ABSTRACT
Asymmetric division is crucial for embryonic development and stem cell lineages. In the one-cell Caenorhabditis elegans embryo, a contractile cortical actomyosin network contributes to asymmetric division by segregating partitioning-defective (PAR) proteins to discrete cortical domains. In the current study, we found that the plasma membrane lipid phosphatidylinositol 4,5-bisphosphate (PIP2) localizes to polarized dynamic structures in C. elegans zygotes, distributing in a PAR-dependent manner along the anterior–posterior (A–P) embryonic axis. PIP2 cortical structures overlap with F-actin, and coincide with the actin regulators RHO-1 and CDC-42, as well as ECT-2. Particle image velocimetry analysis revealed that PIP2 and F-actin cortical movements are coupled, with PIP2 structures moving slightly ahead of F-actin. Importantly, we established that PIP2 cortical structure formation and movement is actin dependent. Moreover, we found that decreasing or increasing the level of PIP2 resulted in severe F-actin disorganization, revealing interdependence between these components. Furthermore, we determined that PIP2 and F-actin regulate the sizing of PAR cortical domains, including during the maintenance phase of polarization. Overall, our work establishes that a lipid membrane component, PIP2, modulates actin organization and cell polarity in C. elegans embryos.
INTRODUCTION
Asymmetric division generates cellular diversity and is prevalent throughout development. During intrinsic asymmetric division, cell polarity is first established and then coupled with spindle positioning, resulting in proper cleavage and partitioning of contents to daughter cells. The evolutionarily conserved partitioning-defective (PAR) proteins are crucial for cell polarity and asymmetric division (reviewed by Goldstein and Macara, 2007; Gönczy, 2008; Knoblich, 2010). In contrast to the wealth of knowledge regarding PAR proteins and interacting components, the involvement of lipid plasma membrane components in these processes is less understood, in particular in developing systems.
The early Caenorhabditis elegans embryo has proven instrumental for uncovering mechanisms governing asymmetric division (reviewed by Hoege and Hyman, 2013; Pacquelet, 2017; Rose and Gönczy, 2014). Shortly after fertilization, the embryo surface exhibits uniform contractions of the cortical actomyosin network located underneath the plasma membrane. These contractions are driven by non-muscle myosin 2 (NMY-2), activated by the Rho GTPase RHO-1 and its guanine nucleotide exchange factor (GEF) ECT-2 (Motegi and Sugimoto, 2006; Schonegg and Hyman, 2006). Thereafter, sperm-derived centrosomes induce the local disappearance of cortical ECT-2 in their vicinity, thereby determining the future embryo posterior. This results in local RHO-1 inactivation and cortical flows away from this region, towards the future embryo anterior (Bienkowska and Cowan, 2012; Cowan and Hyman, 2004; Motegi and Sugimoto, 2006). These flows promote PAR polarity establishment, whereby PAR-3, PAR-6 and atypical protein kinase C (PCK-3) are segregated to the anterior side, whereas PAR-1, PAR-2 and lethal giant larvae-like 1 (LGL-1) occupy the expanding posterior cortical domain (reviewed by Hoege and Hyman, 2013; Pacquelet, 2017; Rose and Gönczy, 2014). The Rho GTPase CDC-42 is also segregated to the anterior, where it stabilizes the actomyosin network and promotes PAR-6 cortical association (Kumfer et al., 2010; Motegi and Sugimoto, 2006; Schonegg and Hyman, 2006).
PAR polarity can also be established in C. elegans zygotes through a partially redundant pathway, whereby microtubules nucleated from centrosomes protect PAR-2 from PKC-3-mediated phosphorylation, allowing PAR-2 association with phospholipids at the embryo posterior (Motegi et al., 2011; Zonies et al., 2010). In other systems, homologs of PAR proteins and interacting components also associate with phospholipids, as exemplified by Drosophila DmPar3 binding to phosphatidylinositol 4,5-bisphosphate [PI(4,5)P2, hereafter PIP2] and phosphatidylinositol 3,4,5-trisphosphate [PI(3,4,5)P3, hereafter PIP3] (Krahn et al., 2010). Furthermore, human Cdc42 binds to PIP2 (Johnson et al., 2012). Overall, phospholipids can bind PARs and interacting proteins in some contexts, but their subcellular distribution and potential function in an asymmetrically dividing system, such as the C. elegans zygote, remain unclear.
PIP2 is one of the most abundant of seven phosphorylated phosphatidylinositols and is present mostly in the inner leaflet of the plasma membrane (reviewed by De Craene et al., 2017). PIP2 is mainly generated from phosphatidylinositol 4-phosphate (PI4P) by Type I PI(4)P5-kinases (PIP5K1), and can be further phosphorylated to PIP3 by phosphatidylinositol 3-kinases (PI3K). Conversely, 5-phosphatases, including OCRL and synaptojanin, can dephosphorylate PIP2. In systems from Saccharomyces cerevisiae to Homo sapiens, PIP2 helps link the F-actin cortical network to the plasma membrane and promotes F-actin assembly and reorganization by stimulating, together with Cdc42, WASP family proteins that activate the actin nucleator Arp2/3 (reviewed by Brown, 2015; De Craene et al., 2017; Di Paolo and De Camilli, 2006; McLaughlin et al., 2002; Wu et al., 2014; Yin and Janmey, 2003; Zhang et al., 2012). Whether PIP2 regulates cortical actomyosin network organization in the C. elegans embryo is not known.
The subcellular distribution of PIP2 in the C. elegans zygote is also unclear. In other systems, PIP2 can distribute unevenly in the plasma membrane, for instance accumulating in macrodomains in nascent phagosomes and membrane ruffles or at the leading edge of motile cells, which all exhibit curved membranes that are sites of actin reorganization (Chierico et al., 2015; McLaughlin et al., 2002; Zhang et al., 2012). Accordingly, PIP2 can stimulate actin polymerization in curved but not flat model membranes (Gallop et al., 2013). Interestingly, PIP2 patches assemble at the leading edge of neuronal PC12 cells before F-actin accumulates, but their formation also depends on F-actin (Golub and Caroni, 2005; Golub and Pico, 2005). Moreover, F-actin enrichment is thought to drive clustering of PIP2-containing macrodomains, which in turn further regulates actin polymerization and branching (reviewed by Chichili and Rodgers, 2009). Overall, PIP2 and F-actin polymerization exhibit reciprocal positive feedback regulation in several systems.
The single C. elegans PIP5K1 is PPK-1, which can synthesize PIP2 from PI4P in vitro and in vivo (Weinkove et al., 2008). Overexpression of PPK-1 in developing neurons increases the level of PIP2 and results in extended filopodial-like structures, probably through changes in the actin cytoskeleton (Weinkove et al., 2008). In the somatic gonad, PPK-1 is important for F-actin organization and gonad contractility (Xu et al., 2007). Moreover, PPK-1 is enriched on the posterior cortex of one-cell embryos and has been implicated in the regulation of spindle positioning, but not of cell polarity (Panbianco et al., 2008). However, in other systems, PIP2 can modulate polarity by recruiting PAR proteins and reorganizing the actin cytoskeleton. Thus, in the Drosophila follicular epithelium, PIP2 recruits DmPar3 to the apical plasma membrane to maintain apical–basal polarity (Claret et al., 2014). Moreover, PIP2 might mediate interactions between PAR proteins, the actomyosin network and the plasma membrane in the fly oocyte (Gervais et al., 2008), as well as regulate apical constriction in the fly embryo (Guglielmi et al., 2015). Motile cells, such as mammalian neutrophils or Dictyostelium discoideum, also rely on PIP2, together with PIP3, for actin network reorganization during migration (reviewed by Wu et al., 2014). To summarize, in many systems, PIP2 is essential for proper F-actin organization and cell polarization, but whether this applies to C. elegans embryos is unknown. In this study we investigated the cortical distribution and function of the plasma membrane lipid component PIP2 in the worm zygote.
RESULTS
The PIP2 biomarker GFP::PHPLC1δ1 is present in dynamic polarized cortical structures in one-cell C. elegans embryos
While monitoring the distribution of components involved in asymmetric division of the C. elegans zygote with confocal spinning disk microscopy, we discovered that the PIP2 biomarker GFP::PHPLC1δ1 (Audhya et al., 2005) formed distinct and dynamic structures at the cell cortex (Fig. 1A-J; Fig. S1A; Movie 1). The pleckstrin homology (PH) domain of mammalian phospholipase C1δ1 (PLC1δ1) binds to PIP2 in vitro and in cells with high specificity and affinity (Garcia et al., 1995; Lemmon et al., 1995; Várnai and Balla, 1998). Moreover, in vertebrate cells, the lateral diffusion of GFP-PHPLC1δ1 resembles that of PIP2 (Golebiewska et al., 2008; Hammond et al., 2009), and membrane domains formed by fluorescently labeled PIP2 overlap with those monitored by GFP-PHPLC1δ1 (Chierico et al., 2015). Therefore, GFP-PHPLC1δ1 is well suited to monitor PIP2 dynamics in live cells.
