Coordination of Embryogenesis by the Centrosome in Drosophila melanogaster

Blake-Hedges, Caitlyn; Megraw, Timothy L.

doi:10.1007/978-3-030-23173-6_12

Caitlyn Blake-Hedges⁴ &
Timothy L. Megraw⁴

Part of the book series: Results and Problems in Cell Differentiation ((RESULTS,volume 67))

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Abstract

The first 3 h of Drosophila melanogaster embryo development are exemplified by rapid nuclear divisions within a large syncytium, transforming the zygote to the cellular blastoderm after 13 successive cleavage divisions. As the syncytial embryo develops, it relies on centrosomes and cytoskeletal dynamics to transport nuclei, maintain uniform nuclear distribution throughout cleavage cycles, ensure generation of germ cells, and coordinate cellularization. For the sake of this review, we classify six early embryo stages that rely on processes coordinated by the centrosome and its regulation of the cytoskeleton. The first stage features migration of one of the female pronuclei toward the male pronucleus following maturation of the first embryonic centrosomes. Two subsequent stages distribute the nuclei first axially and then radially in the embryo. The remaining three stages involve centrosome-actin dynamics that control cortical plasma membrane morphogenesis. In this review, we highlight the dynamics of the centrosome and its role in controlling the six stages that culminate in the cellularization of the blastoderm embryo.

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Asymmetrically dividing Drosophila neuroblasts utilize two spatially and temporally independent cytokinesis pathways

Article Open access 20 March 2015

Link of Zygotic Genome Activation and Cell Cycle Control

Reshaping the Syncytial Drosophila Embryo with Cortical Actin Networks: Four Main Steps of Early Development

1 The Development of the Syncytial Embryo: Six Key Steps

Drosophila early embryo development occurs in a large syncytium in 13 rapid and synchronous nuclear cleavage cycles with 10–13 min separating each mitosis. These divisions occur over approximately 2 h, culminating in roughly 6000 nuclei that cellularize in interphase of cycle 14 to form the cellular blastoderm (Foe et al. 1993; Foe and Alberts 1983). During these early cleavage divisions, the centrosome coordinates cytoskeletal dynamics that are essential for proper development.

The centrosome is the major microtubule organizing center (MTOC) in most animal cells and is composed of two centrioles surrounded by the pericentriolar material (PCM) where microtubule assembly occurs. This coordination of microtubule production results in a polar microtubule array with the minus ends of the microtubules anchored at the centrosomes and plus ends that can rapidly grow and shrink. In the syncytial embryo, the centrosome is the only known MTOC. The centrosome has only recently been identified as an actin filament organizing center (Farina et al. 2016) but whether this is the case in the early embryo remains to be determined.

Here we describe the six key cell biological and developmental stages that rely on the centrosome and cytoskeletal dynamics during early embryo development. The first stage involves the maturation of the two centrioles contributed by the sperm, migration of one female pronucleus toward the male pronucleus, and the first zygotic division (Fig. 12.1a). The second stage consists of nuclear migrations that distribute the nuclei axially (Fig. 12.1b). The third stage is a perpendicular nuclear migration toward the cortex that generates the syncytial blastoderm (Fig 12.1c). The fourth, fifth, and sixth stages involve cortical membrane reorganization around each nucleus to generate cells (Fig. 12.1d–e). Each stage utilizes the centrosome in very different modes to organize the nuclei, assist in mitotic divisions, and/or form the first embryonic cells.

The first stage occurs during the initiation of embryogenesis, triggered by sperm entry through the anterior micropyle during fertilization in the uterus. Two paternally supplied centrioles mature and replicate utilizing maternally supplied PCM and centriolar proteins to form the first two embryonic centrosomes (Blachon et al. 2014). These centrosomes nucleate microtubules, termed the sperm aster, that assist in the migration of one female pronucleus toward the male pronucleus (Fig. 12.1a) (Callaini and Riparbelli 1996; Riparbelli et al. 2000). The first zygotic division is orchestrated by the newly formed centrosome pair, and four subsequent cleavage cycles precede the remaining centrosome-dependent stages.

During the second stage, axial nuclear migration, the early nuclei distribute evenly along the anterior-posterior (A-P) axis during cleavage cycles 4–7 (Fig. 12.1b). Localized actomyosin cortical contractions produce cytoplasmic streaming that assists in this nuclear migration (Royou et al. 2002; von Dassow and Schubiger 1994; Wheatley et al. 1995).

The third stage, cortical nuclear migration, positions the majority of the nuclei evenly along the cortex during cleavage cycles 7–9 (Fig 12.1c). Asymmetric microtubules nucleate preferentially toward the interior of the embryo to facilitate in this nuclear migration (Baker et al. 1993). A subset of nuclei, known as the yolk nuclei, remain in the interior of the embryo, complete error-prone replications that result in polyploid nuclei, and eventually lose their centrosomes (Fig. 12.1d) (Foe et al. 1993; Foe and Alberts 1983). Little is known of the molecular regulators of these nuclear migrations, but their function in positioning the nuclei is necessary for subsequent developmental stages (Niki and Okada 1981; Niki 1984; Okada 1982; Hatanaka and Okada 1991).

In the fourth stage, the nuclei that arrive in the posterior pole plasm during cortical migration are the first to cellularize, doing so during cleavage cycles 9–10. These nuclei cellularize before the remainder of the embryo to form the pole cells (primordial germ cells), the future gametic cells of the adult fly (Fig. 12.1d) (Foe and Alberts 1983). The pole plasm contains germ cell-specific proteins and mRNAs that are localized to the posterior of the oocyte in a microtubule-dependent manner during oogenesis (Lantz et al. 1999; Mahowald 2001). The pole plasm, which is necessary and sufficient to drive pole cell cellularization, is contained in polar granules that transport to the nuclei dependent on centrosomes and microtubules (Illmensee and Mahowald 1974; Lerit and Gavis 2011; Shamanski and Orr-Weaver 1991). The centrosomes coordinate reorganization of the plasma membrane to surround each nucleus as it divides, until the membrane is pinched off to form separate cells (Fig. 12.1d) (Raff and Glover 1989).

During the fifth stage, the remaining cortical nuclei complete four final divisions that utilize centrosome-dependent actin-microtubule dynamics to reorganize the cortical plasma membrane (Fig. 12.1e). The membrane dynamics resemble the organization of the posterior membrane during the fourth stage, but the membrane does not seal or close to form new cells. These membrane arrangements are termed pseudo-cleavage furrows, or Rappaport furrows (Raff and Glover 1988; Ede and Counce 1956; Turner and Mahowald 1976; Foe and Alberts 1983). These final divisions are important for increasing nuclear numbers and priming the embryo for cellularization.

The final stage succeeds the 13th division and occurs in the 70-min-long interphase of cleavage cycle 14 (Foe and Alberts 1983). The cortical nuclei are surrounded by long membrane invaginations rich in actin and cytokinetic components that cleave at the base to form cells (Fullilove and Jacobson 1971; Warn and Robert-Nicoud 1990; Young et al. 1991) (Fig. 12.1f). The centrosomes and microtubules assist in the membrane invaginations and eventual cell formation. This last step transitions the syncytial embryo to the cellular blastoderm (Zalokar and Erk 1976; Foe and Alberts 1983).

2 The Structure of the Embryonic Centrosome and Regulation of Microtubule Assembly

The embryonic centrosome is organized into a pair of centrioles surrounded by the PCM from which microtubules are nucleated and regulated. The embryonic centrioles have a canonical structure similar to differentiated tissue and mammalian centrioles with slight variation in length and the number of radial microtubules. Drosophila centrioles do not contain distal and subdistal appendages, structural features found on vertebrate mother centrioles (Callaini et al. 1997). The structure of the syncytial embryo centrioles remains constant throughout all of embryogenesis and into the larval stages, indicating that the structure is not unique to the specificities of the syncytial embryo (Callaini et al. 1997; González et al. 1998).

The embryonic centrioles are ~0.2 μm wide and long, composed of nine doublet microtubules that are all equal in length (Fig. 12.2) (Debec et al. 1999; Moritz et al. 1995; Lattao et al. 2017). The centrioles contain a “cartwheel” structure with a central hub linked to each peripheral doublet through radial spokes along the entire length (Fig. 12.2) (Debec et al. 1999; Callaini et al. 1997). Differentiated tissues, such as wing epidermal cells (Tucker et al. 1986) and ommatidia sensory bristles (Mogensen et al. 1993), contain microtubule triplets absent of the cartwheel structure, while midgut epithelial cells and rhabdomeric cells contain microtubule doublets with the cartwheel structure (Gottardo et al. 2015). Therefore, over development, some specified cells have different centriolar microtubule compositions. The reasons for these differences and any differential functions they might impart are not known.

The embryo is primed to construct centrioles thanks to the maternal supply of centriolar components, but centriole formation is blocked without the sperm centrioles. This block in activation of centriole biogenesis can be bypassed in unfertilized embryos through overexpression of centriole assembly/replication proteins such as Spindle assembly abnormal 6 (Sas-6) (Peel et al. 2007; Rodrigues-Martins et al. 2007a), Anastral spindle 2 (Ana2), Asterless (Asl) (Stevens et al. 2010), or Polo like kinase 4 (Plk4) (Peel et al. 2007; Rodrigues-Martins et al. 2007b), which drives de novo formation of centrioles.

While embryos are permissive for de novo centriole assembly, ovaries are not. Overexpression of either Spindle assembly abnormal 4 (Sas-4), Sas-6, or Plk4 during oogenesis still allows for the destruction of centrioles in the oocyte, resulting in unfertilized embryos initially absent of centrioles (Peel et al. 2007). De novo centriole formation is never seen in wildtype unfertilized embryos, indicating inhibitory mechanisms that limit centriole formation until fertilization. Dynein plays a negative regulatory role in centriole formation, as a dominant negative form of Dynein Heavy Chain 64C (Dhc64C) in unfertilized embryos causes de novo centriole formation (Belecz et al. 2001).

Most de novo centrioles maintain the typical embryonic architecture, such as during Plk4 overexpression (Rodrigues-Martins et al. 2008), but in some cases the centrioles have abnormal structures, such as during Sas-6 overexpression or in dominant negative Dynein heavy chain Dhc64C embryos. Tube-like structures rather than bona fide centrioles are produced, suggesting a precursory role in centriole biogenesis for Sas-6 and Dynein (Rodrigues-Martins et al. 2007a; Belecz et al. 2001). Although most of these embryos contain a large number of de novo centrioles, it is unclear whether the centrioles can replicate on their own. Rodrigues-Martins et al. showed that de novo centrioles from Plk4 overexpression can form procentrioles (Rodrigues-Martins et al. 2007b), but Peel et al. concluded no replication of de novo centrioles through live imaging of fluorescently tagged and overexpressed Plk4, Sas-4, or Sas-6 embryos (Peel et al. 2007). However, de novo centrioles can recruit PCM and nucleate microtubules (Peel et al. 2007; Rodrigues-Martins et al. 2007a, b, 2008; Stevens et al. 2010; Belecz et al. 2001).

