Multiple in vivo phosphorylated residues are critical for LIN-5 function

Previously, we described that at least 25 residues of LIN-5 are phosphorylated in vivo (Fig 1A) [15]. To acquire insight in which phosphorylations are functionally relevant, we replaced each phosphorylated serine or threonine with an alanine residue that cannot be phosphorylated. The relevant codon alterations were introduced in a cloned genomic lin-5 DNA fragment and subsequently tested for functionally complementing the lin-5(e1348) null mutation in vivo [19]. In the presence of maternal product, lin-5(e1348) mutants fail to undergo chromosome segregation during postembryonic divisions and continue abortive mitoses [19–21]. Transgenes containing wild type lin-5 or gfp::lin-5 coding sequences restored post-embryonic cell divisions in lin-5(e1348) null mutants (Fig 1B). However, these lin-5 transgenes appeared susceptible to germline and somatic silencing, as reliable rescue and GFP-LIN-5 expression was observed only in the F1 generation. Hence, we examined transgenic F1 animals, focusing on vulval development and nuclear divisions in the intestine as a quantitative measure for LIN-5 function (Fig 1B).

Alanine substitutions of threonine 168, serine 659, and serine 662 were the only single amino acid changes that significantly compromised LIN-5 function in vivo. The T168A mutation had the strongest effect and almost completely eliminated the ability to restore intestinal divisions in lin-5(e1348) null mutants (Fig 1B). Interestingly, this strong effect was specific for the intestine: LIN-5^T168A expression allowed lin-5 mutants to develop a normal vulva (S1 Fig). T168 forms part of an ideal consensus phosphorylation site (S/T*-P-x-K/R) for the mitotic cyclin-dependent kinase 1 (CDK-1) [22]. CDK-1 is likely to regulate LIN-5, as multiple CDK-1 consensus sites are present in the LIN-5 N- and C-terminus, and CDK1 phosphoregulation of the NuMA C-terminus has been reported [18,23]. We generated double alanine substitutions of T168 in combination with T181 or S199, two nearby candidate residues for CDK-1 phosphorylation. Strikingly, the transgene encoding LIN-5[T168A,T181A], but not LIN-5[T168A,S199A], completely failed to rescue intestinal mitoses and vulva formation in lin-5(e1348) mutants (Fig 1B and S1 Fig). Because phosphorylation of T181 by itself was not essential for post-embryonic divisions, T168 and T181 phosphorylations likely cooperate to control LIN-5 function.

The individual and combined S659A and S662A substitutions (LIN-5[S659A,S662A]) also reduced lin-5(e1348) complementation. By contrast, simultaneous alanine substitutions of serines 729, 734, 737, and 739 did not prevent LIN-5 function (Fig 1B and S1 Fig). In agreement with the latter result, PKC-3 (aPKC) phosphorylation of these residues inhibits LIN-5 function and is not required for cell division [15]. Our alanine-substitution experiments indicate that in addition to spatiotemporal regulation of LIN-5 by PKC-3, phosphorylation of LIN-5 residues in the dynein-interacting N-terminus and GPR-1/2 binding C-terminus may contribute to LIN-5 regulation in vivo.

In vitro kinase assays reveal candidate LIN-5 kinases

To determine whether CDK1 is indeed able to phosphorylate T168 and T181 of LIN-5, we performed in vitro kinase assays with recombinant GST-LIN-5 expressed in E. coli as a substrate. Indeed, immunopurified human CDK1/cyclin B phosphorylated GST-LIN-5, but not GST alone (S2A and S2B Fig). Analysis of in vitro phosphorylated GST-LIN-5 by mass spectrometry revealed extensive phosphorylation of T168, T181, and S744 of LIN-5 (S2B Fig). Additionally, peptides containing phosphorylated T704 and S756 were also found, and some other phosphopeptides less frequently. Taken together, CDK1/cyclin B phosphorylates LIN-5 in vitro at multiple sites including T168 and T181, and phosphorylation of T168 and T181 in vivo appears to be required for LIN-5 function.

In contrast to T168 and T181, residues S659 and S662 are not part of apparent consensus phosphorylation sites. In our previous in vivo mass-spectrometry data, the S659,S662 double phosphorylated peptides were abundant, while the corresponding unphosphorylated peptides were not detected [15]. This may indicate that S659 and S662 are constitutively phosphorylated in early embryos. To gain insight in which kinases may be involved, we examined LIN-5 phosphorylation in vitro with a series of polarity and cell cycle kinases, followed by mass spectrometry analyses. This revealed several residues that were phosphorylated by multiple kinases in vitro (Fig 2A). In striking contrast, S659 was only phosphorylated by GSK3, and none of the tested kinases phosphorylated S662 (Fig 2A). We considered several potential explanations for this lack of phosphorylation: the responsible kinase(s) may not have been included in the assays, residue S662 may not be accessible in the recombinant protein, or S662 phosphorylation may require a priming event. To test the latter possibility, we performed in vitro kinase assays with synthetic peptides that contain the S659 and S662 residues, either unphosphorylated or phosphorylated at one of the positions. Testing several kinases, we found that casein kinase 1 (CK1) efficiently phosphorylates S662, but only when the peptide contained a phosphorylated S659 residue (Fig 2B). As for the full length protein, only GSK3 phosphorylated S659 in the unphosphorylated peptide. Based on the combined in vitro data, we propose that GSK3 phosphorylation of residue S659 is a priming reaction for CK1 phosphorylation of S662. Highly similar phosphorylation has been reported for the Wnt/Frizzled co-receptor LRP6, with GSK3 priming for CK1 phosphorylation at similar sites [24,25].

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Fig 2. LIN-5 phosphorylation by cell cycle and polarity kinases in vitro.

(A) Graphic overview of mass spectrometry analysis of in vitro kinase assays, revealing multiple target residues in LIN-5 phosphorylated by PAR-1, PKC-3, CK1, CK2, GSK3, CDK1 and Aurora B kinase. Yellow residues indicate in vivo phosphorylated residues, brackets indicate essential residues identified in complementation assay. (B) Radioactive counts (CPM) of in vitro kinase assays with CK1 and GSK3 on a synthetic LIN-5 peptide 654–670 with or without synthetic incorporation of phosphorylated amino acids S659 or S662.

https://doi.org/10.1371/journal.pgen.1006291.g002

In addition to CDK1, GSK3 and CK1 phosphorylation, our analyses revealed phosphorylation of LIN-5 by the polarity kinase PAR-1. While several phosphopeptides were detected, some were rare and the quantitative software program MaxQuant only recognized the S397 and S739 LIN-5 residues as in vitro phosphorylated by PAR-1 (Fig 2A). S397 is located in the LIN-5 coiled coil region and its phosphorylation was previously observed in embryos (Fig 2A) [15]. However, our previous in vivo analysis failed to identify LIN-5 phosphorylations that were diminished after par-1 RNAi [15]. Re-evaluation of the quantitative mass spectrometry data revealed that, although masked by an abundant unrelated peptide, the ratio between the phosphorylated and unphosphorylated S397 peptide was severely reduced in par-1(RNAi) embryos compared to control RNAi embryos (S3 Fig). In contrast, S739 phosphorylation was not significantly affected by par-1 knockdown in vivo [15]. Taken together, we identified multiple phosphorylated LIN-5 residues as well as candidate kinases that could be important in the regulation of LIN-5 function. In addition to four adjoining residues phosphorylated by PKC-3 in the C-terminus, T168 and T181 may be phosphorylated by CDK-1, S397 by PAR-1, and S659 by GSK-3, to prime phosphorylation of S662 by CK1.