The PIP2 biomarker GFP::PHPLC1δ1 is enriched in dynamic cortical structures. (A-J) Differential interference contrast (DIC) (A,C,E,G,I, middle plane) and spinning disk confocal imaging (B,D,F,H,J, cortical plane of a different embryo at the corresponding stages, with boxed regions magnified below) of one-cell C. elegans embryos at indicated stages expressing GFP::PHPLC1δ1. See Movie 1. Here and in subsequent figures, the embryo anterior is to the left, time is indicated in min:s, with 00:00 corresponding to the time of centration/rotation (t0 in K-M). (K-M) Fraction of cell cortex covered (K), average size (L) and Elongation Index (M; larger values correspond to more-elongated shapes) of segmented PIP2 cortical structures over time. The timing of pseudocleavage and mitosis are indicated. Here and in subsequent figures, quantification for anterior (orange) and posterior (green) embryo sides is shown as mean±s.d. n=39 embryos. See Materials and Methods for detailed description of how such graphs were generated. Scale bar: 10 µm.
Initially, when the cortical actomyosin network was contractile throughout the early C. elegans embryo, PIP2 was present weakly and evenly on the cell cortex (data not shown). Thereafter, when the actomyosin network began to move towards the anterior at the onset of polarity establishment, striking elongated cortical structures enriched in PIP2 became apparent, primarily on the anterior side of the embryo (Fig. 1A,B,K; Fig. S1A, top). Such PIP2 cortical structures had an initial average area of ∼2.5 µm2 and elongated as the zygote progressed through the cell cycle (Fig. 1L,M). All elongated PIP2 cortical structures moved anteriorly during polarity establishment, distributing in a clearly polarized manner at pseudocleavage, when they covered ∼15% of the anterior cortical surface (Fig. 1C,D, arrow, K). During the subsequent centration/rotation stage, PIP2 cortical structures decreased in size (Fig. 1E,F, arrowhead), before disappearing almost completely, with some remaining small foci by the time of nuclear envelope breakdown (NEDB) (Fig. 1G,H, arrowheads). A few elongated cortical structures reappeared during cytokinesis, primarily in the anterior (Fig. 1I,J). Unlike the structures visible when imaging the cortical plane, discrete PIP2 entities were barely detectable as slight thickenings of the plasma membrane in the embryo middle plane (Fig. S1A, arrowhead), probably explaining why they were not noted previously (Audhya et al., 2005; Blanchoud et al., 2010; Panbianco et al., 2008).
We aimed to verify the cortical distribution revealed by GFP::PHPLC1δ1 using fluorescently labeled PIP2, delivering Bodipy-FL-PIP2 to embryos the eggshells of which had been permeabilized using perm-1(RNAi) (Carvalho et al., 2011) (Fig. S1B-D). We found that Bodipy-FL-PIP2 provided to otherwise wild-type embryos distributed in dynamic polarized cortical structures akin to those observed with mCherry::PHPLC1δ1 (Fig. S1B,C; Movie 2). Moreover, delivering Bodipy-FL-PIP2 to embryos expressing mCherry::PHPLC1δ1 established that the two components overlapped (Fig. S1D). Overall, we concluded that GFP::PHPLC1δ1 is a faithful marker of PIP2 in one-cell C. elegans embryos, where it is present in dynamic and polarized structures at the plasma membrane.
A–P polarity cues regulate the polarized distribution of PIP2 cortical structures
We aimed to determine what regulates the polarized distribution of PIP2 cortical structures, which is particularly apparent during pseudocleavage. We found that PIP2 cortical structures did not overlap with GFP::PAR-2, which marks the posterior cortical domain (Fig. 2A), but did overlap with GFP::PAR-6, which marks the anterior polarity domain (Fig. 2B; Movie 3). Moreover, we observed that PIP2 cortical structures overlapped with elongated GFP::PAR-6 cortical structures (Fig. 2B, arrow), but not with GFP::PAR-6 foci (Fig. 2B, arrowhead), which are two distinct cortical populations of GFP::PAR-6 (Beers and Kemphues, 2006; Robin et al., 2014; Rodriguez et al., 2017; Wang et al., 2017).
PIP2 cortical structures depend on A–P polarity. (A,B) Dual-color spinning disk confocal cortical imaging of pseudocleavage embryos harboring the indicated pairs of fusion proteins, with high magnification views of the boxed insets. (A) mCherry::PHPLC1δ1/GFP::PAR-2, n=5. (B) mCherry::PHPLC1δ1/GFP::PAR-6, n=5. Elongated cortical structures (indicated by an arrow) but not foci (indicated by an arrowhead) of GFP::PAR-6 overlap with mCherry::PHPLC1δ1. See Movie 3. C,E,G Cortical plane at pseudocleavage and mitosis from spinning disk confocal imaging of control (C; n=39), par-3(RNAi) (E; n=10) or par-2(RNAi) (G; n=11) embryos expressing GFP::PHPLC1δ1. (D,F,H) Fraction of cell cortex covered by segmented PIP2 structures in the conditions corresponding to C,E,G with an expanded view of the evolution during mitosis. The overall cortical area covered at the 250 s time point is shown in a boxplot and compared by statistical analysis (Wilcoxon Rank Sum test/Mann–Whitney test). Scale bars: 10 µm.
We next tested whether the polarized distribution of PIP2 cortical structures depended on A–P polarity cues. Compared with the control condition, we found that, upon par-3(RNAi), PIP2 cortical structures distributed more uniformly (compare Fig. 2C,D with 2E,F, and Fig. S2A,B with S2C,D), except for the very posterior of the embryo (Fig. S2C; Fig. 2E, Pseudocleavage), consistent with the known slight posterior clearing of the actomyosin network upon PAR-3 inactivation (Kirby et al., 1990; Munro et al., 2004). Moreover, we found that, upon par-2(RNAi), PIP2 cortical structures first moved anteriorly (Fig. 2G,H, pseudocleavage, Fig. S2E), but then distributed in a more uniform manner (Fig. 2G,H, Mitosis, Fig. S2F), in line with PAR-2 being dispensable for polarity establishment, but essential for polarity maintenance (Cuenca et al., 2003; Hao et al., 2006; Munro et al., 2004). Furthermore, depletion of PAR-2 or PAR-3 in embryos expressing mNG::PHPLC1δ1 and Lifeact::mKate-2 established that the boundary of the two domains along the A–P axis was similar (Fig. S2A-I, Movie 4). This probably also explains why PIP2 cortical structures do not occupy mutually exclusive territories in par-3(RNAi) and par-2(RNAi) embryos, because their distribution coincided with the essentially uniform presence of cortical F-actin. Together, these findings established that the asymmetric distribution of PIP2 cortical structures is regulated by PAR-dependent A–P polarity cues.
PIP2 cortical structures colocalize partially with actin and fully with ECT-2, RHO-1 and CDC-42
Given that PIP2 and F-actin are interdependent in many systems, we tested whether these two components overlapped at the cell cortex of C. elegans embryos. As shown in Fig. 3A and Movie 5, we found a partial overlap of Lifeact::mKate-2, which monitors F-actin (Reymann et al., 2016), and of PIP2 cortical structures marked by mNeonGreen::PHPLC1δ1 (mNG::PHPLC1δ1). By contrast, we detected no substantial overlap between mCherry::PHPLC1δ1 and GFP::NMY-2 (Fig. 3B; Movie 6). Strikingly, we also found that PIP2 cortical structures marked by mCherry::PHPLC1δ1 fully colocalized with GFP::ECT-2, GFP::RHO-1 and GFP::CDC-42 (Fig. 3C-E; Movie 7).
PIP2 cortical structures overlap with ECT-2, CDC-42 and RHO-1, as well as partially with actin. (A-E) Dual-color spinning disk confocal cortical imaging of pseudocleavage embryos harboring the indicated pairs of fusion proteins, with high magnification views of the boxed regions. (A) mNeonGreen::PHPLC1δ/Lifeact::mKate-2, n=46; see Movie 5. (B) mCherry::PHPLC1δ1/GFP::NMY-2, n=9; see Movie 6. (C) mKate2-PHPLC1δ1/GFP::ECT-2, n=13; (D) mCherry::PHPLC1δ1/GFP::RHO-1, n=9; (E) mCherry::PHPLC1δ1/GFP::CDC-42, n=7; see Movie 7. Scale bars: 10 µm.
We tested whether RHO-1 or CDC-42 are needed for PIP2 cortical structures using partial RNAi-mediated depletion, since full depletion leads to sterility (Schonegg and Hyman, 2006). Upon rho-1(RNAi), we found that PIP2 structures formed normally but distributed symmetrically, consistent with the requirement of RHO-1 for A–P polarity (Fig. S3A,C,J,K,M,N). Thereafter, PIP2 cortical structures became polarized later than normal (Fig. S3B,D,G,H), probably through the PAR-2-dependent pathway (Motegi et al., 2011; Zonies et al., 2010). PIP2 cortical structures also formed upon cdc-42(RNAi), with minor and variable shape and size alterations compared with the control (Fig. S3E,F,L,O). The PIP2 structures first segregated towards the anterior in cdc-42(RNAi) embryos, but then extended posteriorly (Fig. S3E,F,I), consistent with CDC-42 being required for polarity maintenance (Motegi and Sugimoto, 2006; Schonegg and Hyman, 2006).
