Abstract
Asymmetric division is crucial for embryonic development and stem cell lineages. In the one-cell C. elegans embryo, a contractile cortical actomyosin network contributes to anterior-posterior (A-P) polarity and asymmetric division by segregating PAR proteins to discrete cortical domains. Here, we discovered that the plasma membrane lipid phosphatidylinositol 4,5-bisphosphate (PIP2) forms dynamic structures in C. elegans zygotes, distributing in a polarized and PAR-dependent manner along the A-P axis. PIP2 cortical structures overlap with F-actin and coincide with the actin regulators RHO-1, CDC-42 and ECT-2. Particle image velocimetry analysis revealed that PIP2 and F-actin cortical movements are coupled, with PIP2 structures moving slightly ahead. Importantly, we established that PIP2 cortical structures form in an actin-dependent manner and, conversely, that decreasing or increasing the level of PIP2 results in severe F-actin disorganization, revealing the interdependence between these components. Furthermore, we uncovered that PIP2 regulates the sizing of PAR cortical domains. Overall, our work establishes for the first time that a lipid membrane component, PIP2, is a critical modulator of actin organization and cell polarity in C. elegans embryos.
Summary statement PI(4,5)P2 is distributed in dynamic cortical structures and regulates asymmetric division by controlling actin organization and cell polarity in the one-cell C. elegans embryo.
Introduction
Asymmetric division generates cellular diversity and is particularly prevalent during development. During intrinsic asymmetric division, polarity is established and maintained in a mother cell; thereafter, polarity is translated into correct spindle positioning during mitosis along this polarity axis, resulting in the proper cleavage and partition of cellular contents to daughter cells. The extensively studied and evolutionarily conserved partitioning defective (PAR) proteins are critical for cell polarity and asymmetric division (reviewed in Goldstein and Macara, 2007; Gönczy, 2008; Knoblich, 2010). By contrast to the wealth of knowledge regarding PAR proteins and interacting components, the involvement of lipid plasma membrane components in cell polarity is less understood, in particular in developing systems.
The early C. elegans embryo has proven instrumental for dissecting the mechanisms governing asymmetric division (reviewed in Hoege and Hyman, 2013; Pacquelet, 2017; Rose and Gönczy, 2014). Shortly after fertilization, the entire embryo surface exhibits uniform contractions of the cortical actomyosin network located underneath the plasma membrane (Munro et al., 2004). These contractions are driven by non-muscle myosin 2 (NMY-2), which is activated by the Rho GTPase RHO-1 and its guanine nucleotide exchange factor (GEF) ECT-2 (Motegi and Sugimoto, 2006; Schonegg and Hyman, 2006). Sperm-derived centrioles are key for breaking symmetry of this system and for inducing local disappearance of cortical ECT-2 in their vicinity, thereby determining the embryo posterior. This leads to local inactivation of RHO-1 and initiation of cortical flows away from this region, towards the future embryo anterior (Bienkowska and Cowan, 2012; Cowan and Hyman, 2004; Motegi and Sugimoto, 2006). Polarized contractility promotes establishment of PAR polarity, whereby PAR-3, PAR-6, and atypical protein kinase C-like 3 (PCK-3) are segregated to the anterior side, whereas PAR-1, PAR-2, and Larval Giant Larvae-like 1(LGL-1) occupy the expanding posterior cortical domain (reviewed in Hoege and Hyman, 2013; Pacquelet, 2017; Rose and Gönczy, 2014). The RhoGTPase CDC-42 is also segregated to the anterior, where it stabilizes the actomyosin network and promotes PAR-6 association with the cortex (Kumfer et al., 2010; Motegi and Sugimoto, 2006; Schonegg and Hyman, 2006).
PAR polarity can be established in C. elegans zygotes also though a partially redundant pathway, whereby microtubules nucleated from centrosomes protect PAR-2 from PKC-3-mediated phosphorylation, thus allowing PAR-2 association with phospholipids at the embryo posterior (Motegi et al., 2011). In other systems, homologues 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, referred to hereafter as PIP2for simplicity) and phosphatidylinositol 3, 4,5-triphosphate (PI(4,5,6)P3, hereafter PIP3) (Krahn et al., 2010). Furthermore, human Cdc42 binds to PIP2 (Johnson et al., 2012). Overall, whereas it is clear that phospholipids can bind PARs and interacting proteins in some contexts, their subcellular distribution and potential function in an asymmetrically dividing system such as the C. elegans zygote remain unclear.
