, Pio is not properly released into the lumen of Np mutants (as stated by the authors and seen by comparing movies S1 and S4). Furthermore, on minute 0 of the FRAP experiment in Np embryos, there is no detectable Pio in the DT lumen. Therefore, recovery was not expected in Np mutants and should not be claimed as a conclusion for this experiment.
We thank the reviewer for careful reading and apologize our wrong description. We changed it accordingly as follows:
“In contrast to the control, extracellular mCherry::Pio is not released into the tube lumen within 56 min after bleaching in Np mutant embryos (Figure 6C, Video S3).”
8. Brodu et al. (Dev Cell 2010) have shown that Pio is important for cytoskeletal modulation during tracheal maturation. Pio is important for non-centrosomal microtubule (MT) arrays anchored at the tracheal cell apical membranes. In addition, MT disruption in tracheal cells leads to lumen formation defects (Brodu et al., Dev Cell 2010). In the absence of Pio, the tracheal cytoskeleton is altered, and this could explain some of the results observed. Ideally, the work should be complemented with a basic cytoskeletal analysis, but if this is not possible, the authors should discuss some of the phenotypes in light of this Pio function.
Dear reviewer, this is a great idea. Therefore, we analyzed F-actin with Phalloidin and β tubulin (E7 antibody, DSHB) in the dorsal trunk cells of stage 16 control and pio mutant embryos. However, tracheal cells are tiny and only gross irregularities can be realized. So, confocal Z-stack analysis of the stainings did not show gross differences between control and pio mutant embryos. We observe the expected apical subcortical accumulation for the actin and tubulin cytoskeleton in dorsal trunk cells of pio stage 16 mutant embryos which also has been shown for wt embryos elsewhere. These new data are presented in the supplement Figure S7.
Minor comments
The model should not be in supplementary materials and should be moved to the main manuscript.
We thank the reviewer for this suggestion and moved the model to the main part – now Figure 9. As requested by the reviewer 1, we extended the model, showing the timing events of Pio function.
Throughout the manuscript embryonic stages are described using different nomenclature (stage X, stX and st X). Either way is correct, but the same nomenclature should be used throughout.
We apologize for the different nomenclature and use "stage X" in the manuscript and "stX" in the figures for space reasons. Legend 1 clarifies the abbreviation.
In Figure S1 B and C the authors should specify which pio allele is being analysed (as in Figure 7).
The same should be done in the text.
That's a fairly good point. To be clear from the beginning, we now state the following in the first paragraph of the results:
“The tracheal phenotypes of pio5m are shown in the supplement (Figure S1B-F). In the all other Figures, we show phenotypes of the pio17c allele.”
Line 131, it is not correct to say that WGA visualizes cell membranes. WGA marks/stains cell membranes.
Thanks for finding this mistake, it’s now corrected.
Line 165 "leads to excessive tube dilation and length expansion due to strongly reduced luminal chitin" is not correct. Chitin reduction leads to excessive tube dilation but not to length expansion, as reported in the papers cited at the end of the sentence.
Thanks very much for careful reading, we deleted “and length expansion” from the sentence.
Line 220-221, what do authors refer to as "stage 16 wt-like control embryos"?
Thanks for finding these mistakes. We corrected as follows:
“In stage 16 embryos mCherry::Pio puncta….”
Line 221, "some minutes" should be replaced by a specific number of minutes. According to Movie S2 reappearance of tracheal cell Pio happens from minute 16.
We agree with the reviewer to state the time when mCerry::Pio puncta reappear. We observe first large puncta within two minutes after bleaching in tracheal cells at the ROI (Video S2, lower cell row at the movie). We further observe the reappearance of first large puncta at the ROI within 6 minutes in the tracheal tube lumen.
We corrected it as follows: “In stage 16 embryos mCherry::Pio puncta reappeared in tracheal cells within 2 minutes of bleaching and in the tubular lumen within 6 minutes.”
Line 291 "time laps" should be lapse.
Thanks for finding the typo, it is corrected now.
Line 302, "Pio was not shedded into the lumen but remained at the cell" should be "Pio was not shed into the lumen but remained in the cell".
Thanks for finding the typo, it is corrected now.
Referees cross-commenting
I agree. Taken together, all the comments will improve the quality of the work and of a future manuscript. Also, everything seems quite doable and will not present any problems.
Reviewer #2 (Significance (Required)):
The findings shown in this manuscript shed light on the regulation of tubulogenesis by ZP proteins and how their interaction with the ECM can be regulated by proteolysis. It was known that Pio is involved in tracheal development, is secreted into the lumen, regulating tube elongation (Jaźwińska et al., Nat.Cell Biol., 2003) and anchoring MTs to the apical membrane during tubulogenesis (Brodu et al., Dev. Cell 2010). This work provides additional molecular insights into Pio dynamics and regulation during tube maturation.