Overall, we concluded that RHO-1 and CDC-42 are dispensable for PIP2 cortical structure formation and, importantly, that such structures colocalize partially with F-actin, as well as completely with ECT-2, RHO-1 and CDC-42.
PIP2 cortical structures and the F-actin cytoskeleton move in concert
Live imaging of control, par-3(RNAi) and par-2(RNAi) embryos expressing Lifeact::mKate-2 and mNG::PHPLC1δ1 suggested that movements of PIP2 cortical structures and the F-actin network are coupled (Fig. S2A-I, Movie 5). To investigate this potential coupling quantitatively, we used particle image velocimetry (PIV) to simultaneously analyze cortical flows of Lifeact::mKate-2 and mNG::PHPLC1δ1 during polarity establishment (Fig. 4A-C) (Thielicke and Stamhuis, 2014). This analysis revealed highly correlated local flow velocities and directions at each time point (Fig. 4B,C; Fig. S4A,B). Analogous findings were made when comparing mCherry::PHPLC1δ1 and GFP::CDC-42 (Fig. S4C). By contrast, no strong correlation was found between mCherry::PHPLC1δ1 and the caveolin marker CAV-1::GFP (Fig. S4C), which might mark lipid rafts (Kurzchalia and Ward, 2003; Kurzchalia et al., 1999; Merris et al., 2003). Together, these results demonstrated that movements of PIP2 cortical structures and the F-actin network are coupled.
Coupling between PIP2 cortical structures and F-actin. (A) Spinning disk confocal cortical imaging of embryos expressing mNG::PHPLC1δ1 and Lifeact::mKate-2 (first column), corresponding binary thresholded images (second column) and PIV velocity vectors (third column), with high magnification views of boxed regions (arrow direction and length represent flow direction and velocity, respectively). (B) Correlation between mNG::PHPLC1δ1 and Lifeact::mKate-2 velocities in the same position and time, represented by a color scale dependent on their spatial density, from denser to sparser (red, yellow, light blue, respectively). Pearson correlation coefficient: ρ=0.61, P<10E−16 with Matlab precision, unpaired t-test. n=13 embryos for B-D. (C) Angle distribution between flow velocity vectors of mNG::PHPLC1δ1 and Lifeact::mKate-2 in the same position and time. The angle distribution peaks at θ=0° and decays exponentially thereafter (cut-off angle: θ=38°). Two independent velocity fields cannot result in the observed angle distribution (P<10E−16 with Matlab precision, χ2-test). (D) Cross-correlation between thresholded binary movies of mNG::PHPLC1δ1 and Lifeact::mKate-2, shifting Lifeact::mKate-2 with different time intervals relative to mNG::PHPLC1δ1. The boxed region is magnified on the right, showing that maximal overlap is achieved with a time shift of –9.3±1.5 s (mean±s.d.), irrespective of the order in which the two signals are recorded (see Materials and Methods). (E) Embryo expressing mNG::PHPLC1δ1 and Lifeact::mKate-2; the boxed region is magnified on the right and shows snapshots from the corresponding movie, illustrating that a PIP2 cortical structure (mNeonGreen::PHPLC1δ1, indicated by the arrows) moves ahead of polymerizing F-actin (Lifeact::mKate-2). The dot marks the starting point of the PIP2 structure. The brightness of Lifeact::mKate-2 in this image is increased to clearly show the polymerizing F-actin behind the PIP2 structures. (F,H,J) Cortical plane at pseudocleavage from spinning disk confocal imaging of embryos treated as indicated and expressing GFP::PHPLC1δ1. (F) nmy-2(RNAi), (H) act-1(RNAi) and (J) tba-2(RNAi). Both act-1(RNAi) and tba-2(RNAi) are severe but partial depletion conditions, because more-complete depletion results in sterility. (G,I,K) Fraction of cell cortex covered by segmented PIP2 structures in nmy-2(RNAi) [(G) n=7], act-1(RNAi) [(I) n= 8 at pseudocleavage, n=12 at mitosis] or tba-2(RNAi) [(K) n=10] embryos. (J,K) Although PIP2 cortical structures form normally upon tba-2(RNAi), they are slightly more numerous on the embryo anterior during mitosis (compare with Fig. 2D). Scale bars: 10 µm.
We next addressed whether the movements of PIP2 cortical structures and of F-actin are either synchronous or exhibit a time shift, which could suggest that one component leads the other. Close examination of embryos expressing Lifeact::mKate-2 and mNG::PHPLC1δ1 suggested that cortical PIP2 structures are followed by polymerizing F-actin (Fig. 4E). To address this possibility quantitatively, we cross-correlated time-shifted images of Lifeact::mKate-2 and mNG::PHPLC1δ1, and determined that maximal overlap between the two signals occurred when mNG::PHPLC1δ1 was ∼9.3±1.5 s ahead of Lifeact::mKate-2 (Fig. 4D). Overall, we concluded that PIP2 cortical structures move together with, but slightly ahead of, polymerizing F-actin filaments.
Actin polymerization drives the movement and formation of PIP2 cortical structures
Given that PIP2 cortical structures are followed by F-actin filaments, we hypothesized that actin polymerization might push PIP2 cortical structures. Compatible with this possibility, we found that the velocity of PIP2 cortical structures was ∼0.17±0.03 µm/s (Fig. S4D,E), in the range of actin polymerization-driven motility in other systems (Brangbour et al., 2011; Cameron et al., 2000; Carlsson, 2010; Mogilner and Oster, 1996). To test this possibility, we impaired actin polymerization by partially depleting the profilin PFN-1, which binds and functions together with the Formin Homology protein CYK-1 to promote polymer assembly (Severson et al., 2002; Velarde et al., 2007). A disruption of F-actin filaments monitored by Lifeact::mKate-2 was observed in pfn-1(RNAi) embryos (Fig. S5A, top, Movie 9). Importantly, we also found that the overall velocity of PIP2 cortical structures was reduced to ∼0.06±0.04 µm/s in pfn-1(RNAi) embryos (Fig. S5B). Moreover, whereas control embryos exhibited fast and directed movement (Fig. S5C, Movie 8), in half of the pfn-1(RNAi) embryos (n=4/8), which were the least affected, given that cytokinesis was still occurring, PIP2 cortical structures moved in a somewhat directed manner, albeit at lower velocities (Fig. S5,D). In more strongly affected pfn-1(RNAi) embryos (n=4/8), in which cytokinesis did not occur, PIP2 cortical structures moved only very locally and in seemingly random directions with occasional jumps (Fig. S5E, Movie 10). Together, these findings indicated that PIP2 cortical structure movements are driven by actin polymerization.
Given the tight coupling between cortical PIP2 structures and F-actin, we investigated whether the actomyosin network regulates not only the movement, but also the formation of PIP2 structures. We found that, in nmy-2(RNAi) embryos, in which actomyosin network contractility is abolished (Munro et al., 2004), PIP2 cortical structures were present (Fig. 4F,G), but more elongated than usual (Fig. S6A,B). Moreover, PIP2 cortical structures were distributed symmetrically in nmy-2(RNAi) embryos (Fig. 4F,G), as expected from the known requirement of NMY-2 in A–P polarity (Guo and Kemphues, 1996). Therefore, formation of PIP2 cortical structures does not depend on a contractile actomyosin cortex. By contrast, we found that few PIP2 cortical structures form in act-1(RNAi) embryos (Fig. 4G,H,I; Fig. S6C-F). Moreover, acute impairment of F-actin through treatment of perm-1(RNAi) embryos with cytochalasin D led to the disappearance of PIP2 cortical structures (Fig. S6G,H). However, PIP2 cortical structures remained upon depletion of the α-tubulin TBA-2 (Fig. 4J,K), indicating that they form independently of the microtubule cytoskeleton. Overall, we concluded that the formation of PIP2 cortical structures depends on F-actin.
Reducing the cellular level of PIP2 impacts F-actin distribution
We set out to address whether, conversely, PIP2 regulates F-actin organization. If so, then changing the level of PIP2 should alter actomyosin network organization. We tested this prediction first by depleting PIP2. To this end, we activated phospholipase C, an enzyme that cleaves PIP2, by delivering ionomycin and Ca2+ into perm-1(RNAi) embryos (Fig. S7A) (Hammond et al., 2012; Várnai and Balla, 1998). Cleavage of PIP2 at the plasma membrane was monitored by the gradual loss of GFP::PHPLC1δ1 plasma membrane signal, which enabled us to determine the time t1/2 when half of the initial GFP::PHPLC1δ1 plasma membrane fluorescence signal disappeared in the embryo middle plane (Fig. S7B,C). We found that PIP2 removal following ionomycin/Ca2+ treatment during pseudocleavage led to a rapid change in embryo shape on the anterior side, coincident with altered F-actin organization (compare Fig. 5A and Fig. 5B, Fig. S7E). An analogous F-actin alteration was observed following ionomycin/Ca2+ treatment during mitosis (Fig. 5C; Movie 11). Furthermore, we noted that ionomycin/Ca2+-treated embryos exhibited variable spindle positioning compared with the control condition (Fig. S8A-F).