One of the principle components of the PCM, γ-Tubulin, is the main microtubule nucleator at the centrosome. γ-Tubulin is expressed as two isoforms. γTUB37C, a maternal isoform, is expressed only in the ovaries and embryos. γTUB23C is the ubiquitous isoform, but the two isoforms are functionally redundant (Wilson et al. 1997). γ-Tubulin assembles into at least two different complexes that are important for microtubule nucleation and anchoring of the microtubule minus ends to MTOCs. The γ-Tubulin small complex (γ-TuSC) and the γ-Tubulin ring complex (γ-TuRC) are composed of γ-Tubulin complex proteins (GCPs) that contain grip domains which associate with γ-Tubulin and with other GCPs (Gunawardane et al. 2000; Oakley 2000; Farache et al. 2018; Lin et al. 2015; Kollman et al. 2015; Oakley et al. 2015).

γ-Tubulin is essential for syncytial embryo development (Tavosanis et al. 1997). γTUB37C mutants cannot nucleate astral microtubules and PCM recruitment is disrupted although spindles can still form (Wilson and Borisy 1998; Llamazares et al. 1999). γ-Tubulin complexes are recruited to the centrosome by the PCM component Centrosomin (Cnn) by the Centrosomin Motif 1 (CM1) domain of Cnn (Zhang and Megraw 2007; Chen et al. 2017).

In somatic tissues, Cnn is required for γ-Tubulin accumulation at the centrosome as well as astral microtubule production and PCM recruitment (Megraw et al. 1999; 2001; Mahoney et al. 2006). In the embryonic cleavage divisions of cnn maternal-effect mutants, PCM components such as γ-Tubulin are severely depleted at the centrosomes, but the centrioles can support a reduced amount of microtubule assembly, evident by small astral microtubules (Zhang and Megraw 2007). However, once the centrioles are lost, particularly in the later cleavage cycles, the centrioles cannot be properly maintained at the spindle poles, resulting in no detectable microtubule asters (Lucas and Raff 2007). Other PCM components are absent from the centrosome or are transiently recruited in cnn mutants, consistent with its primary role in recruiting PCM components (Zhang and Megraw 2007; Lucas and Raff 2007; Megraw et al. 1999; Vaizel-Ohayon and Schejter 1999).

The microtubules nucleated from γ-Tubulin are stabilized by Transforming acidic coiled-coil protein (Tacc) and Minispindles (Msps), which form a complex that is required for microtubule assembly and regulates astral microtubule length. Tacc-Msps localize at the centrosomes, and Tacc appears to recruit Msps. tacc mutants display a reduction in Msps localization at the centrosomes and a reduction in astral microtubules. Overexpression of Tacc causes a greater density of astral microtubules and more Msps recruitment to the centrosome than wildtype (Gergely et al. 2000a; Lee et al. 2001).

Tacc-Msps localization at the centrosomes is dependent on Aurora A (AurA) and the CM1 domain of Cnn (Gergely et al. 2000b; Barros et al. 2005; Cullen and Ohkura 2001; Lee et al. 2001; Giet et al. 2002; Zhang and Megraw 2007). aurA mutants display less localization of Tacc-Msps and shorter astral microtubules (Giet et al. 2002), while null cnn or CM1 domain mutants (cnn ^Δ1) still partially recruit Tacc-Msps (Zhang and Megraw 2007). The CM1 of Cnn domain also recruits γ-Tubulin, and recent work has revealed that Msps orthologs (Stu2 and Alp14) directly binds to γ-Tubulin complex proteins to assist in microtubule nucleation through their tumor overexpressed gene (TOG) domains (Gunzelmann et al. 2018; Flor-Parra et al. 2018; Nithianantham et al. 2018). Therefore, reduced Tacc-Msps in cnn mutants may be due to reduced γ-Tubulin localization at the centrosomes.

For broader coverage of the centrosome and MTOCs, see Centrosomal and Non-centrosomal Microtubule-Organizing Centers (MTOCs) in Drosophila melanogaster (Tillery et al. 2018).

3 Fertilization and the First Zygotic Division

Embryogenesis of the zygote begins with syngamy of the haploid female and male pronuclei at fertilization. Sperm entry, pronuclear migration, and the first zygotic division all occur within 15 min and rely on the complementary contributions of the paternal centrioles and maternally supplied PCM components (Foe et al. 1993). The sperm enters the egg in the uterus and female meiosis, arrested in metaphase I, is activated by passage through the oviduct (Von Stetina and Orr-Weaver 2011). Meiosis produces four pronuclei that are arranged in a row perpendicular to the cortex (Fig. 12.3a–c). The sperm supplies two centrioles that immediately recruit maternal PCM components and assemble astral microtubules that stretch toward the cortex, termed the sperm aster (Fig. 12.3b–c) (Callaini and Riparbelli 1996; Riparbelli et al. 2000). The female pronucleus farthest from the cortex (and closest to the sperm aster) migrates toward the male pronucleus along the sperm aster until the two pronuclei are in apposition (Fig. 12.3c–d). The first zygotic division proceeds utilizing the newly matured centrioles and their templated daughter centrioles that form the first embryonic centrosomes (Loppin et al. 2015).

Two centrioles are supplied by the sperm: a larger “giant” centriole (GC), derived from the basal body, and the smaller unconventional centriole, referred to as the PCL (proximal centriole-like) (Fig. 12.3a) (Blachon et al. 2009; Blachon et al. 2014). The PCL is unconventional in that it lacks centriolar microtubules (Khire et al. 2016). During centriole maturation, maternal PCM components including Cnn, Asl, γ-Tubulin, Spindle defective 2 (Spd2), Pericentrin-like protein (Plp), and Centrosomal protein 190kD (CP190) localize at the GC and PCL upon sperm entry (Blachon et al. 2014; Callaini et al. 1999; Khire et al. 2016). PCM recruitment to sperm centrioles is reliant on Spd, as spd2 mutants recruit Asl but fail to recruit Cnn and have impaired sperm aster formation and pronuclear migration (Dix and Raff 2007). This requirement of Spd2 for sperm centriole maturation differs from somatic cells where loss of Spd2 only partially impedes PCM recruitment, allowing for significant centrosome activity to remain (Dix and Raff 2007; Giansanti et al. 2008).

Following PCM recruitment/maturation of the sperm centrioles and the completion of female meiosis, the sperm aster facilitates in the migration of one female pronucleus to the male pronucleus (Fig. 12.3c–d) (Loppin et al. 2015). The three remaining female pronuclei become polar bodies, which do not divide and remain throughout the syncytial embryo stages (Fig. 12.3c–d) (Dävring and Sunner 1973).

MTOC function and sperm aster assembly are essential for pronucleus migration. Loss of Asl or Spd2 disrupts sperm aster formation and pronuclear migration fails (Blachon et al. 2014; Varmark et al. 2007; Dix and Raff 2007). Loss of the PCM component, Asp, or the spindle microtubule regulators, Polo or Wispy (Wisp), produces a weaker phenotype where the sperm aster forms but does not fully extend toward the cortex, also preventing pronuclear migration (Riparbelli et al. 2002; Riparbelli et al. 2000; Brent et al. 2000). Loss of Tacc, which regulates microtubule stability, results in pronuclear migration failure. tacc mutants that survive to later stages of embryogenesis display diminished aster and spindle microtubules, suggesting failure of pronuclear migration may be due to a diminished sperm aster (Gergely et al. 2000b).

The Linker of Nucleoskeleton and Cytoskeleton (LINC) complex helps maintain centrosome-nucleus connections (Hieda 2017) and facilitates female pronuclear migration in zebrafish and Caenorhabditis elegans embryos (Lindeman and Pelegri 2012; Malone et al. 2003). However, in Drosophila, Klarischt (Klar) and Muscle-Specific Protein 300 kDa (Msp300), both LINC complex components, are not necessary for pronuclear migration (Technau and Roth 2008). In C. elegans, the LINC complex cooperates with microtubule motor proteins to assist in pronuclear migration (Meyerzon et al. 2009), and while the LINC complex is not necessary in Drosophila, motors are needed. What tethers nuclei to the cytoskeleton during pronuclear migration and other nuclear movements in early Drosophila embryos remains unknown.

The kinesins Non-claret disjunctional (Ncd), Subito (Sub), and Kinesin-like protein at 3A (Klp3A) all play roles in pronuclear migration. The female pronucleus migrates in a minus-end-directed manner, and Ncd, a microtubule minus-end-directed motor, assists in this migration in conjunction with an isoform of α-Tubulin, αTub67C (Komma and Endow 1997). αTub67C is a maternal-specific isoform of α-Tubulin with the special property of conveying faster microtubule assembly (Venkei et al. 2006). Sub is a kinesin involved in antiparallel microtubule bundling and sub mutants display similar phenotypes to polo, wispy, and α-tub67C mutants, where the mitotic spindles do not form properly and the embryos arrest early without any zygotic divisions (Giunta et al. 2002; Cesario et al. 2006). Therefore, Sub may play a role in attaching the female pronucleus to the sperm aster through microtubule interactions. Loss of Klp3A prevents pronuclear migration, but Klp3A is a plus-end-directed motor, implicating an indirect role in pronuclear migration. Klp3A recruits Polo, suggesting that it may regulate the formation of the sperm aster, which is necessary for pronuclear migration (Glover 2005; Williams et al. 1997).

During migration the pronuclei swell until the nuclei are in apposition to one another, resulting in a slightly larger female pronucleus (Fig. 12.3d) (Callaini and Riparbelli 1996). The paternal centrioles separate to opposite poles once the pronuclei are apposed and template daughter centrioles to form two functioning centrosomes that aid in the first zygotic division (Fig. 12.3d–e) (Blachon et al. 2014). This division is gonomeric because the female and male chromosomes remain separated on the metaphase plate until telophase when they join to form two diploid zygotic nuclei (Fig. 12.3e) (Callaini and Riparbelli 1996).

In order to properly replicate and recruit PCM proteins, the PCL requires unique components such as Proteome of centrioles 1 (Poc1). In poc1 mutant testis, the PCL does not assemble and sperm contain only a GC. In poc1 paternal effect mutant embryos, PCM proteins are recruited to the GC only (because no PCL is delivered with the sperm), resulting in monopolar spindles that contain only the GC and its replicated daughter centriole (Khire et al. 2016).

The ultrastructure of each centriole goes through multiple changes, and centriolar components such as Anastral spindle 1 (Ana1), Ana2, Asl, Sas-4, and Sas-6 are stripped away during spermatogenesis (Khire et al. 2016; Blachon et al. 2014). These components are maternally supplied in the embryo, and at least Asl is recruited to the sperm centrioles, while Sas-4 and Sas-6 remain absent from the GC and PCL. However, when the centrioles replicate, Sas-4 and Sas-6 are present at the newly formed daughter centrioles (Blachon et al. 2014).

