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Epidermal PAR-6 and PKC-3 are essential for larval development of C. elegans and organize non-centrosomal microtubules

  1. Victoria G Castiglioni
  2. Helena R Pires
  3. Rodrigo Rosas Bertolini
  4. Amalia Riga
  5. Jana Kerver
  6. Mike Boxem  Is a corresponding author
  1. Division of Developmental Biology, Institute of Biodynamics and Biocomplexity, Department of Biology, Faculty of Science, Utrecht University, Netherlands
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Cite this article as: eLife 2020;9:e62067 doi: 10.7554/eLife.62067

Abstract

The cortical polarity regulators PAR-6, PKC-3, and PAR-3 are essential for the polarization of a broad variety of cell types in multicellular animals. In C. elegans, the roles of the PAR proteins in embryonic development have been extensively studied, yet little is known about their functions during larval development. Using inducible protein degradation, we show that PAR-6 and PKC-3, but not PAR-3, are essential for postembryonic development. PAR-6 and PKC-3 are required in the epidermal epithelium for animal growth, molting, and the proper pattern of seam-cell divisions. Finally, we uncovered a novel role for PAR-6 in organizing non-centrosomal microtubule arrays in the epidermis. PAR-6 was required for the localization of the microtubule organizer NOCA-1/Ninein, and defects in a noca-1 mutant are highly similar to those caused by epidermal PAR-6 depletion. As NOCA-1 physically interacts with PAR-6, we propose that PAR-6 promotes non-centrosomal microtubule organization through localization of NOCA-1/Ninein.

Introduction

Polarity is a near universal property of cells that is essential for establishing proper cellular architecture and function. Epithelial cells – one of the major polarized animal cell types – polarize along an apical–basal axis and establish molecularly and functionally distinct apical, basal, and lateral membrane domains. The boundary between apical and lateral domains is marked by the presence of cell–cell junctions that provide adhesion between cells and prevent unwanted paracellular passage of molecules. The polarization of epithelial cells is orchestrated by conserved cortical polarity regulators that establish opposing membrane domains through mutually antagonistic interactions. In metazoans, the partitioning defective (PAR) proteins Par3, Par6, and atypical protein kinase C (aPKC) play a central role in the establishment of epithelial cell polarity. These highly conserved polarity regulators are essential determinants of apical domain identity, and are required for the positioning, maturation, and maintenance of apical cell junctions (Achilleos et al., 2010; Franz and Riechmann, 2010; Georgiou et al., 2008; Harris and Tepass, 2008; Harris and Peifer, 2005; Harris and Peifer, 2004; Hutterer et al., 2004; Izumi et al., 1998; Joberty et al., 2000; Leibfried et al., 2008; Lin et al., 2000; Totong et al., 2007; Wodarz et al., 2000; Yamanaka et al., 2001).

Par6 and Par3 are both PDZ domain-containing scaffold proteins that can interact with each other, with aPKC, and with numerous other proteins. Par6 and aPKC form a stable subcomplex by interacting through their PB1 domains (Hirano et al., 2005; Wilson et al., 2003). The association of Par6–aPKC with Par3 is dynamic. In C. elegans zygotes, PAR-6/PKC-3 shuttle between a kinase inactive complex with PAR-3 that promotes anterior segregation, and an active complex with the small GTPase CDC-42 (Aceto et al., 2006; Beers and Kemphues, 2006; Rodriguez et al., 2017; Wang et al., 2017). In epithelia, Par3 can promote the apical recruitment of Par6–aPKC (Franz and Riechmann, 2010; Harris and Peifer, 2005; Hutterer et al., 2004; Joberty et al., 2000; Lin et al., 2000; Wodarz et al., 2000). In mature epithelia, however, the bulk of Par3 segregates to the apical/lateral border, where it plays an essential role in the positioning and assembly of apical junctions (Achilleos et al., 2010; Georgiou et al., 2008; Harris and Tepass, 2008; Harris and Peifer, 2004; Izumi et al., 1998; Leibfried et al., 2008; Totong et al., 2007; Yamanaka et al., 2001). The segregation of Par3 from Par6–aPKC in epithelia depends on phosphorylation of Par3 by aPKC, and involves handoff of Par6–aPKC to Cdc42 and the epithelial-specific Crumbs polarity complex (Bilder et al., 2003; Harris and Peifer, 2005; Hong et al., 2003; Krahn et al., 2010; Morais-de-Sá et al., 2010; Nagai-Tamai et al., 2002; Nunes de Almeida et al., 2019; Walther and Pichaud, 2010).

In addition to interactions that mediate the subcellular localization of Par6–aPKC or Par3, both Par6 and Par3 can interact with effector proteins to connect cortical polarity with downstream pathways (McCaffrey and Macara, 2009). For example, Par3 modulates phospholipid levels by recruiting the lipid phosphatase PTEN to cell junctions (Feng et al., 2008; Martin-Belmonte et al., 2007; Pinal et al., 2006; von Stein et al., 2005), inhibits Rac activity by binding to and inactivating the RacGEF Tiam1 (Chen and Macara, 2005, p. 1; Mertens et al., 2005, p. 1), and mediates spindle positioning in Drosophila neuroblasts through recruitment of Inscuteable (Schober et al., 1999; Wodarz et al., 1999). For Par6, fewer downstream targets have been described. In mammals, Par6 can recruit the E3 ubiquitin ligase Smurf1 to promote degradation of the small GTPase RhoA, causing dissolution of tight junctions (Ozdamar et al., 2005; Sánchez and Barnett, 2012, p. 1; Wang et al., 2003). Par6 can also bind to the nucleotide exchange factor ECT2 to regulate epithelial polarization and control actin assembly at metaphase in dividing epithelial cells (Liu et al., 2006; Liu et al., 2004; Rosa et al., 2015). As high-throughput studies have identified multiple candidate binding partners that have not yet been investigated (Boxem et al., 2008; Brajenovic et al., 2004; Giot et al., 2003; Grossmann et al., 2015; Hein et al., 2015; Huttlin et al., 2015; Lenfant et al., 2010; Luck et al., 2020; Waaijers et al., 2016), additional interactions important for the functioning of Par6 and for linking cortical polarity to other processes involved in epithelial polarization likely remain to be discovered.

Despite the conserved requirements for Par6–aPKC and Par3 in epithelial cells there are important context and cell-type dependent differences in the functioning of these polarity proteins (Pickett et al., 2019; St Johnston, 2018). For example, in Drosophila, Bazooka (Par3) is not required for junction positioning or polarization of cells in the follicular epithelium (Pickett et al., 2019; Shahab et al., 2015), and in the adult Drosophila midgut, the canonical Par, Crumbs, and Scribble polarity modules are not essential for apical–basal polarity (Chen et al., 2018). In C. elegans, requirements for PAR-3 and PAR-6 in embryonic epithelia also vary. PAR-6 appears to be required for apical junction formation in all epithelia, including the epidermis, intestine, foregut, and pharyngeal arcade cells (Montoyo-Rosario et al., 2020; Totong et al., 2007; Von Stetina et al., 2017; Von Stetina and Mango, 2015). However, while arcade cells show a complete lack of polarization upon PAR-6 loss, foregut, intestinal, and epidermal epithelial cells still establish an apical domain (Totong et al., 2007; Von Stetina and Mango, 2015). PAR-3 is required for apical junction formation in embryonic intestinal and pharyngeal epithelia, but not in epidermal epithelial cells (Achilleos et al., 2010).

Studies of PAR-6, PKC-3, and PAR-3 in C. elegans have largely focused on embryonic tissues. Here, we make use of targeted protein degradation to investigate the role of PAR-6, PKC-3, and PAR-3 in larval epithelia of C. elegans. Ubiquitous depletion of PAR-6 and PKC-3, but not PAR-3, resulted in a larval growth arrest, demonstrating that these proteins are required for larval development. Through tissue-specific depletion, we identified an essential role for PAR-6 and PKC-3 in the C. elegans epidermis. Depletion in this tissue caused growth arrest, a failure to undergo molting, and severe defects in the division pattern of the epidermal seam cells. We also observed defects in the maintenance of apical cell junctions, and a failure to exclude LGL-1 from the apical domain. Finally, we identified a novel role for PAR-6 in organizing non-centrosomal microtubule arrays in the epidermis. Epidermal depletion of PAR-6 led to defects in the localization of the microtubule organizer NOCA-1/Ninein, as well as of the γ-tubulin ring complex component GIP-1, and of the sole Patronin/CAMSAP/Nezha homolog PTRN-1. Microtubule defects in a noca-1 mutant closely resembled those in PAR-6 depleted animals, including the loss of GIP-1 localization. As NOCA-1 physically interacts with PAR-6, we conclude that PAR-6 likely organizes non-centrosomal microtubule arrays through localization of NOCA-1.

Results

PAR-6 and PKC-3 are essential for larval development

To investigate the role of PAR-6, PKC-3, and PAR-3 in larval development, we made use of the auxin-inducible degradation (AID) system. The AID system enables targeted degradation of AID-degron tagged proteins through expression of the plant-derived auxin-dependent E3 ubiquitin ligase specificity factor TIR1 (Nishimura et al., 2009; Zhang et al., 2015; Figure 1A). Using CRISPR/Cas9, we inserted sequences encoding the AID-degron and the green fluorescent protein (GFP) into the endogenous par-6, pkc-3, and par-3 loci, such that all known isoforms of each protein are tagged (Figure 1B). PAR-6 was tagged at the shared C-terminus, and PKC-3 at the N-terminus. The par-3 locus encodes two groups of splice variants that use two alternative start sites, termed PAR-3L (for long) and PAR-3S (for short) (Achilleos et al., 2010; Li et al., 2010a). PAR-3L isoforms are expressed maternally and in larval stages, but not or at low levels in the embryo, while PAR-3S isoforms are expressed zygotically and in larval stages, but not maternally (Achilleos et al., 2010; Li et al., 2010a). To deplete both PAR-3L and PAR-3S isoforms, we inserted the GFP–AID tag at both start sites. To examine if the presence of the GFP–AID tags interfered with protein function, we examined the growth rates of the tagged strains. Homozygous animals were viable and showed the same growth rates as wild-type, indicating that the proteins are still functional (Figure 1C–E). Each protein was enriched at the apical membrane domain of epithelial tissues, including the pharynx, excretory canal, intestine and epidermis (Figure 1F,G). This matches previous observations in C. elegans larvae (Li et al., 2010a; Li et al., 2010b), and further indicates that the GFP–AID tag does not interfere with protein functioning. In the epidermis, we sometimes observed higher levels of fluorescence at the seam–seam junctions than at the seam–hyp7 junctions. Similar planar polarization of the PAR module was recently observed in the lateral epidermis of the elongating embryo (Gillard et al., 2019). However, in the larval epidermis we only observed planar enrichment in a subset of animals. Whether this pattern is functionally significant remains to be determined. Finally, to further investigate potential isoform-specific expression of PAR-3, we examined the expression of the PAR-3L isoforms alone during larval development (Figure 1—figure supplement 1). PAR-3L was expressed in the intestine, where it localized to the apical domain, but we observed little or no expression in the pharynx or epidermis (Figure 1—figure supplement 1D). Thus, PAR-3S appears to be the predominant isoform group in larval tissues.

Figure 1 with 1 supplement see all
PAR-6 and PKC-3 are essential for larval development.

(A) Overview of the AID system, which enables targeted degradation of AID-tagged proteins by the plant-derived E3 ubiquitin ligase specificity factor TIR1 upon addition of auxin. (B) Schematic representation of endogenous tagging of par-6, pkc-3, and par-3 loci with sequences encoding a green fluorescent protein (GFP) and auxin-inducible degradation degron (AID) tag. (C–E) Growth curves of N2, par-6::aid::gfp, gfp::aid::pkc-3, and gfp::aid::par-3 animals in absence (- aux) or presence (+ aux) of 4 mM auxin from hatching. Data show mean ± SD. Shading indicates the developmental stage of control animals. n = 6, 7, 8, and 8 for N2 - aux; 6, 7, 9, and 9 for N2 + aux; 7, 6, 9, and 9 for PAR-6 - aux; 8, 6, 7, and 9 for PAR-6 + aux; 22, 11, 10, and 14 for PKC-3 - aux; 19, 14, 9, and 10 for PKC-3 + aux; 10, 10, 10, and 10 for PAR-3 - aux, and 10, 10, 10, and 10 for PAR-3 + aux. (F) Graphical representation of larval epithelial tissues in C. elegans. Green indicates localization of PAR-6, PKC-3, and PAR-3. (G) Distribution of GFP::AID-tagged PAR-6, PKC-3, and PAR-3 in different larval tissues in absence (- auxin) or presence (+ auxin) of 4 mM auxin for 1 hr. Images are maximum intensity projections, and images of the pharynx are stitched montages. Dashed lines in - auxin panels outline pharynx (left panel), intestinal lumen (middle panel) or seam cells (right panel). White arrows point to the excretory canals.

Figure 1—source data 1

Source data for Figure 1.

This zip archive contains the data graphed in panels C–E, and the microscopy images shown in panel G. The raw graph data are in Microsoft Excel format, and the summary data and actual graph in Graphpad Prism format. The images are in TIFF and Adobe Photoshop format. The TIFF file is the unadjusted grayscale maximum intensity projection image generated in ImageJ from an image stack. The Photoshop file contains the original image with the adjustment layers used to arrive at the final image displayed in the main figure.

https://cdn.elifesciences.org/articles/62067/elife-62067-fig1-data1-v2.zip

To investigate the role of PAR-3, PAR-6 and PKC-3 in larval development we degraded each protein using a ubiquitously expressed TIR1 under the control of the eft-3 promoter (Zhang et al., 2015). We tested the efficiency of protein degradation by exposing synchronized L3 larvae to auxin and examining protein expression. Apical enrichment of PAR-3, PAR-6, and PKC-3 became indistinguishable from background fluorescence within 1 hr of exposure to 4 mM auxin in the pharynx, excretory canal, intestine, and epidermis (Figure 1G). To examine if the depletion of PAR-6, PKC-3, or PAR-3 affected larval development, we degraded each protein by addition of auxin at hatching and measured animal growth rates. Ubiquitous degradation of PAR-3 did not cause a defect in larval growth, and animals developed into morphologically normal and fertile adults (Figure 1E). To determine if the lack of phenotype was due to an inherent technical problem with our approach, we also depleted PAR-3 in the germline and early embryos using Pgld-1 driven TIR1. Addition of auxin to L4-stage animals resulted in 100% embryonic lethality in the next generation (n = 378), compared to 3.2% in non-auxin-treated controls (n = 591). Thus, the lack of a visible phenotype upon larval degradation indicates that the functions of PAR-3 are not essential for larval development. Alternatively, despite visual absence of GFP::AID::PAR-3, degradation may be incomplete, or animals may express unpredicted non-tagged larval-specific protein isoforms. In contrast to PAR-3, depletion of PAR-6 or PKC-3 from hatching caused a striking growth arrest with animals not developing beyond L1 size (Figure 1C,D). Thus, PAR-6 and PKC-3 are essential for early larval development, and we focused our further analysis on PAR-6 and PKC-3.

PAR-6 and PKC-3 are essential in the larval epidermis, but not in the intestine

We next wanted to determine which larval tissue or tissues are severely affected by the loss of PAR-6/PKC-3 and contribute to the growth arrest. We focused on the two major epithelial organs: the intestine and the epidermis. The intestine is an epithelial tube formed in embryogenesis by 20 cells, which do not divide during larval development. PAR-6 and PKC-3 are highly enriched at the apical luminal domain (Figure 2A). The epidermis consists of two cell types: hypodermal cells and seam cells. The syncytial hypodermal cell hyp7 covers most of the body. Embedded within hyp7 are two lateral rows of epithelial seam cells, which contribute multiple nuclei to hyp7 through asymmetric divisions in each larval stage. PAR-6 and PKC-3 localize to the apical domain of the seam cells and hyp7 and are enriched at the seam–seam and seam–hyp7 junctions (Figure 2B).