PIP2 is the most abundant of seven phosphorylated phosphatidylinositols and is present mostly in the inner leaflet of the plasma membrane, as revealed for instance by the distribution of the pleckstrin homology (PH) domain of mammalian phospholipase C1δ1 (PLC1 δ1), which binds PIP2 in vitro and in cells with high specificity (Garcia et al., 1995; Lemmon et al., 1995; Várnai and Balla, 1998). PIP2 is mainly phosphorylated from phosphatidylinositol 4-phosphate PI(4)P (PIP) by Type I PI(4)P5-kinases (PIP5K1), and can be further phosphorylated to PIP3 by Phosphatidylinositol 3-kinases (PI3K). Conversely, PIP2 can be dephosphorylated by 5-phosphatases, including OCRL and synaptojanin (reviewed in Brown, 2015; De Craene et al., 2017; McLaughlin et al., 2002). Amongst other roles, in systems from S. cerevisiae to H. sapiens, PIP2 helps link the F-actin cortical network to the plasma membrane, as well as stimulate F-actin assembly and reorganization. The latter function is achieved notably by activating, together with Cdc42, WASP family proteins that, in turn, activate the actin nucleator Arp2/3. Moreover, PIP3 further activates WASP family proteins through RhoGTPase GEFs (reviewed in 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 and, if so, how, PIP2 regulates cortical actomyosin network organization in the C. elegans embryo is not known.
Not only is the potential function of PIP2 in the C. elegans zygote not clear, but the same holds for PIP2 subcellular distribution. In other systems, PIP2 can distribute unevenly in the plasma membrane, for instance accumulating in macrodomains in nascent phagosomes, in 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 prior to F-actin patch accumulation, but their formation also depends on F-actin (Golub and Caroni, 2005; Golub and Pico, 2005). Moreover, it was suggested that F-actin enrichment in cell cortices drives clustering of PIP2-containing macrodomains, which in turn further regulate actin polymerization and branching (reviewed in Chichili and Rodgers, 2009). Overall, PIP2and F-actin polymerization function in a positive feedback mechanism in several systems.
The single PIP5K1 in C. elegans is PPK-1, which can synthesize PIP2 from PIP in vitro and in vivo (Weinkove et al., 2008). Overexpression of PPK-1 in developing worm 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 cytoskeletal reorganization and, therefore, gonad contractility (Xu et al., 2007). Moreover, PPK-1 is enriched on the posterior cortex of one-cell embryos and has been reported to be important for asymmetric spindle positioning, but not for cell polarity (Panbianco et al., 2008). In other systems, however, PIP2 can regulate cell polarity through its ability to recruit PAR proteins and reorganize 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 neurophils or Dictyostelium discoideum also rely on PIP2, together with PIP3, for actin network reorganization and polarization (reviewed in Wu et al., 2014). To summarize, in many systems, PIP2 is essential for F-actin reorganization and cell polarization, but it remains to be investigated whether this is the case in the developing C. elegans embryo.
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 found that the PIP2 biomarker GFP::PHPLC1δ1 (Audhya et al., 2005)forms distinct and dynamic structures at the cell cortex (Fig. 1A-J; Fig. S1A; Movie 1). Initially, when the cortical actomyosin network is contractile throughout the embryo, PIP2 is present weakly and evenly on the cell cortex (data not shown). Thereafter, when the actomyosin network begins to retract towards the anterior at the onset of polarity establishment, we observed the appearance of striking elongated cortical structures enriched in PIP2, primarily on the anterior side of the embryo (Fig. 1A, B, 1K; Fig. S1A, top). Such PIP2 cortical structures have an average size of ~2.5 µm2 and further elongate over time as the cell progresses through the cell cycle (Fig. 1L, M). Subsequently, all elongated PIP2 cortical structures move anteriorly so that they become distributed in a clearly polarized manner at pseudocleavage, which marks the end of the polarity establishment phase (Fig. 1C, D, arrow). At this time, PIP2 structures cover ç15% of the anterior cortical surface (Fig. 1K). During the centration/rotation stage that follows, PIP2 cortical structures first decrease in size (Fig. 1E, F, arrowhead), with most of them disappearing completely but some remaining as small foci by the time the cell enters mitosis (Fig. 1G, H, arrowheads). A few elongated cortical structures reappear during cytokinesis, primarily in the embryo anterior (Fig. 1I, J). In contrast to the discrete structures visible when imaging the cortical plane, PIP2 entities are barely detectable in the embryo middle plane (Fig. S1A, bottom), likely explaining why they were not reported previously (Audhya et al., 2005; Blanchoud et al., 2010; Panbianco et al.,2008).
We set out to verify the cortical distribution revealed by GFP::PHPLC1δ1 using fluorescently labeled synthetic PIP2. To this end, we delivered Bodipy-FL-PIP2 lipids to embryos whose eggshell had been permeabilized using perm-1(RNAi) (Carvalho et al., 2011). Although there was a high background of fluorescent lipids outside the embryo, we found that Bodipy-FL-PIP2 distributes in cortical structures marked by mCherry::PHPLC1δ1 (Fig. S1B, C). Overall, we conclude that PIP2 forms dynamic and polarized structures at the plasma membrane of one-cell C. elegans embryos.