For more on Drosophila fertilization, see a recent review (Loppin et al. 2015).

4 The Syncytial Embryo Employs an Adapted Cell Cycle

Due to the accelerated pace of nuclear divisions, the syncytial embryo involves a modified cell cycle that does not utilize gap phases but only S and M phases until cellularization (Glover et al. 1989). Because of the fast transitions from mitosis to interphase, there is also a severe reduction in transcription until the maternal to zygotic transition (MZT) during the tenth cleavage division (Lamb and Laird 1976; McKnight et al. 1977; Zalokar 1976; Edgar and Schubiger 1986). Instead, the embryo relies on the activities of maternally supplied proteins and mRNAs to execute the syncytial nuclear divisions (O’Farrell 2015; Lasko 2012).

The major cell cycle regulators, Cyclin-dependent kinases (Cdks) and Cyclins, are important in managing the timing of mitosis in the syncytial blastoderm. Cdks and Cyclins cooperate to regulate the timing of protein activation during various stages of the cell cycle. Typically, Cdk and Cyclin levels are regulated through temporal expression, but the lack of transcription in the early embryo results in a modified mechanism to support cleavage cycle regulation. In the embryo, Cdks diffuse through the embryo in waves, causing subsequent waves of mitosis (Deneke et al. 2016), whereas cyclins are locally degraded at the centrosomes to prevent global destruction that would halt further mitotic cycles (Huang and Raff 1999; Raff et al. 2002).

The syncytial nuclei divide in a synchronous wave regulated by Cdk1 that propagates along the A-P axis. Cdk1 forms complexes with Cyclin A and B to regulate entry into mitosis. Because diffusion would take too long for mitotic activation in the embryo, Cdk1 propagates throughout the embryo as a wave that signals the nuclei to enter mitosis. Particularly in the later cleavage cycle stages, Cdk1 waves spread throughout the embryo during S phase to trigger mitosis, resulting in a subsequent wave of mitosis of the nuclei (Deneke et al. 2016).

Both Cyclin A and B are involved in regulating the syncytial embryo cell cycle. Cyclin A is localized at the nucleus and regulates the duration of the entire cell cycle, as decreased Cyclin A results in a longer cell cycle, while the mitotic index remains the same (Edgar et al. 1994; Stiffler et al. 1999). Cyclin B plays a more complex role, regulating not only specific mitotic stages but also microtubule length and nuclear velocity in the migration stages. Cyclin B localizes to the spindle microtubules during metaphase and astral microtubules in later mitotic stages (Huang and Raff 1999; Stiffler et al. 1999). Decreasing Cyclin B levels results in longer astral and spindle microtubules, as well as centrosome detachment from their respective nuclei. Increasing Cyclin B levels causes shorter astral and spindle microtubules, resulting in nuclear spacing defects (Stiffler et al. 1999).

Cyclin B is destroyed during metaphase and this destruction is localized to the spindle microtubules, starting at the centrosomes (Huang and Raff 1999) and catalyzed by Cdc20 (Fizzy (Fzy))-dependent Anaphase-promoting complex (APC) activation (Raff et al. 2002; Sigrist et al. 1995). Fzy is localized to the centrosomes, spindle microtubules, and kinetochores during the start of mitosis and begins to disappear during metaphase. The localization of Fzy to the centrosome is microtubule-dependent as colcemid treatment (a microtubule depolymerizer) causes Fzy to localize strictly to the kinetochores. It is hypothesized that the Fzy-APC complexes are activated at the centrosomes and spread to the kinetochores due to the localization of Fzy (Raff et al. 2002) and that centrosome and spindle attachment is necessary for Cyclin B destruction (Wakefield et al. 2000).

When centrosomes detach from the spindle and Cyclin B is not destroyed, spindles arrest in mitosis, which is also seen in sas-4 mutants (spindle arrest, absent of centrioles), supporting a role for the centrosomes and Cyclin B destruction (Stevens et al. 2007; Wakefield et al. 2000). Pan gu (png) forms a complex with Giant nuclei (Gnu) and Plutonium (Plu) to regulate Cyclin B levels, specifically in the early embryo. It is hypothesized that this complex works to stabilize Cyclin B, as Cyclin B levels are decreased in either png, glu, or plu mutants (Fenger et al. 2000). These mutants also display DNA replication without division resulting in polyploid nuclei, as well as centrosome detachment from the spindles. (Freeman et al. 1986; Freeman and Glover 1987; Elfring et al. 1997; Shamanski and Orr-Weaver 1991). This phenotype is also seen in embryos lacking Cyclin B, supporting the idea of localized Cyclin B destruction at the centrosome (Stiffler et al. 1999). The detached centrosomes continue to replicate, uncoupled from the DNA replication cycle, indicating a mechanism by which the centrosome and DNA replication cycles can be uncoupled (Freeman et al. 1986; Freeman and Glover 1987; Elfring et al. 1997; Shamanski and Orr-Weaver 1991).

In aphidicolin-injected embryos (DNA replication inhibitor) that have a prolonged S phase, centrosomes separate from the nuclei, and over-replication of centrosomes occurs (Raff and Glover 1988; Debec et al. 1996). Also in dhc64C mutants, centrosomes separate from the nuclei and continue to replicate, leading to excessive centrosome replication (Belecz et al. 2001). Similarly, in mcph1 mutants where S phase is prolonged causing an increase in the DNA replication cycle length, the length of the centrosome replication cycle stays the same, resulting in excessive centriole replication (Brunk et al. 2007).

Centriole replication throughout the syncytial embryo mitoses is regulated by the same components involved in initial centriole formation during fertilization. The centriole assembly proteins, Sas-4, Sas-6, and Plk4, are required for the formation of the daughter centriole during centrosome replication. In sas-4, sas-6, or plk4 maternal mutants, centrioles are not formed, and embryos arrest early with very few divisions that have abnormally shaped anastral spindles. Additionally, PCM components such as Cnn are not recruited to the spindle poles (Rodrigues-Martins et al. 2008; Stevens et al. 2007). Overexpression of either Sas-6 or Plk4 causes excessive centrosome replication, due to their role in centriole biogenesis (Peel et al. 2007; Rodrigues-Martins et al. 2007b). Excessive centrosome replication due to Sas-6 overexpression is exacerbated by the loss of Centriole Coiled Coil Protein 110 kDa (CP110). cp110 mutants cause excessive centrosome replication when either Asl or Ana2 was overexpressed, which do not display abnormal centrosome replication on their own (Franz et al. 2013). Therefore, CP110 negatively regulates centrosome duplication through Sas-6, Asl, and Ana2.

5 Centrosome-Nucleus Association

As in most cell types and organisms, centrosomes are closely linked to the nuclei of the syncytial embryo through microtubule interactions, motor proteins, and microtubule-associated proteins (MAPs). Close centrosome-nuclear localization allows for rapid assembly of the mitotic spindle during the quick transitions of the cleavage cycle, as well as aiding in force mechanisms for nuclear migration and positioning. Due to the syncytial nature of the embryo, this association is important to prevent centrosomes from drifting away from their respective nuclei, which is less of a concern in the containment of a cell. Free centrosomes can disrupt nuclear divisions as well as prevent proper nuclear positioning; therefore, the syncytial embryo requires unique mechanisms to keep the centrosome-nuclear association intact.

Nuclear envelope breakdown in the embryo deviates from the canonical cell cycle, as the centrosomes remain extremely close to the nuclei. During prophase, the nuclei become indented near the centrosomes; during prometaphase, portions of the nuclear envelope breakdown at these indents, theoretically due to the astral microtubules piercing the nucleus. Remnants of the nuclear envelope continue to surround the spindle until telophase, when it fully breaks down and reforms at the two newly separated nuclei (Stafstrom and Staehelin 1984; Paddy et al. 1996; Rothwell and Sullivan 2000).

The LINC complex is an obvious candidate for nuclear attachment to the centrosome; however, loss of the LINC complex components Klar, Msp300, or Klaroid (Koi) does not display any obvious centrosome-nuclear attachment defects (Archambault and Pinson 2010; Technau and Roth 2008). Rather, the microtubule motor protein dynein and PCM proteins are necessary for this attachment. dhc64c mutants display detached centrosomes from interphase/prophase nuclei as well as spindle poles, and centrosomes often fail to separate properly during prophase (Robinson et al. 1999). Loss of Polo also causes interphase/prophase centrosome detachment, particularly during the cortical migration stage. This may be due to the recapturing of detached centrosomes during spindle formation when they are still close by, but during the migration stage, the nuclei move too far for the centrosomes to be recaptured, resulting in monopolar spindles. Centrosome detachment is not highly penetrant in polo mutants, but this phenotype is exacerbated by the overexpression of Microtubule-associated protein 205 (Map 205) or Greatwall (Gwl) (Archambault et al. 2007; Archambault et al. 2008). Map 205 sequesters Polo to microtubules during interphase, and Gwl antagonizes Polo via inhibition of the regulatory subunit of Protein phosphatase 2A, Twins (Wang et al. 2011).

Mutations in the gene for another MAP, Mars (HURP homolog), shows centrosome detachment from prophase nuclei when depleted, but more often centrosomes detach from the mitotic spindle (Zhang et al. 2009). Centrosome detachment from the spindle is also seen in asp mutants, evident by monopolar spindles (González et al. 1990). A syncytial embryo specific MAP, Toucan (Toc), localizes to the nuclear envelope and centrosomes during interphase and the spindle microtubules during mitosis. Mutant toc embryos display detached centrosomes from spindles and defective spindle formation. Astral microtubules remain intact and these embryos typically arrest early on in a metaphase state, indicating a specialized role for Toc in regulating syncytial mitotic spindles (Debec et al. 2001; Mirouse et al. 2005).

Microcephalin (MCPH1), which localizes to the centrosomes and spindle, is necessary for centrosome-spindle pole attachment as mutant mcph1 embryos display detached centrosomes and monopolar spindles. These mutants arrest early on in a metaphase state but also have a delayed S phase, which results in uncoupling of the centrosome and cell cycles. Desynchronization of these cycles can result in over-replication of centrosomes, which may be the cause of the detached centrosomes (Brunk et al. 2007).

For more on centrosome attachment in the syncytial embryo, see Free Centrosomes: Where Do They Come From? (Archambault and Pinson 2010).

6 Axial Nuclear Migration Distributes Nuclei along the A-P Axis

The first 3–4 cleavage cycles are skewed toward the anterior end of the embryo where syngamy occurs (Karr 1991). Before the nuclei migrate toward the cortex and form the blastoderm, they organize into a uniform distribution along the A-P axis through axial nuclear migration (also known as axial expansion) (Fig. 12.1b) (Hatanaka and Okada 1991). Axial nuclear migration occurs during cleavage cycles 4–7 and relies on cytoskeletal dynamics that are regulated through the cell cycle and the centrosome.