A proper PIP2 cellular level is essential for correct organization of the actin cytoskeleton. (A-I) Confocal spinning disk imaging of embryos expressing GFP::PHPLC1δ1 and Lifeact::mKate-2 (LA::Kate-2, A-C: middle plane, D-I: cortical plane). (A) DMSO-treated perm-1(RNAi) control embryos. (B,C) perm-1(RNAi) embryos treated with ionomycin/Ca2+. t1/2=00:00: time when half of the plasma membrane GFP::PHPLC1δ1 fluorescence disappears; the time stamps are shown in yellow relative to this time here and in subsequent related panels. n=17, all stages combined. (B) t1/2 at pseudocleavage. The absence of coverslip, which is needed to preserve fragile perm-1(RNAi) embryos, prevents their flattening, such that more surface ruffles are apparent. In addition, the pseudocleavage furrow moves towards the anterior and either remains there (n=4, as shown) or relaxes (n=4, not shown). (C) t1/2 at mitosis. See Movie 11. (D-I) Control (D,G) and ocrl-1(RNAi) unc-26(s1710) (E,F,H,I) embryos during pseudocleavage (D-F) or mitosis (G-I). (E,H) Class 1 phenotype (n=43/72 scored with GFP::PHPLC1δ1 or mCherry-PHPLC1δ1 only, n=3/7 scored by Lifeact::mKate-2 only, n=3/7 scored by GFP::-PHPLC1δ1 and Lifeact::mKate-2); arrows indicate immotile structures, arrowhead indicates motile structures. (F,I) Class 2 phenotype (n=29/72 embryos scored with GFP::PHPLC1δ1 or mCherry-PHPLC1δ1 only, n=4/7 scored by Lifeact::mKate-2 only, n=4/7 scored by GFP::-PHPLC1δ1 and Lifeact::mKate-2). Dashed lines in G-I show the positions utilized to create the corresponding kymographs in J-L. See Movie 13. (J-L) Corresponding kymographs aligned at cytokinesis. Arrow indicates immotile structures, arrowhead indicates motile structures. Motile cortical PIP2 structures eventually move towards the cleavage furrow, partly correcting the aberrant PIP2 cortical domain distribution, consistent with a cortical domain correction mechanism operating at this stage (Schenk et al., 2010). Entire duration of kymographs, in min:s: (J) 16:30, (K) 20:00 and (L) 19:00. Scale bars: 10 µm.
To test whether alterations in F-actin organization cause the embryo shape change following ionomycin/Ca2+ treatment, we added latrunculin A to such embryos, thus depolymerizing the F-actin network. As shown in Fig. S7F and Movie 12, we found that this resulted in normally shaped embryos. Therefore, shape changes following PIP2 removal are F-actin dependent. Although we cannot formally rule out that ionomycin/Ca2+ caused these phenotypes for another reason, these results taken together indicated that PIP2 is crucial for proper F-actin distribution and, thus, the shape of the C. elegans zygote.
Increasing the cellular level of PIP2 impacts F-actin distribution via enhanced actin polymerization
We sought to further examine the relationship between PIP2 and F-actin by increasing the level of PIP2. We investigated whether this could be achieved by altering individual enzymes from the PIP2 biosynthetic pathway using RNAi or mutant animals (Fig. S7A, Table S1). Using both RNAi and an age-1(m333) null mutant, we first targeted AGE-1, the catalytic subunit of the sole C. elegans Class I PI3K, which can generate PIP3 from PIP2. We found that age-1(RNAi) embryos did not exhibit PIP2 alterations (Table S1). Likewise, whereas the progeny of age-1 null mutants arrest as dauer larvae (Larsen et al., 1995; Morris et al., 1996), we found no change in PIP2 in the corresponding embryos (Table S1), perhaps because compensatory mechanisms operate to keep the PIP2 level constant (Ayyadevara et al., 2009; Shmookler Reis et al., 2012). Furthermore, we did not find another single RNAi or mutant condition with altered PIP2 (Table S1), possibly because of redundancy among enzymes in PIP2 biosynthesis. With this in mind, we jointly inactivated ocrl-1 and unc-26, for the following reasons. OCRL-1 is an inositol 5-phosphatase that hydrolyzes PIP2 to PI4P, and its depletion leads to increased PIP2 on C. elegans phagosomes (Cheng et al., 2015). Moreover, UNC-26 is homologous to Synaptojanin, a polyphosphoinositide phosphatase that also hydrolyzes PIP2 to PI4P, and the impairment of which results in vesicle trafficking defects and cytoskeletal abnormalities in the worm nervous system (Charest et al., 1990; Harris et al., 2000).
We jointly depleted the function of these two PIP2 phosphatases, using RNAi for ocrl-1 and a mutant for unc-26. By comparing cortical mCherry-PHPLC1δ1 mean intensity values, we found that ocrl-1(RNAi) unc-26(s1710) embryos exhibited an increased overall level of PIP2 (Fig. S9A,B). Importantly, this led to drastic alterations in PIP2 monitored by GFP::PHPLC1δ1 or mCherry-PHPLC1δ1 (compare Fig. 5D and 5E,F top; Fig. S9C-H). First, in addition to motile PIP2 structures, we found immotile PIP2 clusters residing between the eggshell and the plasma membrane (Fig. 5E,H,K, arrows; Fig. S9I, arrowhead). Second, motile PIP2 structures did not disappear as readily after pseudocleavage as they normally do (Fig. 5H,I, compare with 5G; Fig. S9D). Third, we found that motile PIP2 structures exhibited altered distribution in all ocrl-1(RNAi) unc-26(s1710) embryos (Fig. 5E,F; Fig. S9F,H). In some cases (hereafter referred to as class I embryos, n=43/72), anteriorly directed movements of PIP2 cortical structures did not stop at pseudocleavage, but instead continued until the end of mitosis, resulting in a small anterior PIP2 domain [compare Fig. 5G,J (top) with Fig. 5H (top),K; Movie 13]. In class II ocrl-1(RNAi) unc-26(s1710) embryos (n=29/72), weak anteriorly directed movement of cortical PIP2 was initiated, but PIP2 structures then became distributed throughout the cortex by the end of the first cell cycle, except at the very posterior (Fig. 5I,L). Whereas a clear cytokinesis furrow formed in all class 1 embryos, this happened in only 14/29 class 2 embryos; this subset exhibited the most pronounced anteriorly directed movements. Overall, we concluded that depletion of PIP2 5-phosphatases was less pronounced in class 1 than in class 2 embryos, with the severe phenotype in the latter perhaps reflecting an impact on multiple processes. Interestingly, imaging ocrl-1(RNAi) unc-26(s1710) embryos expressing Lifeact::mKate-2 (class 1 n=3/7, class 2 n=4/7) or Lifeact::mKate-2 and GFP::PHPLC1δ1 (class 1 n=3/7, class 2 n=4/7) revealed that F-actin distributed as PIP2 cortical structures (Fig. 5D-L). We also noted that spindle positioning was more variable in both class 1 and class 2 ocrl-1(RNAi) unc-26(s1710) embryos than in the control condition (Fig. S8G-N). These findings established that an increase in PIP2, as achieved in class 1 embryos, leads to sustained cortical flows towards the anterior side. Moreover, although there might also be indirect effects of PIP2 5-phosphatase depletion on phosphoinositides other than PIP2, these results further indicated that PIP2 regulates actin cytoskeletal organization in one-cell C. elegans embryos.
We sought to investigate whether such regulation of PIP2 is exerted through the promotion of actin polymerization. We reasoned that, if this were the case, then enhancing actin polymerization in a different manner might mimic the unc-26(s1710)/ocrl1(RNAi) phenotype. Therefore, we treated perm-1(RNAi) embryos with jasplakinolide, which promotes actin polymerization (Fig. S10). Upon such treatment, large actin clumps formed, sometimes leading to detachment of the actin cytoskeleton from the plasma membrane (n=7/17; data not shown). However, the actin cytoskeleton moved exaggeratedly towards the anterior in most embryos treated with jasplakinolide during pseudocleavage (Fig. S10B,D,H; n=10/17), as in class 1 unc-26(s1710)/ocrl1(RNAi) embryos. To further test the hypothesis that PIP2 regulation is exerted through actin polymerization promotion, we treated permeabilized unc-26(s1710)/ocrl1(RNAi) embryos with low concentrations of cytochalasin D (250-500 nM) (Fig. S10A-F). We reasoned that this should ameliorate the unc-26(s1710)/ocrl1(RNAi) phenotype if it is caused by increased actin polymerization. This expectation was verified: whereas DMSO-treated perm-1(RNAi)/unc-26(s1710)/ocrl1(RNAi) embryos usually exhibited class 1 or class 2 phenotypes (n=6 and n=1, respectively, with one additional embryo exhibiting no apparent phenotype), all cytochalasin D-treated perm-1(RNAi)/unc-26(s1710)/ocrl1(RNAi) embryos polarized normally (Fig. S10A,C,E; n=6). Together, these experiments established that PIP2 regulates F-actin polymerization.
