Two mechanisms drive pronuclear migration in mouse zygotes

Scheffler, Kathleen; Uraji, Julia; Jentoft, Ida; Cavazza, Tommaso; Mönnich, Eike; Mogessie, Binyam; Schuh, Melina

doi:10.1038/s41467-021-21020-x

Download PDF

Article
Open access
Published: 05 February 2021

Two mechanisms drive pronuclear migration in mouse zygotes

Nature Communications volume 12, Article number: 841 (2021) Cite this article

10k Accesses
31 Citations
219 Altmetric
Metrics details

Subjects

Abstract

A new life begins with the unification of the maternal and paternal chromosomes upon fertilization. The parental chromosomes first become enclosed in two separate pronuclei near the surface of the fertilized egg. The mechanisms that then move the pronuclei inwards for their unification are only poorly understood in mammals. Here, we report two mechanisms that act in concert to unite the parental genomes in fertilized mouse eggs. The male pronucleus assembles within the fertilization cone and is rapidly moved inwards by the flattening cone. Rab11a recruits the actin nucleation factors Spire and Formin-2 into the fertilization cone, where they locally nucleate actin and further accelerate the pronucleus inwards. In parallel, a dynamic network of microtubules assembles that slowly moves the male and female pronuclei towards the cell centre in a dynein-dependent manner. Both mechanisms are partially redundant and act in concert to unite the parental pronuclei in the zygote’s centre.

The WD40-protein CFAP52/WDR16 is a centrosome/basal body protein and localizes to the manchette and the flagellum in male germ cells

Article Open access 28 August 2020

Ran-GTP assembles a specialized spindle structure for accurate chromosome segregation in medaka early embryos

Article Open access 01 February 2024

Apicobasal RNA asymmetries regulate cell fate in the early mouse embryo

Article Open access 30 May 2023

Introduction

Fertilization unites the father’s and the mother’s chromosomes to create a genetically unique embryo. Following sperm entry, the egg completes the second meiotic division and the male and female pronuclei subsequently form in proximity of the zygote’s surface^1,2,3, which is lined by an actin cortex. From the zygote’s periphery, they need to move inwards in order to unite the paternal and maternal chromosomes on the first mitotic spindle. Recent studies revealed that incomplete pronuclear migration leads to defects during embryo development in mammals. If the pronuclei are too far apart when they break down, two entirely separate spindles will form^4,5. Zygotes with two separate spindles are more likely to form multiple nuclei upon the first mitotic division, and often fail to develop into healthy embryos^4,6.

Despite the importance of pronuclear migration, the mechanisms that unite male and female pronuclei in mammals are poorly understood. Work that was mostly carried out in non-mammalian species has established essential functions for microtubules and dynein in pronuclear migration^{7,8,9,10,11,12,13,14,15,16}. Dynein anchored in the cytoplasm is thought to move the male pronucleus inwards by pulling on the sperm aster that is associated with the male pronucleus^15,17,18. The female pronucleus recruits dynein on its nuclear envelope and moves towards the male pronucleus along the sperm aster¹⁹. Pronuclear centration is further promoted by pushing of pronucleus-associated microtubules against the zygote’s cortex^8,20. Microtubule-dependent pushing similarly drives nuclear movements in Drosophila oocytes²¹ and fission yeast²².

Although mouse zygotes contain a dense microtubule network and acentriolar microtubule-organizing centres (aMTOCs)^23,24, pronuclear migration in this system is thought to rely more on the actin cytoskeleton^{5,25,26,27,28}. How actin helps to centre pronuclei before cell division is unclear.

Here, we developed an experimental system that allowed us to follow and experimentally probe the entire process of pronuclear migration in live mouse zygotes. By combining it with loss-of-function assays, we identified two distinct mechanisms that drive pronuclear migration. In the first mechanism, the male pronucleus is initially moved inwards by the flattening fertilization cone. At the same time, the actin nucleation factors Formin-2 and Spire are recruited into the fertilization cone in a Rab11a-dependent manner, where they locally nucleate actin, accelerating the pronucleus’ inwards movement and detaching it from the plasma membrane. In the second mechanism, which acts throughout the central region of the zygote, a cytoplasmic microtubule network forms and carries the pronuclei inwards in a dynein-dependent manner. Both mechanisms act in concert and are partially redundant, which increases the likelihood of successful pronuclear migration. Thus, both the actin and microtubule cytoskeletons actively participate in the unification of parental chromosomes in mouse zygotes.

Results

Pronuclear migration is driven by distinct mechanisms in the periphery and in the central region of the zygote

The formation of mammalian pronuclei and the early stages of their migration have not been visualized previously. To observe early events in live zygotes immediately after fertilization (Supplementary Fig. 1a), we microinjected mRNAs encoding fluorescent markers into eggs before in vitro fertilization (IVF), by adapting a particularly gentle and quantitative method for microinjection²⁹. We found this approach is compatible with high fertilization rates and mouse embryo developmental capacity (Supplementary Fig. 11a, b).

Introduction of the markers histone H2B-mCherry and MyrGFP (myristoylated EGFP) into unfertilized eggs allowed us to, respectively, visualize chromatin and the cell surface in zygotes (Fig. 1a). In time-lapse microscopy experiments that captured images from early pronuclear formation to the first mitotic division of the mouse embryo, we recorded high-resolution z-stacks covering the entire zygote. We then reconstructed from fluorescence signals the pronuclei and cell volume in 3D to quantitatively assess pronuclear migration (Fig. 1a, b, Supplementary Fig. 1b, c and Supplementary Movie 1). The distances of the pronuclear centroids to the cell centre were calculated in 3D and then plotted over time (Fig. 1a, b and Supplementary Fig. 1d).

**Fig. 1: Pronuclei migrate by different modes in the periphery and central region of the zygote.**

The male pronucleus formed directly underneath the cell surface, within the fertilization cone, at an average distance of 37.2 ± 2.3 µm from the cell centre (Fig. 1a and Supplementary Fig. 1a, e). During the first 30 min, it moved rapidly inwards with a mean velocity of 0.38 ± 0.06 µm/min (Fig. 1b, c). In the subsequent 30 min, its velocity decreased to 0.10 ± 0.04 µm/min. Over the following 7 h, its velocity decreased further, from ~0.04 to 0.01 µm/min (Fig. 1b, c). In contrast, the female pronucleus formed further inwards than the male pronucleus, at an average distance of 23.7 ± 2.7 µm from the cell centre (Fig. 1a and Supplementary Fig. 1a, e). During the first 30 min upon its formation, the female pronucleus moved with an average velocity of 0.06 ± 0.08 µm/min and was hence around six times slower than the male pronucleus (Fig. 1b, c). Over the following 7.5 h, its speed decreased further from ~0.04 to 0.01 µm/min. From 2 to 8 h upon pronuclear formation, the male and the female pronuclei moved at similar speeds, ranging between ~0.04 and 0.01 µm/min (Fig. 1c).

We repeated this analysis also in the outbred CD-1 mouse strain with similar results, suggesting that this movement pattern is conserved between strains (Supplementary Fig. 1f, g).

The male pronucleus was around six times faster during the initial phase of migration than the female pronucleus. We asked if this disparity is due to the different distances from the cell surface at which the male and female pronuclei were forming. To test this hypothesis, we forced the female pronucleus to form more peripherally (at an average distance of 34.2 ± 5.8 µm from the cell centre) (Fig. 2a, b and Supplementary Movie 2) by partially depolymerizing the spindle remnant that held the female pronucleus away from the surface (Supplementary Fig. 1h). Strikingly, the female pronucleus now showed a similar migratory pattern to the male pronucleus: during the first 30 min, it moved rapidly at an average speed of 0.33 ± 0.17 µm/min before adopting a slower speed, ranging between ~0.08 and 0.03 µm/min from 0.5 to 2 h (Fig. 2c, d). Conversely, in untreated zygotes, male pronuclei occasionally (3/56 zygotes; green dots in Supplementary Fig. 1e) formed at a distance from the cell surface, and then did not display the fast phase of migration (green curves in Supplementary Fig. 1i).

**Fig. 2: Female pronuclei have a pronounced fast migration phase when forced to form close to the cell surface.**

Together, these data establish that pronuclei move distinctly in different regions of the zygote. They adopt a rapid migration mode near the surface, and a slow migration mode throughout the centre. Male pronuclei normally form close to the surface and show both the first fast phase of migration as well as the second slower phase of migration. Female pronuclei typically form deeper inside the zygote and show a less pronounced fast migration phase, in addition to the second slower phase.

Pronuclei migrate inwards in the absence of Rab11a-positive vesicles

What are the mechanisms that drive pronuclear migration in the periphery and in the central region? Nuclei also become centred in unfertilized, prophase-arrested oocytes³⁰ where Rab11a-positive vesicles move via Myosin-5b motors in the cytoplasm³¹ and participate in nuclear movement^30,31 by generating a pressure gradient³⁰. A similar vesicle-dependent gradient could also be involved pronuclear centration in zygotes⁵.

Live and fixed cell analysis showed that Rab11a-positive vesicles were present in early stages of pronuclear migration, but largely depleted during the late stages of pronuclear migration (Supplementary Fig. 2a–f). To test if Rab11a is important for pronuclear migration, we expressed a dominant-negative variant of Rab11a, or treated zygotes with brefeldin A (BFA)³², which depleted the vesicles in the zygote (Supplementary Fig. 2g–j). Interestingly, the male and female pronuclei still moved inwards under both conditions, demonstrating that Rab11a-positive vesicles are not essential for pronuclear migration (Fig. 3a–c, Supplementary Movie 3 and Supplementary Fig. 3a–c). Dominant-negative Rab11a and BFA decreased the velocity of pronuclear migration during the initial fast migratory phase, but not during the second slower phase (Fig. 3d, e and Supplementary Fig. 3d e). Moreover, pronuclear distances to the cell centre at nuclear envelope breakdown (NEBD) were only mildly increased upon expression of the dominant-negative Rab11a^S25N variant (by 30% for the male pronucleus and 10% for the female pronucleus; Fig. 3f).