Actin and non-muscle myosin II (MyoII) localize at the anterior cortex in a cell cycle-dependent manner, where they control physical contractions of the embryo that assist in the axial nuclear migration (Fig. 12.4d) (Royou et al. 2002; von Dassow and Schubiger 1994; Wheatley et al. 1995). It is hypothesized that this contraction causes cytoplasmic streaming that forces the nuclei to migrate along the A-P axis, which is further supported by the cytoplasmic movements during these stages (Fig. 12.4b) (von Dassow and Schubiger 1994). A second hypothesis is that the actin network causes the cytoplasm to become stiffer in the middle of the embryo and looser toward the poles. Interacting microtubules from neighboring centrosomes repel the nuclei away from one another, forcing the nuclei to expand laterally along the A-P axis during these contractions. (Foe et al. 1993; von Dassow and Schubiger 1994). There is support for both of these hypotheses, and both may contribute to the nuclear movements during axial nuclear migration.

Three distinct phases of nuclear movements transpire during each cleavage cycle 4–7 at different stages of the cleavage cycle (Fig. 12.4a). First, the nuclei slowly begin to migrate toward the poles along the A-P axis at the end of interphase. In the second phase, this movement rapidly increases during prophase and early metaphase. Finally, nuclear movement slows down and they regress slightly along the A-P axis, away from the poles, during anaphase and telophase (Baker et al. 1993; von Dassow and Schubiger 1994).

The specificity of cell cycle stages during these nuclear movements suggests a role for cell cycle regulation in axial nuclear migration. In support of this, increasing overall Cyclin B levels in the embryo decreases the velocity of nuclear movements, and decreasing overall Cyclin B levels in the embryo increases the velocity of nuclear movements (Stiffler et al. 1999). Therefore, Cyclin B regulates the nuclear velocity during these three phases of nuclear migration.

The cytoplasm contained inside the entire embryo displays movements that mimic nuclear migration. As the nuclei move toward the pole, the cytoplasm deep in the middle of the embryo moves outward toward the poles as well (Fig. 12.4b). The cytoplasm at the periphery of the cortex flows toward the middle of the embryo, and the two distinct waves, from the posterior and anterior, converge slightly anterior at the cortex, at what is termed the constriction point. Because of the opposing flows of the deep cytoplasm and peripheral cytoplasm, this movement is referred to as fountain streaming. During the final phase of each nuclear movement when the nuclei slightly retract, the cytoplasm also retracts toward the middle of the embryo (Fig. 12.4c) (von Dassow and Schubiger 1994).

Axial nuclear migration relies on actin filaments and not microtubules as colchicine treatment (a microtubule depolymerizer) does not affect A-P nuclear distribution, but cytochalasin (prevents actin polymerization) does inhibit it (Zalokar and Erk 1976; Hatanaka and Okada 1991). Filamentous actin (F-actin) appears to nucleate from the centrosomes starting at metaphase, growing through telophase, and dispersing in interphase. These actin asters are greatest at the outward nuclei, closest to the poles, and weakest at the inward nuclei, furthest from the poles (Fig. 12.4d). Because the outward nuclei move more than the inward nuclei, it is suggested that these actin asters facilitate in axial nuclear migration (von Dassow and Schubiger 1994).

Loss of Grandchildless N26 (Gs(1)N26), Grandchildless N41 (Gs(1)N41), or Paralog (Par), all of which have not been mapped to a physical locus, cause actin to appear as a uniform layer over the cortex with rough aggregates and defective axial nuclear migration (Hatanaka and Okada 1991). This indicates they play a role in regulating actin distribution during axial nuclear migration.

Actin, together with MyoII which is also necessary for axial nuclear migration, shows a distinct localization progressing through cleavage cycles 4–7 (Kiehart et al. 1990; Wheatley et al. 1995; Royou et al. 2002). MyoII localizes at the cortex, slightly anterior, during interphase of the axial nuclear cleavage divisions (Fig. 12.4e). MyoII localizes to the anterior constriction site starting at interphase 4 and increases in intensity during the following interphase cycles until interphase 7, where it starts to disperse along the cortex. By interphase 8, when the nuclei are evenly distributed along the A-P axis, MyoII appears almost entirely disperse at the cortex. Actin also cycles to the cortex in a similar manner (Royou et al. 2002).

The cell cycle-dependent localization of MyoII is regulated by Cdk1 and Cyclin B. Localized degradation of Cyclin B during late anaphase inactivates Cdk1 (Su et al. 1998), and increasing Cyclin B levels prevents MyoII cortical localization, while inhibiting Cdk1 results in abnormal localization of cortical MyoII (Royou et al. 2002). However, cortical MyoII localization is not reliant on actin, as cytochalasin or latrunculin (prevents actin polymerization) injection does not disrupt MyoII localization (Chodagam et al. 2005). Additionally, in mutants for the regulatory light chain of MyoII, Spaghetti squash (Sqh), cortical actin localization is not disrupted, indicating it is independent of MyoII (Royou et al. 2002).

Antimyosin antibody injection or sqh mutants display defective axial nuclear migration (Kiehart et al. 1990; Wheatley et al. 1995), specifically, the primary activating phosphorylation site of Sqh, Serine 21 (Karess et al. 1991), is necessary for this migration. Phosphorylation site mutants that either mimic phosphorylation or prevent phosphorylation of Serine 21 both hinder axial nuclear expansion, indicating proper regulation of Sqh phosphorylation is necessary for axial nuclear migration (Jordan and Karess 1997). This site is phosphorylated by Rho kinase (Rok), and inhibition of Rok with Y-27632 also hinders axial nuclear expansion as well as MyoII distribution (Royou et al. 2002).

The centrosome, apparently acting through the centrosomal protein CP190, also plays a role in regulating axial nuclear migration. cp190 mutants display defective axial nuclear migration and MyoII localization, but actin organization remains intact. The constitutively active Sqh phosphomimetic mutant can partially rescue this phenotype, implicating the centrosome in MyoII regulation. Although originally identified as a centrosomal protein, CP190 is best known as a chromatin insulator (Kellogg et al. 1989; Pai et al. 2004). CP190 localization is regulated during the cell cycle as it localizes to the centrosomes during mitosis and is nuclear during interphase (Chodagam et al. 2005). These localization dynamics may regulate the localization of MyoII during axial nuclear migration. Two unmapped genes regulate axial nuclear migration and affect centrosome localization: shackleton (shkl) and out of sync (oosy) mutants display defective axial nuclear migration, and centrosome loss from the spindles. oosy mutants also display asynchronous cleavage divisions (Yohn et al. 2003). Therefore, the timing of axial nuclear migration relies on cell cycle regulators that may involve the centrosome in an unknown way.

Overall, axial nuclear migration is a poorly understood process that is regulated by Rho1-dependent actin-MyoII dynamics and has an unclear connection to centrosomes. It is critical for the timely delivery of nuclei to the germ plasm during the cycle 10 window of pole cell formation. It also spreads the nuclei along the A-P axis to establish an even migration to the cortex.

7 Cortical Nuclear Migration Positions the Nuclei at the Cortex

During cleavage cycles 7–9, the majority of nuclei migrate from the interior of the embryo to the cortex (Fig. 12.1c) (Foe and Alberts 1983). A subset of nuclei, the yolk nuclei, fall back into the interior of the embryo between cleavage cycles 8 and 9 (Foe et al. 1993). Little is known about the function of these nuclei, but they are involved in yolk digestion (Bownes 1982) as well as the future development of the midgut (Walker et al. 2000).

The yolk nuclei asynchronously divide twice, then complete two rounds of DNA replication without divisions to become polyploid (Zalokar and Erk 1976; Foe and Alberts 1983). During these divisions, the centrosomes display defects in mitotic spindle organization that ultimately leads to centrosome loss at the yolk nuclei. During the first asynchronous division, a majority of the nuclei display defective centrosome separation, resulting in “V”-shaped spindles that resemble monopolar spindles. These aberrant spindles result in defective DNA segregation, yet the centrosomes continue to replicate, and a second round of abnormal divisions occurs. The replicated centrosomes also do not separate, as mother and daughter centrosomes remain close to one another (Callaini and Dallai 1991; Riparbelli and Callaini 2003). By the second yolk nuclei division, most of the centrosomes have detached from the spindles. The centrosomes appear normal, as CP190, Asp, Pavarotti, and γ-Tubulin all remain localized there. However, Cyclin B localization is disrupted as it does not associate with the aberrant spindles but weakly localizes to the centrosomes during anaphase and telophase. It does not localize to the entire spindle pole but rather the inner core of the centrosomes (Callaini and Dallai 1991; Riparbelli and Callaini 2003).

In contrast with axial nuclear migration, cortical nuclear migration relies on microtubules as colchicine treatment inhibits cortical nuclear migration, and does not rely on actin, as cytochalasin treatment does not inhibit cortical nuclear migration (Zalokar and Erk 1976). Centrosomes appear to carry nuclei during this migration, as centrosomes dissociated from their respective nuclei continue to migrate to the cortex (Raff and Glover 1989).

It was initially hypothesized that the centrosomes nucleate microtubules that connect to the cortex to pull the nuclei by forces produced by the microtubules (Wolf 1980). However, analysis of the microtubule arrays during cortical nuclear migration has disputed this theory. The majority of microtubules project inwards, rather than to the cortex, and interact with the microtubules of the yolk nuclei (Fig. 12.5). Shorter microtubules project toward the cortex, but do not reach the membrane (Fig. 12.5). Microtubules of neighboring nuclei, as well as nuclei at the opposite side of the embryo, also interact with one another (Fig. 12.5) (Baker et al. 1993).

The density of astral microtubules is far greater in the cortical nuclear migration stages compared to those during the axial nuclear migration stages (Baker et al. 1993). In tacc mutants, the astral microtubules are reduced or absent, and the nuclei do not migrate to the cortex (Gergely et al. 2000b). The current hypothesis is that antiparallel microtubule interactions provide a pushing force that directs the centrosomes, with their nuclei as passengers, toward the cortex (Fig. 12.5) (Baker et al. 1993; Raff and Glover 1989).

Very little is known about the molecular regulators of cortical nuclear migration, but it is coordinated with the cleavage cycle. The nuclear velocity is greatest during telophase when astral microtubules are most abundant, further supporting their necessity during this migration (Foe and Alberts 1983; Baker et al. 1993). As mentioned earlier, higher levels of cyclin B reduce the speed of nuclear migration, but this could be due to the decrease in microtubule length caused by the increased levels (Stiffler et al. 1999). In oosy mutants, the cleavage divisions are not synchronous and cortical migration is defective, suggesting that this synchrony is also important for proper migration (Yohn et al. 2003).

Overall, while little is understood about how cortical nuclear migration is regulated, it appears to be driven by astral microtubules assembled at centrosomes that grow toward the center of the embryo and which push the nuclei outward toward the cortex as they grow.

