The polarity-induced force imbalance in Caenorhabditis elegans embryos is caused by asymmetric binding rates of dynein to the cortex

INTRODUCTION

Successful symmetric and asymmetric cell division relies on the precise positioning and orientation of the mitotic spindle, which in turn ensures the correct partitioning of chromosomes and cell organelles. This choreography requires cortical pulling forces. From yeast to humans, the molecular motor dynein is key to producing these (Carminati and Stearns, 1997; Shaw et al., 1997; Gonczy et al., 1999; Karki and Holzbaur, 1999; Dujardin and Vallee, 2002; Nguyen-Ngoc et al., 2007; Moore et al., 2008; Markus and Lee, 2011b; Collins et al., 2012; Laan et al., 2012; Kotak and Gonczy, 2013). It localizes at the cell cortex and pulls on the astral microtubules that radiate from the spindle poles (McNally, 2013). Indeed, cytoplasmic dynein (hereafter referred to simply as “dynein”) is minus-end directed, walking toward the spindle poles.

Dynein is a dimer of a multisubunit complex that performs various functions depending on the choice of subunits (Pfister and Lo, 2012). In Caenorhabditis elegans, because only one homologue of the intermediate chain of dynein DYCI-1 exists, we used a labeling of this subunit to reveal dynein in all its functions. Consistently, DYCI-1 depletion results in phenotypes that mostly reflect the loss of the heavy chain, containing the motor domain (Kamath and Ahringer, 2003; Sonnichsen et al., 2005). To understand how dynein-generated forces contribute to spindle positioning, we previously analyzed spindle rocking during mitotic anaphase in C. elegans zygotes. Our results suggest that cortical force generators pull briefly, for 1 s or less (Pecreaux et al., 2006a). This was confirmed by the direct viewing of dynein heavy chain (DHC-1) at the cortex both in nematodes and mammalian cells (Collins et al., 2012; Kiyomitsu and Cheeseman, 2012; Barbosa et al., 2017; Schmidt et al., 2017). So with such a transient residence, how does dynein reach the cortex? In higher eukaryotes, it most likely arrives from the cytoplasm rather than from the pool at the microtubule plus ends, thus relegating plus-end accumulation to a minor role (Splinter et al., 2012; Duellberg et al., 2014; Barbosa et al., 2017; Baumbach et al., 2017; Jha et al., 2017; Schmidt et al., 2017). However, pulling forces may still depend weakly on plus-end tracking protein EBP-2^EB, which is required for dynein accumulation. Therefore, more investigation into dynein accumulation at the plus ends and into dynein’s role in cortical force generation is still needed.

Although it is clear that the cortical forces generated by dynein play an essential role in a broad range of organisms, the mechanistic link between polarity and these forces remains elusive (McNally, 2013). Indeed, in the C. elegans zygote the posterior displacement of the spindle has been attributed to polarity-cue induction of higher forces at the posterior cortex (Grill et al., 2001; Colombo et al., 2003; Grill et al., 2003). This is caused by a doubling of the active force generators at the posterior cortex as compared with the anterior one (Grill et al., 2003; Grill and Hyman, 2005; Pecreaux et al., 2006a; Nguyen-Ngoc et al., 2007; Redemann et al., 2010). Besides an unequal amount of dynein at the cortex, the force imbalance could also be caused by a differential regulation of the intrinsic properties of dynein acting as a molecular motor, or of its dynamics, such as a higher binding rate or a lower unbinding rate on the stronger-force side. Importantly, the mechanism that translates polarity into a force imbalance is involved in the segregation of cell fate determinants and in the balance between proliferation and differentiation (Gonczy, 2008; Moore and Cooper, 2010; Morin and Bellaiche, 2011; McNally, 2013; Rose and Gonczy, 2014; Williams et al., 2014; di Pietro et al., 2016).

To explore the causes of the force imbalance, we first focused on the mechanisms for bringing dynein to the cortex. Combining advanced image processing and fluorescence correlation spectroscopy (FCS), we found that the dynein accumulated at the microtubule plus ends in an EBP-2^EB–dependent manner was not transported to the cortex. However, this dynein pool does contribute slightly to the pulling forces. Most dyneins probably reach the cortex by three-dimensional diffusion from the cytoplasm instead. In both mechanisms, we found that the dynein inflow to the cortex was not polarized, and they could not account for the pulling force imbalance. However, by carefully analyzing the dynamics at the cortex, we did find an asymmetric binding rate of dynein to the force-generating complex. This asymmetry leads to more microtubule pulling events on the posterior side in a polarity-dependent manner.