A 30 amino acid LIN-5 domain that includes S659,S662 mediates GPR-1/2 interaction

Both S659,S662 and the four residues phosphorylated by PKC-3 are located in the LIN-5 C-terminus which mediates GPR-1/2 binding [15,26]. As phosphorylation could affect GPR-1/2 association, we wanted to define which LIN-5 residues are critical for GPR-1/2 binding. Testing deletion constructs in yeast two-hybrid assays confirmed that the LIN-5 C-terminal region is sufficient for GPR-1 association. GPR-1 interaction was observed for all truncated LIN-5 proteins except for those with deletions in the 609–671 amino acid region (Fig 3A). At the same time, including only the 609–671 LIN-5 fragment did not allow growth in this assay, possibly due to an inability of this short fragment to fold properly in yeast (Fig 3A). The essential 609–671 region does not contain serine 729, 734, 737, and 739 phosphorylated by PKC-3 in vivo, in agreement with our previous conclusion that PKC-3 phosphorylation of LIN-5 does not prevent interaction with GPR-1/2 [15].

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Fig 3. Yeast two-hybrid assays identify critical residues in LIN-5 for GPR-1 interaction.

(A) Yeast two-hybrid analysis of DB::GPR-1 (bait) interaction with AD::LIN-5 (prey) fragments of varying sizes on Sc (synthetic complete) −Leu−Trp (+His, control) and Sc −Leu−Trp−His + 3-AT (-His, selection) plates. (B) Yeast two-hybrid analysis of DB::GPR-1 (bait) interaction with AD::LIN-5 (prey) fragments containing single amino acid changes on Sc −Leu−Trp and Sc −Leu−Trp−His + 3-AT plates. (C) Graphical representation of LIN-5 residues required for GPR-1 binding. Size of bars indicates frequency of found mutations, with every bar representing a different amino acid change. (D) Yeast two-hybrid analysis of DB::GPR-1 (bait) interaction with AD::LIN-5 (prey) full length containing single amino acid changes on Sc −Leu−Trp−His + 3-AT plates at 20°C and 30°C.

https://doi.org/10.1371/journal.pgen.1006291.g003

To identify specific LIN-5 amino acids required for GPR-1/2 interaction, we performed “reverse yeast two-hybrid screening”. This method selects mutations that disrupt bait-prey protein interactions, making use of URA3-mediated conversion of 5-fluoroorotic acid (5-FOA) to a toxic product [27]. The normal interaction between LIN-5 and GPR-1 leads to GAL4-controlled URA3 expression in yeast two-hybrid assays, and causes cell death in the presence of 5-FOA. Thus, following mutagenesis of one of the binding partners, interaction-deficient alleles can be recovered from 5-FOA-resistant colonies [28]. We used PCR-based random mutagenesis of LIN-5 prey fragments (amino acids 609–821), and isolated 163 5-FOA resistant yeast colonies in a reverse yeast two-hybrid screen (for details see Materials and Methods, S4A Fig). 89 colonies contained a single missense mutation in the LIN-5 coding sequences, together changing 15 different amino acids. Substitutions of 12 of these 15 individual amino acids caused loss of GPR-1 interaction again in the re-test (Fig 3B and 3C). The 12 affected residues were all located between amino acids 638–667 of LIN-5. Importantly, the interaction-defective alleles included missense mutations of the phosphorylated residues S659 and S662. In fact, S662 was found altered to glycine, cysteine and asparagine (S4B Fig). These data indicate that a 30 amino acid stretch in the LIN-5 C-terminal region, which includes the in vivo phosphorylated S659 and S662 residues, mediates the interaction with GPR-1.

Following up on the interaction defective alleles, we noticed that the effect of missense mutations was substantially reduced when tested in the context of full length LIN-5, compared to the C-terminus only. Western blot analysis did not reveal substantial differences in protein levels compared to wild type (S5A Fig). The LIN-5 coiled-coil region promotes dimerization and is thereby expected to increase GPR-1 binding avidity. Only one of the four most frequently identified mutations, L663S, also interfered with full length LIN-5 binding to GPR-1 (Fig 3D, left panel). However, at a reduced temperature (20°C), this leucine 663 to serine (LIN-5[L663S]) mutation still allowed growth on selective media, indicating that GPR-1 interaction is not completely abolished. We also tested S659 and S662 phosphorylation-site mutants in the context of full length LIN-5. While the single mutations had little effect on GPR-1 binding, replacement of both serine 659 and 662 by alanine reduced GPR-1 interaction in yeast, as detected by lack of growth on -His plates at 30°C (Fig 3D, right panel). Phosphomimetic substitutions (S to D or E) of S659, S662, or both, did not reduce interaction (Fig 3D, right panel). These results are consistent with phosphorylation of S659 and S662 contributing to GPR-1/2 binding, and taking place in yeast as well as C. elegans. Taken together, our forward and reverse yeast two-hybrid assays identified LIN-5 residues that appear to mediate interaction with GPR-1/2, which are located within a 30 amino acid C-terminal domain. This includes S659 and S662, of which the phosphorylation in vivo likely contributes to GPR-1/2 binding.

Examining LIN-5 phosphoregulation in vivo by CRISPR/Cas9-mediated genome engineering

We used CRISPR/Cas9-mediated gene targeting to engineer lin-5 alleles and examine the effects of amino acid substitutions in vivo [29–32]. First, we created the lin-5[L663S] mutation by introducing a single nucleotide alteration in the endogenous lin-5 locus. This resulted in a typical lin-5 loss-of-function phenotype, with homozygous sterile, thin and uncoordinated larvae that fail to undergo chromosome segregation but continue abortive mitoses [19,20]. We determined the number of nuclei in the intestine and ventral cord, following fixation and staining of DNA. In lin-5[L663S] mutants, both tissues contained severely reduced numbers of nuclei compared to the wild type, consistent with a failure to undergo chromosome segregation in most post-embryonic divisions (Fig 4A and 4B, S6A Fig). Thus, a single change of amino acid L663 in the GPR-1-binding motif of LIN-5 results in strong loss-of-lin-5 function. This result confirms the power of reverse yeast two-hybrid screening in identifying amino acids that affect protein-protein interactions in vivo [27].