8 Pole Cells Cellularize Before the Other Nuclei

During cortical nuclear migration, a subset of nuclei reaches the posterior cortex and pole plasm during cleavage cycle 9. These nuclei will cellularize at cycle 10, becoming pole cells, while the majority of nuclei at the cortex continue to divide (Fig. 12.1d) (Foe and Alberts 1983). Pole cells will develop into the germ cells of the fly and contain germ cell-specific components. The germ plasm is localized to the posterior end of the oocyte during oogenesis and is contained in polar granules in the early embryo that are anchored by the actin cytoskeleton (Fig. 12.6a) (Lantz et al. 1999; Mahowald 2001).

The pole plasm is sufficient to initiate pole cell formation, as transplantation of the pole plasm to the anterior cortex stimulates abnormal pole cell formation at the anterior pole (Illmensee and Mahowald 1974). Oskar (Osk), a germ cell-specific protein required to localize other germ cell-specific components, is also sufficient to cause abnormal pole cell formation when transplanted to the anterior pole (Ephrussi and Lehmann 1992). Germ-cell less (Gcl) is necessary for pole cell formation, but is not sufficient for the formation of pole cells when transplanted to the anterior pole (Jongens et al. 1992). However, when transplanted together with Anillin (encoded by scraps), which regulates actomyosin contractile rings and is also necessary for pole cell formation, they can stimulate abnormal pole cell formation at the anterior pole (Cinalli and Lehmann 2013; Field et al. 2005).

Gcl also plays a role in limiting pole cell formation to the posterior pole. Gcl promotes actomyosin organization downstream of the Rho1 pathway to constrict the plasma membrane and form the pole cells at the posterior pole (Cinalli and Lehmann 2013). The Arf-GEF Stepkke (Step) inhibits the Rho1-actomyosin pathway, and during pole cell formation stages, it is equally distributed around the cortical membrane of the embryo, preventing any cellular formation. Loss of Step activity results in abnormal pole cell formation at the anterior cortex due to loss of Rho1 inhibition. However, the posterior pole is distinct from the remainder of the embryonic membrane due to the presence of Gcl, which locally inhibits Step activity to allow for Rho1-mediated actomyosin membrane constriction. Loss of Step in a gcl mutant background allows for proper pole cell formation (Lee et al. 2015).

Another determinant of pole cell formation is proper axial nuclear migration. During interphase of cleavage cycle 9, the first nuclei reach the posterior pole in the proper time window for pole cell formation to begin. If axial nuclear migration is defective and the nuclei do not reach the posterior pole until a time point after cleavage cycle 10, pole cell formation is inhibited (Niki and Okada 1981; Niki 1984; Okada 1982; Hatanaka and Okada 1991). This is likely due to the degradation of germ plasm components before the nuclei can reach the posterior pole. When pole cell formation fails in gs(1)N26 or gs(1)N441 mutants, which display defective axial nuclear migration, the localized pole plasm components either degrade or delocalize from the posterior pole (Iida and Kobayashi 2000). In shkl or oosy mutant embryos, which also display defective axial expansion, pole cell numbers are lower than wildtype, further supporting the necessity of axial nuclear migration and timely arrival of nuclei at the posterior pole plasm for pole cell formation (Yohn et al. 2003).

When the nuclei reach the posterior membrane during cleavage cycle 9, they enter prophase and plasma membrane protrusions, called pole buds, form above each nuclei (Fig. 12.6b) (Foe et al. 2000; Warn et al. 1985; Cinalli and Lehmann 2013). As the nuclei begin to divide, the pole buds protrude farther from the membrane to surround the metaphase spindles (Fig. 12.6b). Membrane furrows, termed the bud furrows (BFs), form at the edge of the pole buds and constrict beneath the chromosomes at the basal membrane (Fig. 12.6b). Once the nuclei progress into anaphase, a second furrow, the anaphase furrow (AF), forms above the spindle opposite to the BF. Similar to the cytokinetic furrow, it forms in between the dividing nuclei (Fig. 12.6b). Once mitosis is complete, the AF constricts to separate the two nuclei into separate cells, while the BF constricts, liberating the nascent pole cells from the embryo (Fig. 12.6b) (Cinalli and Lehmann 2013).

Known cytokinetic factors Anillin, MyoII, Peanut (Pnut), and Rho guanine nucleotide exchange factor 2 (RhoGEF2) all localize to the actin-rich BFs in preparation for cleavage of the plasma membrane (Padash Barmchi et al. 2005; Warn et al. 1985; Field and Alberts 1995; Young et al. 1991). Anillin and MyoII also localize to the actin-rich AFs, but further analysis is required to determine other components of the AF. Rho1 functions upstream of cytokinetic components, and inhibiting Rho1 or Rok prevents pole bud formation, specifically diminishing Anillin localization at the BFs (Cinalli and Lehmann 2013). Anillin is necessary for cleavage of the pole cells, as scraps mutants display BFs that retract and never form pole cells (Field et al. 2005). diaphanous (dia) (a Formin downstream of Rho1) or rhogef2 mutants display defective BF cleavage due to the disruption of the actomyosin contractile ring, as actin and MyoII are absent at the BFs (Afshar et al. 2000; Padash Barmchi et al. 2005).

The rate of BF constriction is regulated by Gcl, as overexpressing Gcl causes over-constriction of the BFs, resulting in the displacement of somatic nuclei and increased pole cell numbers (Cinalli and Lehmann 2013; Jongens et al. 1994). gcl mutants display under-constriction of the BFs, resulting in decreased pole cell numbers (Robertson et al. 1999; Cinalli and Lehmann 2013). In gcl mutants, the AF constricts to separate the cells but the BF never constricts, even though Anillin is present at both, preventing the formation of the pole cells. Therefore, Gcl is necessary for BF cleavage, but not AF cleavage (Cinalli and Lehmann 2013).

Both actin and microtubules are necessary for proper pole cell formation as injection of either Colcemid or cytochalasin inhibits pole cell formation (Raff and Glover 1989; Cinalli and Lehmann 2013). Colcemid treatment prevents AF cleavage but not BF cleavage and the nuclei arrest in metaphase, resulting in large pole cell-like cells with inappropriate DNA content (Cinalli and Lehmann 2013). Cytochalasin injection after pole cell formation causes the cells to collapse, indicating actin is necessary for pole cell stabilization (Raff and Glover 1989).

Centrosomes alone are sufficient to produce pole cells. Active centrosomes dissociated from nuclei during aphidicolin injection still migrate to the posterior pole and produce pole buds and reorganize actin. Pole cells form that are indistinguishable from normal pole cells except that they lack nuclei (Raff and Glover 1989). The centrosomal protein CP110 regulates pole cell formation. Neuralized E3 ubiquitin protein ligase 4 (Neurl4) localizes to the centrosome and downregulates CP110 levels in mammalian cells (Al-Hakim et al. 2012; Li et al. 2012). In neurl4 mutant embryos, CP110 levels are elevated at centrosomes compared to wildtype as well as enriched at foci distinct from centrosomes. neurl4 mutants display reduced pole cell numbers with abnormal morphologies due to CP110 overexpression, as the phenotype was partially rescued in neurl4 mutants with the addition of one mutant copy of cp110, resulting in less abnormal pole cells (Jones and Macdonald 2015). How CP110 impacts the centrosome and affects pole cell formation is unclear.

Centrosome separation during mitosis regulates the formation and size of the BFs. gcl mutants display disrupted centrosome separation specifically in the pole cells and not somatic cells, which leads to shallow BFs (Lerit et al. 2017). These smaller BFs often retract, preventing their cellularization which leads to lower numbers of pole cells (Cinalli and Lehmann 2013). The percentage of pole buds with defective centrosome separation in gcl mutants correlates with the number of gcl mutant embryos that lack pole cells (Lerit et al. 2017; Robertson et al. 1999). Some nuclei overcome this BF defect and form pole cells, but they contain an abnormal number of centrosomes and multipolar spindles (Lerit et al. 2017). The CM1 domain of Cnn and also the Kinesin-5 motor protein (Klp61F), a plus-end-directed motor, are necessary for centrosome separation (Heck et al. 1993; Zhang and Megraw 2007), and cnn ^Δ1 mutants or RNAi-mediated knockdown of Klp61F results in shallow, abnormally shaped BFs and reduced pole cell numbers (Lerit et al. 2017).

Defective centrosome separation also disrupts astral microtubule organization, which assists in the transportation of the polar granules that contain germ cell-specific mRNAs and proteins toward the cortical nuclei (Fig. 12.6a) (Lerit and Gavis 2011; Lerit et al. 2017). Polar granules migrate along the astral microtubules toward the centrosomes during interphase and remain localized around the centrosomes during mitosis until they segregate into separate cells during membrane cleavage (Fig. 12.6a) (Lerit and Gavis 2011). In gcl or Klp61f mutants where centrosomes do not properly separate and microtubules are not properly organized, polar granules still migrate from the posterior cortex, but their distribution around the nuclei is aberrant resulting in a reduced number of granules in pole cells (Lerit et al. 2017).

Microtubules are necessary for polar granule transport, as colcemid treatment prevents their movement, but actin is not as cytochalasin or latrunculin does not disrupt their transport (Lerit and Gavis 2011). In png mutants, active centrosomes travel to the posterior end without their respective nuclei (Shamanski and Orr-Weaver 1991), and polar granules are still trafficked along their astral microtubules, indicating centrosomes and astral microtubules are sufficient for not only pole cell formation but also transport of the polar granules. Cnn, Tacc, and AurA all regulate astral microtubule length and stability (Gergely et al. 2000b; Giet et al. 2002; Megraw et al. 1999), and loss of any of these three proteins results in impaired polar granules transport. Additionally, fewer pole cells are formed, and those that do form contain either a reduced level of polar granules or none at all (Lerit and Gavis 2011).

Polar granules rely on Dynein for trafficking along the astral microtubules in a minus-end-directed manner toward the centrosomes. In dhc64c mutants, astral microtubules remain intact, but the directed movement of polar granules is inhibited, resulting in a reduction of pole plasm in pole cells and reduced pole cell numbers (Lerit and Gavis 2011). Overexpression of Dynactin 2, p150 subunit (DCTN2-p150) inhibits dynein function (Burkhardt et al. 1997), resulting in cessation of polar granule motility. Conversely, the plus-end-directed motor kinesin does not function in trafficking polar granules, as kinesin heavy chain (khc) mutants display proper polar granule movement (Lerit and Gavis 2011). Therefore, the transport of polar granules relies on minus-end motors but not plus-end.

9 The Cortical Cleavage Cycles

After cortical nuclear migration, four final cleavage divisions (cycles 10–13) occur at the cortex before cellularization in interphase of cycle 14. These divisions form actin-rich furrows that resemble pole cell BFs, but the nuclei do not cleave off to form separate cells (Fig. 12.7a) (Raff and Glover 1988; Ede and Counce 1956; Turner and Mahowald 1976; Foe and Alberts 1983). Instead, these actin furrows are called pseudo-cleavage furrows (also known as cortical cleavage or Rappaport furrows). They are analogous to the furrows that form in-between adjacent nuclei from overlapping astral microtubules discovered by Rappaport in sand dollar embryos (Rappaport 1961). The centrosome regulates the formation of these furrows.