**Fig. 3: Rab11a is required for pronuclear migration in the periphery of the zygote.**

If a pressure gradient was the main driving force behind pronuclear migration it should also move oil droplets inwards that have a similar size as pronuclei³³. However, oil droplets ranging in diameter from 9 to 24 µm (female pronuclei grow from ~7 to 21 μm; male pronuclei from ~10 to 24 μm during migration; Supplementary Fig. 1j, k) did not move inwards unlike pronuclei (Supplementary Fig. 3f, g). Only oil droplets that were in close proximity of a male pronucleus as it formed in the fertilization cone rapidly moved inwards right after pronuclear formation, but later remained at their position or only underwent local random movements (Supplementary Fig. 3h–j and Supplementary Movie 4).

Together, these data indicate that pronuclear migration is not driven by global pressure gradients in mouse zygotes. Consistent with our findings, a recent study using microsensors to monitor forces in the cytoplasm of the zygote did not observe global inwards-directed forces acting on polyethylene microspheres or nanodevices³⁴.

The flattening fertilization cone moves the male pronucleus inward and recruits Rab11a, Formin-2 and Spire

Although dominant-negative Rab11a did not block pronuclear migration, it decreased the velocity of the male and female pronuclei during the initial fast migratory phase by ~60–70% (Fig. 3d, e). We thus investigated how Rab11a promotes this phase of migration. High-resolution live microscopy of mScarlet-Rab11a revealed an unexpected Rab11a-positive structure in proximity of the forming male pronucleus (Fig. 4a and Supplementary Fig. 4a): Rab11a was strongly enriched at the cell surface, directly behind the forming male pronucleus (Fig. 4a and Supplementary Movie 5). The enrichment was transient (lasting for 22 ± 15 min; n = 19), coinciding with the fast migration phase. In mouse oocytes, Rab11a-vesicles recruit the actin nucleation factor Spire2 to the plasma membrane³², which acts in concert with Formin-2 to generate actin filaments in mouse oocytes³⁵. We thus asked if Spire2 and Formin-2 are also enriched behind the male pronucleus. Live-cell microscopy revealed that fluorescently tagged versions of the actin nucleation factors Spire2 and Formin-2 also became concentrated in the cell surface region behind the forming male pronucleus, concomitant with the enrichment of Rab11a (Fig. 4b, c, Supplementary Fig. 4b, c and Supplementary Movie 6).

**Fig. 4: Rab11a, Spire, Formin-2 and actin accumulate in the fertilization cone.**

High-resolution 3D microscopy revealed that Spire2 was enriched throughout the entire fertilization cone, and most highly concentrated in a ring at the base of the cone (Fig. 4b and Supplementary Fig. 4f). As the cone flattened, the Spire2 ring contracted and eventually coalesced into a single spot directly behind the male pronucleus.

The flattening fertilization cone moved the male pronucleus inwards. The pronucleus remained in direct proximity of the cell surface as the cone flattened. Only when the boundary of the contracting Spire2 ring reached the area just behind the male pronucleus, the male pronucleus detached and rapidly moved inwards (Fig. 4b and Supplementary Fig. 4f). Formin-2 was initially uniformly located at the cortex and rather depleted in the area of the fertilization cone (Fig. 4c and Supplementary Fig. 4g). As the fertilization cone flattened, Formin-2 became gradually enriched at the base of the contracting fertilization cone and was most highly concentrated behind the male pronucleus as it detached from the plasma membrane. The initial rapid phase of pronuclear migration is hence a two-step process: first, the flattening of the fertilization cone that moves the male pronucleus inwards and second, the launching of the male pronucleus away from the plasma membrane that coincides with an accumulation of the actin nucleation factors Spire2 and Formin-2 behind the male pronucleus.

Finally, we tested if the ablation of the initial fast phase of migration in dominant-negative Rab11a expressing oocytes may be due to a requirement of Rab11a in targeting the actin nucleation factors to the fertilization cone. Indeed, Spire2 was no longer enriched in the cell surface region in proximity of the forming male pronucleus upon expression of dominant-negative Rab11a (Fig. 4d, e).

Actin nucleation by Spire2 and Formin-2 pushes the male pronucleus inwards

Consistent with an accumulation of actin nucleation factors, the cell surface region was an active site of actin nucleation, and F-actin-enriched patches were transiently detected below the cell surface as the pronucleus left the cell surface (Fig. 4f, Supplementary Movie 7 and Supplementary Fig. 4h). A previous study reported that the actin network density towards the cortex is higher than the density towards the centre in the region of the pronuclei⁵, consistent with this observation.

It is well established that local actin nucleation can generate pushing forces, which can for instance promote membrane protrusion or the propulsion of pathogens in a host cell^36,37,38,39. We therefore asked if Spire and Formin-2 in the region of the fertilization cone might similarly cause pushing forces that propel the pronuclei away from the cell surface towards the centre of the zygote. To answer this question, we first overexpressed Spire2 to assess if we could enhance inwards pronuclear movement by increasing local actin nucleation. Indeed, overexpression of Spire2 increased F-actin behind the male pronucleus (Fig. 5a and Supplementary Movie 8), and strikingly, extended the rapid fast inwards movement of the male pronucleus, resulting in a 70–110% increase in the velocity of the male pronucleus between 0.5 and 1.5 h after its formation (Fig. 5b–d, Supplementary Fig. 5a and Supplementary Movie 9). In 14/31 zygotes that expressed very high levels of Spire2, the pronucleus moved beyond the zygote’s centre (Supplementary Movie 10), into the opposite hemisphere of the zygote. In five of these cells, the pronuclei even reached the opposite cortex and bounced back from there, coinciding with a new Spire2 enrichment event (Supplementary Fig. 5b and Supplementary Movie 11).

**Fig. 5: Spire-dependent local actin nucleation drives pronuclear migration in the periphery of the zygote.**

When Spire2 was highly overexpressed, some zygotes formed Spire2 surface patches in regions that were devoid of pronuclei. Interestingly, also these Spire2 surface patches generated inwards-directed forces, as we observed five cases, where oil droplets were located close to these patches and then rapidly accelerated inwards (Supplementary Fig. 5c and Supplementary Movie 12). Together, these data establish that local enrichments of Spire at the cortex cause inwards-directed forces that can move structures such as oil droplets and pronuclei away from the cell periphery. Consistent with inwards-directed forces in the region of the fertilization cone, particle image velocimetry (PIV) analysis revealed local inwards-directed flows in the region of the male pronucleus, but no global large-scale flows throughout the cytoplasm (Supplementary Fig. 5d).

To directly test for pushing forces, we carried out a laser ablation experiment: using a two-photon laser, we cut either behind or in front of the male pronucleus during the fast migration phase (Fig. 5e and Supplementary Fig. 5e). Cutting behind the pronucleus led to a significant ~70% decrease in pronuclear velocity (Fig. 5f, g). In contrast, cutting in front of the pronucleus, did not decrease its velocity (Fig. 5f, g). Together, these data establish that local pushing forces that act behind the male pronucleus move the pronucleus inwards during the initial fast migration phase.

Next, we tested directly if actin nucleation by Spire and Formin-2 is required for the initial rapid phase of pronuclear migration in proximity of the cell surface. To this end, we used a dominant-negative Formin Homology 2 domain (FH2) construct, which blocks the interaction between Spire and Formin-2⁴⁰. Previously, we showed that this FH2 construct inhibits Spire- and Formin-2-dependent nucleation of the cytoplasmic actin network in oocytes⁴⁰, and we here show that it also depletes the cytoplasmic actin network in zygotes (Supplementary Fig. 6c, e). Microinjection of recombinant FH2 domain significantly reduced the mean velocity by 61% during the initial 30 min of male pronuclear migration (Fig. 6a, b, d). Pronuclear migration was not inhibited during the slower, second phase (Fig. 6d), and the pronuclei eventually reached the cell centre (Fig. 6f).

**Fig. 6: The fast migration mode requires actin nucleation in the periphery of the zygote.**

As we had observed that the female pronucleus also adopts a more pronounced rapid initial migratory mode when it is forced to form close to the cell surface (Fig. 2a–d), we next asked if this rapid mode is also due to a local enrichment of Rab11a and Spire2. Indeed, Rab11a and Spire2 were both enriched behind the female pronucleus when it formed beneath the cell surface, in a structure that closely resembled the fertilization cone (Fig. 6g and Supplementary Fig. 4d, e, i). As the fertilization cone-like structure flattened, the female pronucleus moved inwards and prominent actin patches appeared between the cell cortex and the female pronucleus (Fig. 6h and Supplementary Movie 13). The launching of the peripheral female pronucleus was hence very similar to that of the male pronucleus.

Also in control cells, the female pronucleus was slowed down by FH2 (Fig. 6c, e) or dominant-negative Rab11a during the initial phase of migration (Fig. 3c, e). Rab11a and Spire/Formin-2-dependent actin nucleation hence also promotes the early stages of female pronuclear migration in unperturbed cells. Actin and Spire2 were strongly enriched in the cytokinetic furrow (Fig. 6g, h), and also the spindle remnant contained actin (Fig. 6h, top row)⁴¹. Both structures are in close proximity of the female pronucleus. An additional prominent enrichment of Rab11a and Spire/Formin-2 at the cortex was not detectable but might have been shielded by their strong enrichment in the cytokinetic furrow.