Figure 2 with 1 supplement see all
PAR-6 and PKC-3 are essential in the epidermis to support larval growth.

(A, B) Distribution of PAR-6::AID::GFP and GFP::AID::PKC-3 in the intestine (A) and epidermis (B) in absence (- auxin) or presence (+ auxin) of 1 mM auxin for 1 hr. Images are maximum intensity projections of the luminal domain for the intestine, and the apical domain for the epidermis. Drawings are schematic representation of the area imaged, with the localization of PAR-6 and PKC-3 indicated in green shades. Gray indicates absence of PAR-6 and PKC-3. Short colored lines indicate the area quantified in C–F. (C–F) Quantification of apical GFP fluorescence intensity at the intestinal lumen and the hyp7–seam-cell junction in par-6::aid::gfp and gfp::aid::pkc-3 animals in the absence (- aux) or presence (+ aux) of 1 mM auxin for 1 hr. Solid lines and shading represent mean ± SD. For the intestine, n = 10 animals for PAR-6 - aux, PAR-6 + aux, PKC-3 - aux, and PKC-3 + aux. For the epidermis, n = 8 animals for PAR-6 - aux, 6 for PAR-6 + aux, 5 for PKC-3 + aux, and 5 for PKC-3 - aux. (G–J) Growth curves of N2, par-6::aid::gfp, and gfp::aid::pkc-3 animals in absence (- aux) or presence (+ aux) of 4 mM auxin from hatching. Solid lines and shading represent mean ± SD. In G and H, degradation was induced in the intestine, and in I and J in the epidermis. In the intestine, n = 13, 10, 13, 14, and 12 for N2 - aux; 7, 7, 7, 5, and 9 for PAR-6 - aux; 6, 6, 6, 5, and 7 for PAR-6 + aux; 8, 7, 8, 4, and 9 for PKC-3 - aux; and 8, 7, 8, 8, and 8 for PKC-3 + aux. In the epidermis, n = 6, 7, 8, and 8 for N2 - aux; 6, 5, 11, and 8 for PAR-6 - aux; 5, 10, 8, and 9 for PAR-6 + aux; 7, 7, 10, and 8 for PKC-3 - aux; and 8, 7, 12, and 13 for PKC-3 + aux.

Figure 2—source data 1

Source data for Figure 2.

This zip archive contains the microscopy images shown in panel A and B, and the data graphed in panels C–J. The images are in TIFF and Adobe Photoshop format. The TIFF file is the unadjusted grayscale maximum intensity projection image generated in ImageJ from an image stack. The Photoshop file contains the original image with the adjustment layers used to arrive at the final image displayed in the main figure. The raw graph data are in Microsoft Excel format, and the summary data and actual graph in Graphpad Prism format.

https://cdn.elifesciences.org/articles/62067/elife-62067-fig2-data1-v2.zip

To enable tissue-specific depletion of PAR-6 and PKC-3, we generated single-copy integrant lines expressing TIR1 in the intestine and epidermal lineages, using the tissue-specific promoters Pelt-2 and Pwrt-2, respectively. In both tissues, protein depletion occurred within 1 hr of addition of 1 mM auxin (Figure 2A–F). To determine the contribution of the intestine and epidermis to the larval growth defects we observed above, we measured the growth rate of animals depleted of PAR-6 or PKC-3 from hatching in each tissue. Depletion of either protein from the intestine did not result in a growth delay or in obvious defects in morphology of the intestine (Figure 2G,H). These results are in contrast to the embryonic intestine, where PAR-6 has been shown to be required to maintain apical and junctional integrity (Sallee et al., 2020; Totong et al., 2007). Simultaneous depletion of PAR-6 and PKC-3 also did not result in a growth delay or visible abnormalities in the intestine (Figure 2—figure supplement 1A). These data indicate that PAR-6 and PKC-3 are not essential for the functioning and homeostasis of the larval intestine, though we cannot exclude that very low protein levels that we were not able to detect by fluorescence microscopy are sufficient in this tissue.

In contrast to the intestine, depletion of PAR-6 or PKC-3 from hatching in the epidermis caused an early larval growth arrest, as observed with ubiquitous degradation (Figure 2I,J). Thus, PAR-6 and PKC-3 play an essential role in the functioning and/or development of epidermal larval epithelia. We also noticed a small delay in growth in PAR-6::AID::GFP animals not exposed to auxin. One explanation is that TIR1 causes leaky degradation of PAR-6. However, no growth delay was observed in the absence of auxin when using ubiquitously expressed TIR1. Hence, the delayed growth may be due to other differences in genetic background. Finally, animals with ubiquitous PAR-6 or PKC-3 depletion have a more sick appearance than epidermal depleted animals, indicating that the functions of PAR-6 and PKC-3 are not limited to the epidermis. Indeed PAR-6 and PKC-3 were recently shown to be required for lumen formation in the excretory canal using a ZF degron-based protein degradation approach (Abrams and Nance, 2020).

Cell autonomous and non-autonomous roles for PAR-6 and PKC-3 in growth, molting, seam-cell divisions, and seam-cell morphology

To characterize the growth arrest of PAR-6 and PKC-3 depleted animals in more detail, we examined arrested animals by Nomarski differential interference contrast (DIC) microscopy. We observed incompletely released cuticles 30 hr past exposure to auxin, indicative of molting defects (Figure 3A). To examine molting progression in more detail, we used a transcriptional reporter expressing GFP from the mlt-10 promoter (Meli et al., 2010). mlt-10 expression cycles, increasing during molting and decreasing during the inter-molt. Upon epidermal degradation of PAR-6 from hatching, mlt-10 driven GFP levels remained low (Figure 3B,C), indicating that these animals fail to go through the L1/L2 molt.

Figure 3 with 2 supplements see all
PAR-6 and PKC-3 are required in the epidermis for molting and seam-cell development.

(A) DIC microscopy images of molting defects upon epidermal depletion of PAR-6 or PKC-3. Animals were grown in absence (-auxin) or presence (+auxin) of 1 mM auxin from hatching, and images were taken 30 hr after hatching. Dotted lines outline detached but unreleased cuticle in the pharynx and in the tail. Defects are observed in ~50% of the animals. Images are stitched montages. (B) Expression of the molting reporter Pmlt-10::gfp::pest in par-6::aid::gfp animals in the absence (- auxin) or presence (+ auxin) of 1 mM auxin from hatching. Images were taken at 22 hr of post-embryonic development. (C) Quantification of Pmlt-10::gfp::pest expression from 11 hr to 32 hr of post-embryonic development (mean fluorescence intensity ± SD) in par-6::aid::gfp animals in absence (- aux) or presence from hatching (+ aux) of 1 mM auxin from hatching. Measurements were done every hour. Each data point is an average of 3–12 measurements, with an average of 8 measurements per data point. (D) Examples of seam-cell division and morphology defects observed upon depletion of PAR-6::GFP::AID from hatching. Seam cells are visualized by expression of nuclear H2B::GFP and membrane-bound PH::GFP markers (Wildwater et al., 2011). Arrows indicate membrane protrusions and arrowhead indicates abnormal division plane orientation. Images are maximum intensity projections. (E) Seam-cell division pattern in par-6::aid::gfp animals in absence (control, blue) or presence (+ auxin, magenta) of 1 mM auxin. Auxin was added after hatching, before L2 divisions or before L3 divisions. For the control, n = 14, 75, 40, and 28 animals for the L1, L2, L3 and L4 divisions. For before L1, n = 17 animals for the L1 and 143 animals for the delayed L2 divisions. For before L2, n = 91 animals. For before L3, n = 40 animals.

Figure 3—source data 1

Source data for Figure 3.

This zip archive contains the microscopy images shown in panels A, B, and D, and the data graphed in panel C. The images are in Adobe Photoshop format. The Photoshop file contains the original unadjusted image with the adjustment layers used to arrive at the final image displayed in the main figure. The raw graph data are in Microsoft Excel format, and the summary data and actual graph in Graphpad Prism format.

https://cdn.elifesciences.org/articles/62067/elife-62067-fig3-data1-v2.zip

To determine if the growth arrest and molting defects reflect a complete developmental arrest, we next examined the effects of PAR-6 depletion on the stereotypical division pattern of the seam cells. In every larval stage, an asymmetric cell division creates a new seam-cell daughter and a cell that differentiates to form neurons or fuse with hyp7 (Chisholm and Hsiao, 2012a; Figure 3E, blue shaded lineage tree). In the second larval stage, a symmetric division precedes the asymmetric division to double the number of seam cells. Depletion of PAR-6 from hatching did not disrupt the L1 asymmetric divisions, indicating that these animals are not developmentally arrested. As an additional marker of L1 development, we followed outgrowth of the excretory canals. During L1 development, both the anterior and posterior canals elongate from their initial positions at hatching to their final positions near the head and tail (Figure 3—figure supplement 1ABuechner et al., 2020). Canal elongation still took place in PAR-6 depleted animals, with only a minor reduction in final length of the posterior canals (Figure 3—figure supplement 1B). Thus, PAR-6 depleted animals appear to continue the L1 developmental program, despite the lack of growth.

In contrast to L1 seam-cell divisions, the divisions that normally take place in the L2 stage were severely delayed (Figure 3D,E). At the time when control animals were already undergoing the L3 divisions, L2-stage divisions had still not taken place in PAR-6 depleted animals. Eventually, a next round of divisions did take place, but we observed various deviations from the normal L2 division pattern, including division failures and abnormal differentiation and fusion with hyp7. We did not observe any further divisions (Figure 3E). Following the delayed seam-cell divisions, we also observed numerous morphological abnormalities such as membrane protrusions, blebs, and abnormal division plane orientation (Figure 3D). Exposure of synchronized populations to auxin starting after the L1 or L2 divisions resulted in similar defects, indicating that seam-cell divisions require the functioning of PAR-6 throughout development (Figure 3E).

Expression of TIR1 under the wrt-2 promoter results in degradation of target proteins in both the syncytial hypodermis and the seam cells. As the hypodermis is essential for molting and is involved in the control of larval growth (Chisholm and Hsiao, 2012a; Chisholm and Xu, 2012b; Lažetić and Fay, 2017), it is possible that the seam-cell defects are a secondary consequence of defects in the hypodermis. To address this, we expressed an exogenous copy of par-6::mCherry lacking the degron sequence using the hypodermal-specific dpy-7 promoter (Gilleard et al., 1997). In combination with auxin-induced depletion of PAR-6::GFP::AID by Pwrt-2 driven TIR-1, this results in absence of PAR-6 only from the seam cells. Hypodermal-specific expression of par-6::mCherry rescued the molting defects and seam-cell division delay observed upon PAR-6 epidermal degradation, and partially rescued the growth arrest (Figure 3—figure supplement 1A–D). However, seam-cell morphology defects and the abnormal cell division plane were not restored (Figure 3—figure supplement 1C).

To confirm that abnormalities in the hypodermis can affect seam-cell divisions, we used a CRISPR-tagged NEKL-2::AID strain that arrests growth and molting upon auxin addition (Joseph et al., 2020). Indeed, NEKL-2 depletion caused defects in the morphology of seam cells, as well as a partially penetrant reduction in seam-cell divisions, confirming that abnormalities in the hypodermis can affect the seam cells (Figure 3—figure supplement 1E,F).

Taken together, our data show that PAR-6 is essential in the C. elegans hypodermis to support animal growth and molting. Whether the growth and molting phenotypes reflect separate functions of PAR-6, or are caused by the same underlying defect is difficult to establish, as molting is required for animal growth, but has also been reported to be governed by a size threshold (Chisholm and Hsiao, 2012a; Lažetić and Fay, 2017; Uppaluri and Brangwynne, 2015). The seam-cell division timing defects we observed appear to be a secondary consequence of hypodermal or molting defects. However, the fact that the growth arrest and seam abnormalities were not fully rescued by expression of PAR-6 in the hypodermis may indicate cell autonomous roles for PAR-6 in the seam, or alternatively that the Pdpy-7::par-6::mCherry rescue construct is not fully functional.

PAR-6 and PKC-3 mediate apical LGL-1 exclusion and promote junction integrity in the larval epidermis

As one of the major functions of the apical PAR complex is to mediate the exclusion of basolateral proteins from the apical domain, we next examined the effects of PKC-3 depletion on two key aPKC target genes: LGL-1/Lgl and PAR-1. Both proteins are direct aPKC targets in epithelia, and in the C. elegans zygote their anterior exclusion is mediated by PKC-3 (Beatty et al., 2010; Betschinger et al., 2003; Doerflinger et al., 2010; Hoege et al., 2010; Hurov et al., 2004; Motegi et al., 2011; Plant et al., 2003; Ramanujam et al., 2018; Yamanaka et al., 2003). For these experiments we made use of integrated LGL-1::GFP transgene (Waaijers et al., 2015) and an endogenously tagged PAR-1::GFP fusion.

Depletion of PKC-3 in the intestine did not result in apical invasion of LGL-1 (Figure 4—figure supplement 1A). In contrast, degradation of PKC-3 in the epidermis resulted in clear apical LGL-1 localization in the seam cells within 6 hr of auxin addition (Figure 4A,B). Degradation of PKC-3 in the epidermis did not result in apical PAR-1 localization (Figure 4C,D). Instead, prolonged depletion of PKC-3 for 24 hr resulted in fragmentation of the normally contiguous PAR-1 signal at cell junctions, which may reflect an indirect effect of PKC-3 on junction organization (Figure 4C). These results demonstrate that PKC-3 is necessary to maintain the basolateral localization of LGL-1 in the seam cells, but not the intestine. In contrast, the apical exclusion of PAR-1 is not solely mediated by aPKC.

Figure 4 with 2 supplements see all
PKC-3 excludes LGL-1 from the apical cortex and, together with PAR-6, regulates junctions.

(A, B) Distribution and quantification of LGL-1::GFP in the epidermis of lgl-1::gfp animals without auxin and in lgl-1::gfp; gfp::aid::pkc-3; Pwrt-2::tir-1::bfp animals in the presence of 4 mM auxin for 6 hr. Images are maximum intensity projections of the apical domain. Quantifications shows mean apical GFP fluorescence intensity ± SD at the hyp7–seam-cell junction, normalized to background intensity of each animal measured in the hypodermis. n = 7 animals for both conditions. Short colored lines in A indicate the area quantified in B. (C, D) Distribution and quantification of PAR-1::GFP in the epidermis in par-1::gfp animals without auxin and in par-1::gfp; gfp::aid::pkc-3; Pwrt-2::tir-1::bfp animals in the presence of 4 mM auxin for 6 or 24 hr. Images are maximum intensity projections of the apical domain. Quantifications show mean apical GFP fluorescence intensity ± SD at the hyp7–seam-cell junction, normalized to the background intensity of each animal measured in the hypodermis. n = 6 animals for both conditions. Short colored lines in C indicate the area quantified in D. (E, F) Junction organization visualized by DLG-1::mCherry expression in par-6::aid::gfp or gfp::aid::pkc-3 animals in the absence (- auxin) or presence (+ auxin) of 1 mM auxin for 6 (E) or 24 (F) hours. Images are maximum intensity projections of the junctional domain. (G) Graphical representation of junctional defects in the seam cells upon PAR-6 or PKC-3 degradation.

Figure 4—source data 1

Source data for Figure 4.