When nuclei reach the cortex during interphase of cleavage cycle 10, the centrosomes organize the plasma membrane to form a cortical bud, similar to the pole bud but not as protruding (Fig. 12.7a) (Raff and Glover 1988; Ede and Counce 1956; Turner and Mahowald 1976; Foe and Alberts 1983). Filamentous actin is rearranged from an evenly distributed layer along the cortical membrane to highly localized pockets directly over the centrosomes, termed actin caps (Fig. 12.7b) (Karr and Alberts 1986; Kellogg et al. 1988; Warn et al. 1984). Actin caps are necessary for even nuclear distribution along the cortex as disrupted cap formation due to cytochalasin treatment or mutants that affect actin organization leads to abnormal clustering of nuclei (Stevenson et al. 2001; Zalokar and Erk 1976; Callaini et al. 1992). As the nuclei enter mitosis and the centrosomes separate, the actin caps expand and distribute into furrows of invaginated membrane (Fig. 12.7a–b). As mitosis proceeds, the furrows surround the spindles and then begin to recede during telophase to form actin caps over the centrosomes of the newly divided nuclei as they proceed into the next cleavage cycle (Fig. 12.7a–b) (Karr and Alberts 1986; Kellogg et al. 1988; Warn et al. 1984).

Actin cap expansion is necessary for furrow formation and relies on a number of actin nucleators to organize actin at the furrows. The Arp2/3 complex aids in actin nucleation to create branched actin filaments through the regulation of SCAR/WAVE proteins (Pollitt and Insall 2009). Actin-related protein 3 (Arp3), a component of the Arp2/3 complex, localizes to the furrows and between actin caps dependent on the centrosomal protein Scrambled (Sced) (Stevenson et al. 2002). Scrambled localizes to the same locations as Arp3, as well as the centrosomes, independent of microtubules. In sced mutants, the actin caps do not expand preventing furrow formation, which leads to spindle fusion and chromosome segregation errors (Stevenson et al. 2001). Loss of Actin-related protein 2/3 complex, subunit 1 (Arpc1), another component of the Arp2/3 complex, displays the same phenotype as sced mutants, suggesting Sced may recruit Arp2/3 to actin cap margins to aid in actin polymerization at the furrows (Stevenson et al. 2002; Zallen et al. 2002). It is unclear how Sced may recruit Arp2/3, but further studies have revealed that SCAR also localizes to the furrows and is required for furrow assembly (Zallen et al. 2002). Therefore, Sced may interact with SCAR to recruit Arp2/3 to the furrows, implicating a role for the centrosome in actin polymerization at the furrows.

Rho1 inhibition also prevents actin cap expansion and is an upstream regulator of the formin Dia that nucleates actin filaments (Cao et al. 2010; Watanabe et al. 1997). Dia localizes between the actin caps and furrows, specifically the tips of the advancing furrows (Afshar et al. 2000), and dia mutants do not form furrows due to defective actin cap expansion (Cao et al. 2010; Afshar et al. 2000; Webb et al. 2009). Dia is also required for the localization of cytokinetic components such as MyoII, Anillin, Pnut, and Adenomatous polyposis coli 2 (Apc2) to the invaginated furrows, and between the actin caps, as they weakly localize between the caps, but are absent from furrows (Afshar et al. 2000; Webb et al. 2009).

As the cleavage furrows form, new membrane and actin is supplied through recycling endosome (RE)-derived vesicles that are localized at the centrosome and transported along astral microtubules to the growing furrow (Swanson and Poodry 1981; Mermall et al. 1994; Mermall and Miller 1995; Rothwell et al. 1999; Riggs et al. 2003). The recruitment of these vesicles relies on Nuclear fallout (Nuf), which is an adaptor protein that links Rab11 to microtubule-based motors (dynein and Kinesin-1) for trafficking on microtubules (Riggs et al. 2003). Loss of either Nuf or Rab11 disrupts vesicle-based membrane recruitment and transport of furrow components, leading to disrupted actin furrows (Riggs et al. 2003; Rothwell et al. 1999). Nuf/Rab11 complexes localize Discontinuous actin hexagon (Dah), which is required for furrow formation, to the furrows, as nuf or rab11 mutants display abnormal localization of Dah (Riggs et al. 2003; Rothwell et al. 1999; Zhang et al. 2000). Both nuf and rab11 also display disruption in RhoGEF2 localization at the furrow, and injection of active RhoA (the mammalian ortholog of Rho1) in nuf mutants rescues the furrow phenotype. However, in nuf mutants, Rho1 and Dia localization is normal, suggesting their localization does not rely on RE vesicle transport, but RhoGEF2 localization does (Cao et al. 2008).

Endocytosis at the membrane occurs from interphase until metaphase, where it is inhibited to distribute membrane from RE-derived vesicles to the furrows as they are growing. Endocytosis once again occurs during telophase as the furrows are regressing (Sokac and Wieschaus 2008a; Rikhy et al. 2015). Dynamin, encoded by shibire (shi), and Clathrin are involved in endocytic vesicle formation and localize to the cleavage furrows and between actin caps at times of invagination, while localizing to the spindles during metaphase. In shi mutants, furrow formation is disrupted, and vesicles cannot sever from the plasma membrane, while Dia, Anillin, Pnut, involved in actin remodeling, are no longer localized to the furrows (Rikhy et al. 2015).

The cleavage furrows contain similar components to cytokinetic furrows (Miller and Kiehart 1995), including the centralspindlin complex, and yet cytokinesis does not occur (Crest et al. 2012; Minestrini et al. 2003). This is due to the absence of the cytokinetic regulator of Rho1, RhoGEF Pebble (Pbl), at the furrows and central spindle in the syncytial embryo. Instead, RhoGEF2 activates Rho1 locally at the cleavage furrows. Introducing ectopic active RhoA into the embryo during the cortical cleavage stages induces cytokinetic furrow formation over the central spindle, indicating that the machinery is in place but that spatial Rho1 is deterministic of the site of furrow formation (Crest et al. 2012). Rho1 and another major actin regulator Cdc42 play an antagonistic role in furrow formation. Constitutively active Cdc42 or dominant negative Rho1 both disrupt actin furrows and MyoII localization, but the microtubules remain intact (Crawford et al. 1998).

Centrosomes are sufficient for actin rearrangement and furrow ingression. In aphidicolin-injected embryos where only the centrosomes reach the cortex, cortical buds are formed, and actin is rearranged in a cell cycle-dependent manner (Raff and Glover 1989; Yasuda et al. 1991). Microtubules are required for actin reorganization in a specific time window. Colchicine injection during anaphase disrupts actin rearrangement to the furrows, but injection during interphase or telophase does not disrupt actin rearrangement. During telophase, robust astral microtubules of neighboring centrosomes overlap, possibly defining the furrow position for the subsequent cleavage cycle (Riggs et al. 2007). The polar bodies lack centrosomes but form microtubule projections similar to the astral microtubules of the centrosome and when the polar bodies lie close to the cortex, they rearrange the cortical actin (Foe et al. 2000).

There is also an accumulation of actin at the centrosomes that form aster-like filaments as the nuclei divide, appearing to emanate from the centrosome similar to astral microtubules (Fig. 12.7a). These actin asters appear to be reliant on proper centrosome activity and microtubules, as cnn mutants that disrupt the PCM and astral microtubules, or colchicine treatment also disrupts actin aster formation (Riparbelli et al. 2007). However, the role of actin asters in cortical cleavage divisions remains unclear.

The furrows provide a physical barrier between the dividing nuclei that prevents centrosomes from interacting with the spindles of neighboring nuclei (Kotadia et al. 2010; Sullivan et al. 1993b). When furrows are disrupted, mitotic spindles of neighboring nuclei aberrantly interact, which causes centrosomes to dissociate from their respective nuclei. The nuclei recede into the embryo leaving a patch devoid of a nucleus at the cortex (Sullivan et al. 1993a; Sullivan et al. 1993b). This process, which occurs in multiple mutant backgrounds when cleavage furrows are disrupted, is known as nuclear fallout. cnn, arpc1, scar, and dia mutants all display fused spindles and nuclear fallout due to the absence of furrow formation (Stevenson et al. 2002; Zallen et al. 2002; Afshar et al. 2000; Megraw et al. 1999).

The mechanism of nuclear fallout is a protective checkpoint that prevents damaged nuclei from multiplying and integrating into the future embryo by removing them from the cortex (O’Farrell et al. 2004). Due to the absence of gap phases and decrease in transcription, the syncytial embryo has an altered response to cell cycle checkpoint mechanisms. For example, DNA replication arrest due to aphidicolin injection does not block nuclei from entering mitosis (Raff and Glover 1988; Foe and Alberts 1983). This is especially important during the cortical cleavage cycles, due to the increase in cell cycle length and the MZT.

During cleavage cycles 10–13, the cell cycle increases in length from roughly 8 to 21 min (Foe and Alberts 1983). The increase in transcription that starts during cleavage cycle 10 requires a longer cell cycle, especially in S phase (Lamb and Laird 1976; McKnight et al. 1977; Zalokar 1976; Edgar and Schubiger 1986; Shermoen et al. 2010). The increase in length during these cycles is reliant on grapes (grp, Chk1 homolog) and meiotic 41 (mei-41, ATR homolog) as grp or mei-41 mutants the lengthening of the cleavage cycles fails (Sibon et al. 1997; Sibon et al. 1999). As a result, grp or mei-41 mutants also arrest during the cleavage division cycles with damaged nuclei and nuclear fallout (Fogarty et al. 1994; Fogarty et al. 1997). Centrosomes lose their function indicated by the loss of γ-Tubulin and γTuRC components from the centrosomes and the inability to separate chromosomes during mitosis (Sibon et al. 2000). Centrosome inactivation and nuclear fallout also occur during aphidicolin injection as well as a treatment with a variety of DNA-damaging agents (Raff and Glover 1988; Sibon et al. 2000; Takada et al. 2003).

This pathway of centrosome inactivation is regulated by loki (Lok, also known mnk, a Chk2 homolog), which localizes to centrosomes and spindles. During DNA damage, this localization increases, and Lok also accumulates at the nuclei (Takada et al. 2003). Lok causes mRNA nuclear retention after DNA damage, including mRNAs that encode for centrosomal proteins, such as γ-Tubulin ring protein 91 (Grip91); this mRNA nuclear retention causes nuclear fallout (Iampietro et al. 2014). In lok mutants, when DNA is damaged through either Bleomycin (induces DNA damage) injection or in a grp mutant background, centrosomes do not inactivate (Takada et al. 2003).