Together, these data establish a two-step mechanism that drives the rapid inwards pronuclear movement in the cortical region of the zygote. The male pronucleus forms within the fertilization cone, which becomes enriched in Spire and Formin-2 in a Rab11a-dependent manner. First, the cone flattens, thereby moving the pronucleus inwards. Second, Spire and Formin-2 concentrate behind the male pronucleus where they locally nucleate actin. This generates inwards-directed forces that launch the pronucleus away from the cell surface. The female pronucleus can trigger the same launching mechanism when it is forced to form directly underneath the cell surface and is also accelerated inwards by Spire and Formin-2 in unperturbed zygotes, albeit to a lesser degree than the male pronucleus.

Microtubules and dynein drive the slower, second phase of pronuclear migration

We next investigated the slow movement pattern in the central region of the zygote. Neither loss of vesicles by dominant-negative Rab11a nor disruption of the cytoplasmic actin network by FH2 slowed down pronuclei in the central region of the zygote (Figs. 3a–f and 6a–f). Moreover, Rab11a-positive vesicles were strongly depleted from oocytes during the second phase of migration (Supplementary Fig. 2a, b). Together, these data suggest that the second migration phase does not rely on Rab11a-positive vesicles or the cytoplasmic actin network but must be mediated by other cytoskeletal structures instead. Consistent with previous studies^23,42, zygotes assembled a cytoplasmic microtubule network as the pronuclei formed and grew (Fig. 7a and Supplementary Movie 14). Live microscopy of microtubules in zygotes using the fluorescently tagged microtubule binding domain of MAP7/Ensconsin (EMTB) revealed that the microtubule network density increased gradually from formation of the pronuclei onwards (Fig. 7a and Supplementary Fig. 7a). The microtubule network was dynamic and included multiple microtubule asters that were nucleated at aMTOCs (Fig. 7a)⁴². We hence asked if microtubules are involved in the second, slower phase of migration that occurs throughout the centre of the zygote.

**Fig. 7: Microtubule- and dynein-dependent transport drives pronuclear migration throughout the centre.**

Depolymerization of microtubules by nocodazole (Supplementary Fig. 7b) significantly decreased the speed of both the male and the female pronucleus during the second slower phase of migration (Fig. 7b–f and Supplementary Movie 15). Consistently, pronuclear migration was incomplete, because the distance of the male and female pronucleus from the cell centre at NEBD was increased by 50% and 60%, respectively (Fig. 7g). This demonstrates that both pronuclei rely on microtubules to efficiently migrate to the zygote’s centre. By contrast, the fast migration in the periphery of the zygote was not slowed down (Fig. 7c, e), indicating that microtubules are dispensable for the initial fast migration phase.

Next, we investigated, which motor proteins are required for the inwards movement of the pronuclei. Pronuclear migration in several other species is dynein dependent^13,15,43, making dynein a good candidate for inwards movement. Indeed, inhibition of dynein by microinjection of the p150-CC1 peptide⁴⁴ significantly decreased the velocity of the male and female pronucleus during the second period of migration (Fig. 7h–l), while significantly increasing the distance of the pronuclei to the cell centre at NEBD was significantly increased (Fig. 7m).

In further support of inwards-directed forces acting within the microtubule network, 33/47 aMTOC-nucleated microtubule asters (nine cells) moved inwards over 10 h in a microtubule-dependent manner (Fig. 7a, Supplementary Fig. 7e–g and Supplementary Movie 14). Dynein inhibition also blocked the inwards-directed aMTOC movement, and even caused aMTOCs to drift outwards (Supplementary Fig. 7h, i). Moreover, the microtubule network was much less dynamic in 4/4 zygotes microinjected with the p150-CC1 peptide (Supplementary Fig. 7j and Supplementary Movie 16). However, the inwards-directed movement of the male and female pronucleus could occur independently of aMTOCs though, as their removal by Trim-away of Pericentrin^45,46 (Supplementary Fig. 8a–d) did not affect pronuclear migration (Supplementary Fig. 8e–h) or the formation of the microtubule network (Supplementary Fig. 8c, i). Together, these data establish that the second, slow phase of migration is driven by microtubules and dynein.

Microtubule network dynamics during the second phase rely on an intact actin cytoskeleton

This finding might be surprising given that current models for pronuclear migration are focused on actin instead of microtubules^5,25,26. The importance of actin in pronuclear migration has been highlighted by work in which zygotes were treated with the actin depolymerizing drug cytochalasin D^5,25,27. Indeed, cytochalasin D completely blocks pronuclear migration, as confirmed by our experiments (Supplementary Fig. 9a–f). As we outlined above, local actin polymerization in the region of the fertilization cone is essential for the initial fast mode of pronuclear migration, and it is clear why depolymerizing actin disrupts this initial mode. But why is the second, slow phase also disrupted by cytochalasin D, if it is microtubule dependent?

We first tested if cytochalasin D might prevent an early essential step in pronuclear assembly or migration but could in fact be dispensable for pronuclear migration in the second phase. To this end, we added cytochalasin D acutely, after the pronuclei had formed and started to migrate. Cytochalasin D still prevented pronuclear migration though (Fig. 8a–e), suggesting that actin does indeed play an essential role during the second slow phase of migration as well.

**Fig. 8: Inwards-directed microtubule-based transport requires actin.**

Which actin structures are required for the second, slow phase of migration? Our data suggested that the cytoplasmic actin network, which forms on Rab11a-vesicles and is Spire- and Formin-2-dependent, is not essential during this phase (Figs. 3a–e and 6a–e). Zygotes also contain another prominent actin structure that was disrupted by cytochalasin D but not by the FH2 construct—the actin cortex (Fig. 8f and Supplementary Figs. 9g and 6c, d). There is indeed evidence that the actin cortex promotes pronuclear migration in other systems. In worm zygotes, pronuclear migration and spindle centration are thought to rely on pushing of microtubules against the cell cortex^20,47,48. Disrupting actin reduces the efficiency of pushing^{20,49,50,51,52}.

Could pushing against the actin cortex also be important in mouse zygotes? High-resolution imaging of microtubules in live mouse zygotes revealed that microtubules continued to grow as they encountered the cell surface (Fig. 8g and Supplementary Movie 17). Upon hitting the surface, they bent (yellow and green arrowhead, Fig. 8g), or the microtubule end pointing towards the centre of the zygote was pushed inwards⁵³ (magenta and blue arrowheads, Fig. 8g). We also found that microtubule asters nucleated at aMTOCs had microtubules that reached all the way to the cortex and grew against the cortex as the aster moved inwards, consistent with a pushing-based mechanism (Supplementary Fig. 9h and Supplementary Movie 18).

If microtubules, pronuclei and aMTOCs rely on pushing against the cortex to move inwards, disrupting cortical actin should interfere with their movement. To test this possibility, we imaged microtubules and aMTOCs in the presence and absence of cytochalasin D. Strikingly, the microtubule network was static in 7/7 cytochalasin D-treated zygotes (Fig. 8h and Supplementary Movie 19). Moreover, microtubule asters and aMTOCs failed to migrate inwards upon actin depolymerization (Fig. 8h, i and Supplementary Fig. 9j), just like pronuclei (Fig. 8a–c and Supplementary Fig. 9a–c). Taken together, our data show that an intact actin cytoskeleton is essential for physiological microtubule network dynamics as required for the second microtubule-dependent phase of pronuclear migration.

Our work establishes that two partially redundant mechanisms drive pronuclear migration in the mouse zygote: a mechanism that involves the flattening of the fertilization cone and relies on local actin polymerization by Spire and Formin-2 that acts in the periphery of the zygote, and a microtubule-dependent mechanism that acts in the central region of the zygote. Inhibition of either mechanism alone does not entirely prevent pronuclear migration or the unification of the parental genomes (Figs. 3a–f, 6a–f, 7b–g and Supplementary Table 1, column 8), suggesting that both mechanisms act in a redundant manner to promote pronuclear migration. In further support of redundance, the simultaneous inhibition of the microtubule-dependent pathway (by nocodazole) and the Rab11a-Spire-Fmn2-dependent pathway (by Rab11a^S25N) (Fig. 9a, Supplementary Fig. 10a, b and Supplementary Movie 20) prevented pronuclear migration in 11/27 zygotes (Fig. 9a, bottom panel) and led to an additional significant increase in the distance of the pronuclei from the zygote’s centre at NEBD (by 120 or 80% in comparison to the controls, in male and female pronuclei, respectively (Fig. 9b, c)), despite compression of 10/27 zygotes which caused a decrease in the apparent distance of the pronuclei from the cell centre (Fig. 9a, bottom panel). Simultaneous inhibition of both pathways with cytochalasin D also caused the pronuclei to stay at a large distance from each other at NEBD (Fig. 9d and Supplementary Fig. 9f) and lead to the formation of two entirely separate spindles (Supplementary Table 1, column 8). Together, these data establish that the two pathways that drive pronuclear migration in mouse zygotes are partially redundant.

**Fig. 9: Local actin nucleation by Spire and Formin-2 and microtubule-based transport drive pronuclear migration in a partially redundant manner.**

Discussion

In this study, we established methods to follow and experimentally probe the entire process of pronuclear migration in mammalian zygotes. Using these methods, we demonstrate that pronuclear migration is driven by two partially redundant mechanisms and define the molecular basis for these two mechanisms (Fig. 10).

**Fig. 10: Model for pronuclear migration in mouse zygotes.**

The first mechanism is required to launch the pronuclei away from the cell cortex. This mechanism is most important for the male pronucleus because it forms within the fertilization cone, from where it is rapidly accelerated inwards by the flattening fertilization cone. The inwards-directed movement is enhanced by pushing from behind the male pronucleus and depends on local actin nucleation by Spire and Formin-2, which become enriched in the fertilization cone in a Rab11a-dependent manner. This mechanism is highly effective in launching the male pronucleus away from the zygote’s surface, as illustrated by the fact that enhancing actin nucleation from the surface can push pronuclei even beyond the zygote’s centre, into the opposite hemisphere of the zygote (Supplementary Movie 10). The same mechanism accelerates the female pronucleus inwards, but to a lesser degree as the female pronucleus normally forms further away from the cell surface.