This work will be of interest to a broad cell and developmental biology community as they provide a mechanistic advance in ZP proteins involved in morphogenesis. It is of specific interest to the specialized field of tubulogenesis and tracheal morphogenesis.
Field of expertise:
Drosophila, morphogenesis, tracheal tubulogenesis, cytoskeleton
Reviewer #3
Summary
In this manuscript, Drees and colleagues analysed, during the formation and growth of tubular systems, how cells combine forces at the cell membranes while maintaining tubular network integrity. A fundamental question is to understand how cells manage to integrate the axial forces to stabilise the cell membrane and the apical extracellular matrix (aECM).
To address this question, the authors study the formation of the tracheal system in Drosophila embryos, a well-established and detailed model system to investigate formation of tubular networks. In particular, they focused on the formation of the larger tube of the tracheal network, the dorsal trunk. The formation of this tube depends in part of axial extension along the anteroposterior axis.
They concentrated their work on the function of Piopio (Pio), a Zona-Pellucida (ZP)-domain protein. They showed that Pio together with the protease Notopleural (Np) contribute the sense and support mechanical stresses when tracheal tubes elongate, thus ensuring normal membrane -aECM morphology.
Major Comments
In a previous work, Drees et al. (PLOS Genetics 2019), showed the matriptase-prostasin proteolytic cascade (MPPC), is conserved and essential for both Drosophila ECM morphogenesis and physiology.
The functionally conserved components of the MPPC mediate cleavage of zona pellucidadomain (ZP-domain) proteins, which play crucial roles in organizing apical structures of the ECM in both vertebrates and invertebrates. They showed that ZP-proteins are molecular targets of the conserved MPPC and that cleavage within the ZP-domains is a conserved mechanism of ECM development and differentiation.
Here, Drees et al. investigate further how the coupling between membrane and matrix takes place to ensure proper tube growth.
Pio distribution and phenotypes
They first focused on the tracheal phenotypes observed in a pio null mutant context. So far, the only pio mutant characterised was a point mutation in the ZP domain. Using CRISPR/Cas9, they generated new alleles of pio which are lack of function alleles. In the context, Drees and colleagues observed over-elongated dorsal trunk tubes, with bulges appearing at stage 16 between the apical domain of tracheal cells and adjacent extra-luminal matrix.
Additionally, pio mutant embryos showed impaired tube lumen clearance of the some of the aECM components, which prevent gas-filling of the airways.
To detect Pio distribution, the authors used either anti-Pio antibody directed toward a short stretch with the Pio ZP domain or generated a CRISPR/Cas9 piomCherry::pio line.
1. The Pio antibody shows a strong luminal staining as already published. But the authors reported an apical membrane signal in tracheal cells. I find this apical membrane signal really difficult to observe in panel Figure 2B. The overlap between the Pio dots and the apical membrane labelled with Uif showed in Figure 2C can be due to the 3D projection. It is only when endocytosis is unpaired (Suppl Figure 2), that data are more convincing.
We thank the reviewer for this important point, we are sorry for the unconvincing presentation and for having the chance to improve it.
We show the 3D image of Pio puncta as voxels overlapping with Uif at the apical cell membrane. The amount of Pio voxels overlapping with the Uif marked apical cell membrane increased in mega mutant and due to tracheal Cht2 overexpression. This result was indicated by a representative region (frame) and white arrows and is shown now in Figure 2C. We further used orthogonal projections across the tracheal tube of the airyscan Z-stacks. Random usage confirmed that puncta of Pio antibody staining overlap with Uif at the tube lumen. We observed overlap in controls, but increasing overlap in mega mutant and Cht2 overexpressing embryos. This result is shown now in Figure 2E.
However, to overcome any misinterpretations of projections, we used single images of the original airyscan Z-stacks for co-localization analysis with the Zeiss ZEN software (black, 2.3, sp1). We used two available and independent standard methods to compare fluorescence pixel intensities of different channels namely the ZEN co-localization and the ZEN profile tool. Both are described in the Materials section.
With the co-localization tool we compared directly fluorescence pixel intensities of Pio and Uif. Highest overlap of the intensities, shown in the ZEN tool as third quadrant, were set to white for better visualization in the images. These new images are included as Figure 2D and show recurrent overlap of Pio and Uif antibody stainings (punctuate pattern) along the apical cell membrane at the dorsal trunk of stage 16 control embryos. This overlap pattern increased in mega mutant and Cht2 overexpression embryos.