RESULTS

Dynein pools in the cytoplasm are not polarized

To determine whether cytoplasmic dynein pools could contribute to building the force imbalance at the cortex, we first investigated whether soluble dynein was symmetrically distributed in the cytoplasm. We used a strain expressing a fluorescent dynein intermediate chain, DYCI-1, labeled by mCherry expressed under its own promoter and carried by a transgene (Sarov et al., 2006, 2012). During mitosis, this strain displayed no significant phenotype, and the corresponding transgene rescued the dyci-1(tm4732) null allele. So even with possibly altered expression levels, DYCI-1::mCherry performs like the native protein (Supplemental Text 1.1-2). We used spinning disk microscopy to examine the strain in a plane between the coverslip and the spindle, the so-called lower spindle plane (LSP), and at the cortex (Figure 1, A–D). In the LSP, we observed both spindle and central spindle staining, with dynein spots moving toward the cortex during metaphase and anaphase (Figure 1, A and B, Supplemental Figure S1, A and C, and Supplemental Movies S1 and S2). This is consistent with the spindle and dotty cytoplasmic localizations that were previously revealed after both antibody staining and live imaging of the dynein subunit DHC-1 (Gonczy et al., 1999; Nguyen-Ngoc et al., 2007; Barbosa et al., 2017; Schmidt et al., 2017). After confirming our labeling, we then checked whether the cytoplasmic dynein fraction was polarized. FCS measurements of the dynein in the cytoplasm did not show an asymmetry (Figure 1E and Supplemental Text 1.3) (Widengren et al., 1994), and we concluded that equal amounts of soluble dynein are available in both embryo halves, ready to diffuse and bind to the cortex.

The unequal amounts of dynein at each cortex half could be caused by EB homologue proteins and the corresponding accumulation of dynein at the plus ends. To further investigate dynein behavior in the cytoplasm, we denoised the images, tracked the dynein spots in the LSP, and analyzed their dynamics in detail (Supplemental Figure S2, A–C, and Supplemental Text 2) (Sage et al., 2005; Huet et al., 2006; Jaqaman et al., 2008; Coupe et al., 2012). We mostly found directed tracks that moved from the centrosome to the cortex (Figure 1, F and G). Since those moving in the reverse direction were rare (5 ± 3%) and not expected to help bring dynein to the cortex, these directed tracks were not investigated further. In a similar analysis of simulated particles, we observed that short tracks tended to be classified as diffusive-like regardless of their actual motion. So, in processing experimental data, we ignored the diffusive-like tracks, being uncertain of their real motion (Supplemental Figure S3, Supplemental Movie S8 and S9, and Supplemental Text 2.5). We therefore focused only on spots displaying a directed motion toward the cortex. To ascertain that these corresponded to dyneins accumulating at the microtubule plus ends, we confirmed that the tracks depended on end-tracking protein EBP-2^EB, the dynactin subunit DNC-1^p150glued, and LIS-1 but not on CLIP-1^CLIP170, in agreement with previous findings (Supplemental Figure S4, A–F, and Supplemental Text 3.1) (Barbosa et al., 2017; Schmidt et al., 2017). EBP-2 by itself consistently displayed centrifugal spots independently of DNC-1^p150glued and LIS-1, confirming that dynein accumulates at the plus ends by interacting with these proteins, which in turn binds EBP-2 (Supplemental Figure S4G). Furthermore, the dynein spots colocalized with microtubules and EBP-2 (Figure 2, A and B, Supplemental Figure S5, A–D, Supplemental Movies S10 and S11, and Supplemental Text 3.2). Since these spots were therefore proven to correspond to the plus-end dyneins, we checked the symmetry of their distribution. We measured the posterior-to-anterior ratios, which were 0.95 ± 0.09 for the directed tracks (p = 0.7, compared with 1) and 1.0 ± 0.1 for the diffusive-like ones (p = 0.5, N = 7 embryos, 1341 tracks). In both cases, we found no significant asymmetry. Importantly, the spot intensities were similarly distributed between the anterior and posterior (Supplemental Figure S5E, Kolmogorov-Smirnov test, p = 0.27, N = 8 embryos). Thus, the dynein counts in each spot are the same for both the anterior and posterior embryo halves. Therefore, the plus-end accumulation of dynein is not polarized. To conclude, there are equal pools of dynein in the anterior and posterior cytoplasm, resulting in symmetric inflow to the cortex either by cytoplasmic diffusion or by via the microtubule plus ends.

Dyneins are not transported to the periphery by the microtubule plus ends

We next wondered whether dynein accumulation at the microtubule plus ends could have a more subtle contribution to the force imbalance. Besides having larger or more numerous dynein spots on the posterior side, their dynamics could be asymmetrical, for instance, if spots arrive faster to that side. Indeed, while the microtubule growth rates are similar on both sides, their residence time at the cortex was reported to be polarized (Labbe et al., 2003; Srayko et al., 2005). We first checked whether the amount of dynein reaching the cortex through the plus ends is enough to contribute significantly to cortical forces. We used FCS to image the microtubule tips, finding ∼50 DYCI-1::mCherry proteins per plus end in the strain also carrying the endogenous copies. In comparison, we counted three to four times more EBP-2::GFP in each plus end (Supplemental Text 1.4). Previous studies showed 100–200 microtubule contacts at the cortex at any time during anaphase (Pecreaux et al., 2016; Redemann et al., 2017; Bouvrais et al., 2019). Therefore, dynein accumulation at the microtubule plus ends could provide a large excess of dyneins to the cortex, much higher than the previously reported 10–100 active force generators residing 1 s or less on each half cortex (Grill et al., 2003; Pecreaux et al., 2006a; Barbosa et al., 2017; Schmidt et al., 2017).