This zip archive contains the microscopy images shown in panels A, C, E, and F, and the data graphed in panels B, D. The images are in TIFF and Adobe Photoshop format. The TIFF file is the unadjusted grayscale maximum intensity projection image generated in ImageJ from an image stack. The Photoshop file contains the original image with the adjustment layers used to arrive at the final image displayed in the main figure. The raw graph data are in Microsoft Excel format, and the summary data and actual graph in Graphpad Prism format.

https://cdn.elifesciences.org/articles/62067/elife-62067-fig4-data1-v2.zip

In embryonic epithelia, PAR-6 and PKC-3 are essential for proper junction formation, with loss of either protein resulting in fragmented cell junctions (Montoyo-Rosario et al., 2020; Totong et al., 2007). To investigate the requirement for PAR-6 and PKC-3 in junction integrity in larval epithelia, we assessed the localization of an endogenous mCherry fusion of the junctional protein DLG-1/Discs large upon degradation of PAR-6 or PKC-3 from hatching. In control animals not exposed to auxin, DLG-1 displays the typical ladder-like intestinal junction pattern and forms a continuous apical belt around the seam cells (Figure 4—figure supplements 1B and E,F). Upon degradation of PAR-6 in the intestine, we did not observe junctional defects (Figure 4—figure supplement 1B). We also did not observe any changes to the DLG-1 localization pattern in the epidermis after 6 hr of PAR-6 or PKC-3 depletion (Figure 4E). However, after 24 hr of degradation, DLG-1 no longer localized in a uniform band around the seam cells but appeared fragmented, with aggregates of bright DLG-1 interspersed with areas lacking fluorescent signal (Figure 4F). We also observed fluorescent aggregates in the hypodermis (Figure 4F). Thus, as in the embryo, PAR-6 and PKC-3 are essential for junction integrity in the epidermis. The fact that junctional defects took 24 hr to develop, compared to 6 hr for LGL-1 mislocalization, points to an inherent stability of cell junctions.

Finally, we investigated the localization dependencies between PAR-6 and PKC-3. Several studies demonstrated that PAR-6 and PKC-3 co-localize throughout development, and are mutually dependent on each other for their asymmetric localization (Bossinger et al., 2001; Leung et al., 1999; McMahon et al., 2001; Nance et al., 2003; Nance and Priess, 2002; Tabuse et al., 1998; Totong et al., 2007). Moreover, binding of PAR-6 to PKC-3 is required for apical localization of PAR-6, including in larval epithelia (Li et al., 2010b). Degradation of PAR-6 in the intestine resulted in the rapid loss of PKC-3 from the apical membrane domain, and degradation of PKC-3 similarly disrupted PAR-6 localization (Figure 4—figure supplement 2A,B). When we followed the apical loss of PKC-3 in the intestine over time, we observed similar dynamics of PAR-6 depletion and PKC-3 loss (Video 1). In the epidermis, the levels of PAR-6 and PKC-3 are more difficult to determine accurately, due to the low levels of expression of these proteins and the aggregation due to the mCherry reporters used. Nevertheless, depletion of PAR-6 resulted in a loss of the junctional enrichment of PKC-3, and vice versa (Figure 4—figure supplement 2C–D). These disruptions occurred rapidly, within 1 hr of auxin addition. Our results thus confirm the interdependency between PAR-6 and PKC-3.

Video 1
Time-lapse imaging of PAR-6::AID::GFP and PKC-3::mCherry in animals expressing intestine-specific TIR1 upon addition of 1 mM Auxin.

PAR-6 and PKC-3 control the organization of non-centrosomal microtubule arrays in the hypodermis

The loss of PAR-6 or PKC-3 affected several epidermal processes in which cytoskeletal elements play important roles, including molting, seam-cell divisions, and maintaining proper seam-cell morphology. The PAR proteins play essential roles in organizing the actomyosin cytoskeleton and microtubules in different settings, including asymmetric cell division, neuronal differentiation, and epithelial polarization (Goldstein and Macara, 2007; Rodriguez-Boulan and Macara, 2014; St Johnston, 2018). We therefore investigated if PAR-6 degradation affects the organization of actin or microtubules in the epidermis. To assess the organization of the actin cytoskeleton we used an epidermal transgene expressing the actin-binding-domain of vab-10 fused to mCherry (Gally et al., 2009). We depleted PAR-6 from hatching and examined actin organization after 24 hr, when control larvae are in late L2 stage. Consistent with previous observations (Costa et al., 1997), we observed prominent circumferential actin bundles in hyp7, strong enrichment of actin along the hyp7–seam junctions, and largely anterior/posteriorly organized actin within the seam cells of control animals at this stage (Figure 5A). Upon PAR-6 depletion, actin organization appeared largely undisturbed in both the seam cells and hypodermis (Figure 5A), and actin bundles in hyp7 remained perpendicular to the seam cells (Figure 5B). These data indicate that PAR-6 does not play a major role in regulating the actin cytoskeleton in the C. elegans larval epidermis.

PAR-6 and PKC-3 control microtubule organization in the C.elegans epidermis.

(A) Actin organization visualized by the Plin-26::ABDvab-10::mCherry reporter in par-6::aid::gfp animals in absence (- auxin) or presence (+ auxin) of 1 mM auxin for 24 hr. Images are maximum intensity projections. (B) Quantification of actin bundle orientation. Angle is measured relative to the anterior (180°) – posterior (0°) axis. n = 100 bundles in five animals per condition. (C) Microtubule organization of the indicated genotypes visualized by MAPH-1.1::GFP in absence (- auxin) or presence (+ auxin) of 1 mM auxin for 24 hr. Images are maximum intensity projections. (D) Hypodermal microtubule bundle density. n = 13 animals for control, 15 for PAR-6 deg, 14 for PKC-3 deg, 13 for noca-1(ok3692), and 10 for PAR-6 deg. in noca-1(ok3692). Bars show mean ± SD. (E) Microtubule growth visualized by the plus end marker EBP-2::GFP in absence (- auxin) or presence (+ auxin) of 1 mM auxin for 1 hr. Images are a single frame or a 60 s maximum projection (one frame/second). To match the age of animals in (C), we depleted PAR-6 for 1 hr starting with 23 hr old L2 animals. (F) EBP-2 comet density in hyp7 in 24 hr old animals. n = 12 animals for control and PAR-6 deg, 8 for noca-1(ok3692), and 10 for PAR-6 deg in noca-1(ok3692). Auxin was present for 1 hr, from 23 to 24 hr of development. Bars show mean ± SD (G) Microtubule growth rate in 24 hr old animals. n > 400 comets in two animals (seam), four animals (hyp7 control), or five animals (hyp7 + auxin). Auxin was present for 1 hr, from 23 to 24 hr of development. Bars = mean ± SD (H) Quantification of microtubule growth orientation in hyp7 in 24 hr old animals. Auxin was present for 1 hr, from 23 to 24 hr of development. Vertical axis: left/right orientation; horizontal axis: anterior/posterior orientation. n = 150 comets. Bars = mean ± SD. Tests of significance: Tukey’s multiple comparisons test for D, and Dunn’s multiple comparisons test for F and G. ns = not significant.

Figure 5—source data 1

Source data for Figure 5.

This zip archive contains the microscopy images shown in panels A, C, and E, and the data graphed in panels B, D, F, G, and H. The images are in TIFF and Adobe Photoshop format. The TIFF file is the unadjusted grayscale maximum intensity projection image generated in ImageJ from an image stack. The Photoshop file contains the original image with the adjustment layers used to arrive at the final image displayed in the main figure. The raw graph data are in Microsoft Excel format, and the summary data and actual graph in Graphpad Prism format.

https://cdn.elifesciences.org/articles/62067/elife-62067-fig5-data1-v2.zip

We next inspected the organization of the microtubule cytoskeleton using an endogenously GFP tagged variant of the microtubule-binding protein MAPH-1.1 (Waaijers et al., 2016). We degraded PAR-6 in the epidermis from hatching and assessed the organization of epidermal microtubule arrays. In control animals, we observed highly ordered circumferential microtubule bundles in the dorsal and ventral sections of hyp7 underlying the muscle quadrants, and a microtubule meshwork in the lateral sections of hyp7 abutting the seam cells, as previously reported (Chuang et al., 2016; Costa et al., 1997; Taffoni et al., 2020; Wang et al., 2015; Figure 5C). In the seam cells the microtubule network was less well defined but also forms a meshwork (Figure 5C). In PAR-6 depleted animals, after 24 hr of development we observed a significant reduction in the density of circumferential microtubule bundles in the hypodermis (Figure 5C,D). Epidermal depletion of PKC-3 resulted in similar defects (Figure 5C,D). To understand the cause of the reduced microtubule density, we investigated microtubule dynamics using an endogenous fusion of the microtubule plus-end tracking protein EBP-2EB1 to GFP (Videos 2 and 3). In control animals, EB1 comets moved predominantly in a circumferential direction, consistent with the organization of microtubule bundles in the epidermis, and both comet density and growth rates matched previous reports (Figure 5E–H; Chuang et al., 2016; Taffoni et al., 2020; Wang et al., 2015). Already within 1 hr of inducing depletion of PAR-6, we observed reduced microtubule dynamics (Figure 5E–G). The density of growing MTs was reduced by 56% (Figure 5F), and microtubule growth rate was reduced by 14% in hyp7 and by 16% in the seam cells (Figure 5G). These results suggest that the reduced density of microtubule bundles upon depletion of PAR-6 is the result of reduced growth of microtubules. We also observed a defect in the directionality of microtubule growth. While 54% of the comets in control animals travel perpendicular to the seam cells (70–110°), this number is reduced to 24% upon PAR-6 degradation (Figure 5H), consistent with the defects in organization observed with GFP::MAPH-1.1.

Video 2
Time-lapse imaging of EBP-2::GFP in control animals and PAR-6 depleted animals.

Freeze frame circles EBP-2 comets as an example of quantification. Final frames show a time projection as displayed in Figure 5E.

Video 3
Dual-color time-lapse imaging of EBP-2::mKate2 and MAPH-1.1::GFP.

New microtubules largely grow along existing bundles.

PAR-6 controls microtubule organization through its interaction partner NOCA-1/Ninein and the γ-tubulin ring complex

Two large-scale protein–protein interaction mapping studies in C. elegans had identified the microtubule-organizing protein NOCA-1 as an interaction partner of PAR-6 (Boxem et al., 2008; Lenfant et al., 2010). Affinity purification experiments showed that PAR-6 interacts with NOCA-1 through its PDZ domain (Lenfant et al., 2010), and we were able to confirm the PAR-6 PDZ interaction with NOCA-1 by yeast two-hybrid (Figure 6—figure supplement 1). NOCA-1 functions together with γ-tubulin to assemble non-centrosomal microtubule arrays in multiple tissues, including the epidermis, and is thought to be a functional homolog of the vertebrate microtubule organizer Ninein (Green et al., 2011; Wang et al., 2015). NOCA-1 localizes to the apical cortex in seam cells, similar to the localization of PAR-6 (Figures 1G and 6A), but the mechanisms that mediate apical localization of NOCA-1 are currently not known. The physical interaction between PAR-6 and NOCA-1 prompted us to investigate if PAR-6 regulates non-centrosomal microtubule arrays through NOCA-1.

Figure 6 with 1 supplement see all
PAR-6 promotes the localization of its binding partner NOCA-1, as well as of GIP-1 and PTRN-1.

(A, B) Distribution and quantification of NOCA-1de::GFP in the epidermis of noca-1de::gfp animals without auxin, and noca-1de::gfp; par-6::aid::gfp; Pwrt-2::tir-1::bfp animals in the presence of 4 mM auxin for 6 hr. n = 9 animals for Control, and 10 for PAR-6 epid. deg. Short colored lines in A indicate the area quantified in B. (C, D) Distribution and quantification of GIP-1::RFP in the epidermis of gip-1::rfp animals without auxin, and gip-1::rfp; par-6::aid::gfp; Pwrt-2::tir-1::bfp animals in the presence of 4 mM auxin for 6 hr. n = 6 for Control and 6 for PAR-6 epid. deg. Short colored lines in C indicate the area quantified in D. (E, F) Distribution and quantification of PTRN-1::GFP in the hyp7 and seam cells of ptrn-1::gfp animals without auxin, and ptrn-1::gfp; par-6::aid::gfp; Pwrt-2::tir-1::bfp animals in the presence of 4 mM auxin for 6 hr. n = 10 for Control and 10 for PAR-6 deg. Short colored lines in E indicate the area quantified in F. (G, H) NOCA-1de::GFP in the epidermis of noca-1de::gfp animals without auxin, and noca-1de::gfp; gfp::aid::pkc-3; Pwrt-2::tir-1::bfp animals in the presence of 4 mM auxin for 6 hr. n = 10 for Control and 10 for PKC-3 epid. deg. Short colored lines in G indicate the area quantified in H. (I, J) Distribution and quantification of GIP-1::RFP in the epidermis of gip-1::rfp animals and gip-1::rfp; noca-1(ok3692). n = 6 for Control and 6 for noca-1(ok3692). Short colored lines in I indicate the area quantified in J. All images are maximum intensity projections of the apical domain. Quantifications in B, D, H and J show mean apical GFP fluorescence intensity ± SD at the hyp7-seam-cell junction (indicated by colored lines), normalized to background intensity of each animal measured in the hypodermis. Quantification in F shows mean PTRN-1::GFP puncta density ± SD. Tests of significance: unpaired t-test for F.

Figure 6—source data 1

Source data for Figure 6.

This zip archive contains the microscopy images shown in panels A, C, E, G, and I, and the data graphed in panels B, D, F, H, and J. The images are in TIFF and Adobe Photoshop format. The TIFF file is the unadjusted grayscale maximum intensity projection image generated in ImageJ from an image stack. The Photoshop file contains the original image with the adjustment layers used to arrive at the final image displayed in the main figure. The raw graph data are in Microsoft Excel format, and the summary data and actual graph in Graphpad Prism format.

https://cdn.elifesciences.org/articles/62067/elife-62067-fig6-data1-v2.zip

We first examined the effect of epidermal PAR-6 depletion on the localization of NOCA-1. To visualize NOCA-1 we made use of an existing transgenic line that expresses the epidermal-specific NOCA-1d and e isoforms fused to GFP from their endogenous promoter (Wang et al., 2015). In untreated control animals, we observed punctate localization of NOCA-1 in the epidermis, mostly clustered at the seam–seam and seam–hyp7 junctions, as previously observed (Figure 6A; Wang et al., 2015). Addition of auxin to induce epidermal PAR-6 degradation led to a 61% reduction in junctional levels of NOCA-1 within 6 hr (Figure 6A,B). Depletion of PKC-3 resulted in a similar reduction in NOCA-1, (Figure 6G,H). These results demonstrate that PAR-6 and PKC-3 promote the apical localization of NOCA-1. Because of the physical interaction between PAR-6 and NOCA-1, we hypothesize that the loss of PKC-3 indirectly affects NOCA-1 through loss of PAR-6 localization.