PIP2 is needed for RHO-1 and CDC-42 cortical structures
The regulation of actin polymerization and F-actin organization by PIP2 could conceivably be mediated through the actin regulators ECT-2, RHO-1 and CDC-42, with which PIP2 cortical structures coincide (Fig. 3). We explored this possibility for RHO-1 and CDC-42 by depleting PIP2 using ionomycin/Ca2+. This resulted in the loss of both cortical GFP::RHO-1 (n=5) and GFP::CDC-42 (n=7) if PIP2 depletion occurred before pseudocleavage (Fig. S11A-D). Moreover, PIP2 depletion after pseudocleavage resulted in cortical GFP::RHO-1 loss (n=2), but generally not in that of GFP::CDC-42 structures (Fig. S11E, n=5/7).
Given the above, altering PIP2 cortical structure distribution might likewise change that of RHO-1 and CDC-42. To test this possibility, we examined GFP::RHO-1 and GFP::CDC-42 distribution in unc-26(s1710)/ocrl1(RNAi) embryos, in which PIP2 cortical structure organization is altered (Fig. 5). We found that GFP::RHO-1 and GFP::CDC-42 distributions mirrored that of mCherry::PHPLC1δ1 in such embryos (Fig. S11F-K). Overall, these results indicated that PIP2 is needed for RHO-1 cortical structures early and late during the first cell cycle, and also for CDC-42 cortical structures, primarily until the pseudocleavage stage.
An appropriate level of PIP2 is essential for proper PAR polarity establishment and maintenance
It is well known that the actomyosin network is essential for A–P polarity in C. elegans zygotes (Guo and Kemphues, 1996; Hill and Strome, 1990; Munro et al., 2004; Strome and Wood, 1983). Given that a proper level of PIP2 is essential for correct actomyosin network organization, we tested whether it is also important for A–P polarity. We investigated the impact of excess PIP2 on polarity using ocrl-1(RNAi) unc-26(s1710) embryos expressing mCherry::PHPLC1δ1 and GFP::PAR-2 or GFP::PAR-6 (Fig. 6A-L). We found that the distribution of GFP::PAR-2 and GFP::PAR-6 domains changed from early on in development, consistent with alterations in motile PIP2 structures and F-actin. Thus, for GFP::PAR-6, either a small domain formed on the very anterior (Fig. 6B,E; class 1, n=5/9; Movie 14) or the fusion protein remained present over the entire cortex, except on the very posterior (Fig. 6C,F class 2; n=4/9). As expected, GFP::PAR-2 distributed reciprocally, either expanding drastically towards the anterior (Fig. 6H,K; class 1, n=12/22; Movie 15) or remaining restricted to the very posterior (Fig. 6I,L; class 2, n=10/22). Moreover, we observed that spindle positioning was affected in a variable manner in ocrl-1(RNAi) unc-26(s1710) embryos (Fig. S8G-N), as anticipated from altered A–P polarity and potentially other consequences of excess PIP2. Overall, these results indicated that a correct level of PIP2 is needed for proper F-actin network localization and, as a consequence, appropriate PAR polarity and accurate spindle positioning.
Proper PIP2 cellular level is essential for correct PAR polarity. (A-C,G-I) Confocal spinning disk cortical imaging of control (A,G) or ocrl-1(RNAi) unc-26(s1710) (B,C,H,I) embryos expressing mCherry::PHPLC1δ1 (mCherry::PH) and GFP::PAR-6 (A-C, n=5/9 class 1, n=4/9 class 2) or GFP::PAR-2 (G-I, n=12/22 class 1; n=10/22 class 2). Dashed lines indicate positions used to create the corresponding kymographs in D-F. See Movies 14 and 15. (D-F,J-L) Corresponding kymographs aligned at cytokinesis. Entire duration of kymographs, in min:s: (D) 16:40, (E) 11:20 and (F-I) 17:40. (M-O) Images acquired by confocal imaging of embryos expressing GFP::PAR-2, 4.5 µm below the cortical plane. The dashed line marks the boundary of the PAR-2 domain. (M): DMSO-treated control perm-1(RNAi) embryo. (N,O): perm-1(RNAi) embryo treated with ionomycin/Ca2+ (t1/2=00:00) during pseudocleavage (n=6) or mitosis (n=3). See Movies 16-18. Scale bars: 10 µm.
In the above experiments, the level of PIP2 was in excess from the beginning of development, such that the impact on polarity might reflect a role either strictly during the establishment phase or during both the establishment and maintenance phases. We reasoned that one could test specifically a potential role for PIP2 in polarity maintenance by adding ionomycin/Ca2+ during pseudocleavage, after polarity establishment, to perm-1(RNAi) embryos expressing mCherry::PHPLC1δ1 and GFP::PAR-2. We found that GFP::PAR-2 expanded slowly towards the anterior, starting ∼3 min after t1/2 (Fig. 6M-O), with the pseudocleavage furrow initially moving anteriorly, and then either disappearing (Fig. 6N, Movie 16; n=6/14), or remaining at the very anterior (Movie 17; n=8/14). Likewise, GFP::PAR-2 expanded slowly towards the anterior when t1/2 occurred at nuclear envelope breakdown (NEBD) (Fig. 6O, Movie 18). Together, these results indicated that an appropriate level of PIP2 is also essential for proper PAR polarity during the maintenance phase.
In principle, PIP2 could alter PAR polarity during the maintenance phase either through an impact on F-actin organization or via an actin-independent role. Unlike its well-known role during polarity establishment, a potential role of F-actin in polarity maintenance is somewhat controversial (Goehring et al., 2011; Hill and Strome, 1990; Liu et al., 2010; Severson and Bowerman, 2003). Considering our findings related to changes in the PIP2 level, we directly tested whether F-actin plays a role in polarity maintenance. We first added cytochalasin D to perm-1(RNAi) embryos expressing Lifeact::mKate-2 and GFP::PAR-2 after polarity establishment (Fig. 7A,B). Consistent with previous studies (Goehring et al., 2011; Hill and Strome, 1990), we found that cytochalasin D addition at this stage did not significantly alter GFP::PAR-2 distribution (Fig. 7A,B, Movie 19). However, we also found that cytochalasin D did not fully disrupt F-actin, because clumps of Lifeact::mKate-2 remained in the embryo anterior (Fig. 7B). Thus, we turned to inhibiting F-actin polymerization using latrunculin A, which resulted in complete F-actin depletion (Fig. 7C; n=12; Movie 20). We observed membrane invaginations that removed GFP::PAR-2 from the cortex into cytoplasmic aggregates (Fig. 7C, arrowhead), as previously reported (Goehring et al., 2011; Redemann et al., 2010). Importantly, we monitored changes in GFP::PAR-2 distribution as a function not of drug addition time, as done previously (Goehring et al., 2011), but of the time at which half of the Lifeact::mKate-2 fluorescence had disappeared from the membrane. In doing so, we found a decrease of the GFP::PAR-2 domain after t1/2 in all embryos analyzed (Fig. 7C, bottom; n=12) that was highly correlated with Lifeact::mKate-2 disappearance (Fig. S9J), demonstrating that F-actin is crucial for PAR polarity maintenance.
F-actin impairment also affects GFP::PAR-2 during polarity maintenance. (A-C) Confocal spinning disk cortical imaging of perm-1(RNAi) embryos expressing GFP::PAR-2 and Lifeact::mKate-2 (middle plane), treated during early centration/rotation either with DMSO [(A) n=6], cytochalasin D [(B) n=5, movie acquired with binning=2], or latrunculin A [(C) n=18]. Arrowhead points to plasma membrane invagination (Redemann et al., 2010). See Movies 19 and 20. (D) Schematic working model (not to scale). PIP2 is enriched in dynamic and polarized structures at the cortex of one-cell Caenorhabditis elegans embryos, moving ahead of F-actin. These two components exhibit mutually reciprocal requirements: the formation of PIP2 cortical structures requires F-actin and a proper PIP2 level is essential for F-actin organization. Moreover, through its ability to organize the F-actin network properly, PIP2 is essential for proper PAR domain sizing and, thus, A–P polarity. See the main text for further details. For simplicity, only those actin filaments that move in concert with PIP2 cortical structures are represented on the top right. Scale bars: 10 µm.
Overall, we found that an appropriate level of PIP2 is essential for the correct sizing of PAR domains, through the reorganization of F-actin, during not only polarity establishment, but also polarity maintenance.
DISCUSSION
In this work, we demonstrated that PIP2 forms cortical structures in the one-cell C. elegans embryo. We showed that the formation and movement of these structures depend on F-actin and, reciprocally, that PIP2 regulates F-actin organization, revealing an interdependence of these two components in the worm zygote (Fig. 7D). Moreover, probably through its impact on the actin cytoskeleton, PIP2 is also needed for the correct sizing of PAR domains, demonstrating that a plasma membrane lipid component participates in A–P polarity in the C. elegans embryo.