A second approach for comparing fluorescence intensities is the ZEN “profile” tool. Drawing a line across the tube allowed us to compare peak fluorescence pixel intensities of the different channels at distinct regions, such as the apical cell membrane and the tube lumen including the cbp marked chitin cable. This tool detected overlap of peak fluorescence intensities of UIF and Pio antibody staining’s, confirming that Pio is located together with UIF at the apical membrane of dorsal trunk tracheal cells. These new intensity profiles and the corresponding images are presented in the supplement as Figure S4B-D. Quantifications of this method comparing the ration of Pio peak intensities between the apical cell membrane and the tube lumen are presented as Figure 2F (as requested by Reviewer 2).
2. When the author used their CRISPR/Cas9 piomCherry::pio line to characterise Pio distribution (Figure 4), Pio is localised at the apical plasma membrane before stage 16. Only at stage 16, Pio is detected within the lumen. This timing of Pio release in the lumen is critical for the model proposed by Drees at al. This is an important point to assess the difference between the use of the antibody (which mostly label the lumen) while piomCherry::pio line is mostly at the membrane.
We agree with the reviewer that the Pio antibody shows a different pattern within the tube lumen of earlier stages. The Pio antibody shows intense extracellular staining from early stage 12 onwards, presumably due to its early function at dorsal and ventral branches, as shown by Anna Jazwinska (Jazwinska et al., 2003). The intense luminal Pio antibody staining, predominantly at the chitin cable, persist until its disappearance due to airway protein clearance during stage 17. Unfortunately, this strong luminal Pio staining made it impossible to examine the Pio distribution pattern in more detail during stage 16. Nevertheless, Np overexpression experiments indicate that luminal Pio release occurs specifically in stage 16 embryos (Figure S13), which was tested with the Pio antibody, see results, second last paragraph:
“Our data assumes that Np overexpression may enhance Pio shedding in stage 16 embryos, affecting the Pio-mediated ZP matrix function. Upon breathless (btl)-Gal4-mediated expression of UAS-Np in tracheal cells, we observed a high amount of Pio puncta across the entire tracheal tube lumen, specifically in stage 16 embryos but not in earlier stages (Figure S13).”
We further agree with the reviewer that mCherry::Pio was used to characterize in vivo Pio distribution within the dorsal trunk cells and tube lumen during stage 16. The Figure 5A shows apical mCherry::Pio distribution pattern in early and late stage 16 embryos. Importantly, the appearance of luminal mCherry::Pio increased during stage 16 and mainly enriched at late stage 16. See Figure 5A, red arrowheads point to apical Pio and red arrows to luminal Pio staining.
Furthermore, as discussed above and shown by different ZEN tools, such as co-localization and fluorescence intensity profile tools, Pio antibody stainings revealed a punctuate pattern at the apical cell membrane of dorsal trunk cells in stage 16 embryos, which is reflected also by the appearance of apical mCherry::Pio puncta at the membrane surface. Additionally, we observed mCherry::Pio puncta also within the tube lumen (see the new Figures S4B & S8). Thus, subcellular Pio distribution at the apical cell membrane and lumen were observed for both, Pio antibody staining and mCherry::pio pattern.
Nonetheless, there is different luminal appearance between the Pio antibody staining and mCherry::Pio. Pio antibody detects a short stretch at the ZP domain and thus detects all possible Pio variants, uncleaved and cleaved. Due to early tracheal Pio function, Pio enriches within the tube lumen in an intense core-like structure, which is recognized by the Pio antibody and is comparable with the Dpy::eYFP pattern. Also mCherry::Pio labels all Pio variants, uncleaved and cleaved. The spatial temporal mCherry::Pio expression pattern (Figure S5) is comparable with the Pio antibody pattern and the staining at the membrane in stage 16 embryos. However, mCherry::Pio did not enrich in the lumen in a core-like structure, nonetheless, shows overlap with luminal Dpy::eYFP.
Jaswinska showed that Pio antibody staining is intracellular in the trachea of stage 11 pio2R-16
point mutation embryos (Jaswinska et al., 2003; Figure 2d). To understand more about the specificity of the antibody, we performed stainings in the null mutant embryos. In contrast, to the high number of intracellular Pio puncta in pio2R-16 point mutation embryos, Pio stainings were much more reduced in pio5m and pio17c mutants, but a low number of Pio puncta were still detectable in the embryos (Figure S1G,H). It is of note that also dpy mutants showed strongly reduced Pio antibody staining (Figure S10E). Thus, discussing underlying causes of enriched (Pio antibody) versus non-enriched (mCherry::Pio) luminal staining are speculative. However, observations by Jaswinska et al. (2003) and our new observations, investigating the Pio antibody stainings in pio null mutants, dpy mutants, eYFP::Dpy embryos and NP overexpression may hint to the possibility of cross-reactivity of the Pio antibody to other ZP domains which may intensify the appearance of luminal Pio antibody staining in control embryos.