A-P polarity cues regulate the polarized distribution of PIP2 cortical structures
We set out to address what regulates the polarized distribution of PIP2 cortical structures, which is particularly apparent during the pseudocleavage stage, as evidenced also by the fact that they do not overlap with GFP::PAR-2, which marks the posterior cortical domain (Fig. 2A). By contrast, we found that PIP2 cortical structures overlap with the anterior polarity domain harboring GFP::PAR-6 (Fig. 2B; Movie 2). Moreover, we observed that PIP2 cortical structures overlap only 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 previously reported to exist (Beers and Kemphues, 2006; Robin et al., 2014; Rodriguez et al., 2017; Wang et al., 2017).
We next tested whether the polarized distribution of PIP2 cortical structures depends on A-P polarity cues. In contrast to the polarized distribution observed in the control condition, we found that upon par-3(RNAi), PIP2 cortical structures distribute essentially uniformly over the cell cortex (compare Fig. 2C, D with 2E, F), except for the very posterior of the embryo, consistent with the known slight posterior clearing of the actomyosin network upon PAR-3 inactivation (Kirby et al., 1990; Munro et al., 2004). Furthermore, we found that upon par-2(RNAi), PIP2 cortical structures first move anteriorly (Fig. 2G, H), but then become distributed in a more uniform manner (Fig. 2H), 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). Together, these findings establish 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
Since PIP2 and F-actin are interdependent in many systems, we tested whether these two components overlap at the cell cortex of C. elegans embryos. As shown in Figure 3A and Movie 3, we found a partial overlap of Lifeact::mKate-2, which monitors F-actin, 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 4), the non-muscle myosin that powers contractions of the cortical actomyosin network (Guo and Kemphues, 1996; Munro et al., 2004). Strikingly, in addition, we found that PIP2 cortical structures marked by mCherry::PHPLC1δ1 fully colocalize with GFP::ECT-2, GFP::RHO-1 and GFP::CDC-42 (Fig. 2C-E; Movie 5). Overall, we conclude that PIP2 cortical structures colocalize partially with F-actin, as well as completely with the actomyosin network regulators ECT-2, RHO-1 and CDC-42 in the one-cell C. elegans embryo.
PIP2 cortical structures and the F-actin cytoskeleton move in concert
Live imaging of embryos expressing both Lifeact::mKate-2 and mNG::PHPLC1δ1 suggested that movements of PIP2 cortical structures and of the F-actin network are somehow coupled, as drastic changes in PIP2 cortical structures coincide with alterations in the actomyosin network across the first cell cycle (Movie 3). To investigate this potential coupling in a quantitative manner, we used particle image velocimetry (PIV) to analyze cortical flows of Lifeact::mKate-2 and mNG::PHPLC1δ1 in the same embryo during polarity establishment (Fig. 4A-C) (Thielicke and Stamhuis, 2014). This analysis revealed highly correlated local flow velocities at each time point (Fig. 4B; Fig. S2A; Pearson coefficient ρ = 0.61, p<0.0001). Moreover, the direction of flow vectors at each time point is very similar (Fig. 4C; Fig. S2B; θcut off=38° p<0.0001, Material and Methods). Analogous findings were made when comparing mCherry::PHPLC1δ1 and GFP::CDC-42 (Fig. S2C). By contrast, no strong correlation was found between mCherry::PHPLC1δ1 and the caveolin marker CAV-1::GFP (Fig. S2C), which has been suggested to mark lipid rafts in C. elegans (Entchev and Kurzchalia, 2005; Kurzchalia and Ward, 2003; Kurzchalia et al., 1999; Merris et al., 2003). Together, these results demonstrate that cortical movements of PIP2 cortical structures and the F-actin network are coupled.
We next addressed whether the movements of PIP2 cortical structures and of F-actin are synchronous or instead exhibit a time shift that might suggest a potential hierarchy between them, with one component leading the other. Close examination of movies of embryos expressing Lifeact::mKate-2 and mNG::PHPLC1δ1 suggested that PIP2 cortical structures move slightly ahead of cortical F-actin (Fig. 4E; Movie 6). To address this possibility quantitatively, we cross-correlated time-shifted images of Lifeact::mKate-2 and of mNG::PHPLC1δ1 to determine when there is maximal overlap between the two signals. As shown in Figure 4D, this analysis revealed that maximal correlation is achieved when mNeGr::PHPLC1δ1 is on average ~9.3 +/−1.5 seconds ahead of Lifeact::mKate-2. Overall, we conclude that PIP2 cortical structures move together with, but slightly ahead of, F-actin filaments.
What drives these movements of PIP2 cortical structures? The observation that PIP2 cortical structures move slightly ahead of F-actin filaments, with the trailing end of PIP2 cortical structure being seemingly in contact with the leading tip of coupled actin filaments (see Fig. 4E), led us to hypothesize that actin polymerization might push PIP2 cortical structures. Compatible with this possibility, we found that the average velocity of PIP2 cortical structures was ~0.17 +/−0.03 µm/s (Fig. S2D, E), in the range of actin polymerization driven motility in other systems (Brangbour et al., 2011; Cameron et al., 2000; Carlsson, 2003; Carlsson, 2010; Mogilner and Oster, 1996). These findings lead us to propose that the movements of PIP2 cortical structures are driven by actin polymerization.