So does an accumulation of dynein at the microtubule plus ends contribute to a cortical-force imbalance? To explore this, we checked whether the dynein pool is transported to the cell periphery as it is in yeast (Sheeman et al., 2003; Lee et al., 2005; Markus et al., 2009; Markus and Lee, 2011a; Roberts et al., 2014). By FCS, we examined microtubule ends moving across the focal volume (comet tails, see Figure 2C) in a doubly labeled EBP-2::GFP;DYCI-1::mCherry strain. Both EBP-2 and DYCI-1 had similar spot association kinetics, which depend exponentially on local cytoplasmic concentrations of the corresponding proteins (Figure 2D and Supplemental Text 4.1) (Dragestein et al., 2008). Furthermore, the association rates obtained by fitting were similar. Therefore, just like EBP-2^EB, dynein is mostly recruited from the cytoplasm at the plus ends. Interestingly, when examining these two proteins in the cytoplasm by fluorescence cross-correlation spectroscopy, they are not associated, which indicates that dynein attaches to the plus ends in its own way (Supplemental Figure S6A). This is reminiscent of the cortical targeting mechanism of dynein in yeast and is thus compatible with its transport.

In contrast to yeast, dynein plus-end accumulation did not seem to be transported after an in vitro reconstitution using mammalian purified proteins (Duellberg et al., 2014; Baumbach et al., 2017; Jha et al., 2017). The critical question is thus whether dynein stays on the microtubule lattice when EBP-2^EB leaves the plus ends. Indeed, EB is only accumulated there but not transported (Bieling et al., 2007). EBP-2 and DYCI-1:mCherry detachment rates were similar (Figure 2E), suggesting that dynein is not transported. To strengthen this result, we next checked whether dynein displays the same detachment dependence on microtubule growth rates as EB proteins. Indeed, the tail of the EB fluorescent comet at the plus ends grows as the microtubule grows faster. We thus measured (generally in seven embryos, 1500 tracks per condition) the DYCI-1::mCherry comet-tails, varying the growth rates through hypomorphic klp-7(RNAi) and clip-1(RNAi). These showed a linear relation with a slope significantly different from zero: 1.2 ± 0.2 s (p = 0.03) (Figure 2F, Supplemental Figure S6B, and Supplemental Text 4.2). This result shows that most dyneins leave the microtubule plus end similarly to EBP-2^EB. Therefore, like that protein, dynein is only accumulated and not transported at the microtubule plus ends. In conclusion, the force imbalance at the cortex could not be caused by an asymmetry in dynein dynamics at the plus ends.

EBP-2 contributes modestly to cortical pulling forces

Dynein may appear in two pools at the cortex. One is dependent on LIN-5^NuMA, a member of the trimeric complex that generates pulling forces, and the other is dependent on EBP-2^EB and may act as a backup to the first pool to ensure cortical pulling (Schmidt et al., 2017). To see whether this second pool could contribute to force imbalance, we first checked whether EBP-2^EB and dynein accumulated at the plus ends contribute to cortical forces. To do so, we used labeled γ-tubulin (γTUB)::GFP centrosome labeling and analyzed the oscillations of the posterior centrosome on ebp-1/2/3(RNAi) or crossing with null mutation ebp-2(gk756). We found that although its paralogues EBP-1 and EBP-3 do not, EBP-2 contributes a bit to force generation (Figure 3A). Could this be related to dynein accumulating at the plus ends through EBP-2? We measured the oscillation amplitudes in control and ebp-2(gk756) null mutants in the absence of DYCI-1 and found no significant differences (Figure 3A). As a control, we performed the same analysis in ebp-2(gk756) null mutant and found no significant difference with and without clip-1^CLIP170(RNAi) treatment. These experiments suggest that EBP-2^EB and DYCI-1 probably act along the same pathway. We therefore suggest that dynein accumulated at the microtubule plus ends contributes mildly to cortical pulling forces.

To strengthen this conjecture, we went at the question from a different direction, using our previously published “tube” assay. This reports the location of force-generating events by creating cytoplasmic membrane invaginations (Figure 3B) (Redemann et al., 2010). These are rare in nontreated conditions but more numerous on weakening the actin-myosin cortex by nmy-2(RNAi), done only partially to preserve the polarity (Supplemental Figure S6C and Supplemental Text 5.1). PH::GFP membrane labeling reveals these invaginations. Importantly, RNA interference (RNAi) depletion of either cortical force generators or related proteins significantly decreases invagination counts. We added ebp-2(RNAi) to the hypomorphic nmy-2(RNAi), resulting in significantly decreased invaginations as compared with the control (Figure 3C). We thus confirmed that EBP-2 enhances cortical force generation.