PIP2 is present in discrete cortical structures in C. elegans zygotes
The distribution and dynamics of PIP2 at the plasma membrane of early C. elegans embryos were not clear before this work, primarily because only the middle embryo plane was analyzed in past investigations. Here, we assayed subcellular distributions at the cortex, where the function of PAR proteins and components that are crucial for asymmetric division is exerted. Thus, we discovered that PIP2 is present in dynamic polarized cortical structures marked by the PIP2 biomarker GFP::PHPLC1δ1 and Bodipy-FL-PIP2, consistent with recent observations mentioning non-uniform GFP::PHPLC1δ1 distribution (Rodriguez et al., 2017; Wang et al., 2017). Although patches of plasma membrane enriched in PIP2 have been observed elsewhere (Chierico et al., 2015; Golub and Caroni, 2005; McLaughlin et al., 2002; Zhang et al., 2012), the stereotyped progression through the first cell cycle of the large worm zygote enabled us to probe such cortical structures with unprecedented resolution. Why were these structures not observed previously? On top of not being noticeable when imaging the middle plane of the embryo, other plausible reasons are that PIP2 cortical structures appear only transiently during the cell cycle and that they are not preserved upon fixation (data not shown).
How do PIP2 cortical structures assemble? Two reasons lead us to propose that PIP2 cortical structures might form by redistributing existing PIP2 rather than by de novo synthesis through PIP5K1. First, PIP2 in other systems has been suggested to diffuse much faster than it is synthesized (McLaughlin et al., 2002), such that potential local synthesis is unlikely to dictate restricted PIP2 localization. Second, PPK-1, the sole PIP5K1 in C. elegans, is enriched in the posterior of the embryo (Panbianco et al., 2008), seemingly not where most PIP2 cortical structures reside. However, localization of PIP5K1 to PIP2 cortical structures might have been missed by analyzing fixed embryos and imaging the middle plane. Interestingly, we also found that PIP2 cortical structures form and move independently of actomyosin contractions powered by NMY-2. Nevertheless, PIP2 cortical structures might form at membrane protrusions or ruffles, which would be consistent with PIP2 stimulating F-actin polymerization in curved but not flat membranes (Gallop et al., 2013), and accumulating in membrane ruffles, nascent phagosomes and the leading edge of motile cells (reviewed by McLaughlin et al., 2002; Zhang et al., 2012). In other systems, increased PIP2 induces actin polymerization around membrane vesicles, generating actin comets that propel vesicles forward (Ma et al., 1998; Rozelle et al., 2000). We also found occasional vesicles and connected local F-actin accumulation upon increasing PIP2, although no actin comets were observed (data not shown).
In summary, we propose that PIP2 cortical structures form through the redistribution of existing PIP2 at the plasma membrane in the C. elegans zygote, perhaps preferentially at membrane protrusions or ruffles.
Interdependence of PIP2 and F-actin
PIP2 and F-actin exhibit a reciprocal relationship in a number of systems, and we show here that this is also the case in C. elegans. We found that PIP2 cortical structures and F-actin movements are coupled, with PIP2 structures moving slightly ahead, at velocities compatible with actin polymerization driving these movements. Moreover, impairing the profilin PFN-1, which is essential for microfilament assembly (Severson et al., 2002; Velarde et al., 2007), reveals that PIP2 structure movements are actin driven. This leads us to propose that actin polymerization pushes PIP2 cortical structures, reminiscently of other actin-dependent motility processes, such as that of Listeria monocytogenes (reviewed by Mogilner and Oster, 1996). While being pushed ahead of F-actin in C. elegans, PIP2 structures might recruit factors promoting actin polymerization and branching, such as ECT-1, RHO-1 and CDC-42, as in other systems (reviewed by Chichili and Rodgers, 2009). Intriguingly, a biosensor for active CDC-42 distributes on the cortex like PIP2 does (Cheng et al., 2015; Kumfer et al., 2010), indicating that CDC-42 at PIP2 structures is active. By contrast, the distribution of a biosensor for active RhoA overlaps with that of NMY-2 (Reymann et al., 2016; Tse et al., 2012). Given that we showed here that PIP2 cortical structures did not overlap with NMY-2, whereas they overlapped with GFP::RHO-1, the bulk of RHO-1 associated with PIP2 cortical structures might be inactive. Alternatively, given that RHO-1 colocalizes with its activating GEF ECT-2, this RhoA biosensor might not detect all active RHO-1 species. Furthermore, it is interesting that non-muscle myosin 2 contributes to actin network disassembly in fish keratinocytes (Wilson et al., 2010). Extrapolating from this observation, it is tempting to speculate that PIP2, by promoting F-actin assembly, and NMY-2, by promoting F-actin disassembly, in addition to powering network contractility, might together ensure proper F-actin dynamics in C. elegans embryos.
PIP2 and PAR-dependent polarity
PAR proteins are also distributed unevenly within their domain. Thus, cortical PAR-6 exists in two populations, one diffuse that depends on CDC-42 and one punctate that colocalizes with PAR-3 (Beers and Kemphues, 2006; Robin et al., 2014). Moreover, PAR-3 forms clusters that are crucial for proper polarity, and the assembly of which depends on PCK-3, CDC-42, as well as actomyosin contractility (Rodriguez et al., 2017; Wang et al., 2017). Moreover, the putative CDC-42 GAP CHIN-1 forms clusters on the posterior, and the transport of these clusters stabilizes the A–P boundary during polarity maintenance (Sailer et al., 2015). Intriguingly, we found that PIP2 cortical structures colocalized within the more diffuse cortical PAR-6 population, which lacks PAR-3 (Beers and Kemphues, 2006; Robin et al., 2014; Rodriguez et al., 2017; Wang et al., 2017). Moreover, we established that increasing PIP2 augmented the segregation of both PAR-6 populations to the anterior. Given that clustering of PAR-3 depends on actomyosin reorganization (Rodriguez et al., 2017; Wang et al., 2017), we propose that increasing the level of PIP2 might reorganize the actin cytoskeleton in a way that promotes PAR-3 clustering, thereby aiding segregation.
We showed that an appropriate PIP2 level is essential for proper polarity establishment and maintenance. When increasing the level of PIP2, the continued movement of PAR domains towards the anterior until the end of mitosis altered their relative size. This is reminiscent of changes in PAR domain sizing that occur upon RGA-3/4 depletion (Schonegg et al., 2007). However, whereas rga-3/4(RNAi) embryos exhibit a hypercontractile actomyosin network, embryos with increased PIP2 level do not. Thus, we propose that actomyosin contractility regulated by NMY-2 and F-actin organization regulated by PIP2 contribute in concert to the correct movements of the actomyosin network and, therefore, proper sizing of PAR polarity domains.
On the role of the actomyosin network in polarity establishment and maintenance
The actomyosin network plays a well-established role during polarity establishment, whereas its role during polarity maintenance has been debated (Goehring et al., 2011; Hill and Strome, 1990; Liu et al., 2010; Severson and Bowerman, 2003). Our results, together with those of others, indicated that the actomyosin network regulates PAR domains in two ways. First, when the actomyosin network moves anteriorly during the establishment phase, PAR domains alter their distribution accordingly. This relationship was clear before this work, and we reached an identical conclusion here. Second, actin has been suggested to play merely a passive role during the maintenance phase in preventing cortical PAR-2 removal through membrane invaginations driven by microtubules (Goehring et al., 2011). We showed here that this can lead to the near-total disappearance of cortical GFP::PAR-2, further emphasizing the importance of actin also during the maintenance phase.
Overall, our results in C. elegans are consistent with the role of PIP2 in F-actin reorganization and polarity in other organisms. Previous work in C. elegans showed that depletion of CSNK-1, which negatively regulates PPK-1 localization, although perturbing spindle positioning, does not impact polarity (Panbianco et al., 2008), perhaps because PPK-1 plays only a minor role in regulating the level of PIP2 in the zygote. By contrast, we established here that alterations in the level of PIP2 impair polarity establishment and maintenance during the first asymmetric division of C. elegans embryos.
MATERIALS AND METHODS
Worm strains
Nematodes were maintained at 24°C using standard protocols (Brenner, 1974). The following strains were used: GFP::PHPLC1δ1 (OD58, unc-119(ed3) III; ltIs38[pie-1p::GFP::PH(PLC1delta1)+unc-119(+)]) (Audhya et al., 2005); mCherry::PHPLC1δ1 (OD70, unc-119(ed3) III; ltIs44[pie-1p::mCherry::PH(PLC1delta1)+unc-119(+)]V) (Audhya et al., 2005); GFP::PAR-2 (TH129) and GFP::PAR-6 (TH110) (Schonegg et al., 2007); GFP::NMY-2 (LP162, nmy-2(cp13[nmy-2::gfp+LoxP]) I) (Dickinson et al., 2013); CAV-1::GFP (RT688, unc-119(ed3) III; pwIs281[CAV-1::GFP, unc-119(+)]) (Sato et al., 2006); mNeonGreen::PHPLC1δ1 (LP274, cpIs45[Pmex-5::mNeonGreen::PLCδ-PH::tbb-2 3′UTR+unc-119(+)] II; unc-119(ed3) III), mKate-2::PHPLC1δ1 (LP307, cpIs54[Pmex-5::mKate2::PLCδ-PH(A735T)::tbb-2 3′UTR+unc-119(+)] II; unc-119(ed3) III) and mCherry::PHPLC1δ1 (LP308, cpIs55[Pmex-5::mCherry-C1::PLCδ-PH::tbb-2 3′UTR+unc-119(+)] II; unc-119(ed3) III) (Heppert et al., 2016); Lifeact::mKate-2 (strain SWG001) (Reymann et al., 2016); GFP::RHO-1 (SA115, unc-119(ed3) III; tjIs1[pie-1::GFP::rho-1+unc-119(+)]) (Motegi et al., 2006); GFP::CDC-42 (SA131, unc-119(ed3) III; tjIs6[pie-1p::GFP::cdc-42+unc-119(+)]) (Motegi and Sugimoto, 2006); GFP::ECT-2 (SA125, unc-119(ed3) III; tjIs4[pie-1::GFP::ect-2+unc-119(+)]) (Motegi and Sugimoto, 2006); unc-26(s1710) (EG3027, unc-26(s1710) IV) (Charest et al., 1990); and age-1(m333), (DR722, age-1(m333)/mnC1 dpy-10(e128) unc-52(e444) II) (Riddle, 1988). Crosses were performed at 20°C to generate lines that were homozygous for all transgenes, which were then maintained at 24°C. For GFP::RHO-1 and mCherry::PHPLC1δ1 (only in Fig. 3D), as well as GFP::PHPLC1δ1 and Lifeact::mKate-2, worm lines were crossed and F1 progeny heterozygous for both transgenes were imaged.