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Fig 4. In vivo phenotypical analysis shows developmental defects in LIN-5 phosphomutants.

(A) Quantification of intestinal nuclei following propidium iodide staining in homozygous LIN-5 phosphorylation mutants compared to wild type and lin-5(e1348) null animals. Null mutants are expected to contain 20 nuclei, wild type animals 32–34 nuclei. Individual values are plotted, n>11. (B) Quantification of P-cells and juvenile motor neurons in the ventral cord region P2-P10 in propidium iodide stained homozygous LIN-5 phosphorylation mutants compared to wild type and lin-5(e1348) null animals. Nuclei were counted in late larval stages of asynchronous populations of worms, with null mutants expected to have 24 nuclei (9 P cells and 15 juvenile motor neurons) and wild type animals 65 nuclei. Individual values are plotted, n>11. (C) Analysis of hallmarks of the first 2 embryonic divisions in LIN-5 phosphorylation mutant, based on time-lapse DIC microscopy. Oscillations are plotted as percentage of embryo height, elongation and AB size as a percentage of embryo width, flattening and rotation as a total fraction of analyzed embryos. (D, E) Mean peak velocities (μm s−1) of anterior and posterior spindle poles measured in a 12.5 s time frame after spindle severing in one-cell embryos of the indicated genotypes, cultured at 25°C (D) or 20°C (E). Error bars: s.e.m. Statistical analyses: Unpaired Welch Students T-test,*P < 0.01 compared to wild type, ** P < 0.001 compared to wild type, *** P < 0.0001 compared to wild type, ns not significant.

https://doi.org/10.1371/journal.pgen.1006291.g004

Next, we used genome engineering to alter the in vivo phosphorylated residues T168, T181, S397, S659 and S662. For each residue, we created a non-phosphorylatable alanine substitution allele, as well as one or more potentially phosphomimetic alleles that contain aspartic acid or glutamic acid at the relevant positions. Alteration of the PAR-1 phosphorylated S397 residue had no apparent effect. Homozygous S397A and S397E animals were viable and showed normal development. Even close examination of LIN-5-mediated processes did not reveal abnormalities (See below; Fig 4C and 4D, S6A and S6B Fig and S1 Video). Thus, although this phosphorylation occurs in vivo, it is by itself not a major determinant of LIN-5 function.

Compared to our transgene rescue experiments (Fig 1B and S1 Fig), the effect of S659 and S662 alanine substitution mutations in endogenous lin-5 was quite mild. The lin-5[S659A,S662A] double mutant animals were viable, with only a slight reduction in intestinal nuclei number (S6A Fig), but displayed a significant increase in embryonic lethality (3.6±1.0% at 25°C, wild type 0.9±0.4%). The phosphomimetic lin-5[S659E,S662D] mutation did not cause embryonic lethality or larval defects, consistent with constitutive phosphorylation of these residues in early embryos (S6A Fig).

In stark contrast, alteration of the candidate CDK-1 phosphorylated residues in the N-terminus, threonine 168 and 181 to alanine (lin-5[T168A,T181A]), aspartic acid (lin-5[T168D,T181D]) or glutamic acid (lin-5[T168E,T181E]), all resulted in typical lin-5 mutant offspring. Regardless of the mutant combination, homozygous animals derived from heterozygous parents developed into sterile, thin and uncoordinated larvae, and showed severely impaired cell division during larval development (Fig 4A and 4B). Importantly, substitution of threonine 168 and 181 with serine residues (lin-5[T168S, T181S]) did not lead to any detectable phenotype or defects in cell division (Fig 4A and 4B). These observations and the in vivo phosphorylation of T168 and T181 indicates that phosphoregulation of T168 and T181 is critical for LIN-5 function, in agreement with the results of the transgene rescue experiments (Fig 1B and S1 Fig). Together, our targeted genome alterations identified several individual amino acids that are required for the in vivo function of LIN-5, including phosphorylated residues in the N-terminus and residues in the GPR-1 binding domain.

LIN-5 S659S662 phosphorylation contributes to cortical pulling forces

Additional characterizations of the non-phosphorylatable and phosphomimetic mutants revealed insight in the functional contribution of LIN-5 phosphorylation. As the contribution of S659 and S662 phosphorylation appeared quite subtle, we examined the spindle in early embryos with substitutions of these residues in detail. In the wild type, meiosis completes after fertilization and results in the formation of a haploid maternal pronucleus, which migrates to meet the paternal pronucleus in the posterior, after which the adjoined pronuclei and centrosomes migrate to the center, rotate and form a spindle along the long axis of the zygote [1,6] (S1 Video). Observations with differential interference contrast (DIC) microscopy showed that these events all occur normally in lin-5[S659A,S662A] and lin-5[S659E,S662D] mutants. Subsequently, in wild type embryos, the chromosomes become aligned at the metaphase plate and are segregated to opposite poles during anaphase. During spindle elongation, the posterior spindle pole oscillates extensively, while the anterior pole remains relatively steady. This coincides with spindle movement towards the posterior, and is followed by flattening of the posterior pole (S1 Video). Starting in anaphase, mutant embryos with non-phosphorylatable lin-5[S659A,S662A] deviated from the wild type, while lin-5[S659E,S662D] mutants showed no phenotype. Specifically, lin-5[S659A,S662A] mutants showed significantly dampened oscillation of both the anterior and posterior pole, reduced spindle elongation, and nearly absent flattening of the posterior spindle pole (Fig 4C and S6B Fig). Nevertheless, both non-phosphorylatable and phosphomimetic S659,S662 mutants underwent asymmetric division of the zygote as normal, which resulted in the formation of a larger anterior blastomere (AB) and smaller germline precursor cell (P1). The spindle normally rotates by 90 degrees prior to division of the P1 blastomere (S1 Video). This failed to occur or was incomplete in 47.1% of the lin-5[S659A,S662A] two-cell embryos, compared to 6.3% and 7.3% incomplete rotation scored in wild type and lin-5[S659E,S662D] mutant embryos, respectively (Fig 4C and S6B Fig). As protein levels were comparable to wild type (S5B Fig), these results suggest that cortical pulling forces are reduced in lin-5[S659A,S662A] mutants. Interestingly, this does not disrupt the asymmetry of the first division and has only a small effect on viability.