It was recently suggested that EB helps initiate the dynein run, thus in our case generating cortical forces. We therefore decided to address whether EBP-2 promotes dynein targeting to the cortex by measuring dynein after EBP-2 depletion. However, first, we had to ensure that the DYCI-1::mCherry visible as transient spots at the cortex is involved in force-generating events (Figure 1C, Supplemental Figure S1, E and G, and Supplemental Movie S3). We began by using the tube assay and crossed the DYCI-1::mCherry and PH::GFP strains, observing the cortex on partial nmy-2(RNAi) (Supplemental Movie S6 and Figure 3, D and E). We found that half of the invaginations colocalized with DYCI-1::mCherry spots (Figure 3F and Supplemental Text 5.1). We figured that the lack of colocalization in the other half was because of detection limits imposed by high DYCI-1::mCherry cytoplasmic background fluorescence, and indeed the threshold for spot detection over the background fluorescence was estimated at 26 ± 4 dyneins per spot (Supplemental Text 2.6 and Supplemental Figure S8). We also noticed that dynein spots appeared 0.4 s before invaginations, suggesting that pulling forces created by labeled dyneins must generate the invaginations (Figure 3, D and E, and Supplemental Movie S7). To reinforce our hypothesis, we asked whether labeled DYCI-1 colocalizes with the members of the cortical force-generating complex GPR-1/2 (Nguyen-Ngoc et al., 2007). We crossed strains carrying randomly integrated DYCI-1::mCherry and GPR-1::YFP and acquired images of both dyes. We found that 30% of the cortical spots in DYCI-1::mCherry (N = 8 embryos) colocalized with GPR-1::YFP ones (Figure 3G and Supplemental Text 3.2). Because there are a limited number of dynein cortical anchors, not all DYCI-1::mCherry spots are likely to contribute to pulling forces, and thus this colocalization proportion is coherent (Grill and Hyman, 2005; Pecreaux et al., 2006a; Park and Rose, 2008; Riche et al., 2013). Next, we did RNAi to partially deplete the dynein light chain DLI-1, known to be involved in cortical pulling forces, resulting in an almost complete disappearance of the DYCI-1::mCherry spots at the cortex (Figure 3H) (Yoder and Han, 2001; Pecreaux et al., 2006a). Overall, DYCI-1::mCherry thus clearly reveals the dyneins involved in force generation at the cortex.

Having ascertained that cortical DYCI-1::mCherry spots correspond to force generators, we observed them on depleting EBP-2 either by ebp-2(RNAi) or by crossing with the null ebp-2 mutant (Figure 3, I and J). We found a drastic reduction in both diffusive-like and directed cortical track densities (Figure 3K), similarly to what happens in the lower spindle plane on EBP-2 depletion. Together with the results based on the posterior centrosomal oscillations, it suggested that EBP-2 may contribute, albeit modestly, to targeting dynein to the cortex and to pulling force generation there. EBP-2’s limited contribution makes it unlikely that it plays a part in the anteroposterior imbalance of forces at the cortex.

Dynein dynamics at the cell cortex

Before further investigations on the mechanism creating the force imbalance, we analyzed the dynamics of dynein at the cortex as we did in the LSP, finding equal proportions of directed and diffusive-like tracks (Figure 4, A and B). Furthermore, spots of both types spent less than 1 s at the cortex (Figure 4, C and D, and Supplemental Text 6.1). Since dynein spots mostly display a directed motion in the cytoplasm, we reasoned that cortical directed tracks might correspond to dynein spots completing their arrival to the cortex. Indeed, the optical sectioning in spinning disk microscopes may allow for viewing of the subcortical regions. To test this hypothesis, we used RNAi to deplete EFA-6, a putative microtubule regulator whose depletion results in more stable microtubules, which are then more numerous at the cortex. On efa-6(RNAi), we found more directed tracks than in the control, while the diffusive-like population was not significantly affected (Figure 4, E and F). Therefore, we concluded that directed tracks should correspond to dynein at the plus ends of microtubules. In this respect, these tracks belong to the EBP-2–dependent population of cortical dynein (Schmidt et al., 2017). Because the dyneins move in the same way as those in the LSP, which is not polarized, we can conclude that directed-tracks do not contribute to the cortical force imbalance.

Cortical dynein spots with diffusive-like motion display an asymmetric distribution that is polarity dependent

To further understand how polarity translates into force imbalance, we checked whether the population having diffusive-like motion could be dynein residing at the cortex, engaged in pulling-force generation. We analyzed the motion of dynein spots within four regions extending equally along the AP axis and measured the number of tracks per unit of time, also known as the frequency (Figure 5D and Supplemental Text 2.4), which is independent of the dynein residence time at the cortex. To account for dynein expression level variability (Supplemental Text 2.4), we normalized to the average value of the anterior-most region (#1 in Figure 5D) in the control. We observed that only the spots with a diffusive-like motion were polarized (Figure 5, A and B). Because the frequencies in the middle regions were symmetric and forces from these regions were suggested to have smaller contributions to spindle positioning, we concentrated only on the outside regions (1 and 4 in Figure 5D) (Krueger et al., 2010). We reasoned that since GPR-1/2^LGN is posited to be the limiting factor in force generation, its depletion should alter the diffusive-like track counts (Colombo et al., 2003; Park and Rose, 2008; Riche et al., 2013). We measured their frequency in embryos subjected to gpr-1/2(RNAi) and found that it was lower than in the control (Figure 5E). Furthermore, this treatment also suppressed the asymmetry (Figure 5F). Although partial, this treatment was penetrant, since another experiment using differential interference contrast (DIC) microscopy showed a total disappearance of posterior centrosomal oscillations under the same conditions (N = 7 embryos), while they remained in the N = 8 untreated controls as previously observed (Colombo et al., 2003). Importantly, the gpr-1/2(RNAi) treatment preserved a posterior displacement that was indistinguishable from the control, with a final centrosomal position at 76 ± 7% of embryo length (mean ± SD) compared with 79 ± 1% in the control, and thus the positional regulation of forces was normal (Riche et al., 2013; Bouvrais et al., 2019). The following treatments were also partial to preserve a spindle positioning indistinguishable from the control. Overall, we concluded that the diffusive-like population consists of dynein involved in cortical pulling and does reveal force-generating events.