Anyway, we clarify the difference in luminal Pio pattern in the discussion as follows:
“Indeed, the anti-Pio antibody, which detects all different Pio variants, showed a punctuate Pio pattern overlapping with the apical cell membrane markers Crb and Uif at the dorsal trunk cells of stage 16 embryos (Figure 2; Figure S3,S4). Additionally, Pio antibody also revealed early tracheal expression from embryonic stage 11 onwards, and due to Pio function in narrow dorsal and ventral branches, strong luminal Pio antibody staining is detectable from early stage 14 until stage 17, when airway protein clearance removes luminal contents. In the pio5m and pio17c mutants Pio stainings were strongly reduced although some puncta were still detectable in the trachea (Figure S1G,H). Similarly, Pio antibody staining is intracellular in the trachea of stage 11 pio2R-16 point mutation embryos (Jaźwińska et al., 2003). Interestingly, also dpy mutants showed strongly reduced and intracellular Pio antibody staining (Figure S10E).
We generated mCherry::Pio as a tool for in vivo Pio expression and localization pattern analysis during tube lumen length expansion. The mCherry::Pio resembled the Pio antibody expression pattern from early tracheal development onwards. However, luminal mCherry::Pio enrichment occurs specifically during stage 16, when tubes expand. The stage 16 embryos showed mCherry::Pio puncta accumulating apically in dorsal trunk cells. Moreover, mCherry::Pio puncta partially overlapped with Dpy::YFP and chitin at the taenidial folds, forming at apical cell membranes. Supported by several observations, such as antibody staining, Video monitoring, FRAP experiments, and Western Blot studies (Figures 4,5), these findings indicate that Pio may play a significant role at the apical cell membrane and matrix in dorsal trunk cells of stage 16 embryos.”
3. Another important point is to explain the discrepancy between the pio mutant alleles. The allele containing a point mutation in the ZP domain shows no over-elongated tubes (Dong et al. 2014, Jazwinska et al. 2003) while the lack of function alleles does.
The reviewer is correct that the pio2R-16 mutation shows only a disintegration phenotype whereas our pio null mutations show in addition tube length defects. However, Dong et al. showed significantly increased dorsal trunk length in shrub; pio2R-16 double mutant embryos when compared with shrub mutant embryos (Supplemental Figure S4A). Also, the shrub;dpyolvR double mutant embryos revealed increased tube length expansion when compared with shrub mutant embryos. Moreover, their quantifications show that the also dpyolvR mutant embryos revealed significantly increased tube expansion when compared with wt. Altogether these previous findings suggests that Pio and Dpy are involved in controlling tube length control during stage 16.
Furthermore, we generated three independent pio null mutation alleles, which lost all the essential Pio protein domains, and caused all embryonic lethality, gas-filling defects, branch disintegration phenotype and tube length defects (quantifications are shown in Figures 9 and S1). In addition, pio null mutations prevent Dpy::eYFP secretion. Thus, we are confident that the observed tube length defects as well as the air-filling defects are due to the loss of Pio, and in particular since these defects could be rescued by Pio Expression in the pio null mutation background, as shown in Figure 3B.
So, what could make the difference?
The described pio2R-16 mutation allele contains a X-ray induced single point mutation that led to an amino acid replacement (V159D) in the ZP domain. It is not clear how the amino acid exchange affects the protein and the ZP domain. It may hamper pio function and maybe this amino acid replacement is problematic for the early tracheal function but not during stage 16. As stated by Jazwinska et al. 2003 (Figure 2 legend), Pio antibody staining is intracellular in the mutants and extracellular in the trachea of wt at stage 13.
They further speculate that the mutant Pio protein may retain in the secretory pathway, but this is not confirmed with co-markers. As luminal Pio function is required to provide a barrier for autocellular AJ formation, this fails in pio2R-16 mutation. In contrast, it is still possible that Pio interacts and supports Dpy secretion in pio2R-16 mutation and additionally it is thinkable that intracellular Pio may reach to some extend the apical cell membrane in pio2R-16 mutation stage 16 and thus can support tube size control. But these assumptions are speculations.