PIP2 cortical structures depend on F-actin
Given notably the tight coupling between cortical PIP2 structures and F-actin, we investigated whether components of the actomyosin network regulate the presence of PIP2 entities. As shown in Figure 4F, we found that PIP2 cortical structures form and move unabated in nmy-2(RNAi) embryos, in which actomyosin network contractility is abolished, although they distribute symmetrically, as expected from the known requirement of NMY-2 in A-P polarity (Fig. 4F, G). Therefore, formation and movement of PIP2 cortical structures do not depend on a contractile actomyosin cortex. In stark contrast, we found that PIP2 cortical structures hardly form in act-1(RNAi) embryos (Fig. 4G, H, I; Fig. S3A-D). Moreover, acute impairment of F-actin through treatment of perm-1(RNAi) embryos with the actin polymerization inhibitor Cytochalasin D led to the disappearance of PIP2 cortical structures (Fig. S3E, F). By contrast, we found that PIP2 cortical structures are essentially independent of the microtubule cytoskeleton, impaired here using tba-2(RNAi) (Fig. 4J, K). Overall, we conclude that the formation of PIP2 cortical structures depends on F-actin.
Lowering the cellular level of PIP2 impacts F-actin distribution
We set out to address whether, conversely, PIP2 regulates F-actin organization. If this were the case, then changing the level of PIP2 Ishould 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. S4A) (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, in each embryo, a comparable time t1/2 when half of the initial GFP::PHPLC1δ1 plasma membrane fluorescence signal disappeared (Fig. S4B, C). A single plane in the middle of embryos expressing GFP::PHPLC1δ1 and Lifeact::mKate-2 was monitored in these experiments, as this proved most reliable for determining t1/2. In doing so, we found that PIP2 removal following addition of Ionomycin/ Ca2+ during pseudocleavage led to rapid alteration of embryo shape on the anterior side, coincident with altered F-actin organization (compare Fig. 5A and 5B, Fig. S4E). An analogous alteration in F-actin distribution was observed following Ionomycin/ Ca2+ addition during mitosis (Fig. 5C; Movie 7).
To test whether the shape change observed following lonomycin/ Ca2+ addition is caused by alterations in F-actin organization, we combined this treatment with the actin depoiymerizing agent Latruncuiin A. As shown in Figure S4F and Movie 8, we found that this results in normally shaped embryos. Therefore, shape changes following PIP2 removal are F-actin dependent. Together, these results uncover that a normal PIP2 level is critical for the proper distribution of F-actin and thus for proper shape of the C. elegans zygote.
Increasing the cellular level of PIP2 also impacts F-actin distribution
We sought to test the relationship between PIP2 and F-actin further 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 worms, but failed to find a single condition where this would be the case (see Table S1 for genes targeted in this study). Therefore, we set out to jointly inactivate OCRL-1 and UNC-26 for the following reasons (Fig. S4A). OCRL-1 is an inositol 5-phosphates that hydrolyzes PIP2to phosphatidyl 4-phosphate (PIP), and whose depletion leads to increased level of PIP2 on C. elegans phagosomes (Cheng et al., 2015). Moreover, UNC-26 is the C. elegans homologue of Synaptojanin, a polyphosphoinositide phosphatase that can also hydrolyze PIP2 to PIP, and whose impairment 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 an extant mutant for unc-26. Importantly, we found by comparing cortical mCherry-PHPLC1δ1 mean intensity values that ocrl-1(RNAi) unc-26(s1710) embryos exhibit an increased overall level of PIP2 (Fig. S5A, B). Importantly, in addition, we found that this leads to drastic alterations in PIP2 and F-actin cortical structures monitored by GFP::PHPLC1δ1 and Lifeact::mKate-2 (compared Fig. 5D and Fig. 5E-F; Fig. S5C-H). First, in addition to motile PIP2 structures, we found a population of immotile PIP2 clusters residing between the eggshell and the plasma membrane (Fig. 5E, H, K, arrow; Fig. S5I, arrowhead), potentially corresponding to PIP2 boluses removed from the cell in an attempt to return to homeostatic conditions. Second, we found that motile PIP2 structures do not disappear as readily after pseudocleavage as they do normally (Fig. 5H, I, compare to Fig. 5G; Fig. S5D). Third, we found that motile PIP2 structures exhibit altered distribution in all ocrl-1(RNAi) unc-26(s1710) embryos (Fig. 5E, F; Fig. S5F, H). In some (hereafter referred as class I embryos, N=27/48), the anterior movements of PIP2 cortical structures and of the actomyosin network do not stop at pseudocleavage, but instead continue until the end of mitosis, resulting in a very small anterior domain of PIP2 and F-actin cortical structures (compare to Fig. 5G, J to Fig. 5H, K; Movie 9). In class II ocrl-1(RNAi) unc-26(s1710) embryos (N=21/48), weak anteriorly directed movement of cortical PIP2 and F-actin is initiated, but both components become distributed throughout the cortexby 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 was the case in only 8/21 class 2 embryos; this subset exhibited more pronounced anteriorly directed movements than the other 13. Overall, we conclude that the extent of PIP2 5-phosphatases depletion is stronger in class 2 than in class 1 embryos, with the severe phenotype in the former perhaps reflecting an impact on multiple cellular processes. Regardless, these findings establish that an increase in the level of PIP2 as achieved in class 1 embryos leads to sustained cortical flows towards the anterior side. Moreover, these results further demonstrate that PIP2 regulates actin cytoskeletal organization in one-cell C. elegans embryos.