Because equal flows of dynein reach the cortex from the anterior and posterior cytoplasm, only three mechanisms can account for the force asymmetry: 1) the binding rate of dynein to the cortex is higher on posterior side, 2) dynein resides on and pulls longer on posterior side, or 3) dynein is more efficient on posterior side (e.g., develops higher forces on that side). This last option appears unlikely, since it fails to account for the asymmetric frequencies of diffusive-like tracks or for the asymmetric number of active force generators (Figure 5A) (Grill et al., 2003). We wondered which alternative would best explain the differences in the anterior and posterior centrosomal oscillations. We examined nontreated embryos carrying γTUB::GFP and found that the frequencies and amplitudes on the posterior side were both greater than those of the anterior one. We compared these results with the predictions of the model from our previous work (Pecreaux et al., 2006a), using these three possibilities (Supplemental Text 6.2). In fact, we found that the experimental oscillations were consistent with an asymmetry in dynein dynamics, either in the binding or unbinding rates (possibilities 1 and 2). But they were not consistent with asymmetric efficiency (#3) or even with unequal total dynein counts caused by an asymmetric inflow to the cortex. We thus set out to investigate the possibilities tied to dynein dynamics. However, because the model is linearized and the experimental frequencies of centrosomal oscillations are moderately different, we could not yet choose between the other two explanations.

We next used direct viewing of force-generating events to decide whether dynein binds to the cortex at a higher rate on posterior or whether it stays there longer (alternatives 1 and 2, respectively). We measured the dynein-spot residence time and found no significant differences between the posterior and anterior regions for the diffusive-like population (Supplemental Text 6.1 and Figure 5, C and D). This supports an asymmetry in binding rates (alternative #1). Furthermore, we observed that gpr-1/2^LGN(RNAi) suppressed the asymmetry in the diffusive-like track frequencies while only slightly decreasing the residence time. To gain certainty, we targeted the force-generating complex in another manner, partially depleting LIN-5^NuMA by RNAi (Figure 5H) (Gotta et al., 2003; Nguyen-Ngoc et al., 2007). This treatment suppressed the asymmetric frequencies of the tracks having a diffusive-like motion, while not significantly altering their residence times (Figure 5, F and H). However, to ensure correct meiosis, we were limited to a more hypomorphic treatment since LIN-5 is involved in more functions than is GPR-1/2 (van der Voet et al., 2009). As above, we checked that the final spindle positioning was not significantly affected by measuring the position at which the cytokinesis furrow started to ingress at the cortex. This was 55.6 ± 2.1% (mean ± SD, N = 5) on lin-5(RNAi), compared with the control’s 59.0 ± 3.9% (N = 9, p = 0.24). Together, these experiments support the idea that polarity is reflected by an increase in dynein binding to the cortex on the posterior side.

Finally, we asked whether GPR-1/2 enrichment at the posterior pole could cause the enhanced binding rate. We altered its localization and investigated the frequency distribution of the diffusive-like tracks. Indeed, on cnsk-1(RNAi), GPR-1/2 and LIN-5 are enriched at the anterior pole of the embryo during early mitosis and then homogeneously distributed later (Panbianco et al., 2008). On csnk-1(RNAi), hypomorphic to preserve the spindle position, we observed the cortex and analyzed the dynein spots during metaphase. We found a significant increase in the frequency at the anterior pole as compared with the control, canceling the asymmetry (Figure 5G). As expected, the residence times of the same dynein spots were unaffected (Figure 5I). Using DIC at the midplane, we ensured that the treatment did not alter the position for ingression of cytokinesis furrows at the cortex. We found 56.4 ± 2.5% (mean ± SD, N = 4) on csnk-1(RNAi) compared with 59.0 ± 3.9% (N = 9, p = 0.24) in the control. Therefore, the GPR-1/2^LGN and LIN-5^NuMA concentrations are reflected in the dynein-binding rate variations. Overall, direct observation of the dynein population displaying diffusive-like motion led us to conclude that the force imbalance is caused by a larger binding rate on the posterior side. This increase is probably due to the presence of larger concentrations of either GPR-1/2 or LIN-5 at the cortex.

DISCUSSION

Using a fluorescence-labeled DYCI-1 as a bona fide reporter of dynein, we discovered that this molecular motor is not actively transported toward the cell periphery. Instead, it accumulates at the microtubule plus ends and is only briefly immobilized on the lattice. The lack of EBP-2^EB mildly but significantly decreases cortical forces, however. It was proposed that cortical forces may position the spindle independently of dynein using hypomorphic RNAi or temperature-sensitive alleles of DHC-1 (Schmidt et al., 2005). However, some dynein activity may have persisted in these experiments, leaving open the possibility that dynein is strictly necessary to move the spindle to the posterior side. Consistently, a partial dli-1(RNAi) cancels out centrosomal oscillations and strongly reduces posterior displacement, suggesting that dynein contributes to most if not all cortical pulling force generation (this work, Yoder and Han [2001], and Pecreaux et al. [2006a]). More recently, using laser severing of the spindle, a reduced peak velocity was observed on ebp-2(RNAi) (Schmidt et al., 2017). Surprisingly, using an ebp-2 null mutant obtained by CRISPR-Cas9 and tracking the centrosomes in DIC at one frame every 2 s, the same authors failed to observe the subtle anaphase decrease in oscillation amplitude. In contrast, we were able to see it using both ebp-2(RNAi) and the ebp-2(gk756) null mutant. To account for this discrepancy, these authors suggest that a pool of dynein dependent on EBP-2 and located at the microtubule plus ends creates a backup mechanism to ensure cortical pulling force generation. Alternately, this discrepancy might be due to the reduced time resolution of their centrosomal tracking assay. We confirmed our results, observing fewer membrane invaginations on ebp-2(RNAi). In sum, it seems clear that EBP-2 plays a role, although modest, in cortical force generation.