The second mechanism is required to actually centre the pronuclei. In this mechanism, microtubules and dynein move the pronuclei inwards until they meet in the zygote’s centre. This mechanism likely relies on an intact actin cortex, explaining why depolymerization of actin with drugs like cytochalasin D fully blocks pronuclear migration^5,25,26,27.

Why are zygotes employing flattening of the fertilization cone and local actin nucleation by Spire and Formin-2 in addition to the more classical microtubule-dependent mechanism to drive pronuclear migration? There are several possible answers to this question.

First, it may make the mechanism of pronuclear migration more robust and increase the chance that both pronuclei are in immediate proximity of each other when they disassemble at mitotic entry. Our data establish that actin and microtubules act in concert to unite the paternal and maternal chromosomes in mouse zygotes. When microtubules are depolymerized or dynein is inhibited, pronuclei still move inwards, albeit at a slower speed, and are positioned significantly further apart at NEBD (Fig. 7g, m). This suggests that the Rab11a-Spire-Formin-2 mechanism can at least partially rescue pronuclear migration in the absence of microtubules. Conversely, microtubules are able to move pronuclei towards the zygote centre when the Rab11a-Spire-Formin-2 pathway is disrupted. The unification of the parental genomes hence appears to be doubly protected, by the Rab11a-Spire-Formin-2-dependent pathway that acts predominantly in the periphery, and the dynein- and microtubule-dependent mechanism throughout the central region of the zygote.

Interestingly, our data suggest that pronuclei always break down at roughly the same time after their formation (Supplementary Figs. 3k and 7k), independently of whether they successfully migrated inwards (Figs. 3f and 7g). There hence appears to be no checkpoint that monitors pronuclear position, which might be one of the reasons why this process is doubly protected.

Second, the flattening of the fertilization cone and the Rab11a-Spire-Formin-2 mechanism may help to initiate the centration of the pronuclei, while the microtubule system is reorganizing from a meiotic-like state, characterized by the presence of the spindle remnant at the end of meiosis II, to an interphase-like state, characterized by the presence of a microtubule network. The Rab11a-Spire-Formin-2 mechanism could already start to centre the pronuclei, while microtubules are still being reorganized, and thereby accelerate the unification of the parental genomes.

Third, moving the pronuclei away from the cortex before the interphase network is assembled may even be a prerequisite for efficient microtubule pushing and centration of pronuclei. In order for microtubules to push the pronuclei inwards they need to form between the pronucleus and the cell cortex. The initial actin-dependent propulsion of pronuclei may therefore help to generate the space that is required for efficient microtubule pushing.

Finally, it may help to synchronize the movement of the male and female pronucleus. Our data establish that the female pronucleus forms further inwards than the male pronucleus. Moving the male pronucleus rapidly inwards allows for it to catch up, and to reach the zygote’s centre as efficiently as the female pronucleus, as illustrated by Fig. 1b. In support of this notion, we observed that at NEBD the distance of male pronuclei to the cell centre was more severely increased (by 30%) than that of the female pronucleus (by 10%), when the fast actin-dependent launching mechanism of male pronuclei was disabled by expression of dominant-negative Rab11a^S25N (Fig. 3f).

It will be interesting to investigate if pronuclei are also rapidly pushed inwards by flattening of the fertilization cone and local actin-dependent pushing from the surface in other systems.

Generally, microtubules are thought to be the drivers of pronuclear centration. Yet, blocking actin nucleation delays male pronuclear migration in worm zygotes²⁰. An initial, rapid actin-dependent migratory phase that accelerates pronuclei away from the surface may hence be conserved beyond mouse zygotes.

Defects during pronuclear migration lead to the failure of spindle alignment and impair embryonic development⁴. Our study defines two mechanisms that act in concert to achieve this vital task: a Rab11a-, Spire- and Formin-2-dependent mechanism that involves flattening of the fertilization cone and pushes the male pronucleus away from the cell surface by local actin nucleation, and a microtubule–dynein-dependent mechanism that acts throughout the centre to move both pronuclei inwards.

Methods

Preparation, culture and in vitro fertilization of mouse eggs

The maintenance and handling of all mice were performed in the MPI-BPC animal facility according to international animal welfare rules (Federation for Laboratory Animal Science Associations guidelines and recommendations). Requirements of formal control of the German national authorities and funding organizations were satisfied, and the study received approval by the Niedersächsisches Landesamt für Verbraucherschutz und Lebensmittelsicherheit (LAVES). 8–12-week-old C57BL/6J × CBA/CaOlaHsd F1 females (housing conditions: 21 °C ambient temperature, 52–55% humidity and a 14-h light/10-h dark cycle) were superovulated by injection of 5 or 7.5 IU pregnant mare serum gonadotropin (THP Medical Products, #hor-272-a) followed 48 h later by injection of 5 or 7.5 IU human chorionic gonadotropin (Intervet, Ovogest^® 1000). Thirteen hours later, meiosis II arrested eggs were collected from oviducts, cultured and microinjected in HEPES-buffered MEM-α medium (Gibco, #12000-014) supplemented with 5% foetal bovine serum (FBS, Gibco, #16000-044), 4 mM NaHCO₃ (Sigma), 20 mM HEPES (Gibco) and 0.075 g/l potassium penicillin-G (Sigma) and 0.05 g/l streptomycin sulfate (Sigma). Microinjections of eggs were performed as previously described for prophase-arrested mouse oocytes^29,54, but using thinner needles to penetrate the egg membrane more easily without using a piezo⁵⁵ yet with high survival rates (83 ± 7%). Briefly, eggs were microinjected in homemade microinjection chambers using mercury-filled needles for quantitative microinjections. The homemade microinjection chambers were assembled by sticking a small rectangular piece of a coverslip on top of a standard size coverslip with a double-sided sticky tape, as previously described²⁹. In this way, a 100 µm height injection shelf was generated, into which the oocytes were loaded during microinjection. The injected volumes ranged between 5 and 9 pl (3–5% of the egg volume). Concentrations of microinjected mRNAs were calculated by dividing the total volume of injected mRNA by the total volume of the egg (≈200 pl) and are listed here: 5–10 µg/ml MyrGFP, 0.2–0.5 µg/ml H2B-mCherry, 20 µg/ml EGFP-LaminB1, 10 µg/ml Rab11a constructs (SNAP-Rab11a, SNAP-Rab11a^S25N and mScarlet-Rab11a), 5 µg/ml Spire2 constructs for mild overexpression (mClover3-Spire2, SNAP-Spire2, mScarlet-Spire2) or 25 µg/ml EGFP-Spire2 for high overexpression, 90 µg/ml Formin-2-GFP, 80 µg/ml GFP-UtrCH, 60 µg/ml EMTB-mClover, 20 µg/ml Cep192-mScarlet and 6 µg/ml miRFP670-Trim21.

For TRIM-away of Pericentrin, the antibodies mouse IgG (Sigma, #12-371) and anti-pericentrin (BD Biosciences, #611815) were concentrated in PBS at 14,000 × g using Amicon^® Ultra centrifugal filters (Millipore, #UFC51024)⁴⁶. Prior to microinjection, antibodies were diluted to 1 mg/ml in PBS containing a final concentration of 0.03% NP40. Antibodies were microinjected into eggs to a final concentration of 20 µg/ml simultaneously with mRNAs.

After microinjection, eggs were washed into CO₂-dependent MEM-α medium supplemented with 5% FBS, 26 mM NaHCO₃ and 0.075 g/l potassium penicillin-G (Sigma) and 0.05 g/l streptomycin sulfate (Sigma) and cultured at 37 °C in 5% CO₂ for 3–4 h to allow recombinant protein expression. Sperm was collected from >12-week-old C57BL/6J × CBA/CaOlaHsd males and capacitated for 2 h in HTF medium (Millipore, #MR-070-D) supplemented with 2 mM Hypotaurine (Sigma, H1384). Subsequently, IVF of eggs was performed in a 40 µl droplet of HTF medium by adding 2–5 µl of capacitated sperm. After 2 h of IVF, zygotes were washed into HEPES-buffered M2 medium and immediately prepared for live-cell microscopy (start of acquisition within 1–1.5 h post IVF to capture pronuclear formation (Supplementary Fig. 1l) or immunofluorescence).

We confirmed that microinjection into eggs prior to IVF did not prevent efficient fertilization (Supplementary Fig. 11a, c) and supported healthy early development (Supplementary Fig. 11b, d, e).

Preparation of zygotes for live microscopy or immunofluorescence

For drug additions, zygotes were washed into M2 medium containing drugs immediately after IVF. Nocodazole was added just prior to pronuclear formation (about 1 h after IVF) to prevent dispersion of maternal chromosomes. Zygotes were treated with the following drugs: 5 µg/ml cytochalasin D (Sigma, #C8273, dissolved in DMSO at 5 mg/ml), 1 or 10 µM nocodazole (Sigma, #M1404, dissolved in DMSO at 10 mM freshly prepared) and 4 µg/ml BFA (Sigma, #B6542, dissolved in Methanol at 1 mg/ml). For acute cytochalasin D treatment, zygotes were first imaged in M2 medium for 2–2.5 h before washed into M2 medium containing 5 µg/ml cytochalasin D.

For dynein inhibition experiments, zygotes were microinjected with 33 mg/ml purified p150-CC1-6xHis (41 kDa) or MBP-6xHis (43 kDa) in PBS containing 0.03% NP40 and 5 µg/ml Dextran Alexa Flour 680 (Invitrogen, D34680) to a final concentration of 30 µM protein per zygote. Alexa Fluor 680-labelled Dextran was used to verify that protein has entered zygotes. Confirming activity of p150-CC1, meiotic spindles in eggs become abnormally elongated and onset of NEBD was significantly delayed.