Nevertheless, to clarify this point we explain the discrepancy between the pio2R-16 mutation and pio null mutations alleles as follows:
“Using CRISPR/Cas9, we generated three pio lack of function alleles (Figure S1A), all exhibiting embryonic lethality and identical tracheal mutant phenotypes. The tracheal phenotypes of pio5m are shown in the supplement (Figure S1B-F). In all other Figures, we show images of the pio17c allele. The pio17c and pio5m null mutant embryos revealed the dorsal and ventral branch disintegration phenotype known from a previously described pio2R-16 mutation allele which contains a X-ray induced single point mutation that led to an amino acid replacement (V159D) in the ZP domain (Jaźwińska et al., 2003). Additionally, the late stage 16 pio17c and pio5m null mutant embryos showed over-elongated tracheal dorsal trunk tubes (see below).”
4. A minor point, the author should provide hypothesis to explain why only the clearance of CBP, Obstructor-A and Knickkopf are affected in a pio mutant background and not Serpentine and Vermiform.
We thank the reviewer for careful reading and the comment on this point. We would be happy to see such a scenario which could give us a hind of Pio interaction partners at the chitinous matrix. However, we stated that luminal material, such as Obst-A and Knk are removed from the lumen (see Figure S5A). We further describe that in pio mutant embryos, luminal Serp and Verm staining appeared reduced but showed wt-like distribution (see Figure S6) in stage 16 embryos.
We do not show Serp and Verm in stage 17 embryos, but they are removed from the tube lumen (not shown). These data are received from immune-staining’s and confocal analysis. Nevertheless, we also state that pio mutant embryos revealed lumen clearance defects in TEM analysis, of undefined material in the tube lumen (see Figure 1D and Figure S2B).
To clarify this point we state in the results as follows:
“Fourth, ultrastructure TEM images revealed aECM remnants in the airway lumen of pio mutant stage 17 embryos, while control embryos cleared their airways (Figure S2B). Consistently, the in vivo analysis of airways in stage 17 pio mutant embryos revealed lack of tracheal air-filling (Figure 3B). The pan-tracheal expression of Pio in pio mutant embryos rescued the lack of gas filling (Figure 3B). Thus, TEM images suggest that pio mutant embryos showed impaired tube lumen clearance of aECM, which prevented subsequent airway gas-filling. “
And
“Also, the pio mutant embryos showed tracheal lumen clearance defects of chitin fibers in ultrastructure (TEM) analysis (Figures 1D, S2B). In contrast, confocal analysis revealed that wellknown chitin matrix proteins, such as Obstructor-A (Obst-A) and Knickkopf (Knk), are removed from the lumen of pio mutants (Figure S5A). These results suggest that the Pio function did not affect airway clearance of Obst-A and Knk and therefore did not play a central role in airway clearance like Wurst. Nevertheless, airway clearance defects observed in TEM images in pio null mutant embryos and, in addition, defective tube lumen morphology in wurst;pio transheterozygous mutant embryos explain the occurrence of airway gas filling defects.”
5. Pio and Dumpy
Dumpy (Dpy) is another ZP domain protein secreted by the tracheal cells and detected in the lumen. To follow Dpy distribution, Drees and colleagues used a Dpy::eYFP protein trap line, the same used in Dong et al. However, in this latter paper, Dong et al. stated, using a Crb staining, that Dpy is not at the apical cell surface but only in the lumen. However, Drees and colleagues reported (line 227 and Figure 4C) that Dpy appears both at the apical cell surface and in the lumen of the tracheal system. But they did not show a co-localisation with an apical marker.
Furthermore, in their previous work, (Drees et al. 2019) they called the apical staining a "peripheral shell" layer. In addition, in S2R+ cell culture, it is only when Pio and Dpy co-express that Dpy is detected at the cell membrane. The in vivo localisation of Dpy is an important point that needs to be clarified as it is of importance for the final model proposed Supp Figure 9. Drees at al. also performed FRAP experiments on Dpy::eYFP protein trap embryos. As excepted as already shown by Dong et al.
The referee is correct, we state “In stage 16 embryos Dpy::eYFP (Lye et al., 2014) appears at the tracheal apical cell surface and predominantly within the lumen (Figure 4C).” The corresponding Figure 4C reveals Dumpy::eYFP staining overlapping with chitin at two subcellular regions: Dpy is enriched as a core-like structure within the lumen overlapping with the chitin cable of the control embryos. Additionally, Dpy::eYFP overlaps with the chitin part that might be part of the apical cell surface. But this observation is hard to see in images in Figure 4C and we apologize it. We therefore repeated the Dpy::eYFP localization analysis and analyzed in more detail with the ZEN profile tools, which shows peak fluorescence pixel intensities of different channels and provides the possibility to prove, if they overlap in XY axis.