Putative mechanisms for the role of EBP-2^EB

When EBP-2^EB is depleted, there are fewer dynein spots at the cortex. This suggests that this protein contributes to the cortical localization of dynein, consistent with previous reports (Barbosa et al., 2017; Schmidt et al., 2017). However, surprisingly, depleting EBP-2^EB only mildly reduces cortical forces. So how does EBP-2 act? In light of recent studies, its mechanisms are threefold: increasing dynein targeting to the cortex, promoting the assembly of the force-generating complex, and initiating the dynein run to the minus ends.

During microtubule growth, an EBP-2–dependent accumulation of dynein could bias its diffusion toward the periphery. Indeed, when EBP-2 and thus dynein detach from GDP-tubulin behind the cap, dynein affinity for the EBP-2 bound to the GTP-cap may encourage its diffusion toward the plus ends, that is, toward the cell periphery (Rousselet et al., 1994; Preciado Lopez et al., 2014). We consistently found that dynein and EBP-2 did not associate in the cytoplasm. Although not very efficient, a subtle mechanism such as this could promote cortical targeting in the context of fast dynein turnover.

At the cortex, having concentrated dyneins at the microtubule plus ends would facilitate the assembly of the force-generating complex together with GPR-1/2 and LIN-5, and the capture of a microtubule for it to pull on. Such an advantage would be even stronger if the force generators act in clusters, as recently suggested using human purified proteins, and also if dynein targeting to the cortex by diffusion is reduced (Okumura et al., 2018). This reduction could occur when the subcortical dynein concentration decreases as dynein leaves the cortex and runs along the microtubules to the centrosome, as in Madin–Darby canine kidney (MDCK) mammalian cells (Zheng et al., 2013). These findings in other organisms call for further studies to elucidate the details of offloading in the nematode.

Finally, once the complex has assembled and has engaged a microtubule, EBP-2^EB may enhance the initiation of the dynein run and thus force generation. This mechanism was previously reported in vitro using human proteins and in the nematode (Barbosa et al., 2017; Jha et al., 2017). EBP-2 can also promote the tracking of depolymerizing microtubule ends by dynein, a supplemental mechanism for force generation also proposed in C. elegans (Kozlowski et al., 2007; Laan et al., 2012; Duellberg et al., 2014; Baumbach et al., 2017; Okumura et al., 2018). Even disregarding these mechanistic details, it is clear that the majority of the dynein-dependent forces are still present if EBP-2^EB and even all three EBP-1/-2/-3 orthologues are absent (Schmidt et al. [2017] and this study). Therefore, we suggest that neither these mechanisms nor the cytoplasmic diffusion of dynein play a role in building the imbalance of cortical forces reflecting embryo polarity.

The diffusive-like population at the cortex reports force-generating events

We found two populations of dynein at the cortex, with distinct spot dynamics. Only the one with diffusive-like motion is asymmetrically distributed. Using fluorescence recovery after photobleaching, Schmidt and colleagues consistently found that only their LIN-5–­dependent dynein population displayed different dynamics between the anterior and posterior cortices, not their EBP-2–dependent one (Schmidt et al., 2017). These populations may well correspond to our tracks with diffusive-like and directed motions. Furthermore, we found that spots with a diffusive-like motion reveal force-generating events. In support of this hypothesis and in addition to their dependence on GPR-1/2 and LIN-5, their residence times are consistent with the brief cortical stay of microtubules engaged in force generating and with the estimated force-generator runtimes deduced from the modeling of centrosomal oscillations during anaphase (Pecreaux et al., 2006a; Kozlowski et al., 2007; Sugioka et al., 2018; Bouvrais et al., 2019). Second, the number of spots is consistent with the expected active force-generator count: 10–100 per cortex half (Grill et al., 2003; Redemann et al., 2010). Indeed, in the posterior region, we observed ∼0.008 diffusive-like tracks per square micrometer of visible cortex area (instantaneous density). This extrapolates to ∼20 diffusive-like tracks in the posterior half at any instant, which is well within the published range.