RNAi
RNAi-mediated depletion was performed essentially as described elsewhere (Kamath et al., 2001), using bacterial feeding strains from the Ahringer (Kamath et al., 2003) or Vidal libraries (Rual et al., 2004) (gift from Jean-François Rual and Marc Vidal). RNAi for par-2 (Ahringer), par-3 (Ahringer), nmy-2 (Ahringer), act-1 (Vidal), tba-2 (Vidal), rho-1 (Vidal), cdc-42 (Vidal) and ocrl-1 (Ahringer) was performed by feeding L3-L4 animals with bacteria expressing the corresponding dsRNA at 24°C for 16-26 h. RNAi for pfn-1 (Vidal) was performed by feeding L4 and young adults with bacteria expressing dsRNA at 24°C for 72-96 h and imaging embryos of their offspring. PERM-1 is a sugar-modifying enzyme essential for forming the permeability barrier of the eggshell (Carvalho et al., 2011; Olson et al., 2012); RNAi for perm-1 (Ahringer) was performed by feeding L4 and young adults with bacteria expressing dsRNA at 20°C for 12-18 h. Double RNAi for ocrl-1 and perm-1 was performed by mixing bacteria expressing ocrl-1 dsRNA and perm-1 dsRNA in a 1:1 ratio and then feeding them to L3 and L4 young adults for 24-30 h at 24°C. The effectiveness of depletion was assessed phenotypically as follows: par-2(RNAi) and par-3(RNAi): symmetric spindle positioning and equal cell division; nmy-2(RNAi) and act-1(RNAi): absence of cortical ruffles, symmetric spindle positioning, no cytokinesis; tba-2(RNAi): defective pronuclear meeting, no centration/rotation, no spindle assembly, misplaced cytokinesis furrow; ocrl-1(RNAi) [in combination with unc-26(s1710)]: immotile PIP2 structures, class 1 or class 2 phenotype (Fig. 5E,F,H,I; Fig. 6B,C,H,I), altered centrosome and spindle positioning (Fig. S8H-K); perm-1(RNAi): successful action of added drug; pfn-1(RNAi): defective cortical F-actin network; cdc-42(RNAi): partial loss of polarity during the maintenance phase; rho-1(RNAi): absence of cortical ruffles.
Live imaging
Gravid hermaphrodites were dissected in osmotically balanced blastomere culture medium (Shelton and Bowerman, 1996) and the resulting embryos mounted on a 2% agarose pad. DIC time lapse microscopy (Fig. 1A,C,E,G,I) was performed at 25°C±1°C with a 100× (NA 1.25 Achrostigmat) objective and standard DIC optics on a Zeiss Axioskop 2 microscope. All other images were acquired at 23°C using an inverted Olympus IX 81 microscope equipped with a Yokogawa spinning disk CSU-W1 with a 63× (NA 1.42U PLAN S APO) objective and a 16-bit PCO Edge sCMOS camera. Images were obtained using 488-nm and 561-nm solid-state lasers with an exposure time of 400 ms and a laser power of 20-60%. For cortical imaging, three planes at the cell cortex (each 0.25 μm apart) were acquired. Cell cycle stages were determined using transmission light microscopy, imaging the middle plane in parallel (data not shown).
Image processing and analysis
Cortical images of GFP::PHPLC1δ1 used for quantification were processed as follows: the three cortical planes were z-projected using average intensity projection, after which a median filter of 1 pixel was applied. The background of the entire image was subtracted using the measured mean background in each frame. Signal intensity decay because of photobleaching was corrected with the Fiji plugin ‘bleach correction’ using the exponential fitting method (ImageJ Plugin CorrectBleach V2.0.2. Zenodo; https://zenodo.org/record/30769#.WvQPLKQvwuU). The entire cortical region was segmented by applying a binary automated histogram-based threshold, followed by iterated morphological operations. Cortical structures were segmented by applying a binary intensity threshold, calculated by fitting the pixel intensity histogram with a Gaussian function and setting the threshold at 4 σ from the Gaussian peak. The boundary between anterior and posterior domains was determined manually during pseudocleavage formation. Upon depletion of NMY-2 and ACT-1, whereby no pseudocleavage furrow forms, the boundary was set during mitosis. The fraction of the total area in each half covered by the segmented PIP2 cortical structures was determined in each case.
Curves of normalized cortical structures sizes were fitted with a sigmoidal model and synchronized, setting the sigmoid inflection point, which corresponded typically to the time of centration/rotation, as time t=0 s. Curves of normalized cortical structures sizes were aligned manually for act-1(RNAi) and unc-26(s1710) ocrl-1(RNAi) embryos using the clear landmark provided by NEDB as a reference, because a sigmoid function could not be fitted with the PH markers in these cases. Given that the time separating centration/rotation from NEDB is typically 150 s, t=0 was set at –150 s before NEDB for act-1(RNAi) and unc-26(s1710) ocrl-1(RNAi) embryos.
The Elongation Index was calculated using Eqn 1:
(1)
using the MATLAB image processing function ‘regionprops’. The Elongation Index was then normalized by a factor of 1/π, such that a square of 2×2 pixels has an Elongation Index of 1.
For assessing the A–P boundary of the segmented PIP2 cortical structures (GFP::PHPLC1δ1) and of F-actin (Lifeact::mKate-2) (Fig. S2), we automatically determined the extent of the PIP2 and F-actin domains relative to the whole embryo length using a Matlab script. For Lifeact::mKate2, a histogram-based method was used to keep only the 95% brightest pixels.
Cortical images obtained by live confocal spinning disk imaging shown in the figures were processed as follows: the three cortical planes were z-projected using maximum intensity projection, then a median filter of 1 was applied. The gray value fluorescence intensity of some transgenes (GFP::PHPLC1δ1 as heterozygote, GFP::PAR-2, GFP::PAR-6, Lifeact::mKate-2, mCherry::PHPLC1δ1 and mNeonGreen-PHPLC1δ1) was slightly variable, probably resulting from variable expression and/or folding of the fluorescent fusion protein. Therefore, the brightness and contrast of images resulting from embryos expressing these transgenes was adjusted accordingly. Such variability was especially pronounced in unc-26(s1710) ocrl-1(RNAi) embryos. Missing edges or corners in the following figures and movies result from one or multiple rotations to position the anterior of the embryo to the left: Fig. 4A, Fig. 5A,B,F,I, Fig. 6M,O, Fig. S2A,E,F, Fig. S3C,D, Fig. S5A, Fig. S9I, Fig. S10D, Fig. S11D, Movies 5, 9, 11, 12, 20. To compare the intensity of mCherry-PHPLC1δ1 in control embryos and unc-26(s1710) ocrl-1(RNAi) embryos, three cortical planes, acquired as described above, were z-projected by summing the intensity of all slices. The resulting mean intensities were then computed as follows. First, an Otsu threshold was used to retrieve the brightest elements, retaining only the biggest blob, which corresponded to the embryo. Values outside the embryo were averaged to obtain the mean background intensity, which was subtracted from the embryo pixel intensities. Thereafter, embryo pixel values were averaged to obtain the mean pixel intensity value.
Lipid delivery
BODIPY FL phosphatidylinositol 4,5-bisphosphate (Echelon Bioscience, C-45F6) (final concentration 10 µM) was delivered to perm-1(RNAi) embryos by adding it to the buffer in which gravid worms were dissected. Embryos were then mounted on a 2% agarose pad using petroleum jelly as a spacer to decrease the pressure exerted on the fragile perm-1(RNAi) embryos.