We asked first, if cbp (chitin) appears at the apical surface of dorsal trunk cells, when Pio becomes cleaved and released. In mid stage 16 embryos cbp staining appeared in the luminal chitin cable and additionally in a distinctive pattern, which fits to the pattern of taenidial folds that start to form. We therefore used the apical cell membrane marker Crumbs to co-stain cbp. Airycsan microscopy fluorescence intensity profile analysis and corresponding close ups images confirmed the overlap of Crb and cbp stainings at this distinctive pattern indicating this shows the chitin matrix at the apical cell surface (Figure S8A). But there was no overlap of cbp and Crb at the chitin cable structure. Thus, knowing the localization of the apical cell surface chitin matrix, we performed co-stainings of cbp with mCherry::Pio (RFP antibody). This revealed, as expected, overlap of cbp and RFP antibody staining at the apical cell surface chitin matrix (distinct pattern) and with the luminal chitin-cable (Figure S8B,C). Finally we repeated the stainings and analysis with cbp, mCherry::Pio (RFP antibody) and Dpy::eYFP (GFP antibody). First, these results revealed overlap of Dpy::eYFP and cbp at the apical cell surface and in the tube lumen (Figure S8D) and second, overlap of punctuate staining of Dpy::eYFP, cbp and mCherry::Pio at the apical cell surface chitin matrix and also at the luminal chitin cable (Figure S8E). Very obvious from images and Z-projection in Figure 4C is the lack of extracellular Dpy::eYFP staining in pio mutant embryos. Dpy::eYFP enriched intracellularly, and thus, the pio mutant caused Dpy::eYFP mis-expression fits well to our results from S2R+ cell culture. As the reviewer notes, it is only when Pio and Dpy co-express that Dpy is detected at the cell membrane. Altogether, Figure 4C, cell culture experiments and our new stainings support our model, that Pio and Dumpy interact and are co-secreted at the apical cell membrane/surface, where Np mediates Pio cleavage. As requested by reviewer 2, we moved the model to Figure 9. As requested by reviewer 1, we extended the model for timing events.
A minor point, the Dpy::eYFP protein trap line used in this study is not listed in the Materials and methods section of the supplementary data.
Thanks, we included it into the List of sources (Supplement). This YFP-trap line (called CPTI lines) was published by Claire M. Lye et al., Development, 141, 2014. We cite it in our manuscript.
6. The serine protease NP and Pio release.
Drees and colleagues have pervious shown, preforming in vitro studies, that protease Notopleural (Np) cleaves the Pio ZP domain (Drees at al. 2019). Here the authors went a step further in demonstrating that it is also true in vivo at stage 17. In addition, they showed that, in Np mutant embryos, mCherry::Pio is mostly detected within tracheal cells and the luminal staining is strongly reduced. In this mutant context, the authors conducted FRAP experiment on the mCherry::Pio signal even very weak in the lumen. They showed hardly no recovery after photobleaching.
In Drosophila S2 cells, Drees and colleagues showed that co-expression of the catalytically inactive NpS990A with mCherry::Pio in showed as a prominent signal the 90kDa mCherry::Pio variant in the cell lysate (Figure 5B), and live imaging revealed mCherry::Pio localisation at the cell surface (Figure S6B). However, in this inactive form context, a strong signal is also detected at 60kDA corresponding to a cleaved form of the Pio ZP domain (Figure 5B), and Pio localisation at the cell surface appears weaker than in controls. They authors did not consider that another protease could be at play.
On the other hand, in their previous work, Drees et al. identified a mutant form of Pio
(PioR196A) which is resistant to NP cleavage in vitro. It will be a step forward to establish by CRISPR/cas9, as the authors seems to be successful with this technique, a mutant line carrying this point mutation. It will be important to determine whether the observed phenotype resembles that of a mutant Np phenotype.
In their previous work (PLOS Genetics 2019), in Np mutant embryos, Drees et al. did not report "budge-like" deformations from stage 16 onwards leading to the detachment of the tracheal cell from their adjacent aECM. Either the alleles or the allelic combination is different between the two studies which could explain this difference, or it is a new phenotype that has not been previously described. In the latter case, it becomes important to quantify the proportion of segments showing these bubbles. Is this a rare phenotype to observe?
We thank the reviewer for the very interesting comments and the careful reading of our manuscripts and the very useful suggestions. We agree, we cannot exclude the possibility that another protease is involved in the cleavage of Pio. Therefore, we included this important point in the Discussion section as follows:
“Unknown proteases may likely be involved in Pio processing since cleaved mCherry::Pio is also detectable in inactive NpS990A cells.”