The force-generating events come from dynein having diffusive-like motion, and we wonder whether some spots in this population may relate to other mechanisms. First, it is possible that some spots represent stalled dyneins (Laan et al., 2012). This would be consistent with the fact that GPR-1/2 colocalizes with only a third of all dynein spots (equal proportions of diffusive-like and directed, so we would have expected ∼50% colocalization). However, the number of stalled dyneins must be low, as a detailed analysis of centrosomal motion does not detect them (Pecreaux et al., 2016). Second, some tracks might mix motions that are directed (microtubules arriving at the cortex) and diffusive-like (dynein taking part in force generation) and are classified according to where they spent the longest time. These two considerations may account for why we observed a posterior-to-anterior ratio of dynein-track frequencies below 2, when it has been proposed that there are double the active force generators present at the posterior pole (Grill et al., 2003). However, since the diffusive-like population mostly corresponds to force-generating events, this offers a unique opportunity for deciphering the details of the polarity-induced force imbalance.

The mechanism of the polarity-reflecting force imbalance

After studying cytoplasmic dynein, we ruled out that a different number of molecules reaches each half-cortex. We instead suggest that the force imbalance is caused by dynein’s higher binding rate at the posterior cortex (Figure 6). Previous investigations reported an anteroposterior asymmetry in cortical residence time of the microtubules, although the low frame rates used (0.5 and 1.4 Hz, respectively) limited the resolution (Labbe et al., 2003; Sugioka et al., 2018). Furthermore, these authors found surprisingly long residence times as compared with established values (Kozlowski et al., 2007; Bouvrais et al., 2019). It is possible that a low-performance linker in the tracking algorithm would lead to incorrect analysis of closely packed successive tracks, interpreting them as the same spot. Such an artefact would take spot frequency imbalances and present it as residency imbalances.

What mechanism could yield higher binding rates of dynein at the posterior cortex? Certainly higher amounts of GPR-1/2^LGN or LIN-5^NuMA there could displace the association/dissociation balance toward more assembly/force-generating (Figure 6). Such a mechanism does not lead to increased dynein residence at the cortex. Furthermore, it enables fast adaption to internal evolution and to dynamic polarity cues (Thery et al., 2006; Fink et al., 2011; Riche et al., 2013; Bouvrais et al., 2019). The increased binding rate could correspond not just to the kinetics of dynein associating with GPR-1/2 and LIN-5, or to the capture rate of an astral microtubule by the complex, but also to the initiation rate of the dynein force generation. In this respect, it is an effective binding rate.

This mechanism translating polarity into force imbalance is part of the threefold regulation of the forces positioning the mitotic spindle. It occurs as reported here through polarity, through spindle pole positioning (Riche et al., 2013; Bouvrais et al., 2019), and also through force-generator persistence in pulling on microtubules (processivity) (Pecreaux et al., 2006a). This last regulation is visible as the residence time in our experiments and is probably controlled by mitotic progression (McCarthy Campbell et al., 2009). Investigating the one-cell nematode embryo has paved the way to understanding the mechanisms of asymmetrical division (Gonczy, 2008; Morin and Bellaiche, 2011). It would be very interesting to explore whether force polarization is due to asymmetric force generator binding rates in other organisms, since these mechanisms based on dynamics are advantageous, promoting adaptation during mitosis.

MATERIALS AND METHODS

Culturing C. elegans

Caenorhabditis elegans strains were cultured as described in Brenner (1974) and dissected to obtain embryos. All strains containing DYCI-1::mCherry or TBA-2^α-tubulin::YFP were kept at 25°C, while functional experiments (anaphase oscillations) investigating the role of CLIP170, DLI-1, DYCI-1, and EB1 homologues were performed at 18°C. The exception to this were strains carrying clip-1(gk470), kept at 23°C.

Caenorhabditis elegans strains

The standard wild-type strain was Bristol N2 (Brenner, 1974). The following fluorescent strains were used: TH163 (DYCI-1::mCherry) (Sarov et al., 2006), TH27 (GFP::TBG-1^γ-tubulin) (Oegema et al., 2001), TH65 (YFP::TBA-2) (Kozlowski et al., 2007), TH66 (GFP::EBP-2) (Srayko et al., 2005), DE74 (GFP::PLCδ1-PH) (Johnston et al., 2010), TH110 (mCherry::PAR-6) (Schonegg et al., 2007), and TH242 (GPR-1::YFP) (Redemann et al., 2011). Standard genetic crosses were done to generate these multilabeled combinations: JEP2 (DYCI-1::mCherry;YFP::TBA-2), JEP12 (DYCI-1::mCherry;GFP::EBP-2), JEP20 (DYCI-1::mCherry;GFP::PLCδ1-PH), and JEP58 (DYCI-1::mCherry;YFP::GPR-1). To obtain JEP27 and JEP32 carrying the GFP::TBG-1 transgene and the ebp-2(gk756) or clip-1(gk470) mutations, we crossed TH27 with VC1614 or VC1071, respectively (C. elegans Deletion Mutant Consortium, 2012). The strain carrying the dyci-1(tm4732) lethal mutation was provided by the Mitani Lab via the National BioResource Project, and JEP9 was generated by crossing with VC2542 to balance the lethal mutation with the nT1[qIS51] translocation. JEP30 and JEP40 strains homozygous for dyci-1(tm4732) were obtained by double-crossing JEP9 with JEP23 and TH163, respectively (Supplemental Text 1.2). The transgenes encoding the GFP, YFP, and mCherry fusion proteins in all constructs but DYCI-1::mCherry were under the control of the pie-1 promoter.