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 the establishment phase of A-P polarization of the C. elegans zygote (Guo and Kemphues, 1996; Hill and Strome, 1988; Hill and Strome, 1990; Munro et al., 2004; Severson and Bowerman, 2003; Shelton et al., 1999). 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. To this end, 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, respectively (Fig. 6A-L). We found that the distribution of GFP::PAR-2 and GFP::PAR-6 domains changes in a manner consistent with alterations in motile PIP2 structures and the F-actin network, and this from early on (Movie 10). Thus, for GFP::PAR-6, either a small domain formed on the very anterior (Fig. 6B, E; class 1, N=5/9; Movie 10) or else 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 in a reciprocal manner, either expanding drastically towards the anterior (Fig. 6H, K; class 1, N=12/22; Movie 11) or else remaining restricted to the very posterior (Fig. 6I, L; class 2, N=10/22). Together, these results indicate that a correct level of PIP2 is needed for proper F-actin network localization and, presumably as a consequence, appropriate PAR polarity.
In the above experiments, the level of PIP2 was in excess from the beginning of development, such that one could not distinguish whether the impact on polarity reflected a role strictly during the establishment phase or during both establishment and maintenance phases. We reasoned that one could test specifically a potential role for polarity maintenance by adding Ionomycin and Ca2+to perm-1(RNAi) embryos expressing mCherry::PHPLC1δ1 and GFP::PAR-2 after the establishment phase. We found that the GFP::PAR-2 domain is unaltered at t1/2 (compare Fig. 6M and 6N, O), even though F-actin is already changed at that moment (see Fig. 5B, C). Importantly, in addition, we found that the GFP::PAR-2 domain expands slowly towards the anterior starting ~3 min thereafter (Fig. 6N, bottom two panels), with the pseudocleavage furrow moving anteriorly initially, and then either retracting (Fig. 6M, N; Movie 12, N=6/14), or else remaining at the very anterior until the end of the first cell cycle Movie 13; N=8/14). Mirroring the findings at pseudocleavage, we found that if t1/2 occurs at nuclear envelope breakdown (NEBD), the GFP::PAR-2 domain likewise expands slowly towards the anterior (Fig. 6O; Movie 14). Together these results indicate that an appropriate level of PIP2 is essential for proper PAR polarity also during the maintenance phase.
In principle, PIP2 could alter PAR polarity during the maintenance phase through an impact on F-actin organization or else via an actin-independent role. A potential role of F-actin in polarity maintenance is somewhat controversial, in contrast to its well known role during polarity establishment (Goehring et al., 2011; Hill and Strome, 1990; Liu and Fletcher, 2006; Severson and Bowerman, 2003). In the light of our findings with PIP2 level alterations, we set out to directly test whether F-actin plays a role in polarity maintenance, first by adding 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 does not alter the GFP::PAR-2 domain in a major manner (Fig. 7A, B; Movie 15). However, we found also that Cytochalasin D treatment does not fully disrupt F-actin, as clumps of Lifeact::mKate-2 remain present on the embryo anterior (Fig. 7B). We hence turned to inhibiting F-actin polymerization using Latrunculin A, which resulted in a total depletion of F-actin (Fig. 7C; N=12; Movie 16). We observed membrane invaginations that remove GFP::PAR-2 from the cortex into cytoplasmic aggregates (Fig. 7C, arrowhead), as reported by others (Goehring et al., 2011; Redemann et al., 2010). Importantly, in addition, we monitored changes in GFP::PAR-2 distribution not as a function of drug addition time, as in earlier work (Goehring et al., 2011), but of the time t1/2 at which half of the Lifeact::mKate-2 fluorescence disappeared from the membrane. In doing so, we found that the size of the GFP::PAR-2 domain decreased significantly after t1/2 in all embryos analyzed (Fig. 7C, bottom; N=12). As reported in Figure S5J, we found that the t1/2 of Lifeact::mKate-2 and of GFP::PAR-2 disappearance are highly correlated (Pearson coefficient p=0.86; p=0.0014). This finding reinforces the conclusion that F-actin is critical not only for the establishment, but also for the maintenance of PAR polarity.
Overall, we uncovered that a proper level of PIP2 is essential for correct sizing of PAR domains presumably through reorganization of F-actin, not only during polarity establishment but also during polarity maintenance phase.