For Formin-2 inhibition, zygotes were microinjected with 5–6 mg/ml purified FH2-6xHis (51 kDa) or MBP-6xHis (43 kDa) in buffer (20 mM potassium phosphate pH 7.4, 300 mM NaCl, 0.1 mM EDTA and 5% sucrose) containing 0.03% NP40 and 5 µg/ml Dextran Alexa Fluor 680 to a final concentration of 3 µM protein per zygote.

To stain DNA in live cells, zygotes were washed and imaged in M2 medium containing 250 nM SiR-DNA (Spirochrome, dissolved in DMSO at 1 mM).

Live-cell confocal and super-resolution microscopy

Images were acquired with Zeiss LSM800 and LSM880 confocal laser-scanning microscopes at 37 °C equipped with a ×40 C-Apochromat 1.2 NA water-immersion objective and controlled by the software ZEN Black 2.1 (LSM880) or ZEN Blue 2.3 (LSM800) with multi-position mode. For live imaging, zygotes were cultured in drops of M2 medium with or without drugs under mineral oil in MatTek-35 mm glass bottom dishes. Super-resolution time-lapse images of zygotes expressing EMTB-mClover or GFP-UtrCH were acquired using the Airyscan module on the Zeiss LSM800 microscope and processed post-acquisition using ZEN Blue.

For pronuclear migration, images were typically acquired at a temporal resolution of 15 min covering an 81 µm range (every 3 µm) with an optical slice thickness of 3 µm for a total duration of 18 h. mScarlet-Rab11a images were acquired covering an 81 µm range (every 1.5 µm) with an optical slice thickness of 2.5 µm. To monitor localization of Rab11a, Spire2 and Formin-2 at the fertilization cap, images were acquired at a temporal resolution of 3 min covering a 20-µm-thick subvolume centred around the pronuclei (every 1.5 µm) with an optical slice thickness of 2 µm for a total duration of 3–4 h. For aMTOC tracking, images were acquired at a temporal resolution of 10 min covering an 81 µm range (every 2.5 µm) with an optical slice thickness of 2 µm for a total duration of 18 h. GFP-UtrCH airyscan images were acquired at a temporal resolution of ~3 or ~30 s at the central plane of the pronucleus. EMTB-mClover airyscan images were acquired at a temporal resolution of either 15 min covering a 40-µm-thick subvolume centred around the middle slice of the cell at 4-µm intervals, or 3 or 5 min covering 3 µm at 1–1.5-µm intervals. Settings of the 488, 561 and 633 nm lasers were similar to verified live imaging conditions for mouse embryos in a previous report⁴² and did not exceed 0.2% laser intensity to minimize light-induced phototoxic stress.

Laser ablation

Zygotes were imaged in M2 medium containing 250 nM SiR-DNA (Spirochrome, dissolved in DMSO at 1 mM) and 1X CellMask Green (Invitrogen C37608, dissolved in DMSO) at 20-s intervals prior to and after laser ablation. Once male pronuclei had moved roughly 1 µm away from the cell surface, a region of 8 × 0.7 × 7 µm (eight sections at 1 µm) was two-photon laser ablated at 790 nm with 18% laser power. The ablated region was manually defined to perform eight cuts in z using the LSM980 confocal laser-scanning microscope. The ablated regions were either located between cell surface and male pronucleus (behind the male pronucleus) or in the cytoplasm in front of the male pronucleus. After ablation, imaging of zygotes was restarted within 70–100 s.

Analysis of cytoplasmic streaming

Zygotes and their pronuclei (SiR-DNA) were imaged in 3–10-s intervals. Transmission images were used to detect and analyse cytoplasmic streaming using the ImageJ PIV Plugin by Qingzong Tseng⁵⁶. The following parameters were used: piv1 = 128, sw1 = 256, vs1 = 64, piv2 = 64, sw2 = 128, vs2 = 32, piv3 = 0, sw3 = 0, vs3 = 0. Single z-slices were analyzed using an in-house ImageJ macro to process full timeseries in one go. The results were averaged over 5-min periods to highlight relevant flows using Python 3.7. Plotting of the vectors was conducted in Matlab 2018b due to a bug affecting short vectors in the original plugin. Vectors outside of the cell were removed.

Immunofluorescence microscopy

Zygotes were fixed at time points indicated in hours post insemination (hpi, start of IVF). We experimentally determined that pronuclear formation occurs 3.9 ± 0.5 hpi (Supplementary Fig. 1l), which was used as a reference time point for immunofluorescence studies to fix zygotes close to the time of pronuclear formation in Supplementary Figs. 1h, 2b, g, i and 4h. Zygotes were treated for 5–10 s in 2.5% Triton X-100 diluted in water before they were fixed for 25 min at 37 °C with 100 mM HEPES, 50 mM EGTA, 10 mM MgSO₄, 2% formaldehyde and 0.4% Triton x-100 and extracted in PBS supplemented with 0.5% Triton X-100 at 4 °C overnight. Antibody incubation, F-actin and DNA staining were performed in PBS, 3% BSA and 0.5% Triton X-100 for 2–4 h at room temperature, except Rab11a staining was performed in PBS, 12% BSA and 0.5% Triton X-100 overnight at 4 °C. Primary antibodies used in this study: rat anti-α-tubulin (Serotec, MCA78G, 1:500), rabbit anti-Rab11a (Abcam, ab65200, 1:2000) and mouse anti-pericentrin (BD Biosciences, #611815, 1:1000). Alexa Fluor-488-labelled anti-mouse, Alexa Fluor-546-labelled anti-rabbit and Alexa Fluor-647-labelled anti-rat (Molecular Probes, 1:400) were used as secondary antibodies. DNA was stained with 25 µg/ml Hoechst 33342 (Molecular Probes); F-actin was stained with Alexa Fluor-488-phalloidin (Molecular Probes, 1:20). Images were acquired with LSM800 and LSM880 confocal microscopes equipped with a ×40 C-Apochromat 1.2 NA water-immersion objective at room temperature. Immunofluorescence images were typically acquired at an optical slice thickness of 2 µm confocal sections covering a subvolume or the entire oocyte at 1-µm intervals if not indicated otherwise. For visualizing F-actin, single optical sections of F-actin in the equatorial region of zygotes were acquired using the Airyscan module on the Zeiss LSM800 microscope. Images in control and perturbed situations were acquired with identical imaging conditions. Care was taken that images were not saturated during acquisition (except for co-imaging of the meiosis II spindle and the interphase microtubule network).

Image analysis and statistics

Microscopic images were processed in ZEN Blue 2.3 and ImageJ (version 1.52p). 3D reconstructions and tracking of pronuclei and aMTOCs were performed using Imaris 9.2 and 9.3 (Bitplane). Cells were reconstructed as using the ImarisCell Module based on plasma membrane labelling. Pronuclei were tracked automatically using the Imaris tracking function and verified manually. The distance between cell centre and pronuclei centroids was calculated from the centres of homogenous mass in Microsoft Excel for every time point as illustrated in Supplementary Fig. 1d. Average velocities were obtained by averaging instantaneous velocities within a specified time-interval for each pronucleus. aMTOCs were detected using the Imaris spot detection and tracked manually.

Pronuclear migration was defined as the period from pronuclear formation (time point 0 h) until 15 min prior to onset of chromosome condensation at NEBD. Pronuclear formation, as detected by the emergence of the nuclear envelope labelled by GFP-LaminB1, coincided with the onset of chromosome decondensation characterized by the sudden volume increase of the H2B-mCherry-labelled DNA mass (Supplementary Fig. 1a–c), which was thereafter used as determinant of pronuclear formation. In Supplementary Fig. 1b, c, pronuclear volumes were normalized to the respective volume at the end of pronuclear migration. Plots for pronuclear migration were aligned relative to pronuclear formation, the time we referred to as time 0 h. Images are displayed as single sections for the cell surface (MyrGFP, equatorial plane) and as a maximum intensity projection of the entire cell for chromosomes (H2B-mCherry) in Figs. 1a, 2a, 3a, 5a, 6a, 7b, h, 8a and 9a. Distinct phases of migration for male and female pronuclei were defined based on typical pronuclear migratory behaviour with the following speeds: fast (>0.1 µm/min), medium (0.05–0.1 µm/min) and slow (<0.05 µm) (Fig. 1a–c).

Only zygotes were analyzed whose pronuclei had not yet formed at the start of acquisition and that progressed to the two-cell stage during the first 18 h of acquisition as these were considered healthy, except for in the cases of treatment with cytochalasin D or p150-CC1, which both caused a delay of NEBD. Furthermore, zygotes were excluded that contained more or less than two pronuclei (parthenogenesis, polyspermy) or male pronuclei forming at a distance from the cell surface (in untreated zygotes 3/56 zygotes)and that exhibited cortical deformations after microinjection of SNAP-Rab11a^S25N, precluding an accurate determination of the centre of the zygote as required for pronuclear tracking (25/65 zygotes).

To confirm reproducibility between repetitions and experiments, mean distances over time from independent repetitions were plotted (Supplementary Fig. 12a–q).