To determine cortical pulling forces more directly, we performed spindle severing assays with a UV laser beam [13]. Confirming our DIC analyses, the peak velocities of spindle pole movements were significantly reduced in lin-5[S659A,S662A] embryos (anterior pole 20.5%, posterior pole 18.4% reduced compared to wild type) (Fig 4D, S2 and S5 Videos). Similar experiments performed with lin-5[S659E,S662D] mutant embryos and PAR-1 phosphorylation site mutants (S397A and S397E) did not reveal significant divergence from the wild type (Fig 4D, S3, S4 and S6 Videos). These data support the conclusion that phosphorylation of S659 and S662 contributes to cortical pulling forces, both in the anterior and posterior, and thereby to spindle pole oscillation, spindle elongation, posterior pole flattening and spindle rotation in P1. Moreover, the finding that pulling forces, albeit reduced, remained asymmetric in lin-5[S659A, S662A] mutants explains why these mutants show normal asymmetry of the first division, and normal sizes of the AB and P1 blastomeres.

We wondered whether Wnt-signaling could locally control GSK-3 kinase activity to affect LIN-5 S659, S662 phosphorylation and asymmetric cell division. In the EMS blastomere of the 4-cell embryo, the spindle rotates from a left/right to anterior/posterior position to correctly specify and position the E and MS daughter cells [33]. This rotation is redundantly controlled by MES-1/SRC-1 and MOM-2/MOM-5 Wnt/Frizzled signaling pathways [34]. We examined whether the Wnt pathway contributes to EMS spindle rotation through phosphorylation of LIN-5[S659,S662]. Making use of a mes-1(bn74ts) mutant strain expressing GFP-β-tubulin, we observed normal spindle rotation in lin-5[S659A,S662A] mutant embryos, with only one of 13 embryos showing a tilted spindle angle in the EMS blastomere (S6C Fig). mes-1(bn74ts); lin-5[S659E,S662D] mutant embryos showed an occasional failed rotation or tilted spindle angle. In control mes-1(bn74ts); gsk-3(RNAi) mutants, the EMS spindle failed to rotate in 9/11 embryos (S6C Fig). This clear difference in phenotype shows that LIN-5 S659 phosphorylation is not the major contribution of GSK-3 in EMS spindle rotation. Asymmetric divisions of epithelial seam cells in the C. elegans epidermis also depend on a Wnt-β- catenin asymmetry pathway [35,36], and remained normal in lin-5[S659A,S662A] and lin-5[S659E,S662D] mutants. Thus, evidence for developmental regulation of LIN-5–GPR-1/2 interaction through Wnt-signaling was not obtained. Instead, absence of unphosphorylated S659,S662 peptides in our mass spectrometry analyses, and the wild type appearance of phosphomimetic mutants point to constitutive phosphorylation of the S659,S662 residues.

GPR-1 fails to localize to the centrosome in LIN-5[S659A,S662A] mutants

Our yeast two-hybrid data showed reduced interaction between LIN-5[S659A,S662A] and GPR-1 compared to wild type, which likely explains the reduced pulling forces observed in vivo. We examined whether the colocalization between LIN-5 and GPR-1 in vivo depends on LIN-5 phosphorylation. Hereto, we generated strains with lin-5[S659,S662] double phosphorylation-site alterations in combination with egfp::gpr-1, a CRISPR/Cas9-mediated knockin allele of the endogenous gpr-1 locus. Immunohistochemical detection of eGFP and LIN-5 showed normal colocalization of LIN-5 and GPR-1 in phosphomimetic lin-5[S659E,S662D] mutants at the centrosomes and cell cortex (Fig 5 and S7 Fig; note that LIN-5 becomes clearly visible at the cortex only after the one-cell stage). In contrast, in lin-5[S659A,S662A] mutant embryos, GPR-1 localized to the cortex but no longer accumulated at the centrosomes (Fig 5). Notably, GPR-1/2 localization at the cortex primarily depends on association with the GOA-1 and GPA-16 Gα proteins and is required for pulling forces, whereas ASPM-1–LIN-5 anchors GPR-1/2 at the centrosome without early embryonic requirement [21,37]. Thus, while the loss of centrosomal GPR-1 appears to confirm a reduced binding affinity for LIN-5[S659A,S662A] compared to wild type LIN-5, the reduced pulling forces likely result from a similarly reduced affinity between these proteins at the cortex. Nevertheless, LIN-5 still localized to the cortex in lin-5[S659A,S662A] mutants (S7 Fig). This likely reflects different dynamics of the two complexes; with rapid exchange of LIN-5 at the cortex while centrosomal GPR-1/2 accumulation likely depends on prolonged LIN-5 association. The combined observations in yeast two-hybrid assays, phenotypic analyses, and protein localization studies strongly indicate that phosphorylation of LIN-5 residues S659 and S662 contributes to the affinity of the LIN-5/GPR-1/2 interaction in vivo.

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Fig 5. Localization of LIN-5 and GPR-1 in phosphorylation mutants.

Immunohistochemical staining of embryos expressing wild type or phosphomutant lin-5 and endogenously tagged egfp::gpr-1. Representative images of one-cell embryos in anaphase, stained with anti-LIN-5 (red) and anti-eGFP (green) antibodies, and DAPI to visualize DNA. All images taken with same exposure time, objective and magnification. Anterior to the left, scale bars 10 μm.

https://doi.org/10.1371/journal.pgen.1006291.g005

A potential LIN-5 phosphorylation-dephosphorylation cycle in dynein recruitment

Characterization of the CDK-1 phosphorylation site mutants required a different strategy, as homozygous lin-5[T168A,T181A] and lin-5[T168D,T181D] mutants are fully sterile. To be able to examine the effects of these mutations in early embryos, we created trans-heterozygotes carrying these mutations and egfp::lin-5, a functional CRISPR/Cas9-generated knockin allele of endogenous lin-5. The egfp::lin-5 allele served both as a visible balancer for the lin-5 phosphorylation site mutations, and allowed selective knockdown of functional lin-5 by RNAi against egfp. This strategy allowed us to obtain and characterize early embryos with CDK1-phosphorylation site alterations in LIN-5.

Control immunohistochemical staining experiments confirmed that egfp RNAi treatment of homozygous egfp::lin-5 adults completely removed LIN-5 and eGFP from the offspring (S8 Fig). Following egfp RNAi treatment of heterozygous animals with wild type lin-5 (lin-5(+) / egfp::lin-5), LIN-5 localized normally, but the eGFP staining was lost (Fig 6). These results demonstrate that the RNAi effect remains specific for egfp::lin-5 and does not carry over to the untagged lin-5 allele. Testing balanced lin-5[T168A,T181A] and lin-5[T168D,T181D] animals the same way, we observed that the mutant LIN-5 proteins are expressed and localize as normal to the cortex and centrosomes, while the early embryonic divisions were clearly defective (Fig 6 and S5C Fig). Interestingly, lin-5[T168A,T181A] and lin-5[T168D,T181D] showed similar abnormalities, emphasizing the critical role for the in vivo phosphorylated threonine residues at these positions.