Cortical flow measurement, correlation analysis and PIP2 structure tracking
For particle image velocimetry (PIV) analysis, cortical image sequences of mNeonGreen::PHPLC1δ1 and Lifeact::mKate-2 were prepared by performing a maximum intensity z-projection of a stack of two planes (0.25 µm apart) and applying a median filter of 1 pixel. PIP2 cortical structures and the F-actin network were then segmented as follows: the embryo was first extracted from the background using a histogram-based automated threshold, keeping only blobs of a size superior to one-third of the biggest blob. The resulting binary images were deemed to be the embryo area. We applied a morphological erosion to the mNeonGreen::PHPLC1δ1 movies with a large structuring element (a disk 30 pixels in radius) to calculate the average value of the pixels not corresponding to PIP2 cortical structures; the PIP2 cortical structures were then segmented as the pixels of intensity higher than the computed average value multiplied by a scaling factor determined empirically (1.7). The extraction of the F-actin network was achieved by determining a histogram-based automated threshold on the morphological top-hat of the F-actin image. F-actin filaments and PIP2 cortical structures were segmented before PIV analysis to ensure that only flow fields in the region of interest were measured.
PIV was then performed to measure cortical flows using the MATLAB based PIVlab toolbox (Thielicke and Stamhuis, 2014), which splits each image of a movie into a regular grid, for which the size of grid cells is given by the user. The position of each grid cell in the next image is estimated by finding the maximum normalized cell-to-cell cross-correlation of equivalent sizes in a geometrical neighborhood called the ‘interrogation area’. Such PIV analysis was applied to mNeonGreen::PHPLC1δ1 and Lifeact::mKate-2 separately, after segmentation of the corresponding cortical structures. The choice of the grid cell sizes and interrogation areas resulted from a balance between two criteria: smaller cells allow one to compute displacements with high spatial resolution, but excessively small cells do not contain enough information to be reliably correlated with other cells; thus, the estimation of displacements of bigger cells is more reliable, but computed with lower resolution. We found empirically 32×32 pixels for cell sizes, and 64×64 pixels for interrogation areas, to be a good compromise.
The PIV velocity fields output for both mNeonGreen::PHPLC1δ1 and Lifeact::mKate-2 signals were compared in terms of angles between colocalized features and correlation of the norms. For each movie, angles between velocity vectors of colocalized features were computed and plotted on a histogram. The average angle value for each time point and each movie was also computed to monitor the coherence between the two vector fields over time. Similarly, we computed the correlations of the norms of all velocities in the two movies, for the whole movie, and also for specific times. The cut-off angle was defined as the θ 0 parameter of the curve of Eqn 2 fitted to the histogram:
(2)
Cross-correlation analysis was performed as follows. Movies used to calculate the cross-correlation were acquired by alternating the acquisition order of the red and green channels to prevent introducing a bias through the order of image acquisition. The colocalization of the thresholded PIP2 cortical structures and F-actin network for a variety of time shifts was computed considering a time shift Δt (positive or negative), the colocalization of the segmented PIP2 image at time t, and the segmented F-actin image at time t–Δt using Eqn 3:
(3)
Colocalization was computed in this manner from Δt=–(T–1) to Δt=(T–1), where T is the total duration of the movie. The Δt for which colocalization was maximal represented the time shift between PIP2 and F-actin. The mean time shift and its error were computed as follows. We fitted a parabola of the following equation:
(4)
to the location correlation as a function of the time shift. We calculated the best a, b and t0 parameters using a least-squares method, and input the standard deviations of the correlations to create a weight matrix used during the adjustment. The results were the mean time shift t0=9.3 s and the standard deviation sigma_t0=1.5 s.
To track PIP2 structures (Fig. 4E, Fig. S4D,E), embryos expressing mNG-PHPLC1δ1 were imaged with an exposure time of 50 ms, a laser power of 60% and a 70-ms frame rate. PIP2 structures were tracked manually on maximum intensity z-projection of the images containing the moving PIP2 structures of interest. The length of the track was obtained by re-slicing it using the Fiji plugin ‘Reslice’. The velocity was calculated from the corresponding number of time points and track length.
Spindle pole tracking
Spindle positioning tracks were generated by manually tracking spindle poles from NEBD to cleavage furrow formation using a MATLAB script that also computed the distance from the first to the last tracked point, providing the corresponding x and y coordinates and the maximum velocity in µm/s. Tracks were automatically placed into ellipses fitted around the embryo. The end positions of the spindle poles were determined at cytokinesis onset.
Drug addition
The eggshell was permeabilized by performing perm-1(RNAi) as described above. Gravid hermaphrodites were dissected in a cell culture dish with a glass bottom, and the resulting embryos imaged with an inverted confocal spinning disk microscope (see above). Drugs were added under the microscope while imaging to precisely control the timing of drug addition. The following drugs and concentrations in solution were utilized: 30 µM ionomycin (Calbiochem, 407950), 3-5 mM CaCl2 (Sigma-Aldrich, C5080), 20 µM cytochalasin D (AppliChem, 22144-77-0) and 12.5 µM latrunculin A (Sigma-Aldrich, 76343-93-6). For control movies, dimethyl sulfoxide (DMSO) at a concentration equivalent to the final DMSO concentration in the drug solutions was added to the buffer before dissection. For the drug delivery experiments shown in Fig. S10, gravid hermaphrodites were dissected in either 250 nM or 500 nM cytochalasin D (AppliChem, 22144-77-0) or 2.5 µM jasplakinolide (AdipoGen, AG-CN2-0037). Embryos were then imaged, acquiring 17 z-planes 0.5 µm apart starting every 30 s. For the experiments reported in Figs 5 and 6, whenever embryos were dissected in the drug-containing solution, we ensured that the action of the drug (as monitored by t1/2) took place >6 min after polarity was established.
Successful drug action was determined for each embryo by the disappearance of the PHPLC1δ1 fluorescence signal from the plasma membrane (ionomycin/Ca2+ and latrunculin A) and of Lifeact::mKate-2 from the cell cortex (cytochalasin D and latrunculin A). The time between drug addition and drug action was variable, probably because of variations in eggshell permeability upon perm-1(RNAi). Therefore, as a comparable reference time between embryos, we determined the time t1/2 (t inflection) when half of fluorescence at the plasma membrane has disappeared, as follows: the total cortical region of the embryo was segmented by applying a binary automated histogram-based threshold; fluorescent values at a distance of 20 pixels from the edge were measured, and their mean fluorescence values plotted over time; the inflection point of a fitted sigmoid function was then determined as t1/2.
After addition of ionomycin/Ca2+ to embryos expressing mCherry::PHPLC1δ1/GFP::RHO-1 or mCherry-PHPLC1δ1/GFP::CDC-42, the disappearance of the mCherry-PHPLC1δ1 signal from the plasma membrane was imaged over time, whereas GFP::RHO-1 and GFP::CDC-42 were not imaged continuously to prevent photobleaching. Instead, only one end point image was taken by acquiring nine z-planes that were 0.5 µm apart.
Statistical analyses
The software packages JMP 13.2.0 (SAS Institute GmbH) and MATLAB 2016 were used to perform statistical analyses. Normal distribution of the data was tested using the Shapiro–Wilk test. Unless stated otherwise, statistical analysis was performed using nonparametric tests assuming non-normal distribution using the Wilcoxon Rank Sum test/Mann–Whitney test for two-group comparisons, and the Kruskal–Wallis test with a pairwise Wilcoxon Rank Sum test as a post-hoc test for three-group comparisons. In Fig. 4B, unpaired t-test was utilized to test the probability that two independent velocity fields result in the observed angle distribution. Values of P≤0.05 were considered statistically significant.
Acknowledgements
We thank Sachin Kotak and Kalyani Thyagarajan for initial observations of PIP2 cortical structures, Olivier Burri (BioImaging and Optics Platform, BIOP, School of Life Sciences, EPFL) for help in developing the script for preprocessing of embryos with ImageJ, as well as the BIOP at large for microscopy support. We thank Aitana Lebrand (Neves) for writing the Matlab script used for tracking spindle poles. For strains, we thank Jon Audyha, Daniel Dickinson, Bob Goldstein, Barth Grant, Stephan Grill, Anthony Hyman, Karen Oegema and Anne-Cécile Reymann, as well as the Caenorhabditis Genetics Center (CGC), which is funded by NIH Office of Research Infrastructure Programs (P40 OD010440). We are also grateful to Alexandra Bezler, Radek Jankele and Kerstin Klinkert for comments on the manuscript.
Footnotes
Competing interests
The authors declare no competing or financial interests.
Author contributions
Conceptualization: M.J.S., P.G.; Methodology: M.J.S., P.G.; Software: K.S.B., A.D.S.; Validation: M.J.S.; Formal analysis: M.J.S., K.S.B.; Investigation: M.J.S., M.B., C.C.N.S., R.W.; Resources: P.G.; Data curation: M.J.S.; Writing - original draft: M.J.S., P.G.; Writing - review & editing: M.J.S., K.S.B., P.G.; Visualization: M.J.S.; Supervision: P.G.; Project administration: M.J.S., P.G.; Funding acquisition: P.G.
Funding
This work was supported by the Swiss National Science Foundation (Schweizerischer Nationalfonds zur Förderung der Wissenschaftlichen Forschung) (31003A_155942).
Supplementary information
Supplementary information available online at http://dev.biologists.org/lookup/doi/10.1242/dev.164988.supplemental
- Received February 26, 2018.
- Accepted April 19, 2018.
- © 2018. Published by The Company of Biologists Ltd
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