Measurements of average velocities suggested that different manipulations affect different portions of pronuclear migration. To confirm the results of our approach, we also used curve fitting with a piecewise linear function to estimate slopes in different portions of the curve after concatenation of all data points (OriginLab). The function of the piecewise linear function can be expressed as follows:

$$y = \left\{ {\begin{array}{*{20}{c}} {\frac{{{y}_1\left( {{x}_3\, -\, x} \right)\, +\, {y}_3\left( {x\, -\, {x}_1} \right)}}{{{x}_3 - {x}_1}},{\rm{if}}\,x\, <\, x_3} \\ {\frac{{{y}_3\left( {{x}_3\, -\, x} \right)\, +\, {y}_3\left( {x\, -\, {x}_1} \right)}}{{{x}_2\, -\, {x}_3}},{\rm{if}}\,x \ge x_3} \end{array}} \right.$$

To ensure to stay within the time window, in which pronuclei move freely, we performed the fitting between 0 and 5 h, whereby x₁ = 0 and x₂ = 5 were fixed and x₃ is the intersection time point. Results of our analysis are summarized in Supplementary Fig. 13a–q and Supplementary Table 2 and confirm our previous approach.

In Supplementary Fig. 8b, d, microtubules asters/aMTOCs were manually counted. For aMTOC tracking, at least six aMTOCs per zygote were tracked using Imaris from the onset to the end of pronuclear migration. Microtubule density in Supplementary Fig. 7a was measured as an average of four regions per cell over time from a maximum projection after drift correction in Imaris and then normalized to pronuclear formation (0 h). Oil droplets were tracked manually using ImageJ (TrackMate Plugin). Temporal-colour coded images in Fig. 8h and Supplementary Fig. 9h were generated in ImageJ.

Average velocities of pronuclei and aMTOCs towards the cell centre were calculated from instantaneous velocities, which were averaged for each pronucleus/aMTOC within the specified interval and plotted as individual dots in the average velocity-over-time-scatter plots.

In live-cell experiments of zygotes expressing mScarlet-Rab11a, vesicles were counted manually in the lower half of the zygote. In immunofluorescence with anti-Rab11a, vesicles were counted automatically in a segment of 25 × 25 × 16.5 µm³ around the equatorial plane of the cell excluding regions overlapping with pronuclear positions using Imaris spot detection (different spot sizes with region growing and an estimated diameter of 0.3 µm, filtered volume of spots > 0.0283 µm³). In immunofluorescence, spots were categorized depending on their volume. Spots above 0.5 µm³ were clearly discernible from background and considered as vesicles. Spots below 0.5 µm³ were difficult to discriminate from background signal and were therefore excluded.

To quantify the density of the cytoplasmic actin network, the mean intensity of Alexa Fluor-488-phalloidin was measured in the cytoplasm (average from 6 to 12 regions per cell) and in a region outside the zygote for background subtraction using ImageJ. The disruption of the actin cortex was determined by eye, and the number of holes in the actin cortex was counted manually (areas of >1 µm in the cortex that were devoid of phalloidin).

Graphs and plots were created using GraphPad Prism 5.0.3 and OriginPro 9.1 (OriginLab). Statistical plots show mean (horizontal middle lines), and standard deviation (whiskers). The box plot shows median (line), mean (small square), 5th, 95th (whiskers) and 25th and 75th percentile (boxes enclosing 50% of the data). Mean, standard deviation and statistical significance based on two-tailed Student’s t test (assuming unequal variances) were calculated in GraphPad Prism. p values are designated as * (p < 0.05), ** (p < 0.005) and *** (p < 0.0005). Nonsignificant values are indicated as n.s. Experiments were performed at least three times. Figures were assembled in Adobe Illustrator.

Expression constructs and mRNA synthesis

To generate a membrane-anchored GFP, a sequence encoding for a myristoylation signal (MGCIKSKRKDNLNDDEAA⁵⁷) was inserted N-terminally in frame with mEGFP by PCR using primers forward (GCTAGCATGGGCTGCATCAAGAGCAAGCGCAAGGACAACCTGAACGACGACGAGGCCGCCATGGTGAGCAAGGGCGAGGAGC) and reverse (GAATTCGAAGCTTGAGCTCG) from pmEGFP-C1 (Addgene, #227464) as a template. The NheI-EcoRI flanked fragment was subsequently cloned into pGEMHE to generate pGEMHE-MyrGFP. To generate pGEMHE-mScarlet-Rab11a, mScarlet was first cloned into the NheI-XhoI of pGEMHE by PCR with pmScarlet-C1 (Addgene #85042) as a template. Subsequently, the Rab11a sequence from pGEMHE-SNAP-Rab11 was inserted into the HindIII site via Gibson Assembly (New England Biolabs). pGEMHE-mClover-Spire2 was obtained from pGEMHE-mCherry-Spire2³⁵ by switching mCherry with mClover via NheI-XhoI by PCR from pKanCMV-mClover3-18aa-TUBA (gift from Michael Lin, Addgene #74253). To generate pGEMHE-SNAP-Spire2 and pGEMHE-mScarlet-Spire2, SNAPf from pSNAPf (New England Biolabs) or mScarlet was cloned into the NheI-BamHI site of pGEMHE-mClover3-Spire2. For pGEMHE-Fmn2-EGFP, Fmn2-EGFP was cloned into BglII-NotI site of pGEMHE from pCS2-Fmn2-EGFP⁵⁸. For pGEMHE-EGFP-UtrCH, EGFP-UtrCH was cloned into the AgeI-NotI site of pGEMHE from pCS2-EGFP-UtrCH⁵⁹. For pGEMHE-EMTB-mClover containing the microtubule binding domain of MAP7/Ensconsin⁶⁰, the 3xEGFP sequence in pGEMHE-EMTB-3xEGFP (gift from Jan Ellenberg) was switched with the mClover sequence from pTALYM3B15 (Addgene #47878) via BamHI-NotI site. The previously published sequence EGFP-LaminB1⁶¹ was inserted into pGEMHE to generate pGEMHE-EGFP-LaminB1. For pGEMHE-miRFP670-Trim21, the EGFP sequence in pGEMHE-EGFP-Trim21⁴⁶ was replaced by the miRFP670 sequence from pmiRFP670-N1 (gift from Vladislav Verkhusha, Addgene # 79987) via AgeI-BglII site. Expression constructs pGEMHE-H2B-mCherry, pGEMHE-H2B-mEGFP⁴⁶, pGEMHE-SNAP-Rab11a, pGEMHE-SNAP-Rab11a^S25N ⁶², pGEMHE-GFP-Spire2³⁵ and pGEMHE-Cep192-mScarlet⁶³ were generated in previous studies as indicated. All primers used in this study to generate the above-named plasmids are listed in Supplementary Table 3.

For in vitro mRNA transcription, pGEMHE plasmids were linearized with AscI, NheI or SphI and capped mRNA was synthesized with T7 polymerase (Ambion mMessage mMachine T7 Kit) according to manufacturer instructions. mRNA concentrations were determined using NanoDrop.

Protein expression and purification of His-tagged proteins

All constructs were expressed in E. coli BL21-CodonPlus (DE3) (Agilent Technologies #230245) from pET-28a with a C-terminal His-tag. pET-28a-p150-CC1-His (gift from Jan Ellenberg) was generated in a previous study⁴⁴. To generate pET-28a-FH2-His and as a negative control pET-28a-MBP-His, p150-CC1 was replaced by the coding sequences of FH2 and MBP amplified from pGEX-6P1-FH2 (gift from Marie France-Carlier⁴⁰) and pET-21a-MBP-His (Addgene # 38006) and inserted via XbaI-XhoI.

For p150-CC1-6xHis purification, bacterial cultures were induced with 1 mM isopropyl-β-d-thiogalactoside at 37 °C overnight. Bacteria were lysed in binding buffer (50 mM NaPO₄ buffer, pH 8.0; 300 mM NaCl; 2 mM β-mercaptoethanol; 15% glycerol) by sonication. p150-CC1-6xHis and MBP-6xHis were bound to Ni-NTA resin (Qiagen) and eluted with 250 mM imidazole in binding buffer. Buffer exchange to PBS was carried out using Slide-A-Lyzer Dialysis Cassettes (ThermoFisher Scientific). Proteins were concentrated using Amicon Ultra centrifugal filter units (Millipore) at 14,000 × g, aliquoted, flash frozen in liquid nitrogen and stored at −80 °C. All purification was carried out at 4 °C to minimize protein aggregation. Proteins were analyzed by SDS-polyacrylamide gel electrophoresis and Coomassie staining (Biorad) and protein concentration was determined using BSA (NEB) as a reference (Supplementary Fig. 7c, d).

FH2-6xHis and MBP-6xHis were purified using Ni-NTA as described for p150-CC1-6xHis, but with the following adaptations: bacterial cultures were induced with 1 mM isopropyl-β-d-thiogalactoside at 16 °C overnight. Bacteria were lysed in binding buffer (20 mM KPO₄ buffer, pH 7.4; 30 mM NaCl, 3 mM DTT, 5% sucrose and 0.1 mM EDTA). After elution, buffer was exchanged to binding buffer without DTT (Supplementary Fig. 6a, b).

A presentation of the uncropped full scan gels is presented in the Source Data file provided with this paper.

Reporting summary

Further information on research design is available in the Nature Research Reporting Summary linked to this article.

Data availability

All relevant and raw data supporting the finding of this study are available from the corresponding author upon request for the following figures: Figs. 1b, 2c, 3b, c, 5c, 6b, c, 7c, d, i, j, 8b, c, i, S1f, j, S3b, c, g, j, S5c, e, S6c, e, S7a, g, i, S8b, e, f, S9b, c, i, S10a, b. The primary microscopy data were not uploaded to a data repository due to their large size, but are available from the authors upon request. Source data are provided with this paper.