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Fig 6. T168 and T181 mutants show normal subcellular LIN-5 localization.

Immunohistochemical staining of embryos heterozygous for egfp::lin-5 and wild type or phosphomutant lin-5 as indicated. L4 animals were treated with feeding RNAi against control or egfp RNAi to specifically remove egfp::lin-5 function, 48 hours before fixation of their embryos. Representative fluorescence microscopy images of embryos stained with anti-LIN-5 (red) and anti-GFP (green) antibodies, and DAPI to visualize DNA. All images taken with same exposure time, objective and magnification. Anterior to the left, scale bars 10 μm.

https://doi.org/10.1371/journal.pgen.1006291.g006

Using the above-described method, we also performed live imaging by time-lapse DIC microscopy and spindle severing experiments with lin-5[T168A,T181A] and lin-5[T168D,T181D] mutant embryos. Again, the defects observed in both mutants resembled lin-5 strong loss-of-function [19,26], and cortical pulling forces were greatly reduced in both mutants (Fig 4E and S7 and S8 Videos). In contrast, homozygous lin-5[T168S,T181S] mutants showed normal spindle pulling forces (Fig 4E and S9 Video). This indicates that the two threonine residues are not essential per se, but phosphorylation and de-phosphorylation at these sites is likely critical. The lin-5[T168S,T181S] mutants did show dampened spindle oscillation, which might result from somewhat different kinetics of threonine versus serine phosphorylation and dephosphorylation in CDK1 substrates [38].

Because the N-terminus of LIN-5 is implicated in the recruitment of dynein [9], we crossed both mutants with an mCherry::dhc-1 strain, in which the mCherry tag was introduced into the endogenous dynein heavy chain gene by CRISPR/Cas9-mediated knockin. This homozygous mCherry::dhc-1 strain is viable and develops as normal. mCherry::DHC-1 was diffusely detected in the cytoplasm, and distinctly localized at the nuclear envelope, kinetochores, astral microtubules, spindle poles and cell cortex. Localization of dynein was dynamic during all stages of mitosis, but cortical dynein was barely detectable at the one-cell stage. However, following treatment of permeabilized embryos with nocodazole to depolymerize microtubules, mCherry::DHC-1 accumulated on the cell cortex of one-cell embryos in metaphase and anaphase (Fig 7A and S9 Fig). Strikingly, this cortical dynein localization was abolished by lin-5 RNAi, and did not occur in lin-5[T168A,T181A] and lin-5[T168D,T181D] mutant embryos (Fig 7A). Since these mutant LIN-5 forms localize to the cell cortex, T168 and T181 are critical for the function of LIN-5 as a cortical dynein anchor.

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Fig 7. DHC-1 fails to accumulate at meiotic spindle poles in LIN-5 T168, T181 phosphosite mutants.

(A) Representative still images of mcherry::dhc-1; lin-5[T168,T181] one-cell embryos treated with perm-1 RNAi, egfp RNAi and 1 μM nocodazole, and imaged by spinning disk confocal microscopy. The number of embryos with cortical mCherry::DHC-1 is indicated, with “parents” indicating the number of animals from which embryos were analyzed. All images were taken with the same objective, magnification, and exposure time. Anterior to the left, scale bars 10 μm. (B) Representative snapshots of live imaging of mcherry::dhc-1; tubulin::gfp; lin-5[T168,T181] meiotic embryos treated with egfp RNAi, and imaged by wide field microscopy. Time interval between prophase and anaphase is 100 s. All images were taken with same exposure time, objective and magnification. Anterior to the left, scale bars 2 μm.

https://doi.org/10.1371/journal.pgen.1006291.g007

In addition to cortical localization of dynein in mitosis, LIN-5 is also required for dynein recruitment to the poles of the meiotic spindle [37]. Accumulation of dynein at the spindle poles, as well as the cell cortex, occurs coincident with anaphase onset of meiosis I and II, and is needed for spindle rotation and expulsion of chromosomes into a polar body [37,39,40]. While homozygous lin-5[T168S,T181S] mutants showed normal meiosis, we observed polar body absence and abnormally large polar bodies in eGFP::LIN-5-depleted lin-5[T168A,T181A] and lin-5[T168D,T181A] embryos, consistent with lin-5 loss of function. To examine meiotic spindle rotation and dynein localization in such embryos, we combined the lin-5 mutations, balanced by egfp::lin-5, with homozygous gfp::tbb-2 β—tubulin and mCherry::DHC-1 dynein reporters (Fig 7B and S10–S21 Videos). In a control strain with wild type LIN-5, spindle rotation and dynein accumulation occurred in 10 of 11 embryos (the one exception showed rotation but only weak mCherry::DHC-1 accumulation) (Fig 7B left and S10–S12 Videos). Examination of egfp RNAi treated egfp::lin-5 embryos with combined DIC and fluorescence microscopy revealed normal diffuse association of DHC-1 with the meiotic spindle in meiotic prophase, followed by gradual loss of mCherry::DHC-1 from the anaphase spindle, rather than accumulation of dynein at the poles. The failure in dynein localization coincided with failure to rotate the meiotic spindle (Fig 7B). These results agree with our previously reported meiotic lin-5 RNAi phenotype [21,37], although this time we also observed abnormally elongated meiotic spindles in meiosis II in a subset of the embryos, as has been reported for dynein complex subunits [40]. eGFP::LIN-5-depleted lin-5[T168A,T181A] and lin-5[T168D,T181D] embryos were indistinguishable from lin-5 knockdown mutants (Fig 7B and S13–S21 Videos).

In conclusion, substitution of LIN-5 T168 and T181 with non-phosphorylatable alanine or phosphomimetic aspartic acid residues creates a severe defect in LIN-5-mediated dynein recruitment. In contrast, replacement of the same residues with phosphorylatable serine residues did not compromise LIN-5 function (Fig 4A, 4B and 4E). Combined with the available literature [37,39,41], these data point to CDK-1-mediated phosphorylation and subsequent dephosphorylation of the LIN-5 N-terminus as a critical step in dynein recruitment to the meiotic spindle and cell cortex (see below).

C. elegans strains

Strains were cultured on nematode growth medium plates, seeded with Escherichia coli OP50 as previously described [56]. Animals were maintained at 20°C, unless stated otherwise. All strains used in this study are found in S1 Table. Genome modifications in strains SV1568, SV1569, SV1586, SV1588, SV1589, SV1590, SV1600, SV1619, SV1621, SV1622, SV1695 and SV1901 were introduced by making use of CRISPR/Cas9 genome editing as described below.