We think the generation of the pioR196A mutant to address Pio localization and tracheal phenotypes is a great idea, which we would like to address in future experiments. Unfortunately, the production of this fly line with such a specific point mutation at this position will take several months, not included the subsequent evaluation and phenotypic analysis of this fly line and mutants. Therefore, we apologize that we cannot pursue this question experimentally.
Nevertheless, mentioning the possibility and the requirement of such an experiment is important and we discuss it as follows:
“Previously we identified a mutation at the Pio ZP domain (R196A) resistant to NP cleavage in cell culture experiments (Drees et al., 2019). Establishing a corresponding mutant fly line would be essential in determining whether the observed phenotype resembles the phenotype of the Np mutant embryos.”
However, knowing that we are not able to provide a new mutant fly line to evaluate the formation of the dorsal tube when an NP non-cleavable form of Pio is expressed, we sought to use an alternative approach by overexpressing Np in the trachea with btl-Gal4. This shows a clear pairing of Np overexpression and Pio release specifically at stage 16 dorsal trunk and associated tube overexpansion.
Finally, the reviewer is correct, we did not mention the appearance of bulges in Np mutant tracheal dorsal trunk cells in our previous publication. We used that same Np alleles in 2019 and a closer look at the publication of 2019 likewise shows the appearance of bulges in Np mutant embryos, e.g. Figure 1B (red-dextran, left part of the tracheal lumen shows bulges) and even the Dpy::YFP matrix tear off at the site of bulges (Figure 4F’’, above the arrowhead). But we did not know at the time the link with Pio and Dumpy.
However, we agree, it is important to know more about the appearance of the phenotype by means of quantifications. The quantifications of bulges per dorsal trunk (n=16) is shown in Figure 7B.
7. Minor point: I don't understand what the authors are trying to show in supplementary Figure 8. Tracheal cells detach and are found in the lumen?
We are sorry for the unclear description in the legend. We corrected it as follows in the legend of Figure S12:
“This indicates disintegration of apical cell membrane at bulges and subsequent leaking of cellular content into the lumen.”
8. Np function conserved matriptase.
In this work, Drees and colleagues showed that Np controls in vivo the cleavage of the Pio ZP domain.
Dumpy and Piopio are not conserved in vertebrates but they both contain a ZP domain which is conserved. The authors tested if other ZP proteins can be cleaved by Np or the human homolog Matriptase. The authors tested in cell culture the ability of the type III Transforming growth factor-β receptor which contains a ZP domain to be cleaved either by Np or Matriptase. This could be a general mechanism that needs to be extended to other ZP domain proteins and that could be at play to structure the matrix and give it its physical properties.
However, as it is all speculative, I find the Discussion section related to these data, for too long and that does not help to understand better the work done in the formation of the tracheal tubes of the Drosophila embryo.
We show that Np mediates cleavage of the Pio ZP domain in vitro and in vivo in Drosophila embryos. We further showed that also the human matriptase was able to cleave the Pio ZP domain. To understand if this is a more general mechanism, we extended our studies with the human TβIII and its ZP domain. These data show that both Drosophila and human matriptases are able to cleave ZP domains of different proteins from different species. These data suggest that Matriptase-mediated ZP domain cleavage is not a Drosophila specific mechanism. We cannot follow the argumentation of the referee to state it all speculative. Nevertheless, we agree that it will need follow up studies to show that the mechanism is more general than two different species and ZP domain proteins. Anyway, as requested by the referee, we deleted the following sentences of the paragraph, since they are speculative in the context of our manuscript and do not directly describe a potential matriptase and ZP domain function:
“Matriptase degrades receptors and ECM in pulmonary fibrinogenesis in squamous cell carcinoma (Bardou et al., 2016; Martin and List, 2019). TβRIII is a membrane-bound proteoglycan that generates a soluble form upon shedding (López-Casillas et al., 1991), a potent neutralizing agent of TGF-β. Expression of the soluble TβRIII inhibits tumor growth due to the inhibition of angiogenesis (Bandyopadhyay et al., 2002). Idiopathic pulmonary fibrosis (IPF) is associated with a progressive loss of lung function due to fibroblast accumulation and relentless ECM deposition (King et al., 2011; Loomis-King et al., 2013). “
However, the comparisons of the tubular organ and the phenotypic expressions of the bulging membrane and the aortic aneurysm appear to us as an important element of the article. In both cases, cell membrane loses its integrity and can break in tubular networks. Thus, with our findings on the modification of extracellular ZP proteins, we offer a potential new molecular approach even for clinical investigation.
9.Minor points: Pio and cytoskeleton organisation.