References

Clift, D. & Schuh, M. Restarting life: fertilization and the transition from meiosis to mitosis. Nat. Rev. Mol. Cell Biol. 14, 549–562 (2013).
Article CAS PubMed PubMed Central Google Scholar
Bianchi, E. & Wright, G. J. Sperm meets egg: the genetics of mammalian fertilization. Annu. Rev. Genet. 50, 93–111 (2016).
Article CAS PubMed Google Scholar
Trebichalska, Z. & Holubcova, Z. Perfect date-the review of current research into molecular bases of mammalian fertilization. J. Assist. Reprod. Genet. 37, 243–256 (2020).
Article PubMed PubMed Central Google Scholar
Reichmann, J. et al. Dual-spindle formation in zygotes keeps parental genomes apart in early mammalian embryos. Science 361, 189–193 (2018).
Article ADS CAS PubMed Google Scholar
Chaigne, A. et al. F-actin mechanics control spindle centring in the mouse zygote. Nat. Commun. 7, 10253 (2016).
Article ADS CAS PubMed PubMed Central Google Scholar
Simerly, C. et al. The paternal inheritance of the centrosome, the cell’s microtubule-organizing center, in humans, and the implications for infertility. Nat. Med. 1, 47–52 (1995).
Article PubMed Google Scholar
Bestor, T. H. & Schatten, G. Anti-tubulin immunofluorescence microscopy of microtubules present during the pronuclear movement of sea urchin fertilization. Dev. Biol. 88, 80–91 (1981).
Article CAS PubMed Google Scholar
Reinsch, S. & Gonczy, P. Mechanisms of nuclear positioning. J. Cell Sci. 111(Pt 16), 2283–2295 (1998).
CAS PubMed Google Scholar
Reinsch, S. & Karsenti, E. Movement of nuclei along microtubules in Xenopus egg extracts. Curr. Biol. 7, 211–214 (1997).
Article CAS PubMed Google Scholar
Riparbelli, M. G., Callaini, G. & Glover, D. M. Failure of pronuclear migration and repeated divisions of polar body nuclei associated with MTOC defects in polo eggs of Drosophila. J. Cell Sci. 113(Pt 18), 3341–3350 (2000).
CAS PubMed Google Scholar
Schatten, G. The movements and fusion of the pronuclei at fertilization of the sea urchin Lytechinus variegatus: time-lapse video microscopy. J. Morphol. 167, 231–247 (1981).
Article PubMed Google Scholar
Strome, S. & Wood, W. B. Generation of asymmetry and segregation of germ-line granules in early C. elegans embryos. Cell 35, 15–25 (1983).
Article CAS PubMed Google Scholar
Gonczy, P., Pichler, S., Kirkham, M. & Hyman, A. A. Cytoplasmic dynein is required for distinct aspects of MTOC positioning, including centrosome separation, in the one cell stage Caenorhabditis elegans embryo. J. Cell Biol. 147, 135–150 (1999).
Article CAS PubMed PubMed Central Google Scholar
Lindeman, R. E. & Pelegri, F. Localized products of futile cycle/lrmp promote centrosome-nucleus attachment in the zebrafish zygote. Curr. Biol. 22, 843–851 (2012).
Article CAS PubMed PubMed Central Google Scholar
Wuhr, M., Tan, E. S., Parker, S. K., Detrich, H. W. 3rd & Mitchison, T. J. A model for cleavage plane determination in early amphibian and fish embryos. Curr. Biol. 20, 2040–2045 (2010).
Article PubMed PubMed Central CAS Google Scholar
Meaders, J. L. & Burgess, D. R. Microtubule-based mechanisms of pronuclear positioning. Cells 9, 505 (2020). https://doi.org/10.3390/cells9020505.
Kimura, K. & Kimura, A. Intracellular organelles mediate cytoplasmic pulling force for centrosome centration in the Caenorhabditis elegans early embryo. Proc. Natl Acad. Sci. USA 108, 137–142 (2011).
Article ADS CAS PubMed Google Scholar
Shinar, T., Mana, M., Piano, F. & Shelley, M. J. A model of cytoplasmically driven microtubule-based motion in the single-celled Caenorhabditis elegans embryo. Proc. Natl Acad. Sci. USA 108, 10508–10513 (2011).
Article ADS CAS PubMed Google Scholar
Malone, C. J. et al. The C. elegans hook protein, ZYG-12, mediates the essential attachment between the centrosome and nucleus. Cell 115, 825–836 (2003).
Article CAS PubMed Google Scholar
Xiong, H., Mohler, W. A. & Soto, M. C. The branched actin nucleator Arp2/3 promotes nuclear migrations and cell polarity in the C. elegans zygote. Dev. Biol. 357, 356–369 (2011).
Article CAS PubMed PubMed Central Google Scholar
Zhao, T., Graham, O. S., Raposo, A. & St Johnston, D. Growing microtubules push the oocyte nucleus to polarize the Drosophila dorsal-ventral axis. Science 336, 999–1003 (2012).
Article ADS CAS PubMed PubMed Central Google Scholar
Tran, P. T., Marsh, L., Doye, V., Inoue, S. & Chang, F. A mechanism for nuclear positioning in fission yeast based on microtubule pushing. J. Cell Biol. 153, 397–411 (2001).
Article CAS PubMed PubMed Central Google Scholar
Schatten, H., Schatten, G., Mazia, D., Balczon, R. & Simerly, C. Behavior of centrosomes during fertilization and cell division in mouse oocytes and in sea urchin eggs. Proc. Natl Acad. Sci. USA 83, 105–109 (1986).
Article ADS CAS PubMed Google Scholar
Schatten, G., Simerly, C. & Schatten, H. Microtubule configurations during fertilization, mitosis, and early development in the mouse and the requirement for egg microtubule-mediated motility during mammalian fertilization. Proc. Natl Acad. Sci. USA 82, 4152–4156 (1985).
Article ADS CAS PubMed Google Scholar
Chaigne, A., Terret, M. E. & Verlhac, M. H. Asymmetries and symmetries in the mouse oocyte and zygote. Results Probl. Cell Differ. 61, 285–299 (2017).
Article CAS PubMed Google Scholar
Bone, C. R. & Starr, D. A. Nuclear migration events throughout development. J. Cell Sci. 129, 1951–1961 (2016).
Article CAS PubMed PubMed Central Google Scholar
Maro, B., Johnson, M. H., Pickering, S. J. & Flach, G. Changes in actin distribution during fertilization of the mouse egg. J. Embryol. Exp. Morphol. 81, 211–237 (1984).
CAS PubMed Google Scholar
Schatten, G. et al. Latrunculin inhibits the microfilament-mediated processes during fertilization, cleavage and early development in sea urchins and mice. Exp. Cell Res. 166, 191–208 (1986).
Article CAS PubMed Google Scholar
Jaffe, L. A. & Terasaki, M. Quantitative microinjection of oocytes, eggs, and embryos. Methods Cell Biol. 74, 219–242 (2004).
Article PubMed PubMed Central Google Scholar
Almonacid, M. et al. Active diffusion positions the nucleus in mouse oocytes. Nat. Cell Biol. 17, 470–479 (2015).
Article CAS PubMed Google Scholar
Schuh, M. An actin-dependent mechanism for long-range vesicle transport. Nat. Cell Biol. 13, 1431–1436 (2011).
Article CAS PubMed PubMed Central Google Scholar
Holubcova, Z., Howard, G. & Schuh, M. Vesicles modulate an actin network for asymmetric spindle positioning. Nat. Cell Biol. 15, 937–947 (2013).
Article CAS PubMed PubMed Central Google Scholar
Colin, A. et al. Active diffusion in oocytes nonspecifically centers large objects during prophase I and meiosis I. J Cell Biol 219, 1–15 (2020).
Duch, M. et al. Tracking intracellular forces and mechanical property changes in mouse one-cell embryo development. Nat. Mater. 19, 1114–1123 (2020).
Article ADS CAS PubMed Google Scholar
Pfender, S., Kuznetsov, V., Pleiser, S., Kerkhoff, E. & Schuh, M. Spire-type actin nucleators cooperate with Formin-2 to drive asymmetric oocyte division. Curr. Biol. 21, 955–960 (2011).
Article CAS PubMed PubMed Central Google Scholar
Borisy, G. G. & Svitkina, T. M. Actin machinery: pushing the envelope. Curr. Opin. Cell Biol. 12, 104–112 (2000).
Article CAS PubMed Google Scholar
Mogilner, A. On the edge: modeling protrusion. Curr. Opin. Cell Biol. 18, 32–39 (2006).
Article CAS PubMed Google Scholar
Lambrechts, A., Gevaert, K., Cossart, P., Vandekerckhove, J. & Van Troys, M. Listeria comet tails: the actin-based motility machinery at work. Trends Cell Biol. 18, 220–227 (2008).
Article CAS PubMed Google Scholar
Footer, M. J., Kerssemakers, J. W., Theriot, J. A. & Dogterom, M. Direct measurement of force generation by actin filament polymerization using an optical trap. Proc. Natl Acad. Sci. USA 104, 2181–2186 (2007).
Article ADS CAS PubMed Google Scholar
Montaville, P. et al. Spire and Formin 2 synergize and antagonize in regulating actin assembly in meiosis by a ping-pong mechanism. PLoS Biol. 12, e1001795 (2014).
Article PubMed PubMed Central CAS Google Scholar
Mogessie, B. & Schuh, M. Actin protects mammalian eggs against chromosome segregation errors. Science 357, (2017). https://doi.org/10.1126/science.aal1647.
Courtois, A., Schuh, M., Ellenberg, J. & Hiiragi, T. The transition from meiotic to mitotic spindle assembly is gradual during early mammalian development. J. Cell Biol. 198, 357–370 (2012).
Article CAS PubMed PubMed Central Google Scholar
Payne, C., Rawe, V., Ramalho-Santos, J., Simerly, C. & Schatten, G. Preferentially localized dynein and perinuclear dynactin associate with nuclear pore complex proteins to mediate genomic union during mammalian fertilization. J. Cell Sci. 116, 4727–4738 (2003).
Article CAS PubMed Google Scholar
Zhang, Q. et al. Nudel promotes axonal lysosome clearance and endo-lysosome formation via dynein-mediated transport. Traffic 10, 1337–1349 (2009).
Article CAS PubMed Google Scholar
Ma, W. & Viveiros, M. M. Depletion of pericentrin in mouse oocytes disrupts microtubule organizing center function and meiotic spindle organization. Mol. Reprod. Dev. 81, 1019–1029 (2014).
Article CAS PubMed PubMed Central Google Scholar
Clift, D. et al. A method for the acute and rapid degradation of endogenous proteins. Cell 171, 1692–1706.e18 (2017).
Article CAS PubMed PubMed Central Google Scholar
Pecreaux, J. et al. The mitotic spindle in the one-cell C. elegans embryo is positioned with high precision and stability. Biophys. J. 111, 1773–1784 (2016).
Article ADS CAS PubMed PubMed Central Google Scholar
Howard, J. & Garzon-Coral, C. Physical limits on the precision of mitotic spindle positioning by microtubule pushing forces: mechanics of mitotic spindle positioning. Bioessays 39 (2017).
Thery, M. & Bornens, M. Get round and stiff for mitosis. HFSP J. 2, 65–71 (2008).
Article PubMed PubMed Central Google Scholar
Salbreux, G., Charras, G. & Paluch, E. Actin cortex mechanics and cellular morphogenesis. Trends Cell Biol. 22, 536–545 (2012).
Article CAS PubMed Google Scholar
Gilden, J. & Krummel, M. F. Control of cortical rigidity by the cytoskeleton: emerging roles for septins. Cytoskeleton 67, 477–486 (2010).
CAS PubMed Google Scholar
Charras, G. T., Hu, C. K., Coughlin, M. & Mitchison, T. J. Reassembly of contractile actin cortex in cell blebs. J. Cell Biol. 175, 477–490 (2006).
Article CAS PubMed PubMed Central Google Scholar
Vleugel, M., Kok, M. & Dogterom, M. Understanding force-generating microtubule systems through in vitro reconstitution. Cell. Adh. Migr. 10, 475–494 (2016).
Article CAS PubMed PubMed Central Google Scholar
Schuh, M. & Ellenberg, J. Self-organization of MTOCs replaces centrosome function during acentrosomal spindle assembly in live mouse oocytes. Cell 130, 484–498 (2007).
Article CAS PubMed Google Scholar
Yoshida, N. & Perry, A. C. Piezo-actuated mouse intracytoplasmic sperm injection (ICSI). Nat. Protoc. 2, 296–304 (2007).
Article CAS PubMed Google Scholar
Tseng, Q. et al. Spatial organization of the extracellular matrix regulates cell-cell junction positioning. Proc. Natl Acad. Sci. USA 109, 1506–1511 (2012).
Article ADS CAS PubMed Google Scholar
Trichas, G., Begbie, J. & Srinivas, S. Use of the viral 2A peptide for bicistronic expression in transgenic mice. BMC Biol. 6, 40 (2008).
Article PubMed PubMed Central CAS Google Scholar
Leader, B. et al. Formin-2, polyploidy, hypofertility and positioning of the meiotic spindle in mouse oocytes. Nat. Cell Biol. 4, 921–928 (2002).
Article CAS PubMed Google Scholar
Burkel, B. M., von Dassow, G. & Bement, W. M. Versatile fluorescent probes for actin filaments based on the actin-binding domain of utrophin. Cell Motil. Cytoskeleton 64, 822–832 (2007).
Article CAS PubMed PubMed Central Google Scholar
Faire, K. et al. E-MAP-115 (ensconsin) associates dynamically with microtubules in vivo and is not a physiological modulator of microtubule dynamics. J. Cell Sci. 112(Pt 23), 4243–4255 (1999).
CAS PubMed Google Scholar
Daigle, N. et al. Nuclear pore complexes form immobile networks and have a very low turnover in live mammalian cells. J. Cell Biol. 154, 71–84 (2001).
Article CAS PubMed PubMed Central Google Scholar
Cheeseman, L. P., Boulanger, J., Bond, L. M. & Schuh, M. Two pathways regulate cortical granule translocation to prevent polyspermy in mouse oocytes. Nat. Commun. 7, 13726 (2016).
Article ADS CAS PubMed PubMed Central Google Scholar
Clift, D. & Schuh, M. A three-step MTOC fragmentation mechanism facilitates bipolar spindle assembly in mouse oocytes. Nat. Commun. 6, 7217 (2015).
Article ADS PubMed PubMed Central Google Scholar