Multiple PCM proteins also play an important role in regulating cleavage furrow formation, and their dysfunction can result in fused spindles, aneuploid nuclei, and nuclear fallout. cnn mutants are maternal-effect lethal, arresting in the cleavage furrow stages and displaying a failure in furrow ingression that leads to fused metaphase spindles and colliding nuclei in telophase and inevitable nuclear fallout (Megraw et al. 1999; Vaizel-Ohayon and Schejter 1999). In cnn ^Δ1 mutants where astral microtubules can still form, some furrows still ingress (Zhang and Megraw 2007), but in CM2 domain cnn mutants (cnn ^b4), astral microtubules are present and furrow ingression was severely impaired. The CM2 domain of Cnn interacts with Centrocortin (Cen), and cen mutants display aberrant actin organization at the furrows as well as spindle fusions, indicating it interacts with cnn to regulate actin organization at the cleavage furrow (Kao and Megraw 2009).

sponge (spg, a Rho GEF family protein) mutants have defective actin caps and furrows leading to aberrant spindle interactions (Postner et al. 1992; Riparbelli et al. 2007). Loss of Eb1, which binds the plus ends of microtubules, does not result in defective actin localization at the furrows. Instead, furrows are partially invaginated, resulting in severe spindle defects and loss of nuclei due to nuclear fallout (Rogers et al. 2002; Webb et al. 2009). Nuf, which supplies membrane and other components to the invaginating furrows, localizes to the centrosome during prophase dependent on microtubules (Rothwell et al. 1998; Riggs et al. 2003). Because REs display a pericentriolar accumulation, this localization of Nuf is important for vesicle transport. nuf mutants display incomplete actin furrows, which results in spindle fusions (Rothwell et al. 1998).

10 Centrosome Separation During the Cortical Cleavage Cycles

As nuclei enter mitosis, the centrosomes begin to separate at prophase to form bipolar spindles. Centrosome separation is especially important in the cortical cleavage cycles due to the membrane invagination and spreading of actin that is orchestrated in conjunction with centrosome movement. It was hypothesized that centrosome separation guides actin cap expansion and furrow invagination as multiple centrosomal components are necessary for proper actin reorganization. Free centrosomes are sufficient to reorganize actin at the membrane (Raff and Glover 1989; Yasuda et al. 1991), and the cycling of actin structures parallels centrosomes’ movements (Karr and Alberts 1986). However, in colchicine-treated embryos, centrosome separation fails, but actin caps still expand (Stevenson et al. 2001; Cao et al. 2010). Therefore, it appears that actin spreading may guide the centrosomes, as either latrunculin, cytochalasin, or jasplakinolide (actin filament stabilizer) injection both inhibit centrosome separation (Cao et al. 2010; Stevenson et al. 2001). Jasplakinolide also shrinks the actin caps, indicating actin turnover is required for centrosome separation (Cao et al. 2010).

It is hypothesized that the astral microtubules of centrosomes interact with actin to allow for proper centrosome separation. In support of this, Apc2, which stabilizes microtubules with Eb1 through Dia (Wen et al. 2004) and interacts with actin at actin-microtubule interaction sites (McCartney and Peifer 2000), plays a role in centrosome separation. Apc2 localizes to the actin caps and furrows where astral microtubules are interacting with the actin furrows (McCartney et al. 2001; Webb et al. 2009). apc2 mutants display centrosome separation defects as well as defective furrow formation (Webb et al. 2009; Buttrick et al. 2008). Apc2 localization is reliant on Akt1 and Shaggy (Sgg, a Zw3 homolog), which is upstream of Akt1 (Shaw et al. 1997). Akt1 mutants have normal actin and microtubule organization, yet they display centrosome separation defects and nuclear fallout due to disruption in Apc2 localization. Introduction of one copy of a sgg mutant allele into akt1 mutants rescues this phenotype. Therefore, Apc2 regulates centrosome separation through microtubule-actin interactions, dependent on Akt1 and Sgg (Buttrick et al. 2008).

αTub67C is enriched at the interpolar microtubules that embrace the nuclear envelope, and αTub67C mutants display shorter microtubules. αTub67C mutants arrest in mitosis due to defective centrosome separation. These interpolar microtubules must rapidly assemble, and αTub67C appears to be necessary for this rapid growth as cells depleted of αTub67C will eventually grow long microtubules from other isoforms of α-Tubulin. Therefore, αTub67C is hypothesized to quickly nucleate these interpolar microtubules to push the centrosomes apart during centrosome separation (Venkei et al. 2006).

Centrosome separation also relies on multiple microtubule-based motor proteins to provide the force that moves them apart. Dynein is required for proper centrosome separation, as dhc64c mutant embryos display centrosomes that do not fully separate (Robinson et al. 1999). Klp61f is required to keep the centrosomes separated during metaphase but is not required for their initial separation. klp61f mutants display normal centrosome separation in prometaphase, but the centrosomes slide back together before the metaphase spindles can form. However, in ncd klp61f double mutant embryos, the centrosomes separate and remain separated until telophase indicating an opposing force on the minus- and plus-end-directed motors, respectively, that keeps the metaphase centrosomes separated. However, in these double mutant backgrounds, the distance between daughter nuclei is abnormally short, indicating a role for these motors in internuclear spacing as well as centrosome separation (Sharp et al. 1999).

During centrosome separation, Cnn fibers, hypothesized to be intercentrosomal microtubules, connect the centrosomes and persist into late anaphase, while cnn ^Δ1 mutants display defects in centrosome separation (Zhang and Megraw 2007). In sced or sponge mutants, where furrows do not form, centrosome separation is also delayed (Stevenson et al. 2001; Postner et al. 1992). arpc1 or dia mutants as well as Rho1 inhibition by C3 exotransferase, which display abnormal furrow formation, all result in defective centrosome separation (Cao et al. 2010). Loss of Daughterless-like (Dal), a protein of unknown function, also results in defective centrosome separation resulting in spindle fusion and nuclear fallout, but only during the cortical cleavage cycles (Sullivan et al. 1990).

In summary, centrosome separation in the early embryo is necessary for proper cleavage furrow formation during the cortical cleavage cycles. Centrosome separation is reliant on actin, microtubules, microtubule motors, and a variety of proteins involved in cleavage furrow formation. The mechanisms that force the centrosomes apart during mitosis still remain unclear, but the variety of factors involved suggest it is a complicated process that requires further investigation.

11 Cellularization Transitions the Syncytial Embryo to the Cellular Blastoderm

After 13 successive cleavage divisions, the cortical nuclei remain in interphase for roughly 70 min as cellular membranes surround each nucleus to form the multicellular embryo. The process of cellularization begins identically to the cortical cleavage cycles: the membrane protrudes forming a cortical bud and actin cap above each nucleus and its associated pair of centrosomes (Fig. 12.8a) (Foe and Alberts 1983). The actin cap is localized to microvillar projections that densely decorate the cortical buds at the onset of cellularization (Fig. 12.8a) (Turner and Mahowald 1979; Fullilove and Jacobson 1971). Furrows begin to form, and the hairpin-shaped tip of each furrow, the furrow canal (FC), is enriched in components necessary for cellularization (Fig. 12.8b) (Fullilove and Jacobson 1971; Warn and Robert-Nicoud 1990; Young et al. 1991). Immediately after furrow ingression, basal cell junctions assemble below the FC composed of E-cadherin, α-catenin, and β-catenin in preparation for cell formation (Hunter and Wieschaus 2000; Müller and Wieschaus 1996).

As the furrow ingresses, the nuclei elongate and extend into the embryo, resulting in oblong nuclei (Lecuit and Wieschaus 2000; Knoblich 2000). This change in nuclear shape is due to a basally extending microtubule basket formed by the astral microtubules (Fig. 12.8b–e) (Callaini and Anselmi 1988; Kellogg et al. 1991). Nocodazole treatment prevents nuclear shape change, supporting a role for these microtubule baskets in nuclear elongation during furrow ingression (Brandt et al. 2006). These microtubule baskets also rely on actin, as cytochalasin treatment disrupts microtubule basket organization (Edgar et al. 1987).

Unlike the cortical cleavage cycles, cellularization requires the nuclei to be present as aphidicolin-injected embryos do not cellularize (Raff and Glover 1989). This is most likely due to the requirement of zygotic transcription that starts during the cortical cleavage cycles and decreases at the end of cellularization (Lamb and Laird 1976; McKnight et al. 1977; Zalokar 1976; Edgar and Schubiger 1986) as α-amanitin injection (a transcription inhibitor) blocks cellularization (Edgar et al. 1986). Most gene products required for cellularization are maternally supplied but some necessary proteins are zygotically transcribed, indicating they have a specific role in this process (Mazumdar and Mazumdar 2002).

The furrows slowly progress until they reach the basal end of the nuclei, where progression then rapidly increases with furrows ultimately reaching a depth of approximately 35 μm (Lecuit and Wieschaus 2000; Foe and Alberts 1983). An actomyosin contractile ring forms at the FC as it invaginates and begins to contract once the furrows have passed the nuclei. This results in a change from the hexagonal array of actin that surrounds the nuclei to a ring shape localization in preparation for cell closure (Fig. 12.8f–g) (Theurkauf 1994). The contractile rings then pinch off the membranes at the end of cellularization to form single cells (Fig. 12.8e).

Because furrow invagination is so extensive and demanding of membrane resources, the membrane present at the cortex does not provide a sufficient supply needed for cellularization, and new sources of membrane are required (Figard et al. 2013). Two different deposits of membrane are supplied to the furrows during different time points of cellularization.

The first deposit is derived from the microvillar actin projections on the surface of the cortex above each nucleus (Fig. 12.8a–d). These microvillar projections assemble before cellularization, and are depleted by its completion (Fabrowski et al. 2013; Fullilove and Jacobson 1971). It is estimated that the microvilli contain about half of the membrane necessary for furrow formation (Figard et al. 2013). New membrane is added at the apical end of the furrows and old membrane is pushed basally into the furrows, rather than the addition of membrane to the FC, supporting the role for microvillar membrane addition (Lecuit and Wieschaus 2000). Microvillar depletion also mimics the kinetics of furrow ingression, starting slow as the furrows reach the bottom of the nuclei and speeding up toward the end of cellularization. Therefore, the microvilli are unfolded and pulled directly into the invaginating furrows (Figard et al. 2013).

Abl tyrosine kinase (Abl) is involved in the regulation of microvillar density and length as abl mutants display longer microvilli than wildtype at the onset of cellularization which do not diminish over time as in wildtype. Actin also abnormally accumulates at the apical end of the furrows due to enrichment of the actin nucleators, Arp3 and Dia, resulting in excessive F-action nucleation and abnormal furrow formation. These defects are due to abnormal localization of Enabled (Ena) at the apical cortex and abnormal actin accumulation, which is normally disperse during cellularization, as ena abl mutants have less cortical actin accumulation. abl mutants also display disrupted microtubule baskets, a further indication that actin regulation plays a role in their formation (Grevengoed et al. 2003).