Post-embryonic analysis of LIN-5 phosphomutants

For functional analysis of wild type and phosphomutant LIN-5, 5 ng/μl Plin-5::gfp::lin-5 DNA, together with 5 ng/μl Psur-5::dsRed and 25 ng/μl Lambda DNA (Fermentas), was injected into the gonad of SV918 young adults. Psur-5::dsRed positive F1 progeny were selected making use of a fluorescence stereo microscope (Leica, MZ16F). After this, lin-5(e1348) homozygous animals were selected based on absence of pharyngeal Pmyo-2::gfp, expressed from the mIn1 balancer chromosome. Rescue analysis of lin-5 null animals was performed by Differential Interference Contrast and fluorescence microscopy, using a Zeiss Axioplan microscope. Intestinal nuclei were counted only in animals expressing Psur-5::dsRed in all intestinal nuclei. Vulval development was assayed for all animals L4 and older.

For quantification of cell numbers in CRISPR/Cas9 knockin mutants, asynchronous populations of animals were fixed, DNA stained with propidium iodide and intestinal and ventral cord nuclei were counted using a Zeiss Axioplan fluorescence microscope. Cells were counted at late larval stages. For the ventral cord, all nuclei of the P2-to-P10 daughter cells and juvenile motor neurons in the region between these cells were counted.

In vitro and in vivo kinase assays

For in vitro CDK1 kinase assays, immunoprecipitations were performed from mitotic lysates of HeLa cells using mouse monoclonal anti cyclin B1 (GNS1) or beads alone as negative control. Immunoprecipitations were incubated for 30 min at 30°C with either Histone H1, bacterially produced GST or GST–LIN-5 in kinase buffer containing 50 mM HEPES at pH 7.5, 5 mM MgCl₂, 2.5 mM MnCl₂, 1 mM dithiothreitol, 50 μM ATP and 2.5 μCi [γ-³²P] ATP. Reactions were terminated by the addition of SDS (5x sample buffer). For mass spectrometry analysis, no [γ-³²P] ATP was added to the kinase assays and incubation time was prolonged to 2 hours at 30°C.

For in vitro GSK3 and CK1 kinase assays, peptides (RRRIRCGSPDSLPDFLADN) containing either unphosphorylated, phosphorylated S659 or phosphorylated S662 were used. Kinases were incubated for 30 min at 25°C with synthetic peptide in kinase buffer containing 200 μM ATP, 50 mM HEPES at pH 7.5, 10 mM MgCl₂, 1 mM EGTA, 2 mM dithiothreitol, supplemented with 20 μCi [γ-³²P] ATP for radioactive kinase assays. Reactions were terminated by the addition of SDS (4x sample buffer).

All other in vitro kinase assays were performed as previously described [15].

In short, kinases were incubated for 30 min at 25°C with bacterially produced GST or GST–LIN-5 in kinase buffer containing 200 μM ATP, 50 mM HEPES at pH 7.5, 10 mM MgCl₂, 1 mM EGTA, 2 mM dithiothreitol, supplemented with 20 μCi [γ-³²P] ATP for radioactive kinase assays. Reactions were terminated by the addition of SDS (4x sample buffer). For mass spectrometry analysis, no [γ-³²P] ATP was added to the kinase assays and incubation time was prolonged to 2 hours at 25°C. Kinases used in this study were: recombinant C. elegans PAR-1 (a kind gift from Erik Griffin and Geraldine Seydoux), and mammalian Aurora B (a kind gift from Susanne Lens), CK1 (New England Biolabs), CK2 (New England Biolabs), and GSK3 (New England Biolabs).

In-gel digestion and phosphopeptide enrichment

Gel bands were cut and processed for protein in-gel digestion as described elsewhere [15]. Briefly, proteins were reduced with dithiothreitol and then alkylated with iodoacetamide. Trypsin was added at a concentration of 10 ng/μl and the mixture was digested overnight at 37°C. Subsequently, peptides were collected from the supernatants and a second extraction using 10% formic acid was performed. Phosphopeptides from LIN-5 were enriched using TiO₂ chromatography [57]. Basically, home-made GELoader tips (Eppendorf, Hamburg, Germany) were packed with TiO2 beads (5 μm, INERTSIL). Peptides were loaded in 10% formic acid and subsequently washed with 20 μl of 80% acetonitrile, 0.1% trifluoroacetic acid (Fluka, Sigma-Aldrich). Phosphopeptides were then eluted twice with 20 μl of 1.25% ammonia solution (Merck, Germany), pH 10.5, and 3 μl of 100% formic acid was finally added to acidify the samples.

Mass-spectrometry analysis

Nanoflow LC-MS/MS was carried out by coupling an Agilent 1100 HPLC system (Agilent Technologies, Waldbronn, Germany) to an LTQ-Orbitrap XL mass spectrometer (Thermo Electron, Bremen, Germany). Peptide samples were delivered to a trap column (AquaTM C18, 5 μm (Phenomenex, Torrance, CA); 20 mm x 100-μm inner diameter, packed in house) at 5 μl/min in 100% solvent A (0.1 M acetic acid in water). Next, peptides eluted from the trap column onto an analytical column (ReproSil-Pur C18-AQ, 3μm (Dr. Maisch GmbH, Ammerbuch, Germany); 40 cm x 50-μm inner diameter, packed in house) at ~100 nl/min in a 90 min or 3 h gradient from 0 to 40% solvent B (0.1 M acetic acid in 8:2 (v/v) acetonitrile/water). The eluent was sprayed via distal coated emitter tips butt-connected to the analytical column. The mass-spectrometer was operated in data-dependent mode, automatically switching between MS and MS/MS. Full-scan MS spectra (from m/z 300 to 1500) were acquired in the Orbitrap with a resolution of 60,000 at m/z 400 after accumulation to target value of 500,000 in the linear ion trap. The five most intense ions at a threshold above 5000 were selected for collision-induced fragmentation in the linear ion trap at a normalized collision energy of 35% after accumulation to a target value of 10,000. Peak lists were created from raw files with MaxQuant⁴³. Peptide identification was carried out with Mascot (Matrix Science) against a Caenorhabditis elegans protein database (http://www.wormbase.org) supplemented with all the frequently observed contaminants in MS (23,502 protein sequences in total). The following parameters were used: 10 ppm precursor mass tolerance, 0.6 Da fragment ion tolerance, up to 3 missed cleavages, carbamidomethyl cysteine as fixed modification, oxidized methionine, phosphorylated serine, threonine and tyrosine as variable modifications. Alternatively, MaxQuant and its search engine Andromeda was also employed for peptide identification and quantification. Data are available via ProteomeXchange with identifier PXD004906