Discussion
In this work, we demonstrate that PIP2 forms cortical structures in the one-cell C. elegans embryo. We show that 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, likely through its impact on the actin cytoskeleton, PIP2 is also needed for the correct sizing of anterior and posterior PAR domains, demonstrating for the first time that a plasma membrane lipid component participates in setting 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 prior to this work, primarily because the middle embryo plane has been analyzed in most past investigations. Here, we developed cortical imaging conditions to assay subcellular distributions at the cortex, the very location where the function of PAR proteins and components critical for asymmetric division is exerted. In doing so, we discovered that PIP2 is present in dynamic polarized cortical structures marked by the PIP2 biomarker GFP::PHPLC1δ1, in line with recent observations mentioning a non-uniform distribution of this fusion protein (Rodriguez et al., 2017; Wang et al., 2017). Although patches of plasma membrane enriched in PIP2 have been observed in other systems (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 C. elegans zygote enabled us to uncover their distribution and dynamics with unprecedented resolution. Why were such remarkable structures not observed in previous studies in the worm? In addition to the fact that they are not noticeable when imaging the middle plane of the embryo, other plausible reasons include 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? We hypothesize that PIP2 cortical structures form by redistribution of existing PIP2 rather than by de novo synthesis through the PIP5K1 PPK-1, and this for two reasons. 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), away from the location where most PIP2 cortical structures are. Interestingly, we find also that PIP2 cortical structures form and move independently of actomyosin contractions, as evidenced by their unchanged presence and behavior upon NMY-2 depletion (data not shown). Nevertheless, it remains possible that PIP2 cortical structures form at membrane protrusions or ruffles, which would be consistent with the finding that PIP2 can stimulate F-actin polymerization in curved but not in flat membranes (Gallop et al., 2013), and with PIP2 accumulating in membrane ruffles, nascent phagosomes and the leading edge of motile cells (reviewed in McLaughlin et al., 2002; Zhang et al., 2012). Overall, we propose that, in the C. elegans zygote, PIP2 cortical structures form through the redistribution of existing PIP2 at the plasma membrane, 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 uncover here that this is the case also in C. elegans embryos. We found that PIP2 cortical structures and F-actin movements are coupled, with PIP2 structures moving slightly ahead of F-actin filaments, at velocities compatible with actin polymerization driving their movements. This leads us to propose that actin polymerization pushes PIP2 cortical structures, in a manner reminiscent of other actin-dependent motility processes such as that of Listeria monocytogenes (reviewed in Mogilner and Oster, 1996). While being pushed ahead of F-actin filaments in C. elegans, PIP2 structures might recruit factors promoting actin polymerization and branching, such as ECT-1, RHO-1 and CDC-42, thus guiding proper F-actin network reorganization, in line with suggestions in other systems (reviewed in Chichili and Rodgers, 2009). Intriguingly, the distribution of a biosensor that detects active RhoA overlaps with that of NMY-2 foci (Reymann et al., 2016; Tse et al., 2012). Given that we show here that PIP2 cortical structures do not overlap with NMY-2, while they do overlap with GFP::RHO-1, perhaps the bulk of RHO-1 associating with PIP2 cortical structures is not active. Alternatively, given that we show here that RHO-1 colocalizes with its activating GEF ECT-2, perhaps the RhoA biosensor used previously does not detected all active RHO-1 species. Furthermore, it is interesting to note that non-muscle myosin 2 plays a role in 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 its role in network contractility, may together ensure proper F-actin dynamics in the early C. elegans embryo.
PIP2 and PAR-dependent polarity
PAR proteins are also distributed unevenly within their cortical domain. For instance, PAR-6 exists in two cortical 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 that assemble in a manner dependent on PCK-3, CDC-42, as well as actomyosin contractility (Rodriguez et al., 2017; Wang et al., 2017). Intriguingly, we find that PIP2 cortical structures colocalize within the more diffuse cortical PAR-6 protein population, the one lacking PAR-3 (Beers and Kemphues, 2006; Robin et al., 2014; Rodriguez et al., 2017; Wang et al., 2017). We establish here that increasing the level of PIP2 augments the segregation of both PAR-6 populations to the embryo anterior, potentially because cortical clustering of PAR-3 depends on the actomyosin cytoskeleton and cortical tension (Beers and Kemphues, 2006; Robin et al., 2014; Rodriguez et al., 2017; Wang et al., 2017). Overall, we propose that increasing the level of PIP2 might augment cortical tension, which could in turn promote PAR-3 clustering and thereby aid segregation.