Gene silencing by RNA interference

Except when otherwise stated, embryonic RNAi was done by feeding, using both the Ahringer library (Fire et al., 1998; Kamath and Ahringer, 2003) and clones ordered from Source BioScience. However, the clone targeting ebp-1/3 was made in the lab. To do so, N2 genomic DNA was used to amplify a region from the target gene (see Table 1). This amplicon was then cloned into the L4440 RNAi feeding vector and transformed into HT115 bacteria. For ebp-1, a region corresponding to exons 2 and 3 after splicing was amplified using four long primers and fused by PCR amplification before L4440 cloning. Table 1 lists the primers used for amplification.

For ebp-1 and ebp-1/3 RNAi treatment, quantitative reverse transcription PCR (Q-RT-PCR) measurements showed a 40–60% reduction in the number of transcripts without changes in the ebp-2 mRNA levels. Total RNA was extracted from ∼20 worms using a Direct-zol RNA MicroPrep kit (Zymo Research). Production of cDNA was done with a ProtoScript II First Strand cDNA Synthesis kit (New England Biolabs). For Q-PCR, Power SYBR Green PCR Master Mix (Thermo Fisher Scientific) was used with a 7900HT Fast Real-Time PCR System (Applied Biosystems).

In the strains where the DYCI-1::mCherry was randomly integrated, its expression levels varied. To account for this, each RNAi experiment was compared or normalized to nontreated or L4440-treated control embryos imaged on the same day.

Except where otherwise stated, RNAi was partial, and observation was performed 23–25 h after plating the worms. To avoid too-strong or unrelated phenotypes, we used the following treatment durations when observing the TH163 randomly integrated DYCI-1::mCherry strain: lin-5(RNAi), 17 h; gpr-1/2(RNAi), 48 h; lis-1(RNAi), 16-18 h; dnc-1(RNAi), 16-18 h; ebp-2(RNAi), 20 h; dyci-1(RNAi), 16 h; dli-1(RNAi) 24 h; and csnk-1(RNAi), 8 h. To perform the “tube assay” (Redemann et al., 2010), embryos were treated by nmy-2(RNAi) for 24 h. In the case of double RNAi experiments for invagination counting, JEP20 L1 or L2 worms were fed over 32 h by bacteria with the plasmid for ebp-2(RNAi) or with L4440 empty vector (as a control) and then plated on a bacterial culture containing the two clones to perform either ebp-2(RNAi)/nmy-2(RNAi) or L4440(RNAi)/nmy-2(RNAi) by feeding for an additional 16 h.

Live imaging

Embryos were dissected in M9 buffer and mounted on pads (2% wt/vol agarose, 0.6% wt/vol NaCl, 4% wt/vol sucrose). We imaged one-cell C. elegans embryos during metaphase and anaphase. Dynein/EBP-2 tracking was performed on a LEICA DMi6000/Yokogawa CSU-X1 M1 spinning disk microscope with an HCX Plan APO 100×/1.40 oil objective. Illumination was done with a homemade setup based on a Fianium white light laser conveniently filtered around 488 and 561 nm (Roul et al., 2015). Except when otherwise stated, images were acquired with a 200-ms exposure time (5 Hz) using a Photometrics Evolve Camera (Roper) and MetaMorph software (Molecular Devices) without binning. During the experiments, the embryos were kept at 24°C. To image embryos at the LSP, we typically moved the focus between 3 and 5 µm below the spindle plane (Supplemental Figure S1D). For invagination analysis, DYCI-1::mCherry and GFP::PLCδ1-PH signals were acquired with a 168-ms exposure time at a final frequency of 2.5 Hz for each color using a Photometrics Evolve Camera on a LEICA DMi8/Yokogawa CSU-X1 M1 spinning disk microscope and Inscoper software (Inscoper, France).

Image processing

The standard deviation maps were generated with Fiji’s ZProject plugin for ImageJ, specifying a “SD” over 6 s of the time-lapse image sequence (Rostampour et al., 1988; Cai et al., 2007).

The tracking of labeled centrosomes and analysis of trajectories were performed using custom tracking software (Pecreaux et al., 2006a) and developed with Matlab (The MathWorks). Tracking of –20°C methanol-fixed γTUB::GFP embryos indicated an accuracy of 10 nm (Pecreaux et al., 2016). Embryo orientations and centers were obtained by cross-correlating embryo background cytoplasmic fluorescence with artificial binary images mimicking embryos or by contour detection of the cell using the background fluorescence of cytoplasmic γTUB::GFP with the help of an active contour algorithm (Pecreaux et al., 2006b). We averaged the results over all of the replicas for each condition.

Invaginations were counted manually using ImageJ for 2 min after anaphase onset, in turn obtained by monitoring the spindle using DYCI-1::mCherry labeling.

Statistics

The displayed center values are the means except when otherwise stated. Averaged values were compared using a two-tailed Student’s t test with the Welch-Satterthwaite correction for unequal variance except if stated otherwise. For the sake of simplicity, we encoded confidence levels using stars as follows: ♦, p < 0.1; *, p ≤ 0.05; **, p ≤ 0.01; ***, p ≤ 0.001; ****, p ≤ 0.0001; and ns, nonsignificant, p > 0.1. The ns indication may be omitted for clarity’s sake.

FOOTNOTES

ACKNOWLEDGMENTS

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