NOCA-1 was reported to work together with γ-tubulin and redundantly with Patronin/PTRN-1 in controlling circumferential microtubule bundle organization in the hypodermis (Wang et al., 2015). We therefore examined the effect of PAR-6 depletion on the localization of PTRN-1 and GIP-1, a core component of the γ-tubulin ring complex (γ-TuRC) required to localize other γ-TuRC components to the apical non-centrosomal microtubule-organizing center (ncMTOC) in the embryonic intestine (Sallee et al., 2018). To visualize PTRN-1 and GIP-1 we used endogenous PTRN-1::GFP and RFP::GIP-1 fusion proteins. GIP-1 localized in a punctate pattern at the seam–seam and seam–hyp7 junctions, similar to NOCA-1 (Figure 6C; Sallee et al., 2018; Wang et al., 2015). PTRN-1 also localized in a punctate pattern, but dispersed through the epidermis and lacking the junctional enrichment seen for NOCA-1 and GIP-1 (Figure 6E; Wang et al., 2015). Upon PAR-6 degradation, junctional GIP-1 levels were strongly reduced (Figure 6C,D), similarly to NOCA-1. We also observed that PAR-6 depletion resulted in a decrease in the number of PTRN-1 puncta in the epidermis (Figure 6E,F). As NOCA-1 is a direct interaction partner of PAR-6, we examined if the loss of GIP-1 is due to the loss of NOCA-1 localization. Indeed, in a noca-1(ok3692) deletion mutation GIP-1 levels were significantly reduced (Figure 6I,J), suggesting that NOCA-1 acts upstream of GIP-1 in the C. elegans larval epidermis.

Finally, we examined if the failure to properly localize NOCA-1 could explain the microtubule defects we observed upon PAR-6 depletion. We determined microtubule bundle density, EB1 comet density, and microtubule growth rate in noca-1(ok3692) animals. In the noca-1(ok3692) deletion mutant, we observed a significant reduction in the density of circumferential microtubule bundles in the hypodermis (Figure 5C,D). As reported in a previous study (Wang et al., 2015), we also observed reduced microtubule dynamics, with the density of growing MTs reduced by 65%, and microtubule growth rates reduced by 65% (Figure 5E,G). These values are all very similar to those we observed upon PAR-6 depletion. To further determine if par-6 and noca-1 act in a linear pathway, we degraded PAR-6 in noca-1(ok3692) mutant animals. Both microtubule bundle density and microtubule dynamics were reduced to a similar extent as in PAR-6 depleted or noca-1 mutant animals alone (Figure 5C–F). These data are consistent with a model in which the microtubule defects caused by PAR-6 depletion are a result of the requirement of PAR-6 in localizing NOCA-1. The effects on PTRN-1 may be a secondary consequence of microtubule defects caused by NOCA-1 loss.

Discussion

Par6 and aPKC are essential for apical–basal polarization across animal species. Most studies of the apical PAR proteins in C. elegans have focused on embryonic tissues, and their roles during postembryonic development remain unclear. Here, we used inducible protein degradation to identify essential roles for PAR-6 and PKC-3 in larval development. The depletion of PAR-6 or PKC-3 caused several developmental defects. When depleted from hatching, the first abnormality we observed is a severe growth defect, with animals barely increasing in length beyond their size at hatching. Surprisingly, the growth arrest is not the result of a complete developmental arrest, as the L1-stage seam-cell divisions take place at their normal time of development and the excretory canals still elongate. The next developmental defect to become apparent was the failure to complete the L1/L2 molt, indicated by incompletely released cuticles and the lack of expression of the molting marker Pmlt-10::gfp::pest. Both the growth defect and molting defect were rescued by expressing a non-degradable copy of PAR-6 in the hypodermis, demonstrating that PAR-6 plays an essential role in this cell type required for organismal growth and molting. Determining the cause or causes underlying the molting and growth defects will require further study. Following the L1/L2 molt we observed severe defects in the normally stereotypical pattern of seam-cell divisions, including a long delay before the next round of cell divisions. However, these defects are likely in large part a secondary consequence of the growth and molting defect, as expression of PAR-6 in the hypodermis was sufficient to restore the timing of divisions, and inducing growth and molting defects by degradation of NEKL-2 similarly induced seam-cell division defects.

In addition to these developmental defects we found that PAR-6 regulates the assembly of microtubule bundles through its interaction partner NOCA-1/Ninein. Already within 1 hr of inducing PAR-6 degradation, we observed reduced numbers of growing microtubules. This makes it unlikely that the microtubule defects are a secondary consequence of the growth or molting defects. Vice versa, a noca-1 mutant displaying very similar microtubule defects does not display the developmental defects observed upon PAR-6 depletion. Hence, the microtubule defects are most likely independent of the growth and molting defects. The finding that depletion of PAR-6 or PKC-3 causes multiple defects likely reflects the versatility of the PAR polarity in coordinating polarity with other cellular pathways.

An essential role in postembryonic development for PAR-6 or PKC-3 has not been described. Depletion of par-6, pkc-3, or par-3 by RNAi in larval stages caused defects in polarization of spermathecal cells and in ovulation, but not in larval development (Aono et al., 2004). Similar results were recently observed using a temperature sensitive pkc-3 allele grown at non-permissive temperature (Montoyo-Rosario et al., 2020). More severe phenotypes were observed in hatching progeny (escapers) of par-6, pkc-3, or par-3 RNAi-treated mothers, which showed partially penetrant defects in outgrowth of vulval precursor and seam cells, migrations of neuroblasts and axons, and the development of the somatic gonad (Welchman et al., 2007). The lack of a growth arrest phenotype in these studies presumably reflects incomplete gene inactivation.

Auxin-inducible protein depletion of PAR proteins

The auxin-inducible degradation approach allowed us to bypass embryonic requirements and examine the roles of PAR-6, PKC-3, and PAR-3 in specific epithelial tissues during larval development. Despite these advantages, one drawback of all protein degradation approaches is that it remains difficult to draw conclusions from negative results. Although we tagged all known PAR-3 protein isoforms and observed efficient protein depletion, ubiquitous depletion of PAR-3 did not cause obvious defects in larvae. Thus, PAR-3 may not be essential in larval tissues, and the L1 lethality previously observed for the par-3(tm2010) null allele (Achilleos et al., 2010; Li et al., 2010a) may be the consequence of defects in embryonic development. In support of this interpretation, PAR-3 was recently found to be largely dispensable for lumen extension of the excretory canals, in contrast to PAR-6 and PKC-3 (Abrams and Nance, 2020). Nevertheless, it is possible that very low levels of PAR-3 are sufficient for its functioning, or that unpredicted splicing events cause the expression of non-degron tagged PAR-3 isoforms. One approach to counteract the latter possibility would be to replace the endogenous gene with a re-engineered copy that is unlikely to express alternative splice variants, for example by replacing natural introns with artificial ones and removing internal promoters. However, removing this level of regulation and expressing only one isoform may affect the functioning of par-3 and cause unintended side effects. We also did not detect phenotypes upon depletion of PAR-6 or PKC-3 in the larval intestine. Similar caveats as for PAR-3 depletion apply here, though PAR-6 depletion did lead to complete loss of PKC-3 from the apical domain, and epidermal depletion caused severe phenotypes. These observations make it less likely that the lack of a phenotype is due to the expression of unknown isoforms. PAR-6 and PKC-3 are likely to play non-essential or redundant roles in the intestine, as a previous study found that PAR-6 contributes to endosome positioning in this tissue (Winter et al., 2012).

Another advantage of inducible protein degradation is that the time it takes for defects to appear can give information on the role of the targeted protein in a particular process or structure. Processes highly dependent on the degraded protein likely show defects sooner after auxin addition than processes in which low levels of the protein suffice. Similarly, the speed at which molecular assemblies display defects will depend on whether the targeted protein is a core component of the assembly, or only required for its initial formation. For example, upon depletion of PAR-6, we observed defects in microtubule growth within 1 hr using a plus-end-binding marker, while defects in circumferential microtubule bundles visualized with a microtubule-binding protein took ~24 hr to become apparent. This indicates that PAR-6 regulates the formation of new microtubules but is not essential for the maintenance of already existing microtubule bundles. Similarly, junctional defects in the epidermis appeared ~24 after PAR-6 or PKC-3 depletion started, indicating that PAR-6 and PKC-3 are important for the assembly of new junctions, but are not integral components.

Roles of PAR-6 and PKC-3 in junction formation and cell polarity

Depletion of PAR-6 and PKC-3 in the epidermis resulted in a fragmented appearance of the hyp7–seam and seam–seam junctions (Figure 4E,F), similar to previous observations in embryonic epithelia (Montoyo-Rosario et al., 2020; Totong et al., 2007; Von Stetina et al., 2017; Von Stetina and Mango, 2015). Localization of PAR-6 and PKC-3 were mutually dependent in both the epidermis and intestine. This result was not surprising, as Par6 and aPKC act as a dimer and have been shown to be mutually dependent in other C. elegans tissues (Hung and Kemphues, 1999; Li et al., 2010b; Nance et al., 2003; Tabuse et al., 1998; Totong et al., 2007). We also examined if PKC-3 functions to exclude the basolateral polarity proteins PAR-1 and LGL-1 from the apical domain. In the epidermis, PKC-3 depletion caused a rapid invasion of LGL-1 in the apical domain of the seam cells, while PAR-1 remained junctional and basal. Thus PKC-3 functions to exclude LGL-1 in the seam cells. A recent study found that LGL-1 can suppress sterility of a temperature sensitive pkc-3 allele, further indicating that the interaction between LGL-1 and PKC-3 is functionally relevant (Montoyo-Rosario et al., 2020).

In contrast to the epidermis, LGL-1 localization in the intestine remained unchanged upon PKC-3 depletion, and we observed no obvious abnormalities in the intestine upon PAR-6 or PKC-3 depletion. Thus, while PAR-6 and PKC-3 are essential for development of the embryonic intestine (Totong et al., 2007), they do not appear to be essential in the larval intestine. Other cellular systems, such as polarized protein trafficking, may suffice to maintain cell polarity in the absence of the apical PAR proteins (Shafaq-Zadah et al., 2012; Zhang et al., 2012; Zhang et al., 2011). An analogous situation exists in the Drosophila midgut, where integrins, but not the apical PAR proteins, are essential for polarization (Chen et al., 2018). The lack of LGL-1 mislocalization also points to the existence of possible redundancies in polarization of cortical polarity regulators, which may be uncovered through enhancer screens in PAR-6 or PKC-3 depleted backgrounds.

In embryonic epithelia, the requirements of the apical PAR proteins also vary between tissues. Intestinal and epidermal cells depleted of PAR-6 or PKC-3 using the ZF1 system still show apicobasal polarization, as evidenced by apical localization of junctional and cytoskeletal proteins (Montoyo-Rosario et al., 2020; Totong et al., 2007). However, in the arcade cells of the pharynx, most PAR-6 depleted animals show no apical enrichment of junctional or apical cytoskeletal markers (Von Stetina and Mango, 2015). These data further highlight that the requirements for PAR-6 and PKC-3 can vary between tissues.

A novel role for PAR-6 in epidermal microtubule organization

Epidermal-specific depletion uncovered a novel role for PAR-6 in organizing non-centrosomal microtubule bundles. In epithelial cells, apical ncMTOCs assemble apical–basal microtubule arrays. ncMTOCs contain proteins and complexes involved in microtubule anchoring, microtubule stabilization, and microtubule nucleation — such as the γ-tubulin ring complex (γ-TuRC) (Sanchez and Feldman, 2017). How apical ncMTOCs are organized is not well understood, but several studies indicate an important role for apical PAR proteins in this process. In the cellularizing Drosophila embryo, aPKC is required for the transition from centrosome emanated asters to non-centrosome associated apical–basal bundles (Harris and Peifer, 2007). In the developing embryonic intestine of C. elegans, PAR-3 is needed for the redistribution of γ-tubulin and other microtubule regulators from the centrosomes to the apical domain of the cell (Feldman and Priess, 2012). A role for Par6 in regulating microtubule-organizing centers may not be limited to epithelial ncMTOCs. For example, in several mammalian cultured cell lines Par6 is a component of centrosomes and regulates centrosomal protein composition (Dormoy et al., 2013; Kodani et al., 2010).

Epidermal depletion of PAR-6 resulted in reduced numbers of circumferential microtubule bundles, as well as a reduced microtubule growth rate and EB1 comet density. Moreover, depletion of PAR-6 led to a loss of apical NOCA-1 enrichment at seam–seam and seam–hyp7 junctions. The effects of PAR-6 depletion on microtubule organization and dynamics are very similar to those we observed in a noca-1 mutant, and their severity did not increase when combining PAR-6 depletion with the noca-1 mutant. While other models are possible, these data are consistent with PAR-6 acting through NOCA-1 to control microtubule organization in the epidermis. The reduced microtubule growth rate and EB1 comet density we observed in noca-1 mutant animals have been reported previously (Wang et al., 2015). However, no defects in circumferential microtubule bundle density were observed in that study, despite using the same noca-1(ok3692) allele. The observed difference may be a result of a difference in exact experimental procedure or the precise genetic background used. For example, whereas we used the microtubule-binding protein GFP::MAPH-1.1 to label microtubules, the study by Wang et al. used a GFP::β-tubulin fusion. However, it is not immediately clear how the markers would differentially affect microtubule density in noca-1(ok3692) animals, as both appear to label all microtubules, and in control animals we observe a similar microtubule density as observed by Wang et al.

We also found that, in the epidermis, the localization of GIP-1 is dependent on NOCA-1. The relationship between NOCA-1 and γ-TuRC components has been examined previously in two different tissues (Sallee et al., 2018; Wang et al., 2015, p. 1). In the germline, NOCA-1 co-localizes with γ-tubulin to non-centrosomal microtubule arrays but is not required for the localization of γ-tubulin (Wang et al., 2015). In fact, in this tissue the localization of a short NOCA-1 protein lacking isoform-specific N-terminal extensions is dependent for its localization on γ-tubulin. The longer NOCA-1h isoform, however, localizes independently of γ-tubulin, indicating the presence of multiple NOCA-1 localization signals (Wang et al., 2015). In the embryonic intestine, the localization of NOCA-1 was not altered by the depletion of GIP-1 (Sallee et al., 2018). However, microtubule organization in the intestine is regulated differently from the epidermis, as apical microtubule organization was largely normal even in ptrn-1 mutant animals depleted of intestinal NOCA-1 and GIP-1 (Sallee et al., 2018). Thus, differential effects of γ-TuRC component loss may reflect differences in the mechanisms of microtubule regulation. Whether PAR-6 plays a role in ncMTOC assembly and microtubule organization in tissues other than the epidermis remains to be investigated.

In addition to the effects on NOCA-1 and GIP-1, PAR-6 depletion resulted in a reduced number of PTRN-1 puncta in the epidermis. PTRN-1 is a member of the Patronin/CAMSAP/Nezha family of minus end-associated proteins, which stabilize and protect uncapped microtubule minus ends (Atherton et al., 2019; Goodwin and Vale, 2010; Hendershott and Vale, 2014; Jiang et al., 2014). NOCA-1 was previously shown to act in parallel with PTRN-1 in organizing circumferential microtubule arrays in the C. elegans epidermis (Wang et al., 2015). The mechanistic details of the relationship between NOCA-1 and PTRN-1 have not been resolved, but their distinct localization patterns suggest that they act on distinct pools of microtubules. Our data does not reveal why PAR-6 depletion results in a reduced number of PTRN-1 foci.

Mechanisms of larval growth arrest and molting defects

The depletion of PAR-6 or PKC-3 in the epidermis led to a rapid growth arrest and failure to molt. What causes these dramatic effects? The junctional defects we observed are unlikely to be the primary consequence of either the growth or molting defects, as effects on cell junctions appeared only after 24 hr of exposure to auxin. The effects on LGL-1 were more rapid but are also not likely to explain the defects, as lgl-1 mutants are viable (Beatty et al., 2010; Hoege et al., 2010, p. 1). The effects on the microtubule cytoskeleton are likely to contribute to the growth arrest and molting defects. However, noca-1 mutants displayed similar microtubule defects as PAR-6 depletion yet develop to adulthood. Interestingly, noca-1; ptrn-1 double mutant animals do grow slowly and frequently die before reaching adulthood (Wang et al., 2015). Thus, the combined defects in NOCA-1 and PTRN-1 localization we observed upon PAR-6 depletion may partially explain the growth defects. The roles of PTRN-1 may not be limited to microtubule regulation, as a recent study demonstrated that PTRN-1 stimulates actin polymerization during endocytic recycling in the intestine (Gong et al., 2018, p. 1).