We show that a correct PIP2 level is essential for proper polarity establishment and maintenance trough correct positioning of GFP::PAR-2 and GFP::PAR-6 domains. When increasing the level of PIP2, the continued movement of PAR domains towards the anterior until the end of mitosis alters their relative size. This is reminiscent of changes in the size of PAR domains that occur upon RGA-3/4 depletion (Schonegg et al., 2007). However, although rga-3/4(RNAi) embryos exhibit a hypercontractile actomyosin network, this is not the case of embryos with increased PIP2 level. We thus propose that actomyosin contractility regulated by NMY-2 and F-actin organization regulated by PIP2 contribute in concert to 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 less clear (Goehring et al., 2011; Hill and Strome, 1990; Liu and Fletcher, 2006; Severson and Bowerman, 2003). Our results, together with that of others, indicate that the actomyosin network regulates the size and localization of PAR domains in two ways. First, when the actomyosin network moves along the A-P embryonic axis, PAR domains alter their distribution accordingly. This relationship was clear prior to this work for the polarity establishment phase, and we show here that this is also the case during polarity maintenance. 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 reveal here that the lack of this function can lead to the near total disappearance of cortical PAR-2, emphasizing the critical importance of actin also during the maintenance phase.
Overall, our results in the C. elegans zygote 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, does not influence polarity at the end of the first cell cycle (Panbianco et al., 2008). Perhaps PPK-1 distribution plays only a minor role in regulating the cellular level of PIP2 in the zygote, being dispensable for PIP2 cortical structure formation. In this case, depleting an enzyme such as CSNK-1 that negatively regulates PPK-1 localization would not be expected to influence PIP2 cortical distribution. Here, by contrast, we establish unequivocally that alterations in the level of PIP2 impairs polarity establishment and maintenance during the first asymmetric division of C. elegans embryos.
Material and Methods
Worm Strains
Nematodes were maintained at 24°C using standard protocols (Brenner, 1974). The following worm 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[p/e-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; tjls1[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; tjls4[pie-1::GFP::ect-2+unc-119(+)]) (Motegi and Sugimoto, 2006); unc-26(s1710) (EG3027, unc-26(s1710) IV) (Charest et al., 1990); age-1(m333), (DR722, age-1(m333)/mnC1 dpy-10(e128) unc-52(e444) II) (Riddle, 1988). Crosses of worm strains were performed at 20°C to generate lines homozygote for all transgenes, which were then maintained at 24°C. For GFP::RHO-1 and mCherry::PH PLC1δ1, as well as GFP::PHPLC1δ1 and Lifeact::mKate-2, worm lines were crossed and F1 progeny heterozygote for both transgenes imaged.
RNAi
RNAi-mediated deletion was performed essentially as described (Kamath et al., 2001), using bacterial feeding strains either from the Ahringer (Kamath et al., 2003) or the Vidal library (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), and ocrl-1 (Ahringer) was performed by feeding L3-L4 animals with bacteria expressing the corresponding dsRNA at 24°C for 20-26 hours. RNAi for perm-1 (Ahringer) was performed by feeding L4 and young adults with bacteria expressing perm-1 dsRNA at 20°C for 12-18 hours. The effectiveness of the deletion was screened phenotypically as follows: par-2(RNAi) and par-3(RNAi) -symmetric spindle positioning and 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 specification; ocrl-1(RNAi) -see results, perm-1(RNAi): successful action of added drug.
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.1 A,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 using an inverted Olympus IX 81 microscope equipped with a Yokogawa spinning disk CSU - W1 with a 63 (NA 1.42 U PLAN S APO) objective and a 16-bit PCO Edge sCMOS camera at 23°C. 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, 3 planes at the cell cortex (each 0.25 µm apart) were acquired. Cell cycle stages were determined using transmission light microscopy by 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 3 cortical planes were z-projected using average intensity projection, then 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 due to photobleaching was corrected with the Fiji plugin "bleach correction" using the exponential fitting method. 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 sigma from the Gaussian peak. The size of cortical structures was normalized to the total cortical area.
Curves of normalized cortical structures sizes were fit 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 Nuclear Envelope Breakdown (NEDB) as a reference, because a sigmoid function could not be fit with the PH markers in these cases. Since the time separating centration/rotation from NEDB is typically150 seconds, t=0 was set at −150 seconds prior to NEDB for act-1(RNAi) and unc-26(s1710) ocrl-1(RNAi) embryos.
The Elongation Index was calculated as follows: ((perimeter˄2)/area)/4π using the MATLAB image processing function "regionprops". We then normalized the Elongation Index by a factor of 1/π such that a square of 2 × 2 pixels has an Elongation Index of 1.
Cortical images obtained by live confocal spinning disk imaging and shown in the figures were processed as follows: the 3 cortical planes were z-projected using maximum intensity projection, then a median filter of 1 was applied. The grey value fluorescence intensity of some transgenes (GFP::PHPLC1δ1 as heterozygote, GFP::PAR-2, GFP::PAR-6, Lifeact::mKate-2, mNeonGreen-PHPLC1δ1) was slightly variable likely resulting from variable expression/folding of the fluorescent fusion protein, which may stem for several reasons, including F1 heterozygosity, temperature shifting, silencing during crossing. The brightness and contrast of images resulting from embryos expressing these transgenes was therefore adjusted accordingly. The fluorescence intensity of mCherry::PHPLC1δ1 was also variable in some cases; therefore, the brightness and contrast of images from embryos expressing mCherry::PHPLC1δ1 were therefore adjusted accordingly as well. Such variability was especially pronounced in UNC-26 and OCRL-1 depleted embryos. To compare the intensity of mCherry-PHPLC1δ1 in control embryos and unc-26(s1710) ocrl-1(RNAi) embryos, 3 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, a Otsu threshold was used to retrieve the brightest elements - including the embryo - of the image, retaining only the biggest blob, corresponding to the embryo. Values outside the embryo were averaged to obtain the mean background intensity value, which was subtracted from the embryo pixels. Thereafter, embryo pixel values were averaged to obtain the mean pixel intensity value.