Yeast two-hybrid fragment analysis

Full length gpr-1 was PCR amplified from the ORFeome library (kind gift from Marc Vidal) using KOD polymerase (Novagen) and cloned into bait vector pPC97. Fragments of lin-5 were PCR amplified from the ORFeome library with KOD polymerase (Novagen) and cloned into prey vector pPC86-AN [58]. DB::GPR-1- and AD::LIN-5-encoding plasmids were transformed sequentially into yeast strain Y8930 [58]. Positive interactions were identified on the basis of the activation of the HIS3 and ADE2 reporter genes, indicated by growth on synthetic complete –leucine −tryptophan −histidine + 2 mM 3-Amino-1,2,4-triazole (Sc −leu−trp−his + 3-AT) and synthetic complete –leucine −tryptophan −adenine (Sc −leu−trp−ade) plates.

Generation and verification of yeast two-hybrid lin-5 mutants

To generate mutant clones, PCR was performed on pVP054 (pPC86-AN containing nucleotides 1821–2466 encoding amino acid 609–821 of lin-5) with increased MgCl₂ concentration of 7 mM. PCR products were cloned into pPC86-AN and transformed to DH5α competent cells. Bacterial colonies were collected and the DNA was isolated using a Nucleobond Xtra DNA purification kit (Macherey-Nagel). Bacterial clones were transformed into MB004 (Y8930 in which ADE2 is replaced by URA3 by homologous recombination) containing DB::GPR-1 and plated on synthetic complete –leucine −tryptophan + 2 g/l 5-fluoroacetic acid (Sc −leu−trp + FOA). Colonies were picked and spotted to synthetic complete –leucine −tryptophan (Sc −leu−trp) plates for validation, PCR and sequencing. Clones with single amino acid changes were re-tested by PCR amplification of the fragment and re-cloning into pPC86-AN. Interaction deficient alleles were identified on the basis of no activation of the HIS3 or URA3 reporter gene, indicated by absence of growth on Sc −leu−trp−his + 3-AT plates and synthetic complete –leucine −tryptophan –uracil (Sc −leu−trp−ura) plates.

Validation of yeast two-hybrid full length mutants

For generation of full length mutant clones, plasmids containing fragment clones with selected point mutations were digested and cloned into pVP055 (pPC86-AN containing nucleotides 1–2466 encoding amino acid 1–821 of lin-5). Phosphomutants were generated by either site-directed mutagenesis of pVP055 or Gibson assembly into pVP055 of short regions of lin-5 with point mutations carried in the overlapping region. DB::GPR-1 and AD::LIN-5-encoding plasmids were transformed sequentially into yeast strain MB004. Interaction deficient alleles were identified on the basis of no activation of the HIS3 or URA3 reporter gene, indicated by absence of growth on Sc −leu−trp−his + 3-AT plates and Sc −leu−trp−ura plates.

Generation of CRISPR/Cas9 repair templates

CRISPR repair constructs were inserted into the pBSK vector using Gibson Assembly (New England Biolabs). Homologous arms of at least 1500 bp upstream and downstream of the CRISPR/Cas9 cleavage site were amplified from either cosmid C03G3 (for lin-5 constructs) or C. elegans genomic DNA using KOD Polymerase (Novagen). Linkers containing the altered cleavage sites and point mutations were synthesized (Integrated DNA technologies). For fkbp::egfp::gpr-1, codon-optimized fkbp was synthesized (Integrated DNA technologies) and codon-optimized egfp was amplified from pMA-egfp (a kind gift from Anthony Hyman). For egfp::lin-5, codon-optimized egfp was amplified from pMA-egfp. For mcherry::dhc-1, codon-optimized mcherry was amplified from TH0563-PAZ-mCherry (a kind gift from Anthony Hyman). Mismatches were introduced in the sgRNA target site to prevent cleavage of knockin alleles. All plasmids and primers used for cloning are available upon request.

CRISPR/Cas9 genome editing

Young adults were injected with a solution containing the following injection mix: 30–50 ng/μl Peft-3::Cas9 (Addgene 46168; [59], 30–100 ng/μl u6::sgRNA with appropriate target for dhc-1, gpr-1 and lin-5, 30–50 ng/μl repair template and 2.5 ng/μl pmyo-2::tdTomato. Progeny of animals that express tdTomato were picked to new plates 3–4 days post injection. PCRs with primers diagnostic for homologous recombination at the endogenous locus were performed on F2-F3 populations, where one primer targeted the altered basepairs in the sgRNA site, point mutation or fluorescent tag and the other targeted a region just outside the homology arm. All primers used for genome editing are available upon request.

Preparation of protein lysates for western blot analysis

For yeast protein lysates, cultures were grown overnight at 30°C. Yeast cells corresponding to 4 OD of culture were harvested, treated with Sodium hydroxide and resuspended in 100 2X sample buffer containing β-mercaptoethanol [60].

For C. elegans protein lysates, strains SV1569, SV1663 and SV1664 were grown at 20°C one generation on NGM plates seeded with HB101, followed by a second generation in S-Medium with HB101 bacteria. Gravid adults were harvested and embryos were isolated by hypochlorite treatment. Embryo pellets were snap frozen in liquid nitrogen, grinded using mortar and pestle and resuspended in 5 ml fresh lysis buffer (containing 20 mM Tris-HCl pH 7.8, 250 mM NaCl, 15% glycerol, 0.5% IGEPAL, 0.5 mM EDTA, 50 mM Sodium fluoride, 1 mM β-mercaptoethanol and protease inhibitors (Roche complete, Mini, EDTA-free)). The suspension was passed through a French press 3 times, and the lysate was cleared at 13,000 rpm for 15 min at 4°C.

Western blot analysis

Protein samples were separated on gradient acrylamide gels and subjected to western blotting on polyvinylidene difluoride membrane (Immobilon-P, Millipore). Membranes were blocked with 5% skim milk in PBST for 1 hour at room temperature, or overnight at 4°C for stripped blots. For protein detection, primary antibodies used in this study were: mouse anti–LIN-5 (1:1000) [19] and rabbit anti-Tubulin (1:1000, Abcam) for stripped blots. Secondary antibodies used were: donkey anti-mouse HRP (1:5000, Abcam) and goat anti-rabbit HRP (1:5000, Jackson Immunoresearch). Proteins were dectected with Signalfire Plus chemiluminescent detection (Cell Signaling Technologies) and a Chemidoc MP Imager (Bio-Rad).