The second source of membrane is supplied by Golgi-derived vesicles and REs. Centrosomes assist in trafficking these vesicles which contain the proteins and membrane needed for furrow invagination along the microtubules (Mazumdar and Mazumdar 2002). Centrosomes are necessary for furrow invagination and loss of astral microtubules disrupts furrow invagination (Zalokar and Erk 1976; Foe and Alberts 1983). During early cleavage cycles, the Golgi is localized in disperse puncta at the cortex of the embryo in close proximity to the ER (Ripoche et al. 1994; Stanley et al. 1997). Once the nuclei reach the cortex and transcription increases, separate ER and Golgi structures segregate to single nuclei, with each nucleus containing a single ER/Golgi system. This segregation is dependent on microtubules, as nocodazole treatment disrupts ER/Golgi compartmentalization, indicating a role for the centrosome (Frescas et al. 2006). Golgi-derived vesicles either fuse to the apical membrane and endocytose to be sorted by the RE for transport, or traffic directly to the membrane (LaLonde et al. 2006; Lee and Harris 2014). Golgi-derived vesicle and RE transport rely on separate trafficking pathways to transport different components to the invaginating furrows.

Golgi-derived vesicles supply membrane and components that are required for the rapid extension of the furrows. Brefeldin A, which inhibits Golgi vesicle transport, inhibits furrow progression in the final, fast stage of cellularization (Sisson et al. 2000; Frescas et al. 2006). Transport of these vesicles is dependent on microtubules, as colcemid or colchicine treatment during the slow cellularization stage stalls vesicle movements at the basal end of the nuclei, preventing furrow invagination (Lecuit and Wieschaus 2000; Sisson et al. 2000; Foe and Alberts 1983; Zalokar and Erk 1976). Golgi vesicle transport is not dependent on actin as cytochalasin injection does not disrupt Golgi vesicle movement during cellularization (Sisson et al. 2000).

The dynein-dynactin complex in association with Cytoplasmic linker protein 190 (CLIP190), which links vesicles to microtubules (Lantz and Miller 1998), assists in transporting Golgi-derived vesicles along the microtubules. dhc64C mutants do not form furrows and Golgi-derived vesicle movement is blocked, while the microtubules remain intact (Papoulas et al. 2005). Dhc64C, DCTN2-p150, and CLIP190 all associate at the Golgi and specifically bind the Golgi-associated protein Lava Lamp (Lva), which acts as an adaptor for Dynein-Dynactin vesicular trafficking (Sisson et al. 2000; Papoulas et al. 2005). In support of this, lva mutants that cannot bind Dhc64C, DCTN2-p150, or CLIP190 display impaired furrow progression and inhibition of Golgi vesicle movement. Lva is necessary for CLIP190-dependent microtubule-vesicle attachment, as lva mutants that cannot bind dynein inhibit CLIP190 localization to the Golgi and FC, resulting in impaired furrow formation (Papoulas et al. 2005).

The second class of vesicles, REs, are necessary for all stages of cellularization. REs localize intermediately near the centrosomes as the vesicles are being sorted to transport membrane and protein to the furrows (van Ijzendoorn 2006). Rab5 and Dynamin are necessary for the initial endocytosis and budding of REs from the apical membrane, as overexpressing a dominant negative variant of Rab5 or shi mutant backgrounds display impaired furrow ingression (Pelissier et al. 2003). An intermediate stage of the RE pathway displays tubular membrane projections from the FC, but only at the onset of cellularization (Fig. 12.8a) (Sokac and Wieschaus 2008a). In shi mutants, these endocytic tubules are longer than normal, indicating stalled trafficking of REs (Sokac and Wieschaus 2008a; Su et al. 2013; Sherlekar and Rikhy 2016).

The regulation of the REs follows similar dynamics to RE trafficking during the cortical cleavage cycles. Rab11, Dynamin, and Nuf are required for vesicle trafficking and dominant negative Rab11, shi, and nuf mutants all display impaired furrow invagination, while shi mutants display Rab11 vesicles halted at the centrosome (Pelissier et al. 2003; Rothwell et al. 1998). Rab11 and Nuf also localize RhoGEF2 to the furrows to drive furrow invagination through the same mechanisms outlined in the cortical cleavage cycles section. (Riggs et al. 2003; Cao et al. 2008).

Rab11, Nuf, and Dynamin regulate the localization of Slow as molasses (Slam), which is both maternally supplied and zygotically transcribed, to the FCs (Acharya et al. 2014). Slam is necessary for Rho1, RhoGEF2, and MyoII localization to the furrows and slam mutants display disrupted furrow invagination (Wenzl et al. 2010; Acharya et al. 2014). Centrosomes are also necessary for Slam localization to the furrows as nuclei-less centrosomes still localize Slam, and ablating centrosomes disrupts Slam localization (Acharya et al. 2014). Therefore, Slam utilizes centrosome-dependent RE vesicle transport for localization at the FCs.

Slam plays a redundant role with Nullo, another zygotically transcribed gene product that is necessary for furrow invagination and stabilization, in Dia localization (Sokac and Wieschaus 2008a, b; Hunter and Wieschaus 2000). slam nullo double mutants, but not single mutants of either, display disruption of Dia localization at the FCs (Acharya et al. 2014). Dia localization is also dependent on RhoGEF2, both of which localize to the FCs before invagination, suggesting they aid in FC formation. Mutants for rhogef2 and dia display areas absent of furrows and misshapen FCs. When either rhogef2 or dia mutants are combined with nullo mutants, the furrow defects are stronger, suggesting Nullo acts in a pathway separate from RhoGEF2 and Dia (Großhans et al. 2005). Therefore, Nullo may coordinate furrow invagination through F-actin regulation by Slam, RhoGEF2, and Dia.

Rho1, Rok, and RhoGEF2 are all required for the final stage of cellularization. RhoGEF2 localizes Rho1 to the furrows for localized Rok activation (Padash Barmchi et al. 2005). rhogef2 and rok mutants display similar defects in cellularization, defective furrow invagination (Dawes-Hoang et al. 2005), and dominant negative rho1 mutants do not cellularize (Crawford et al. 1998). The Rho1 pathway regulates MyoII activation, which is necessary for contractile ring constriction (Xue and Sokac 2016). Anillin is known to link Rho1, actin, and MyoII during cytokinesis (Piekny and Glotzer 2008) and is necessary for MyoII localization as well as Pnut localization to the FCs. In anillin mutants, FC morphology is abnormal, contractile rings fail to form, and cellularization does not occur, indicating it may link Rho1, actin, and MyoII during cellularization (Field et al. 2005; Thomas and Wieschaus 2004).

Bottleneck (Bnk), a zygotically transcribed gene product, regulates the constriction of the contractile ring toward the end of cellularization. Bnk localizes to the FCs during the slow phases of cellularization and disappears during the fast stage when constriction begins. In bnk mutants, the hexagonal actin rings constrict before the furrows are past the nuclei resulting in bottle-shaped nuclei (Schejter and Wieschaus 1993; Theurkauf 1994). Therefore, bnk is a negative regulator of actomyosin constriction during cellularization.

For more on Drosophila cellularization, see How one becomes many: blastoderm cellularization in Drosophila melanogaster (Mazumdar and Mazumdar 2002).

12 Summary

The centrosome plays an important role in regulating the dynamics of early development in the Drosophila syncytial embryo at multiple stages. The centrosome regulates the cytoskeletal elements, microtubules and actin, in varying contexts to allow for each step of embryogenesis to properly occur. The first three centrosome-dependent stages of embryogenesis are nuclear migrations that utilize cytoskeletal components. The remaining three stages utilize centrosome-dependent actin regulation to form furrows that separate the nuclei as they divide and eventually form cellular membranes. Defective centrosomes can disrupt the cell cycle, mitotic cleavage divisions, and nuclear positioning, resulting in morphogenic defects and embryonic lethality. The syncytial embryonic cell cycle is modified to address the rapid nuclear divisions, resulting in changes to the regulation of the centrosome. The central roles of the centrosome in the early embryo contrast with the ability of zygotic development to be accomplished successfully without functional centrosomes. All of the processes controlled by the centrosome in the early embryo are not well understood, so much remains for investigators to discover.

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Department of Biomedical Sciences, College of Medicine, Florida State University, Tallahassee, FL, USA
Caitlyn Blake-Hedges & Timothy L. Megraw

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Department of Surgery, Houston Methodist Hospital, Houston, TX, USA
Malgorzata Kloc

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Blake-Hedges, C., Megraw, T.L. (2019). Coordination of Embryogenesis by the Centrosome in Drosophila melanogaster . In: Kloc, M. (eds) The Golgi Apparatus and Centriole. Results and Problems in Cell Differentiation, vol 67. Springer, Cham. https://doi.org/10.1007/978-3-030-23173-6_12

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DOI: https://doi.org/10.1007/978-3-030-23173-6_12
Published: 22 August 2019
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Coordination of Embryogenesis by the Centrosome in Drosophila melanogaster

Abstract

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Asymmetrically dividing Drosophila neuroblasts utilize two spatially and temporally independent cytokinesis pathways

Link of Zygotic Genome Activation and Cell Cycle Control

Reshaping the Syncytial Drosophila Embryo with Cortical Actin Networks: Four Main Steps of Early Development

1 The Development of the Syncytial Embryo: Six Key Steps

2 The Structure of the Embryonic Centrosome and Regulation of Microtubule Assembly

3 Fertilization and the First Zygotic Division

4 The Syncytial Embryo Employs an Adapted Cell Cycle

5 Centrosome-Nucleus Association

6 Axial Nuclear Migration Distributes Nuclei along the A-P Axis

7 Cortical Nuclear Migration Positions the Nuclei at the Cortex

8 Pole Cells Cellularize Before the Other Nuclei

9 The Cortical Cleavage Cycles

10 Centrosome Separation During the Cortical Cleavage Cycles

11 Cellularization Transitions the Syncytial Embryo to the Cellular Blastoderm

12 Summary

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Coordination of Embryogenesis by the Centrosome in Drosophila melanogaster

Abstract

Similar content being viewed by others

Asymmetrically dividing Drosophila neuroblasts utilize two spatially and temporally independent cytokinesis pathways

Link of Zygotic Genome Activation and Cell Cycle Control

Reshaping the Syncytial Drosophila Embryo with Cortical Actin Networks: Four Main Steps of Early Development

1 The Development of the Syncytial Embryo: Six Key Steps

2 The Structure of the Embryonic Centrosome and Regulation of Microtubule Assembly

3 Fertilization and the First Zygotic Division

4 The Syncytial Embryo Employs an Adapted Cell Cycle

5 Centrosome-Nucleus Association

6 Axial Nuclear Migration Distributes Nuclei along the A-P Axis

7 Cortical Nuclear Migration Positions the Nuclei at the Cortex

8 Pole Cells Cellularize Before the Other Nuclei

9 The Cortical Cleavage Cycles

10 Centrosome Separation During the Cortical Cleavage Cycles

11 Cellularization Transitions the Syncytial Embryo to the Cellular Blastoderm

12 Summary

References

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