Line 78-79, the authors wrongly quoted a work from Brodu et al. (2010). Pio does not anchor the microtubule severing enzyme Spastin. Instead, Spastin releases the microtubule-organising centre from its centrosomal location, then Pio contributes to its apical membrane anchoring. It can therefore be assumed that the organisation of the microtubule network is affected in a pio null mutant. In addition, ZP proteins have been shown to link the aECM to the actin cytoskeleton. Therefore, it would be interesting to look at the organisation of the actin and microtubule cytoskeletons in a pio mutant context in which enlarged apical cell surface area are observed.
We are very thankful for finding this mistake in the introduction. We corrected it as follows: “Further, Pio is involved in relocating microtubule organizing center components γ-TuRC (γtubulin and Grips; γ-tubulin ring proteins). This requires Spastin-mediated release from the centrosome and Pio-mediated γ-TuRC anchoring in the apical membrane.”
Studying cytoskeleton in pio mutant embryos is a helpful idea. Therefore, we analyzed F-actin with Phalloidin and β tubulin (E7 antibody, DSHB) in the dorsal trunk cells of stage 16 control and pio mutant embryos. However, tracheal cells are tiny and only gross changes can be realized. The confocal Z-stack analysis of the stainings did not show gross differences between control and pio mutant embryos. We observe the expected apical subcortical accumulation for the actin and tubulin cytoskeleton in dorsal trunk cells of pio stage 16 mutant embryos which also has been shown for wt embryos elsewhere. These new data are presented in the supplement Figure S7.
https://doi.org/10.7554/eLife.91079.sa2Referees cross-commenting
I have just read the comments of the other two reviewers, who like me are specialists in the formation of the tracheal system in the Drosophila embryo.
I find the comments very fair and balanced. They are in the same spirit as my comments and are very complementary. I hope that all our comments will be constructive for the authors and will improve the quality of their work.
Reviewer #3 (Significance (Required)):
Overall, the methodology is sound, the quality of the data is good and the paper is very well written. Authors combine in vivo, in vitro studies as well a cell culture approach. Using CRISPR/Cas9, they generated a large number of new tools allowing in vivo studies. Drees and colleagues generated new alleles of pio which are lack of function alleles. They described a new phenotype for pio mutant embryos, namely over-elongated tubes. But they authors do not comment on why these new alleles reveal a new phenotype. Furthermore, using their piomCherry::pio line, the authors state that Pio is localised to the plasma membrane. This location is very difficult to assess. Both new results require clarification.
The authors had already demonstrated that Np cleaves the ZP domain of Pio in vitro. Here they demonstrate this in vivo. It appears important to evaluate the formation of the dorsal tube when an NP non-cleavable form of Pio is expressed.
Finally, the model proposing a coupling between the extracellular matrix and the membrane of tracheal cells is very interesting. The demonstration that cleavage of Pio by Np could participate in this coupling is very interesting for those interested in the integration of mechanical stress and cellular deformation. However, such a model has already been discussed in Dong et al. (2014). In this article, Dong et al. proposed that a "coupling of the apical membrane and Dpy matrix core is essential for tube length regulation".
The audience for this article should be specialised and oriented towards basic research. It may be of interest to people working on tubular systems or working on ZP proteins.
My field of expertise is cell biology and developmental biology in Drosophila and formation of tubular networks.
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No external funding was received for this work.
Acknowledgements
We are very grateful to Markus Affolter, Bernard Moussian, and Stefan Luschnig for sharing generously antibodies. We thank Christian Wolf for critical reading and helpful discussion. We thank Jochen Rink and the department of Tissue Dynamics and Regeneration for generous support of our study. The FlyBase, NCBI, and SMART internet tools were always important sources. Stocks obtained from the Bloomington Drosophila Stock Center (NIH P40OD018537), the Vienna Drosophila Resource Center (VDRC), and from DGRC (NIH grant 2P40OD010949) were used in this study. MB expresses his gratitude to Johannes Kacza for his support with Imaris. We acknowledge support from Leipzig University for Open Access Publishing. We appreciate the University Leipzig BioImaging Core Facility (BCF equipment INST 268/230-1; INST 268/293-1; SFB-TR67; EFRE 100192650, 100195814, 100144684) for assistance.
Senior Editor
- Claude Desplan, New York University, United States
Reviewing Editor
- Elisabeth Knust, Max-Planck Institute of Molecular Cell Biology and Genetics, Germany
Version history
- Preprint posted: July 14, 2023 (view preprint)
- Received: July 18, 2023
- Accepted: September 20, 2023
- Version of Record published: October 24, 2023 (version 1)
Copyright
© 2023, Drees 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.
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