Time-lapse imaging and live-cell imaging

DIC time-lapse imaging was performed on strains N2, SV1568, SV1588, SV1590 and SV1600. Animals were grown overnight at 25°C. RNAi feeding of N2 against gsk-3 was performed with the bacterial clone from the Orfeome-based RNAi Library [61,62]. L4 animals were grown for approximately 32 hours at 15°C before shifting overnight to 25°C for imaging, except SV1901 which was kept at 20°C. Embryos were dissected from adults in a solution of 0.8x egg salt (containing 94 mM NaCl, 32 mM KCl, 2.7 mM CaCl₂, 2.7 mM MgCl₂, 4 mM HEPES, pH 7.5; [63]) on coverslips and mounted on slides with 3% agarose prepared with egg salt. Embryos were imaged with 5s time intervals with a 100x/1.4 NA lens on a Zeiss microscope at 20°C. Relative positions of the spindle and furrow were analyzed manually using ImageJ.

Live-cell imaging of EMS rotation was performed on strains SV1783, SV1784 and SV1785. L4 animals were grown overnight at 20°C. Embryos were dissected from young adults as above and imaged with a 100x/1.4 NA lens on a Zeiss microscope at 20°C. Spindle rotation in EMS was followed over time with images taken at several time points.

Live-cell imaging of microtubule depolymerization upon nocodazole treatment was performed on strain AZ244. Young adult animals were injected with dsRNA [64] against perm-1 and grown for 20 hours at 15°C. Embryos were dissected from young adults in a solution of 0.8x egg salt containing 1 μM nocodazole on coverslips and mounted on concave slides. Embryos were imaged with a 60x/1.4 NA lens on a Nikon Eclipse Ti microscope with Perfect Focus System and Yokogawa CSU-X1-A1 spinning disk confocal head at room temperature.

Live-cell imaging of mitotic DHC-1 localization was performed on strains SV1619, SV1635, SV1638 and SV1639. Young adult animals were injected with dsRNA [64] against perm-1 and egfp and grown for 20 hours at 15°C. Embryos were dissected from young adults in a solution of 0.8x egg salt containing 1 μM nocodazole on coverslips and mounted on concave slides. Still images of mitotic embryos in metaphase were taken within minutes after nocodazole addition. Eliminating nonspecific toxic effects, embryos on the same slide at the same time continued nuclear envelope degradation, and perm-1(RNAi) embryos continued embryonic development in the absence of nocodazole.

Embryos were imaged with a 60x/1.4 NA lens on a Nikon Eclipse Ti microscope with Perfect Focus System and Yokogawa CSU-X1-A1 spinning disk confocal head at room temperature. mCherry::DHC-1 localization was analyzed in mitotic one-cell embryos after nuclear envelope breakdown. Mitotic embryos were also identified based on presence of a polar body, enlarged centrosomes and remnant of the mitotic spindle.

Live-cell imaging of meiotic DHC-1 localization was performed on strains SV1702, SV1898, SV1899 and SV1902. For RNAi treated animals, young adult animals were injected with dsRNA [64] against egfp and grown for 24 hours at 15°C. Embryos were dissected from young adults as above and imaged with 10s time intervals with a 100x/1.4 NA lens on a Zeiss microscope at 20°C. For images presented in Fig 7B and S12, S15, S18 and S21 Videos, images were processed by subtracting a Gaussian-blur filtered image (Sigma(Radius): 20) using ImageJ. S10, S11, S13, S14, S16, S17, S19 and S20 Videos represent the unprocessed files.

UV laser spindle ablation

Spindle severing with a UV laser microbeam was performed on strains SV1585, SV1594, SV1596, SV1618, SV1700 and SV1701 essentially as previously described [13]. RNAi feeding of SV1700 and SV1701 against egfp was performed with a bacterial clone containing full length egfp in the L4440 double T7 plasmid [61]. L4 animals were grown on RNAi for approximately 48 hours. For analysis of spindle pulling forces, animals were kept at 25°C for 24 h before ablations. Spindle ablations were carried out at 25°C (Fig 4D) or 20°C (Fig 4E) on a spinning disk confocal microscope. The spindle midzones were severed at anaphase onset and images of GFP-β-tubulin were taken at 0.5 s intervals. For analysis, the position of the spindle poles was automatically tracked using the MTrack2 plugin in ImageJ. Peak velocities of the anterior and posterior spindle poles were determined within a 12.5 s time frame after ablation. Representative videos for every strains are shown in S2–S9 Videos.

Microscope setup: Nikon Eclipse Ti microscope with Perfect Focus System, Yokogawa CSU-X1-A1 spinning disk confocal head, S Fluor 100x N.A. 0.5–1.3 objective (at 1.3), Photometrics Evolve 512 EMCCD camera, Cobolt Calypso 491 nm (100 mW) and Teem Photonics 355 nm Q-switched pulsed lasers, ILas system (Roper Scientific France/ PICT-IBiSA, Institut Curie) to control the UV laser, ET-GFP (49002) filter, ASI motorized stage MS-2000-XYZ with Piezo Top Plate with Tokai Hit INUBG2E-ZILCS Stage Top Incubator (controlled at 25°C), controlled by MetaMorph 7.7 software.

Antibodies and immunohistochemistry

For immunostaining, embryos were dissected from adults in 8 μl of water on poly-L-lysine–coated slides. Embryos were freeze-cracked and fixed for 5 min in methanol at −20°C and then for 20 min in acetone at −20°C. After fixation, embryos were rehydrated in phosphate-buffered saline (PBS) containing 0.05% Tween-20 (PBST) and blocked with blocking solution (PBST containing 1% bovine serum albumin and 1% goat serum [Sigma-Aldrich]) for 1 h. Embryos were stained with primary and secondary antibodies for 1 h and washed after each incubation with PBST four times, 15 min each time. Finally, the embryos were embedded in ProLong Gold Antifade containing 4′,6-diamidino-2-phenylindole (DAPI). Primary antibodies used in this study were: mouse anti–LIN-5 (1:10; [19] and rabbit anti-GFP (1:500, Life Technologies). Secondary antibodies were used at a concentration of 1:500. Secondary antibodies used were: goat anti-rabbit Alexa Fluor 488, and goat anti-mouse Alexa Fluor 568 (Invitrogen). Images were taken with a 63x/1.4 NA lens on a Zeiss confocal microscope.

Immunohistochemistry quantification of LIN-5 levels

Embryos were dissected and stained with antibodies as described above. Images were taken with a 63x/1.4 NA lens on a Zeiss confocal microscope using identical microscope setting for all images taken for every secondary antibody. Mean intensity of LIN-5 was measured using ImageJ by selecting fixed size regions that depended on the developmental stage. For every embryo 2 centrosomes and cytoplasmic regions were quantified.