Download references

Acknowledgements

We thank the staff of the MPI-BPC animal facilities for technical assistance, Katharina Schücker for help with microscopy, Christiane Heuchel, Tina Daniel, Silke Schlott and Kevin Rentsch for assistance with superovulations and members of the Schuh laboratory for discussions. Research was funded by the Max Planck Society and supported by the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation) by a Leibniz Prize to M.S. (SCHU 3047/1-1) and under Germany’s Excellence Strategy—EXC 2067/1-390729940. K.S. was further supported by an EMBO long-term postdoctoral fellowship and a Wellcome Trust and Royal Society Sir Henry Dale Fellowship to B.M. (Grant No. 213470/Z/18/Z). I.J. was supported by a Boehringer Ingelheim Fonds Ph.D. fellowship. Plasmids are available from M.S. under a material transfer agreement with the Max Planck Society.

Funding

Open Access funding enabled and organized by Projekt DEAL.

Author information

Authors and Affiliations

Department of Meiosis, Max Planck Institute for Biophysical Chemistry, Göttingen, Germany
Kathleen Scheffler, Julia Uraji, Ida Jentoft, Tommaso Cavazza, Eike Mönnich & Melina Schuh
School of Biochemistry, University of Bristol, Bristol, UK
Kathleen Scheffler & Binyam Mogessie

Authors

Kathleen Scheffler
View author publications
You can also search for this author in PubMed Google Scholar
Julia Uraji
View author publications
You can also search for this author in PubMed Google Scholar
Ida Jentoft
View author publications
You can also search for this author in PubMed Google Scholar
Tommaso Cavazza
View author publications
You can also search for this author in PubMed Google Scholar
Eike Mönnich
View author publications
You can also search for this author in PubMed Google Scholar
Binyam Mogessie
View author publications
You can also search for this author in PubMed Google Scholar
Melina Schuh
View author publications
You can also search for this author in PubMed Google Scholar

Contributions

K.S. and M.S. conceived the project, and designed experiments and methods for data analysis. J.U., I.J. and T.C. contributed to the design of some experiments and methods for data analysis. K.S. performed and analyzed the experiments in Figs. 1, 2, 4b–f, 5a–d, 7a–g, 8f–h, 9, S1, S3a–e, h–j, S4f–l, S5a, b, S6, S7, S8a–d, I, S9g–h, S10; K.S. and J.U. the experiments in Figs. 3, 6a–f, g, 7h–m, 8i, S2c, S2g, S8e–h, S9a–f; J.U. the experiments in Figs. 4a, 8a–e, S2d–f, i, j, S4d, e; I.J. and T.C. the experiments in Figs. 5e–g, 6h and Supplementary Figs. 3f, g, 5d, e, 11, which were subsequently analyzed by K.S. (Fig. 5e–g and Supplementary Fig. 3f, g) and E.M. (Fig. 5e–g and Supplementary Fig. 5e). E.M. and K.S. performed the curve fitting analysis (Fig. S13). K.S. prepared all figures with input from M.S. and B.M. K.S. and M.S. wrote the paper. All authors discussed the results and edited the paper. M.S. supervised the study.

Corresponding author

Correspondence to Melina Schuh.

Ethics declarations

Competing interests

The authors declare no competing interests.

Additional information

Peer review information Nature Communications thanks the anonymous reviewer(s) for their contribution to the peer review of this work.

Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Supplementary information

Supplementary Information

Description of Additional Supplementary Files

Supplementary Movie 1

Supplementary Movie 2

Supplementary Movie 3

Supplementary Movie 4

Supplementary Movie 5

Supplementary Movie 6

Supplementary Movie 7

Supplementary Movie 8

Supplementary Movie 9

Supplementary Movie 10

Supplementary Movie 11

Supplementary Movie 12

Supplementary Movie 13

Supplementary Movie 14

Supplementary Movie 15

Supplementary Movie 16

Supplementary Movie 17

Supplementary Movie 18

Supplementary Movie 19

Supplementary Movie 20

Reporting Summary

Source data

Source Data

Rights and permissions

Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The images or other third party material in this article are included in the article’s Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this license, visit http://creativecommons.org/licenses/by/4.0/.

Reprints and permissions

About this article

Cite this article

Scheffler, K., Uraji, J., Jentoft, I. et al. Two mechanisms drive pronuclear migration in mouse zygotes. Nat Commun 12, 841 (2021). https://doi.org/10.1038/s41467-021-21020-x

Download citation

Received: 26 October 2019
Accepted: 28 December 2020
Published: 05 February 2021
DOI: https://doi.org/10.1038/s41467-021-21020-x

This article is cited by

Dysfunction of DMT1 and miR-135b in the gut-testis axis in high-fat diet male mice
- Yanru Zhang
- Ruike Ding
- Enqi Liu
Genes & Nutrition (2024)
The evolution of centriole degradation in mouse sperm
- Sushil Khanal
- Ankit Jaiswal
- Tomer Avidor-Reiss
Nature Communications (2024)
Actin-driven chromosome clustering facilitates fast and complete chromosome capture in mammalian oocytes
- Katarina Harasimov
- Julia Uraji
- Melina Schuh
Nature Cell Biology (2023)
Aneuploidy in mammalian oocytes and the impact of maternal ageing
- Chloe Charalambous
- Alexandre Webster
- Melina Schuh
Nature Reviews Molecular Cell Biology (2023)

Comments

By submitting a comment you agree to abide by our Terms and Community Guidelines. If you find something abusive or that does not comply with our terms or guidelines please flag it as inappropriate.