Wheather and how the growth and molting defects are related is difficult to establish. The molting defect may contribute to the growth arrest, as failure to molt can cause a growth arrest (Brooks et al., 2003; Lažetić and Fay, 2017; Russel et al., 2011; Yochem et al., 1999). PAR-6 and PKC-3 could affect molting through intracellular trafficking. Molting requires the coordinated activity of the endocytic and exocytic machineries (Lažetić and Fay, 2017), and several links between cortical polarity regulators and the polarized trafficking machinery have been uncovered (Rodriguez-Boulan and Macara, 2014). In C. elegans, par-3, par-6, and pkc-3 were all found to be required for endocytic trafficking in oocytes, and RNAi for par-3 and par-6 causes scattering of multiple endosome types in the intestine (Balklava et al., 2007; Winter et al., 2012). It is possible, therefore, that PAR-6 and PKC-3 regulate vesicle trafficking in molting as well. Such regulation may be indirect, through regulation of cytoskeletal components, or through more direct mechanisms remaining to be uncovered. However, the fact that animals in which PAR-6 or PKC-3 is depleted from hatching lack any cell growth, rather than arresting at the L1 molt, suggests that the molting defect is not the sole cause of the growth defect. Alternatively, the growth defect may contribute to the molting defect. Diet restricted animals that grow very slowly delay the L1–L2 molt until a certain body size is reached, suggesting that molting is subject to a size threshold (Uppaluri and Brangwynne, 2015).

In summary, our data supports that PAR-6 and PKC-3 have multiple roles in the epidermis that support larval development and molting. We have also uncovered an important role for PAR-6 in regulating the microtubule cytoskeleton, while additional mechanisms through which PAR-6 and PKC-3 control growth and/or molting likely remain to be discovered.

Materials and methods

Key resources table
Reagent type (species) or resourceDesignationSource or referenceIdentifiersAdditional information
Strain, strain background(C. elegans)BOX289This paperpar-6(mib30[par-6::aid::egfp-loxp]) I; ieSi57[eft-3p::TIR1::mRuby::unc-54 3'UTR + Cbr-unc-119(+)] II
Strain, strain background(C. elegans)BOX570This paperpkc-3(mib78[egfp-loxp::aid::pkc-3]) II; ieSi57[eft-3p::TIR1::mRuby::unc-54 3'UTR + Cbr-unc-119(+)] II
Strain, strain background(C. elegans)BOX292This paperieSi57[eft-3p::TIR1::mRuby::unc-54 3'UTR + Cbr-unc-119(+)] II; par-3(mib68[eGFP-Lox2272::AID::par-3b+eGFP(noIntrons)-LoxP::AID::par-3g]) III
Strain, strain background(C. elegans)BOX276This paperpar-3b(mib65[eGFP-Lox2272::AID::par-3b]) III
Strain, strain background(C. elegans)BOX667This paperpar-3(mib68[eGFP-Lox2272::AID::par-3b+eGFP(noIntrons)-LoxP::AID::par-3g]) III; ieSi64 [gld-1p::TIR1::mRuby::gld-1 3’UTR + Cbr-unc-119(+)] II
Strain, strain background(C. elegans)BOX409This paperpar-6(mib30[par-6::aid::egfp-loxp]) I; mibIs49[Pwrt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)] IV
Strain, strain background(C. elegans)BOX607This paperpkc-3(mib78[egfp-loxp::aid::pkc-3]) II; mibIs49[Pwrt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)] IV
Strain, strain background(C. elegans)BOX444This paperpkc-3(mib78[egfp-loxp::aid::pkc-3]) II; mibIs48[Pelt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)] IV
Strain, strain background(C. elegans)BOX285This paperpar-6(mib30[par-6::aid::egfp-loxp]) I; mibIs48[Pelt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)] IV
Strain, strain background(C. elegans)BOX506This paperpar-6(mib30[par-6::aid::egfp-loxp]) I; mibIs49[Pwrt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)] IV; dlg-1(mib23[dlg-1::mCherry-LoxP]) X; mgIs49[mlt-10::gfp-pest]
Strain, strain background(C. elegans)BOX412This paperpar-6(mib30[par-6::aid::egfp-loxp]) I; mibIs49[Pwrt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)] IV; heIs63[Pwrt-2::GFP::PH, Pwrt-2::GFP::H2B, Plin-48::mCherry]V
Strain, strain background(C. elegans)BOX490This paperouIs10[Pscm::NLS::tdTomato(pAW584)+Pwrt2::GFP::PH(pAW561)+Pdpy-7::2xNLS::YFP(pAW516)] I; par-6(mib30[par-6::aid::egfp-loxp]) I; mibIs49[Pwrt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)] IV
Strain, strain background(C. elegans)BOX041This papermibIs23 [lgl-1::GFP-2TEV-Avi 10 ng + Pmyo-3::mCherry 10 ng + lambda DNA 60 ng] V
Strain, strain background(C. elegans)BOX553This paperpkc-3(mib78[egfp-loxp::aid::pkc-3]) II; mibIs49[Pwrt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)] IV; mibIs23 [lgl-1::GFP-2TEV-Avi 10 ng + Pmyo-3::mCherry 10 ng + lambda DNA 60 ng] V
Strain, strain background(C. elegans)BOX554This paperpkc-3(mib78[egfp-loxp::aid::pkc-3]) II; mibIs49[Pwrt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)] IV; par-1(it324[par-1::gfp::par-1 exon 11a])
Strain, strain background(C. elegans)BOX493This paperpkc-3(mib78[egfp-loxp::aid::pkc-3]) II; mibIs49[Pwrt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)] IV; dlg-1(mib23[dlg-1::mCherry-LoxP]) X
Strain, strain background(C. elegans)BOX402This paperpar-6(mib30[par-6::aid::egfp-loxp]) I; mibIs49[Pwrt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)] IV; dlg-1(mib23[dlg-1::mCherry-LoxP]) X
Strain, strain background(C. elegans)BOX494This papermcIs40 [Plin-26::ABDvab-10::mCherry + Pmyo-2::GFP]; par-6(mib30[par-6::aid::egfp-loxp]) I; mibIs49[Pwrt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)] IV; heIs63[Pwrt-2::GFP::PH, Pwrt-2::GFP::H2B, Plin-48::mCherry] V
Strain, strain background(C. elegans)BOX483This paperpar-6(mib30[par-6::aid::egfp-loxp]) I; maph-1.1(mib12[egfp::maph-1.1]) I; mibIs49[Pwrt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)] IV; dlg-1(mib23[dlg-1::mCherry-LoxP]) X
Strain, strain background(C. elegans)BOX505This papermaph-1.1(mib12[egfp::maph-1.1]) I; pkc-3(mib78[egfp-loxp::aid::pkc-3]) II; mibIs49[Pwrt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)] IV; dlg-1(mib23[dlg-1::mCherry-LoxP]) X
Strain, strain background(C. elegans)BOX592This papermaph-1.1(mib12[egfp::maph-1.1]) I; noca-1(ok3692)V/nT1[qIs51](IV;V)
Strain, strain background(C. elegans)BOX658This papermaph-1.1(mib12)I; par-6(mib24[par-6::egfp-loxp] I; mibIs49[Pwrt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)]) IV; noca-1(ok3692)V/nT1[qIs51](IV;V)
Strain, strain background(C. elegans)BOX487This paperpar-6(mib25[par-6::mCherry-LoxP]) I; ebp-2(he293[ebp-2::egfp]) II; mibIs49[Pwrt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)] IV
Strain, strain background(C. elegans)BOX580This paperebp-2(he293[ebp-2::egfp]) II; noca-1(ok3692)V/nT1[qIs51](IV;V)
Strain, strain background(C. elegans)BOX659This paperpar-6(mib24[par-6::egfp-loxp] I; ebp-2(he293[ebp-2::egfp]) II; mibIs49[Pwrt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)]) IV; noca-1(ok3692)V/nT1[qIs51](IV;V)
Strain, strain background(C. elegans)BOX567This paperpar-6(mib30[par-6::aid::egfp-loxp]) I; ltSi540[pOD1343/pSW160; Pnoca-1de::noca-1de::sfGFP; cb-unc-119(+)]II; unc-119(ed3)III; mibIs49[Pwrt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)] IV
Strain, strain background(C. elegans)BOX355This paperpar-6(mib30[par-6::aid::egfp-loxp]) I; ltSi540[pOD1343/pSW160; Pnoca-1de::noca-1de::sfGFP; cb-unc-119(+)]II; unc-119(ed3)III; ieSi57[eft-3p::TIR1::mRuby::unc-54 3'UTR + Cbr-unc-119(+)] II
Strain, strain background(C. elegans)BOX568This paperpar-6(mib30[par-6::aid::egfp-loxp]) I; gip-1(wow25[tagRFP-t::3xMyc::gip-1]) III; mibIs49[Pwrt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)] IV
Strain, strain background(C. elegans)BOX502This paperpar-6(mib30[par-6::aid::egfp-loxp]) I; mibIs49[Pwrt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)] IV; dlg-1(mib23[dlg-1::mCherry-LoxP]) X; ptrn-1(wow4[PTRN-1::GFP]) X
Strain, strain background(C. elegans)BOX657This paperpkc-3(mib78[egfp-loxp::aid::pkc-3]) II; Pnoca-1de::noca-1de::superfolderGFP; cb-unc-119(+)II; unc-119(ed3)III; mibIs49[Pwrt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)] IV
Strain, strain background(C. elegans)BOX579This papergip-1(wow25[tagRFP-t::3xMyc::gip-1]) III; noca-1(ok3692)V/nT1[qIs51](IV;V)
Strain, strain background(C. elegans)BOX561This paperpar-6(mib30[par-6::aid::egfp-loxp]) I; mibIs49[Pwrt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)] IV; mibEx221(Pdpy-7::par-6::mch)
Strain, strain background(C. elegans)BOX563This paperpar-6(mib30[par-6::aid::egfp-loxp]) I; mibIs49[Pwrt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)] IV; heIs63[Pwrt-2::GFP::PH, Pwrt-2::GFP::H2B, Plin-48::mCherry] V; mibEx222(Pdpy-7::par-6::mch; Pmyo-2::egfp)
Strain, strain background(C. elegans)BOX608This paperpw27[nekl-2::aid];pwSi10[phyp7::bfp::tir-1];pw17[chc-1::GFP]; mibEx223(Pwrt-2::mCh::H2B; Pwrt-2::mCh::PH)
Strain, strain background(C. elegans)BOX447This paperpkc-3(mib78[egfp-loxp::aid::pkc-3]) II; mibIs48[Pelt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)] IV; mibIs23 [lgl-1::GFP-2TEV-Avi 10 ng + Pmyo-3::mCherry 10 ng + lambda DNA 60 ng] V
Strain, strain background(C. elegans)BOX431This paperpar-6(mib30[par-6::aid::egfp-loxp]) I; mibIs48[Pelt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)] IV; dlg-1(mib23[dlg-1::mCherry-LoxP]) X
Strain, strain background(C. elegans)BOX406This paperpar-6(mib30[par-6::aid::egfp-loxp]) I; pkc-3(mib80[mcherry-loxp::pkc-3]) II; mibIs48[Pelt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)] IV
Strain, strain background(C. elegans)BOX653This paperpar-6(mib24[par-6::egfp-loxp] I; pkc-3(mib78[egfp-loxp::aid::pkc-3]) II; mibIs48[Pelt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)]) IV
Strain, strain background(C. elegans)BOX411This paperpar-6(mib30[par-6::aid::egfp-loxp]) I; pkc-3(mib80[mcherry-loxp::pkc-3]) II; mibIs49[Pwrt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)] IV
Strain, strain background(C. elegans)BOX578This paperpar-6(mib30[par-6::aid::egfp-loxp]) I; par-3(it300[par-3::mcherry]) III; mibIs49[Pwrt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)] IV
Strain, strain background(C. elegans)BOX484This paperpar-6(mib25[par-6::mCherry-LoxP]) I; pkc-3(mib78[egfp-loxp::aid::pkc-3]) II; mibIs49[Pwrt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)] IV
Strain, strain background(C. elegans)BOX485This paperpkc-3(mib78[egfp-loxp::aid::pkc-3]) II; par-3(it300[par-3::mcherry]) III; mibIs49[Pwrt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)] IV
Strain, strain background(C. elegans)BOX486This paperpar-6(mib25[par-6::mCherry-LoxP]) I; par-3(mib68[eGFP-Lox2272::AID::par-3b+eGFP(noIntrons)-LoxP::AID::par-3g]) III; mibIs49[Pwrt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)] IV
Strain, strain background(C. elegans)BOX492This paperpkc-3(it309[GFP::pkc-3]) II; par-3(mib68[eGFP-Lox2272::AID::par-3b+eGFP(noIntrons)-LoxP::AID::par-3g]) III; mibIs49[Pwrt-2::TIR-1::tagBFP2-Lox511::tbb-2–3'UTR, IV:5014740–5014802 (cxTi10882 site)] IV
Strain, strain background(C. elegans)AW1015Hughes et al., 2014RRID:WB-STRAIN:WBStrain00042230ouIs10[Pscm::NLS::tdTomato(pAW584)+Pwrt2::GFP::PH(pAW561)+Pdpy-7::2xNLS::YFP(pAW516)] I
Strain, strain background(C. elegans)BOX188Waaijers et al., 2016maph-1.1(mib12[egfp::maph-1.1]) I
Strain, strain background(C. elegans)CA1200CGCRRID:WB-STRAIN:WBStrain00004055ieSi57[eft-3p::TIR1::mRuby::unc-54 3'UTR + Cbr-unc-119(+)] II; unc-119(ed3) III
Strain, strain background(C. elegans)GR1395Hayes et al., 2006RRID:WB-STRAIN:WBStrain00007913mgIs49 [mlt-10::GFP-pest; ttx-1::GFP]
Strain, strain background(C. elegans)JLF15Jessica Feldmanptrn-1(wow4[PTRN-1::GFP]) X
Strain, strain background(C. elegans)JLF173Jessica Feldmangip-1(wow25[tagRFP-t::3xMyc::gip-1]) III
Strain, strain background(C. elegans)KK1218CGCRRID:WB-STRAIN:WBStrain00023582par-3(it300[par-3::mCherry]) III
Strain, strain background(C. elegans)KK1228CGCRRID:WB-STRAIN:WBStrain00023583pkc-3(it309[GFP::pkc-3]) II
Strain, strain background(C. elegans)KK1262CGCRRID:WB-STRAIN:WBStrain00023586par-1 (it324[par-1::gfp::par-1 exon 11a])
Strain, strain background(C. elegans)ML916CGCRRID:WB-STRAIN:WBStrain00026581mcIs40 [Plin-26::ABDvab-10::mCherry + Pmyo-2::GFP]
Strain, strain background(C. elegans)OD1652Karen OegemaRRID:WB-STRAIN:WBStrain00044359ltSi540[pOD1343/pSW160; Pnoca-1de::noca-1de::sfGFP; cb-unc-119(+)]II; unc-119(ed3)III
Strain, strain background(C. elegans)RT3638David Faypw27[nekl-2::aid];pwSi10[phyp7::bfp::tir-1];pw17[chc-1::GFP]
Strain, strain background(C. elegans)SV1009Wildwater et al., 2011RRID:WB-STRAIN:WBStrain00034608heIs63[Pwrt-2::GFP::PH, Pwrt-2::GFP::H2B, Plin-48::mCherry]V
Strain, strain background(C. elegans)SV1937Sander van den Heuvelebp-2(he293[ebp-2::egfp]) II
Strain, strain background(C. elegans)VC2998CGCRRID:WB-STRAIN:WBStrain00037614noca-1(ok3692)V/nT1[qIs51](IV;V)
Strain, strain background(C. elegans)CA1352RRID:WB-STRAIN:WBStrain00004071ieSi64 [gld-1p::TIR1::mRuby::gld-1 3’UTR + Cbr-unc-119(+)] II
Strain, strain background(C. elegans)STR320Martin Harterinkmaph-1.1(mib15[GFPKI]);hrtEx110[Pptrn-1::ebp-2::mKate2; Pmyo-2::tdTom]
Recombinant DNA reagentPlasmid: pJJR82Addgene#75027EGFP^SEC^3xFlag vector with ccdB markers for cloning homology arms
Recombinant DNA reagentPlasmid: pJJR83Addgene#75028mCherry^SEC^3xFlag vector with ccdB markers for cloning homology arms
Recombinant DNA reagentPlasmid: pMLS257Addgene#73716SapTrap destination vector for building repair template only vectors
Recombinant DNA reagentPlasmid: pDD379Addgene#91834SapTrap destination vector for building combined sgRNA expression + repair template vectors, using the F+E sgRNA scaffold
Recombinant DNA reagentPlasmid: pJJR50Addgene#75026U6 promoter driven flipped + extended sgRNA expression vector
Recombinant DNA reagentPlasmid: Peft-3::cas9Addgene#46168codon optimized Cas9_SV40 NLS with intron
Recombinant DNA reagentPlasmid: Pdpy-7::par-6::mCherryThis paperPlasmid for expression of PAR-6::mCherry in the hypodermis (Figure 3—figure supplement 2). Full sequence in Supplementary file 1.
Recombinant DNA reagentPlasmid: PAR-6 PDZ in pMB28This paperYeast expression plasmid of PAR-6 PDZ fused to Gal4 DNA binding domain (Figure 6—figure supplement 1). Full sequence in Supplementary file 1.
Recombinant DNA reagentPlasmid: NOCA-1d in pMB29This paperYeast expression plasmid of NOCA-1d fused to Gal4 activation domain (Figure 6—figure supplement 1). Full sequence in Supplementary file 1.
Sequence-based reagentpar-6 sgRNAgacgcaaatgacagtgatagTGGsgRNA target site used to engineer the par-6 locus. PAM site in uppercase.
Sequence-based reagentpkc-3 sgRNAtgggtctccgacatcattagAGGsgRNA target site used to engineer the pkc-3 locus. PAM site in uppercase.
Sequence-based reagentpar-3 sgRNA 1tttcagatcgatcatcatgtCGGsgRNA target site used to target the par-3 locus. PAM site in uppercase.
Sequence-based reagentpar-3 sgRNA 2cacatgcataacggtcgtggTGGsgRNA target site used to target the par-3 locus. PAM site in uppercase.
Sequence-based reagentdlg-1 sgRNAgccacgtcattagatgaaatTGGsgRNA target site used to target the dlg-1 locus. PAM site in uppercase.
Sequence-based reagentmos IV 5013690.5015700 sgRNAagctcaatcgtgtacttgcgTGGsgRNA target site for LG IV position 5013690.5015700, used to insert TIR-1 expression cassette. PAM site in uppercase.
Sequence-based reagentebp-2 sgRNA 1gcaggcaaatctggacgataCGGsgRNA target site used to edit the ebp-2 locus.
Sequence-based reagentebp-2 sgRNA 2tacggggataggataagcaaTGGsgRNA target site used to edit the ebp-2 locus.
Sequence-based reagentPdpy-7_FThis paperPCR primersTGTAATACGACTCACTATAGGGCGAATTGGctcattccacgatttctcgc. See Materials and methods section ‘PAR-6::mCherry transgenic array’ for usage details.
Sequence-based reagentPdpy-7_RThis paperPCR primerstctggaacaaaatgtaagaatattc See Materials and methods section ‘PAR-6::mCherry transgenic array’ for usage details.
Sequence-based reagentpar-6_F1This paperPCR primerstttaagaatattcttacattttgttccagaATGTCCTACAACGGCTCCTA See Materials and methods section ‘PAR-6::mCherry transgenic array’ for usage details.
Sequence-based reagentpar-6_R1This paperPCR primersGGCCATGTTGTCCTCCTCTCCCTTGGACATGTCCTCTCCACTGTCCGAAT See Materials and methods section ‘PAR-6::mCherry transgenic array’ for usage details.
Sequence-based reagentpar-6_UTR_FThis paperPCR primersCACTCCACCGGAGGAATGGACGAGCTCTACTGAaaaactcttttcagcca See Materials and methods section ‘PAR-6::mCherry transgenic array’ for usage details.
Sequence-based reagentpar-6_UTR_RThis paperPCR primersTAAAGGGAACAAAAGCTGGAGCTCCACCGCgaaataaataatttattctc See Materials and methods section ‘PAR-6::mCherry transgenic array’ for usage details.
Sequence-based reagentmCherry_FThis paperPCR primersTCCAAGGGAGAGGAGGACAA See Materials and methods section ‘PAR-6::mCherry transgenic array’ for usage details.
Sequence-based reagentmCherry_RThis paperPCR primersGTAGAGCTCGTCCATTCCTC See Materials and methods section ‘PAR-6::mCherry transgenic array’ for usage details.
Sequence-based reagentpar-6_F2This paperPCR primersggaggcgcgccATGATTGTGCCAGAAGCTCATCG See Materials and methods section ‘yeast two-hybrid’ analysis for usage details.
Sequence-based reagentpar-6_R2This paperPCR primersggagcggccgcTCAGGCGTTCGGTGTTCCTTGTT See Materials and methods section ‘yeast two-hybrid’ analysis for usage details.
Sequence-based reagentnoca-1d_FThis paperPCR primersggaggcgcgccATGAATATTTGTTGTTGTGG See Materials and methods section ‘yeast two-hybrid’ analysis for usage details.
Sequence-based reagentnoca-1d_RThis paperPCR primersggagcggccgcCTATTGAACTCTGCATACAT. See Materials and methods section ‘yeast two-hybrid’ analysis for usage details.