Cortical flow measurement, correlation analysis, and PIP2 structures 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 2 planes (0.25 µm apart) and applying a median filer of 1 pixel. PIP2 cortical structures and the F-actin network were then segmented using the following procedure: 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, times a scaling factor determined empirically (1.7). The extraction of the F-actin network was achieved simply by determining a histogram-based automated threshold on the morphological top-hat of the F-actin image. F-actin filaments and PIP2 cortical structures where segmented prior to PIV analysis to ensure that only flow fields in the region of interest are measured.
PIV was then performed to measure cortical flows using the MATLAB based PIVlab toolbox (Thielicke and Stamhuis, 2014); this 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 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 interrogation area. PIV was applied to mNeonGreen::PHPLC1δ1 and Lifeact::mKate-2 separately, after segmentation of the corresponding cortical structures. The choice of the sizes of the cells and interrogation areas was a balance between two criteria: smaller cells allow to compute displacements with high spatial resolution, but excessively small cells do not contain enough information to be reliably correlated to other cells; the estimation of displacements of bigger cells is hence more reliable, but are computed with lesser resolution. We found empirically that 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, so as 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 movies, and also time-wise. The cut-off angle is defined as the θ 0 parameter of the curve of equation y = a exp(-θ/θ 0) fitted to the histogram.
Cross-correlation analysis was performed as follows. Movies used to calculate the cross-correlation were acquired alternating acquisition of red or green channel first 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 the following formula:
Colocalization = (PIP2(t) ∩ F-actin(t-Δt)/ PIP2 (t)
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 is maximal represents the time shift between PIP2and F-actin. The mean time shift and its error were computed as follows: we fitted a parabola of equation y = a + (t-t0)˄2 + b 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 = 9.3s and the standard deviation sigma_t0 = 1.5s.
To track PIP2 structures, embryos expressing mNG-PHPLC1δ1 were imaged with an exposure time of 50 ms, laser power of 60% and 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 reslicing it using the Fiji plugin “Reslice”. Velocity was calculated from the corresponding number of time points and track length.
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 were utilized: 30 µM Ionomycin (Calbiochem, 407950), 3-5 mM CaCl2 (Sigma-Aldrich, C5080), 20 µM Cytochalasin D (AppliChem, 22144-77-0), 12.5 µM Latrunculin A (Sigma-Aldrich, 76343-93-6). For control movies, DMSO at a concentration equivalent to the final DMSO concentration in the drug solutions was added to the buffer prior to dissection.
Successful drug action was determined for each embryo by the disappearance of the PHPLC1δ1 fluorescence signal from the plasma membrane (lonomycin/ Ca2+, Latrunculin A) and of Lifeact::mKate-2 from the cell cortex (Latrunculin A, Cytochalasin D). The time between drug addition and drug action was variable, probably due to variations in eggshell permeability upon perm-1(RNAi). As a comparable reference time between embryos, we therefore determined the time t1/2 (t inflection) when half of fluorescence at the plasma membrane has disappeared. t1/2 was determined 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.
Lipid delivery
BODIPY® FL Phosphatidylinositol 4,5-bisphosphate (Echelon Bioscience, C-45F6) (end concentration: 2 µM) was delivered to perm-1(RNAi) embryos by adding it to the buffer in which gravid worms were dissected. It typically helped to premix BODIPY® FL Phosphatidylinositol 4,5-bisphosphate (100 µM) with a carrier histone (P-9C2; Echelon Bioscience) (300 µM) (Ozaki et al., 2000). Phosphoinositide-histone complexes were formed as described (Kotak et al., 2014): both components were mixed by vigorous vortexing and then incubated for at 15 min at room temperature.
Statistical analyses
The software package JMP 13.2.0 (SAS Institute GmbH) and MATLAB 2016 were used to perform statistical analysis. Two-group comparisons were performed using Student’s t-test. Results with values of p≤0.05 were considered statistically significant.
Acknowledgments
We thank Kaiyani Thyagarajan and Sachin Kotak for initial observations of PIP2 cortical structures, Olivier Burri (Biolmaging 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. For strains, we thank John 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). This work was supported by the Swiss National Science Foundation (31003A_155942). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Footnotes
Author Contributions: M.S. and P.G. designed the project; M.S. conducted experiments with support from M.B., C.N., and R.W; M.S. and P.G. analyzed the data; M.S. and K.B. performed PIV analysis, M.S., K.B. and A.D. developed image processing and analysis scripts; M.S. and P.G. wrote the manuscript.