C. elegans strains

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All C. elegans strains used in this study are derived from the N2 Bristol strain, and are listed in the Key resources table. All strains were maintained at 20°C on Nematode Growth Medium (NGM) plates seeded with Escherichiae coli OP50 bacteria under standard conditions (Brenner, 1974).

CRISPR/Cas9 genome engineering

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All gene editing was done by homology-directed repair of CRISPR/Cas9-induced DNA double-strand breaks, using plasmid-based expression of Cas9 and sgRNAs. All edits were made in an N2 background, with the exception of 2x(egfp::aid)::par-3, for which egfp::aid::par-3 was used as the starting background. All fusions were repaired using a plasmid-based template with 190–600 bp homology arms and containing a self-excising cassette (SEC) for selection (Dickinson et al., 2015). The homology arms included mutations of the sgRNA recognition sites to prevent re-cutting after repair. The par-6::aid::egfp, par-6::mCherry, dlg-1::mCherry and ebp-2::egfp vectors were cloned using Gibson assembly and vector pJJR82 (Addgene #75027) (Gibson et al., 2009; Ramalho et al., 2020) as the backbone. The 2x(egfp::aid)::par-3, Pwrt-2::tir-1::bfp and Pelt-2::tir-1::bfp vectors were cloned using SapTrap assembly into vector pMLS257 (Addgene #73716) (Schwartz and Jorgensen, 2016), and the egfp::aid::pkc-3 and mCherry::pkc-3 vectors were cloned using SapTrap assembly into vector pDD379 (Addgene #91834) (Dickinson et al., 2018). The sgRNAs were expressed from a plasmid under control of a U6 promoter. To generate sgRNA vectors, antisense oligonucleotide pairs were annealed and ligated into BbsI-linearized pJJR50 (Addgene #75026) (Waaijers et al., 2016), with the exception of the pkc-3 fusions, in which the sgRNA was incorporated into assembly vector pDD379 using SapTrap assembly. The targeted sequences can be found in Table 2. Injection mixes were prepared in MilliQ H2O and contained 50 ng/μl Peft-3::cas9 (Addgene ID #46168) (Friedland et al., 2013), 50–100 ng/μl U6::sgRNA, 50 ng/μl of repair template, with the exception of the pkc-3 fusions, in which the sgRNA-repair-template vector was used at a concentration of 65 ng/μl. All mixes also contained 2.5 ng/μl of the co-injection pharyngeal marker Pmyo-2::GFP or Pmyo-2::tdTomato to aid in visual selection of transgenic strains. Young adult hermaphrodites were injected in the germline using an inverted micro-injection setup (Eppendorf FemtoJet 4x mounted on a Zeiss Axio Observer A.1 equipped with an Eppendorf Transferman 4 r). Candidate edited progeny were selected on plates containing 250 ng/µl of hygromycin (Dickinson et al., 2015), and correct genome editing was confirmed by Sanger sequencing (Macrogen Europe) of PCR amplicons encompassing the edited genomic region. From correctly edited strains, the hygromycin selection cassette was excised by a heat shock of L1 larvae at 34°C for 1 hr in a water bath. Correct excision was confirmed by Sanger sequencing. Sequence files of the final gene fusions in Genbank format are in Supplementary file 1.

C. elegans synchronization

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In order to obtain synchronized worm populations, plates with eggs were carefully washed with M9 (0.22 M KH2PO4, 0.42 M Na2HPO4, 0.85 M NaCl, 0.001 M MgSO4) buffer in order to remove larvae and adults but leave the eggs behind. Plates were washed again using M9 buffer after 1 hr, to collect larvae hatched within that time span.

Auxin-inducible degradation

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Auxin treatment was performed by placing synchronized populations of worms on NGP plates seeded with E. coli OP50 and containing 1 or 4 mM auxin. To prepare plates, auxin (Alfa Aesar A10556) was diluted into the autoclaved NGM agar medium after cooling to 60°C prior to plate pouring. Plates were kept for a maximum of 2 weeks in the dark at 4°C.

C. elegans growth curves

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To measure growth curves, L1 animals synchronized as described above were placed on NGM plates seeded with E. coli OP50 and either lacking auxin or containing 4 mM auxin. Images were taken in 24 hr intervals up to 96 hr, using a Zeiss Axio Zoom.V16 equipped with a PlanNeoFluar Z 1x/0.25 objective and Axiocam 506 color camera, driven by Zen Pro software. Animal length was quantified in ImageJ(FIJI) software by drawing a spline along the center line of the animal (Rueden et al., 2017; Schindelin et al., 2012).

Molting assay

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Synchronized L1 animals were placed on NGM plates seeded with E. coli OP50 and either lacking auxin or containing 1 mM auxin. Fluorescence images were taken in 1 hr intervals from 11 hr to 32 hr of development, using a Zeiss Axio Zoom.V16 equipped with a PlanNeoFluar Z 1x/0.25 objective and Axiocam 506 color camera, driven by Zen Pro software. Expression levels of the Pmlt-10::gfp::pest reporter were quantified in ImageJ(FIJI) software (see image analysis).

Microscopy

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Live imaging of C. elegans larvae was done by mounting larvae on 5% agarose pads in a 10 mM Tetramisole solution in M9 buffer to induce paralysis. DIC imaging was performed with an upright Zeiss AxioImager Z2 microscope using a 63 × 1.4 NA objective and a Zeiss AxioCam 503 monochrome camera, driven by Zeiss Zen software. Spinning disk confocal imaging was performed using a Nikon Ti-U microscope driven by MetaMorph Microscopy Automation and Image Analysis Software (Molecular Devices) and equipped with a Yokogawa CSU-X1-M1 confocal head and an Andor iXon DU-885 camera, using 60 × or 100 × 1.4 NA objectives. All stacks along the z-axis were obtained at 0.25 μm intervals, and all images were analyzed and processed using ImageJ(FIJI) and Adobe Photoshop. For quantifications, the same laser power and exposure times were used within experiments.

Quantitative image analysis

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All image analysis was done in using ImageJ (FIJI). For intensity profile measurements of spinning disk microscopy data, background values were subtracted from the intensity measurements. Mean background intensity was quantified on a circular region in an area not containing any animals, except in quantifications in Figure 4A; Figure 4C; Figure 6A; Figure 6G, where background intensity was quantified on a circular region in an area with no fluorescence inside the worm.

For the intensity profiles in the epidermis, except those of RFP::GIP-1, a 10 px-wide line was drawn in the apical focal plane, from the hyp7 cytoplasm to the seam-cell cytoplasm. The position of the line was chosen to avoid fluorescent signals present in neighboring tissues, notably the intestine and excretory canal. Additionally, mCherry tagged proteins tend to aggregate, as is evident from comparison with the otherwise identical GFP tagged variants. Hence, mCherry intensity profile lines were positioned to avoid aggregates. The RFP::GIP-1 fusion proteins localize in a punctate pattern. To accurately capture the average intensity of this marker protein, we drew 10 separate 20-px wide lines per cell, which covers 25–50% of the total seam-cell circumference. Intensity values were manually aligned at their peak values, and then averaged to obtain a single intensity profile per cell. For the intensity profiles in the intestine, we drew 8 separate 50 px-wide lines from the intestinal lumen to the cytoplasm of the intestinal cells, which were aligned at their peak values and averaged to obtain a single value per worm. The intensity profiles from multiple animals were manually aligned at the peak values for analysis and display.

To quantify the fluorescence intensity for the molting assay, whole worm fluorescence was quantified. A region of interest (ROI) of each whole worm was created by drawing a freehand line around the worm using the transmitted light channel. The corresponding fluorescence of the ROI was measured in the GFP channel.

Microtubule bundles were counted manually as follows: a 5-px-wide freehand line was drawn through an ~80 µm stretch of microtubule bundles at the dorsal or ventral region of an animal, and the intensity profile was plotted. The number of fluorescent peaks was counted, and the microtubule bundle density was calculated by dividing the number of peaks by the measured distance.

EBP-2::GFP comet counting was done manually as follows: an ROI was drawn around the area of hyp7 visible in the camera field of view (corresponding to 300–500 µm2). The entire width of hyp7 was included, from the outside of the animal up to (but excluding) the seam cells. Either the ventral or dorsal hypodermis was analyzed. Comets were counted manually within the ROI, and density was calculated by dividing the number of EBP-2::GFP comets by the surface of the area analyzed.

PTRN-1::GFP puncta counting was done manually. The entire epidermal area visible in the camera field of view was analyzed, and puncta in both the seam cells and the hypodermis were counted. Puncta density was calculated by dividing the number of PTRN-1::GFP puncta by the surface of the area analyzed.

Microtubule growth rate was calculated in an automated manner using the ImageJ plug-in ‘TrackMate’ (Tinevez et al., 2017). An ROI was drawn around either the seam cells or hyp7 visible in the camera field of view. For hyp7 either the ventral or dorsal area was analyzed. The following parameters were chosen: estimated blob diameter = 0.700 um; threshold = 200,000; simple LAP tracker; linking max distance = 1.5 um; gap-closing max distance = 1.5 um; gap-closing max frame gap = 3; duration of track = 10. The mean speed of the comets was averaged to obtain the average microtubule growth rate. Comets in both the seam cells and the hypodermis were measured and represented separately.

To determine the directionality of the actin bundles and microtubule growth, images or movies were rotated to orient the seam cells horizontally. Lines were drawn along the microtubule or actin bundles, and the angle of these lines was calculated relative to the horizontal axis. Per animal, an area containing 20 actin bundles or 30 microtubule bundles was analyzed (all bundles in the area were analyzed). Movies of EBP-2 were used to calculate the directionality of microtubule growth, where the direction of growth of individual comets was annotated manually. Maximum intensity projections of EBP-2 movies were used to calculate the directionality of microtubule growth. Rose plots were generated using MatLAB.

Relative excretory canal outgrowth

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To quantify relative canal outgrowth in the excretory canal cell, synchronized animals were placed on NGM plates seeded with E. coli OP50 and either lacking auxin or containing 4 mM auxin. Animals were placed on plates immediately after hatching. The distance between the cell body and either the anterior distal body tip or the anus was determined by tracing a segmented line along the center of the animal. The length of each individual canal was measured with a segmented line from the anterior-posterior bifurcation points close to the cell body until the canal tip. Relative outgrowth was calculated as the fraction of canal length over the distance between the cell body and the anterior distal tip or the anus.

Seam lineage analysis

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To generate the seam-cell lineage, synchronized animals were placed on NGM plates seeded with E. coli OP50 and either lacking auxin or containing 1 mM auxin. Animals were placed on plates immediately after hatching (before L1 degradation), at 7 hr of development (before L2 degradation) or at 19 hr of development (before L3 degradation). At 1 hr intervals, 5–10 animals were randomly picked and transferred to a microcopy slide. The number of seam cells and hyp7 nuclei were determined manually based on expression of the dual-color marker ouIs10[scmp::NLS::tdTomato; dpy-7p::2xNLS::YFP;wrt-2p::GFP::PH] that marks the seam nuclei in red and the hypodermal nuclei in green. Divisions of V5 were excluded from the analysis as V5 follows a different division pattern at the L2 stage, in which the anterior daughter becomes a neuroblast that generates a sensory structure termed the posterior deirid sensillium. V5 cells are readily recognized based on their position in the row of the seam cells, and in L2 stage additionally on their division pattern. Animals were classified according to showing a wild-type seam-cell division pattern, having developmental defects such as delayed or arrested seam-cell divisions, or having inappropriate seam-cell differentiation. Control animals were classified at each larval stage. PAR-6 depleted animals were classified after they had undergone the delayed L2-stage divisions. From the total number of worms analyzed, the percentages of worms in each category were calculated.

PAR-6::mCherry transgenic array

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The Pdpy-7::par-6::mCherry plasmid used for PAR-6 hypodermal rescue was cloned into the pBSK(+) vector using Gibson assembly. The promoter of dpy7, which is expressed in hyp7 but not in the seam cells (Gilleard et al., 1997; Myers and Greenwald, 2005), was amplified from C. elegans genomic DNA using primers Pdpy-7_F and Pdpy-7_R. A fragment of 5.3 kb containing the entire genomic sequence of par-6 and a fragment of 402 bp of the par-6 3’ UTR were amplified from C. elegans N2 genomic DNA using primers par-6_F1 and par-6_R1, and par-6_UTR_F and par-6_UTR_R, respectively. mCherry was amplified from pJJR83 (Addgene #75028) using primers mCherry_F and mCherry_R. Correct amplification and assembly were confirmed by Sanger sequencing. The plasmid generated can be found in Supplementary file 1. See Key resources table for primer sequences. To generate transgenic lines young adult hermaphrodites were injected in the germline with 30 ng/μl of Pdpy-7::par-6::mCherry. mCherry fluorescence was used to select stable transgenic lines.

Yeast two-hybrid analysis

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Sequences encoding the PAR-6 PDZ domain and full-length NOCA-1d were PCR amplified from a mixed-stage cDNA library using primers par-6_F2 and par-6_R2, and noca-1d_F and noca-1d_R. See Key resources table for primer sequences. PCR products were digested with AscI and NotI, and cloned into Gal4-DB vector pMB28 and Gal4-AD vector pMB29, respectively (Koorman et al., 2016). The resulting plasmids were transformed into Saccharomyces cerevisiae strains Y8930 (MATα) and Y8800 (MATa) (Yu et al., 2008) using the Te/LiAc transformation method (Schiestl and Gietz, 1989). DB::PAR-6/AD::NOCA-1 diploid yeast was generated by mating, and plated on synthetic defined (SD) medium plates lacking leucine, tryptophan, and histidine containing 2 mM 3-Amino-1,2,4-triazole (3-AT); and lacking leucine, tryptophan, and adenine to assess the presence of an interaction, and on an SD plate lacking leucine and histidine containing 1 μg/ml cycloheximide to test for self-activation by the DB::PAR-6 plasmid in the absence of the AD::NOCA-1 plasmid. Controls of known reporter activation strength and behavior on cycloheximide were also added to all plates.

Statistical analysis

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All statistical analyses were performed using GraphPad Prism 8. For population comparisons, a D'Agostino and Pearson test of normality was first performed to determine if the data was sampled from a Gaussian distribution. For data drawn from a Gaussian distribution, comparisons between two populations were done using an unpaired t-test, with Welch’s correction if the SDs of the populations differ significantly, and comparisons between >2 populations were done using a one-way ANOVA, or a Welch’s ANOVA if the SDs of the populations differ significantly. For data not drawn from a Gaussian distribution, a non-parametric test was used (Mann-Whitney for two populations and Kruskal-Wallis for >2 populations). ANOVA and non-parametric tests were followed up with multiple comparison tests of significance (Dunnett’s, Tukey’s, Dunnett’s T3 or Dunn’s). Tests of significance used and sample sizes are indicated in the figure legends. No statistical method was used to pre-determine sample sizes. No samples or animals were excluded from analysis. The experiments were not randomized, and the investigators were not blinded to allocation during experiments and outcome assessment.

References

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    Journal of Cell Science 114:2265–2277.
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Decision letter

  1. Oliver Hobert
    Reviewing Editor; Howard Hughes Medical Institute, Columbia University, United States
  2. Suzanne R Pfeffer
    Senior Editor; Stanford University School of Medicine, United States

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

Thank you for submitting your article "Epidermal PAR-6 and PKC-3 are essential for larval development of C. elegans and organize non-centrosomal microtubules" for consideration by eLife. Your article has been reviewed by three peer reviewers, and the evaluation has been overseen by a Reviewing Editor and Suzanne Pfeffer as the Senior Editor. The reviewers have opted to remain anonymous.

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

The editors have judged that your description of postembryonic roles of PAR proteins are of interest. However, additional experiments are required before the manuscript can be accepted for publication. As you will see described in detail below, a number of issues were raised about the way that some of the phenotypes were scored. In addition, genetic support for the PAR/Noca interaction is a critical, missing piece of information. Together with a number of other points, these requests should be straight-forward to address and all reviewers agree on their importance. Note that because of the Covid-19 situation, we do not require that revised manuscripts are returned within 2 months.

Reviewer #1:

In this study, Castiglioni and colleagues use the AID system to examine the roles of Par proteins Par3, Par6 and aPKC during larval growth. They find that epidermal expression of PAR-6 and PKC-3 (but not PAR-3 as tagged) is required for larval development and molting, and for the pattern of seam-cell divisions. Epistasis experiments using the tagged alleles support some previously published findings, but also suggest possible complexities that were not previously appreciated (such as regulation of PAR-3 localization by PKC-3 but not PAR-6 in seam cells). The most novel aspect of the paper is the finding that PAR-6 is important for non-centrosomal MT regulation, and may do so through its binding partner NOCA-1, although this aspect of the paper would benefit from additional experimentation.

Overall, the data presented are rigorous and I agree with most interpretations, save a few below that are surprising and would benefit from additional experimental tests. Much of the paper reveals findings that are not unexpected based on previous studies in the embryo or other model systems. The most novel and exciting aspect of the paper is the connection between PAR-6 and non-centrosomal MTs, which the authors propose occurs through NOCA-1 recruitment (which the Boxem lab revealed as a PAR-6-binding protein in a previous study). This conclusion should be strengthened by additional genetic tests, such as simultaneous removal of PAR-6 and NOCA-1 to compare phenotypes to the single mutants. In addition, the impact of the paper to the field would be increased if it were shown how PAR-6 interfaces with NOCA-1. Is it simply recruiting NOCA-1 to the apical surface? Does PKC-3 phosphorylation contribute as well, or is the role of PKC-3 in this context limited to localizing PAR-6?

Reviewer #2:

In this manuscript, Castiglioni and colleagues analyzed the role of PAR proteins during larval development in C. elegans. While the function of PARs in embryonic development is well established, their role during larval development is poorly characterized. The authors developed an auxin-inducible depletion system of PAR proteins and showed that PAR-6 and PKC-3 are required in the epidermis for larval development, molting, proper polarity and cell division patterns. They also observed that PAR-6 regulates the pattern of microtubules via its direct interaction with NOCA-1/Ninein. Overall, this is a high quality study. The conclusions are well supported by the experiments and the manuscript is nicely written. The novel interaction between PAR-6 and NOCA-1 is of broad interest and can justify publication in eLife.

There is one point of interest that the authors did not really discuss. In the hyp7 epidermal cell, microtubules are oriented circumferentially (perpendicular to the anteroposterior axis). They observed that PAR-6 depletion perturbed this orientation and the direction of microtubule growth (Figure 5C, 5H). This suggests that, in addition to its role in apicobasal polarity and junction organization, PAR-6 could also have a role in the planar polarity of epidermal cells. Could that be due to PAR-6 enrichment at the hyp7-seam borders ? It would be interesting to discuss how PAR-6 could instruct the microtubule orientation.

Reviewer #3:

This paper describes requirements of PAR proteins in C. elegans larval epithelium using degron system. It shows that PAR-6 and PKC-3 in epidermis are necessary for growth of animals, molting, junctional protein localizations and organizations of non-centrosomal microtubules. However, the manuscript contains a number of problems. Substantial revision and additional experiments are necessary to understand the function of par proteins in epidermis.

1) PAR-6 or PKC-3 depletion causes various defects, however, relationship among them are not clear and difficult to understand, especially since different time points after the auxin treatment were used for each analysis: 24, 48 and 72hrs for growth, 5 and 12hrs for junction, 6 and 24hrs for junctional protein localizations, 1hr for PAR protein localizations, 24hrs for MT density and 1hr for ebp-1 signals. Phenotypes at the same time point (e.g. 6hrs) should be shown in addition to other time points and thoroughly describe and discuss order of their appearance after the treatments.

2) For the growth defects, according to the graphs and raw data, sizes of the depleted animals after 24hrs are similar to those at 1-2.5 hrs hatching (very strangely, except for Figure 3—figure supplement 1B that contradicts with Figure 2I) which indicate that animals stop growing just after the treatment, raising the possibility that most other defects are secondary consequences of the former. In addition, molting defect is likely to affect phenotypes beyond the L2 stage (e.g. 24hrs). It is surprising that animals start molting and undergo L2 stage seam-cell divisions without any growth. Although Figure 3E shows L2 seam divisions, it clearly contradicts with no increase in seam-cell numbers shown in Figure 3—figure supplement 1D. Therefore, I wonder if the lineaged animals (Figure 3E) and animals with molting defects (Figure 3A) are rare escaper from the growth defects. Most of the depleted animals may just arrest at the L1 stage. Since dpy-7::PAR-6 can rescue growth defects but not seam-cell morphology, lineage should be analyzed in this rescued animals to know functions of PAR-6 in seam cells.

3) In the figure legends of many images used for quantification (Figure 2 AB, Figure 4AC, Figure 4—figure supplement 2A-F, Figure 6A), no explanation about the colored lines, "distance" in the graphs and how the quantification was done. Part of such information can only be found in Materials and methods section. But no explanation how positions of the lines are determined. I guess the authors intentionally decide positions of lines (at least this possibility cannot be excluded). Some of the images (Figure 4—figure supplement 2A-D) are quite dirty with background(?) signals. The authors appears to avoid those signals. But not clear how real and background signals are distinguished. They should have fair criteria about positions of the lines, e.g. above the center of seam-cell nuclei. They can avoid background signals only when there are clear reasons to do so. In addition, although it was described "The intensity profiles were manually aligned at the apical peak value" in the Materials and methods section, this is not the case at least for Figure 4B (I checked the source file only for this).

https://doi.org/10.7554/eLife.62067.sa1

Author response

Reviewer #1:

In this study, Castiglioni and colleagues use the AID system to examine the roles of Par proteins Par3, Par6 and aPKC during larval growth. They find that epidermal expression of PAR-6 and PKC-3 (but not PAR-3 as tagged) is required for larval development and molting, and for the pattern of seam cell divisions. Epistasis experiments using the tagged alleles support some previously published findings, but also suggest possible complexities that were not previously appreciated (such as regulation of PAR-3 localization by PKC-3 but not PAR-6 in seam cells). The most novel aspect of the paper is the finding that PAR-6 is important for non-centrosomal MT regulation, and may do so through its binding partner NOCA-1, although this aspect of the paper would benefit from additional experimentation.

Overall, the data presented are rigorous and I agree with most interpretations, save a few below that are surprising and would benefit from additional experimental tests. Much of the paper reveals findings that are not unexpected based on previous studies in the embryo or other model systems. The most novel and exciting aspect of the paper is the connection between PAR-6 and non-centrosomal MTs, which the authors propose occurs through NOCA-1 recruitment (which the Boxem lab revealed as a PAR-6-binding protein in a previous study). This conclusion should be strengthened by additional genetic tests, such as simultaneous removal of PAR-6 and NOCA-1 to compare phenotypes to the single mutants.

Examining the double inactivation is a great suggestion, and one we should have thought of for the initial submission. We have now performed PAR-6 depletion in the noca-1 mutant background and have included the results in the manuscript. We do not see an increase in the severity of microtubule disorganization (using both the MAPH-1.1 and EBP-2 markers). These results strengthen our conclusion that the microtubule disorganization upon PAR-6 depletion is due to the loss of NOCA-1 localization.

In addition, the impact of the paper to the field would be increased if it were shown how PAR-6 interfaces with NOCA-1. Is it simply recruiting NOCA-1 to the apical surface? Does PKC-3 phosphorylation contribute as well, or is the role of PKC-3 in this context limited to localizing PAR-6?

We liked this line of thought a lot and wanted to test this by inactivating the kinase function of PKC-3. In a paper from the Goehring lab (Rodrigues et al., 2017), the authors used a pkc-3 temperature sensitive allele and a drug to inactivate the kinase function of pkc-3. We thought either approach would be ideal to address this point, as neither approach disrupted membrane localization of PAR-6 and PKC-3 in the one-cell embryo. We contacted the drug manufacturer via several channels, but did not receive any replies. We also crossed the ts allele to our microtubule organization markers and to the LGL-1 marker strain. Unfortunately, at the restrictive temperature, we did not see any apical displacement of LGL-1 (in contrast to our depletion results). We also saw no effect on microtubule organization using MAPH-1.1 as the marker, but without displacement of LGL-1 this result is meaningless as we have no indication that the kinase function of PKC-3 is actually disrupted. As pkc-3(ts) mutants become sterile adults, a much milder phenotype than what we observe by protein degradation, it seems likely that the ts allele indeed does not fully disrupt PKC-3 kinase activity. We therefore were not able to address this question.

Reviewer #2:

In this manuscript, Castiglioni and colleagues analyzed the role of PAR proteins during larval development in C. elegans. While the function of PARs in embryonic development is well established, their role during larval development is poorly characterized. The authors developed an auxin-inducible depletion system of PAR proteins and showed that PAR-6 and PKC-3 are required in the epidermis for larval development, molting, proper polarity and cell division patterns. They also observed that PAR-6 regulates the pattern of microtubules via its direct interaction with NOCA-1/Ninein. Overall, this is a high quality study. The conclusions are well supported by the experiments and the manuscript is nicely written. The novel interaction between PAR-6 and NOCA-1 is of broad interest and can justify publication in eLife.

There is one point of interest that the authors did not really discuss. In the hyp7 epidermal cell, microtubules are oriented circumferentially (perpendicular to the anteroposterior axis). They observed that PAR-6 depletion perturbed this orientation and the direction of microtubule growth (Figure 5C, 5H). This suggests that, in addition to its role in apicobasal polarity and junction organization, PAR-6 could also have a role in the planar polarity of epidermal cells. Could that be due to PAR-6 enrichment at the hyp7-seam borders ? It would be interesting to discuss how PAR-6 could instruct the microtubule orientation.

We did notice the PAR-6 enrichment and the hyp7-seam borders, which is consistent with the recent observation by the Michaux lab that the apical PAR module is planar polarized in the lateral embryonic epidermis of the embryo. In the larval stages, however, the planar polarization of PAR-6 was not consistent, and whether it localized planar or more uniformly did not appear to correlate with any particular stage in the seam lineage pattern. For this reason we did not highlight it in the manuscript. Whether the PAR module has a role in planar polarity is difficult to say for sure. The seam cell defects we observed, in particular the orientation defects, do hint at a problem with determining anterior/posterior direction, but could also be a consequence of defects in another cellular process. The microtubule network in the seam cells is not regularly organized, nor visibly disrupted upon loss of PAR-6. We felt that a discussion of planar polarity would therefore be highly speculative. Similarly, we have not data to indicate that, in addition to NOCA-1 recruitment, PAR-6 directly instructs microtubule orientation in the hypodermis. We have now mentioned the planar localization of PAR-6 with a reference to the data from the Michaux lab, but have not added an in depth discussion. We hope the reviewer will agree with us that such a discussion would be too speculative.

Reviewer #3:

This paper describes requirements of PAR proteins in C. elegans larval epithelium using degron system. It shows that PAR-6 and PKC-3 in epidermis are necessary for growth of animals, molting, junctional protein localizations and organizations of non-centrosomal microtubules. However, the manuscript contains a number of problems. Substantial revision and additional experiments are necessary to understand the function of par proteins in epidermis.

1) PAR-6 or PKC-3 depletion causes various defects, however, relationship among them are not clear and difficult to understand, especially since different time points after the auxin treatment were used for each analysis: 24, 48 and 72hrs for growth, 5 and 12hrs for junction, 6 and 24hrs for junctional protein localizations, 1hr for PAR protein localizations, 24hrs for MT density and 1hr for ebp-1 signals. Phenotypes at the same time point (e.g. 6hrs) should be shown in addition to other time points and thoroughly describe and discuss order of their appearance after the treatments.

We understand that the occurrence of multiple different phenotypes upon PAR-6 or PKC-3 depletion can be confusing. Most likely, and in line with current thinking in the field, PAR-6/PKC-3 are involved in multiple processes, and it is expected that multiple phenotypes are uncovered. We have tried to clarify the potential relationships among the phenotypes by opening the discussion with a summary of the effects observed, and which ones we think are secondary and which are independent. We also extensively rewrote the section “Cell autonomous and non-autonomous roles for PAR-6 and PKC-3 in molting, seam cell divisions and seam cell morphology”. We emphasize that the growth defect does not correspond to a general developmental arrest, and added new data showing that excretory canal outgrowth during L1 continues. We also highlight the possible relationships between growth and molting, as molting defects block growth, but molting itself has also been reported to require growth. We already had written and discussed that the seam cell division defects largely are a secondary consequence. Finally, we tried to be careful in the manuscript text to describe the time points analyzed and times at which auxin was added. We have gone through the text and added further time point statements.

With regards to the analysis timepoints, we based these either on the time in development that a certain event occurs (e.g. seam division, the first molt, or appearance of circumferential actin bundles), or on the earliest moment the defect appears (e.g. 1 h for ebp-1 defects and 24 h for MT density defects). Picking a single time point is not possible, as either the event we want to show is not taking place at that time, or the defect is not evident yet. We feel it is important to show the earliest time point at which a defect becomes apparent, as it is in fact informative with regards to the role of the targeted protein. The speed at which a defect appears after the targeted protein is depleted gives an indication on whether the degraded protein is an integral component of the pathway or structure examined, or is required for assembly but not functioning of a structure. For example, upon depletion of PAR-6, we observed defects in microtubule growth or nucleation within 1 h, while defects in circumferential microtubule bundles took ~24 h to become apparent. This indicates that PAR-6 regulates the formation of new microtubules but is not essential for the maintenance of already existing microtubule bundles. Similarly, junctional defects in the epidermis appeared ~24 after PAR-6 or PKC-3 depletion started, indicating that PAR-6 and PKC-3 are important for the assembly of new junctions, but are not integral components. We have added a discussion of this topic to the Discussion section.

2) For the growth defects, according to the graphs and raw data, sizes of the depleted animals after 24hrs are similar to those at 1-2.5 hrs hatching (very strangely, except for Figure 3—figure supplement 1B that contradicts with Figure 2I) which indicate that animals stop growing just after the treatment, raising the possibility that most other defects are secondary consequences of the former. In addition, molting defect is likely to affect phenotypes beyond the L2 stage (e.g. 24hrs). It is surprising that animals start molting and undergo L2 stage seam cell divisions without any growth. Although Figure 3E shows L2 seam divisions, it clearly contradicts with no increase in seam cell numbers shown in Figure 3—figure supplement 1D. Therefore, I wonder if the lineaged animals (Figure 3E) and animals with molting defects (Figure 3A) are rare escaper from the growth defects. Most of the depleted animals may just arrest at the L1 stage. Since dpy-7::PAR-6 can rescue growth defects but not seam cell morphology, lineage should be analyzed in this rescued animals to know functions of PAR-6 in seam cells.

Below we address each of the individual points. Overall, we have rewritten this section of the Results extensively to clarify the questions of the reviewer, and clarify which aspects of the observed phenotypes we think are secondary consequences. We also elaborate on the possible relationships between growth and molting defects, in both the Results and Discussion, where we are now much more careful in describing that growth and molting are interrelated.

– The animals do indeed stop growing just after the treatment. The interpretation that Figure 3—figure supplement 1B contradicts with Figure 2I is, however, not correct. Figure 3—figure supplement 1B shows animals in which we exogenously express wild-type PAR-6 in hyp7. These animals only lack PAR-6 in the seam cells and do indeed grow. This experiment is part of our evidence that the growth defect is due to a role of PAR-6 in the hypodermis.

– Figure 3E also does not contradict with Figure 3—figure supplement 1D. In Figure 3—figure supplement 1D, we show the 24h timepoint. To clarify, we have added timing marks in hours to Figure 3E. Wild-type animals at 24h point have completed the L2 divisions that cause a doubling of cell number. PAR-6 depleted animals have such a long delay that there has been no doubling division. This is why control animals in Figure 3—figure supplement 1D show 10-20 cells, while the PAR-6 depleted animals show 5 seam cells. The third bar shows that expression of PAR-6 in hyp7 is sufficient to rescue the seam cell division timing. The reason for the spread from 10-20 cells is that after division, anterior seam cells fuse with the hypodermis. Hence immediately after division there are 20 cells, and after fusion is complete there are 10 cells again. At the 24h timepoint some cells have fused and some have not.

– We have certainly not analyzed rare escapers. For the growth assays, a number of animals are placed on a plate, and all animals are analyzed at each timepoint. For the lineaging, synchronized populations are washed off the plate, placed on a microscopy slide, and analyzed in the order they are spotted under the microscope.

– We do not think that PAR-6 depletion causes a full L1 developmental arrest, as aspects of development clearly continue despite the growth arrest. In all animals, the L1 seam cell divisions take place at the normal time. We have now also measured elongation of the excretory canals during L1 development as another marker of development. The visible molting defects in Figure 3A are not rare: 30-50% of animals show molting defects indicating that they reached the first molt but fail to complete the molt.

– We do agree that these findings are surprising: that aspects of development continue to take place despite the growth arrest. We have rewritten the section “Cell autonomous and nonautonomous roles for PAR-6 and PKC-3 in molting, seam cell divisions and seam cell morphology” to more clearly state that this is the case, and include the new excretory canal data.

– We completely agree that the growth and molting defects likely cause secondary defects. This is why we performed the experiments shown in Figure 3—figure supplement 1. They show that when we express non-degradable PAR-6 in hyp7, we not only restore the growth and molting defects, but also rescue most of the seam cell lineage defects. Moreover, arresting growth and molting by another method (nekl-2 depletion) causes a defect in seam cell divisions similar to PAR-6 depletion. We have rewritten the results to clarify that the seam cell division defects likely are a secondary defect, and added an extensive discussion of the order in which phenotypes appear and their relationships.

– Finally, while we did not perform a full lineaging analysis of dpy-7::PAR-6 rescued animals, Figure 3—figure supplement 1D shows that in these animals, L2 divisions take place as normal. Hence the severe seam cell lineage defects are due to loss of PAR-6 in hyp7, and PAR-6 is dispensable in the seam cells (at least for cell division timing). A full lineage analysis would be very time consuming and would not give additional information on the functioning of PAR-6 in the seam cells.

3) In the figure legends of many images used for quantification (Figure 2 AB, Figure 4AC, Figure 4—figure supplement 2A-F, Figure 6A), no explanation about the colored lines, "distance" in the graphs and how the quantification was done. Part of such information can only be found in Materials and methods section. But no explanation how positions of the lines are determined. I guess the authors intentionally decide positions of lines (at least this possibility cannot be excluded). Some of the images (Figure 4—figure supplement 2A-D) are quite dirty with background(?) signals. The authors appears to avoid those signals. But not clear how real and background signals are distinguished. They should have fair criteria about positions of the lines, e.g. above the center of seam cell nuclei. They can avoid background signals only when there are clear reasons to do so. In addition, although it was described "The intensity profiles were manually aligned at the apical peak value" in the Materials and methods section, this is not the case at least for Figure 4B (I checked the source file only for this).

We have expanded the Materials and methods section to include more details on quantification. Lines were drawn manually rather than at a predetermined location to avoid two types of aberrant signal that would distort the measurements. (1) areas where the cell being quantified was abutting another cell showing fluorescence. The seam cells, hypodermis, excretory canal, and intestine are all close to each other and all express polarity proteins. It is common therefore to observe fluorescence from multiple cells in one imaging plane. (2) Known background mCherry aggregation signals. As reviewer 1 mentions, mCherry is unfortunately prone to aggregation. However, because we also have the corresponding GFP fusions we know that the aggregates are indeed background. We fully agree with the reviewer that the “dirty” mCherry signals complicate analysis. We address this point in detail in our response to reviewer 1’s questions regarding Figure 4—figure supplement 2.

Most of the markers we used have a uniform localization pattern, and the measurements use a 10 px wide line to average out minor fluctuations in fluorescence (the indicative lines in the drawing are only ~1 px wide). Combined with measuring multiple animals for each experimental condition, this should yield an accurate representative value. The only marker we measured that does not have a uniform localization is GIP-1, which localizes in a punctate pattern. For this marker we drew extra wide lines (20 px) and drew 10 non-overlapping lines per cell, thus covering 25-50% of the total cell circumference.

We have added an explanation of the colored lines to all legends. We have expanded the Materials and methods with the above information on how the position of the lines was determined. We are unsure what kind of explanation the reviewer would like to see for an explanation of distance. The graphs already indicate that the distance is in micrometer, and the short, colored lines indicate where the measurements were taken.

We have double checked the source file for Figure 4B, but it is aligned properly. The values for each line measured are in the columns, and each column was shifted up or down such that the maximum values (reddest color) of all columns align horizontally. The shifting is clearly visible at the top and bottom of the sheet.

https://doi.org/10.7554/eLife.62067.sa2

Article and author information

Author details

  1. Victoria G Castiglioni

    Division of Developmental Biology, Institute of Biodynamics and Biocomplexity, Department of Biology, Faculty of Science, Utrecht University, Utrecht, Netherlands
    Contribution
    Conceptualization, Formal analysis, Investigation, Visualization, Methodology, Writing - original draft, Writing - review and editing
    Contributed equally with
    Helena R Pires
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-9884-2537
  2. Helena R Pires

    Division of Developmental Biology, Institute of Biodynamics and Biocomplexity, Department of Biology, Faculty of Science, Utrecht University, Utrecht, Netherlands
    Contribution
    Conceptualization, Formal analysis, Investigation, Visualization, Methodology, Writing - original draft, Writing - review and editing
    Contributed equally with
    Victoria G Castiglioni
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-8351-0812
  3. Rodrigo Rosas Bertolini

    Division of Developmental Biology, Institute of Biodynamics and Biocomplexity, Department of Biology, Faculty of Science, Utrecht University, Utrecht, Netherlands
    Contribution
    Formal analysis, Investigation
    Competing interests
    No competing interests declared
  4. Amalia Riga

    Division of Developmental Biology, Institute of Biodynamics and Biocomplexity, Department of Biology, Faculty of Science, Utrecht University, Utrecht, Netherlands
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  5. Jana Kerver

    Division of Developmental Biology, Institute of Biodynamics and Biocomplexity, Department of Biology, Faculty of Science, Utrecht University, Utrecht, Netherlands
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  6. Mike Boxem

    Division of Developmental Biology, Institute of Biodynamics and Biocomplexity, Department of Biology, Faculty of Science, Utrecht University, Utrecht, Netherlands
    Contribution
    Conceptualization, Supervision, Funding acquisition, Visualization, Writing - original draft, Project administration, Writing - review and editing
    For correspondence
    M.Boxem@uu.nl
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-3966-4173

Funding

Nederlandse Organisatie voor Wetenschappelijk Onderzoek (824.14.021)

  • Mike Boxem

Nederlandse Organisatie voor Wetenschappelijk Onderzoek (016.VICI.170.165)

  • Mike Boxem

H2020 Marie Skłodowska-Curie Actions (ITN 675407)

  • Mike Boxem

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We thank R Schmidt and S van den Heuvel for sharing strain SV1937, S van den Heuvel for strain SV1009, J Feldman for strains JLF15 and JLF173, K Oegema for strain OD1652, A Woollard for strain AW1015, A Frand for strain GR1395, and D Fay for strain RT3638. We thank S van den Heuvel, M Harterink, D Fay and members of the S van den Heuvel and M Boxem groups for helpful discussions, M Harterink for critical reading of the manuscript, J Sepers for help generating PAR-3 strains, and J Cravo for generating the rose plots. We also thank Wormbase (Harris et al., 2020) and the Biology Imaging Center, Faculty of Sciences, Department of Biology, Utrecht University. Some strains were provided by the Caenorhabditis Genetics Center, which is funded by NIH Office of Research Infrastructure Programs (P40 OD010440). This work was supported by the Netherlands Organization for Scientific Research (NWO)-ALW Open Program 824.14.021 and NWO-VICI 016.VICI.170.165 grants to M Boxem, and the European Union’s Horizon 2020 research and innovation programme under the Marie Skłodowska-Curie grant agreement No. 675407 – PolarNet.

Senior Editor

  1. Suzanne R Pfeffer, Stanford University School of Medicine, United States

Reviewing Editor

  1. Oliver Hobert, Howard Hughes Medical Institute, Columbia University, United States

Publication history

  1. Received: August 12, 2020
  2. Accepted: December 9, 2020
  3. Accepted Manuscript published: December 10, 2020 (version 1)
  4. Version of Record published: December 22, 2020 (version 2)

Copyright

© 2020, Castiglioni 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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