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Ascidians as excellent models for studying cellular events in the chordate body plan.

Abstract. The larvae of non-vertebrate chordate ascidians consist of countable numbers of cells. With this feature, ascidians provide us with excellent models for studying cellular events in the construction of the chordate body. This review discusses the recent observations of morphogenetic movements and cell cycles and divisions along with tissue specifications during ascidian embryogenesis. Unequal cleavages take place at the posterior blastomeres during the early cleavage stages of ascidians, and the structure named the centrosome-attracting body restricts the position of the nuclei near the posterior pole to achieve the unequal cleavages. The most-posterior cells differentiate into the primordial germ cells. The gastrulation of ascidians starts as early as the 110-cell stage. During gastrulation, the endodermal cells show two-step changes in cell shape that are crucial for gastrulation. The ascidian notochord is composed of only 40 cells. The 40 cells align to form a single row by an event named the convergent extension, and then the notochord cells undergo vacuolation to transform the notochord into a single hollowed tube. The strictly restricted number of notochord cells is achieved by the regulated number of cell divisions coupled with the differentiation of the cells conducted by a key transcription factor, Brachyury. The dorsally located neural tube is a characteristic of chordates. During the closure of the ascidian neural tube, the epidermis surrounding the neural plate moves toward the midline to close the neural fold. This morphogenetic movement is allowed by an elongation of interphase in the epidermal cell cycles.

Introduction: Embryogenesis of Ascidians Described at the Cellular Level

A primary goal in the field of developmental biology is to determine how cell behaviors are controlled to form highly organized multicellular organisms. Cells, which have the ability to behave individually, act in a highly coordinated manner to maintain the order of multicellular systems. Regarding the mechanisms underlying this coordination, the regulation of cellular processes such as the cell cycle, cell division, and morphogenetic movement of multicellular bodies must be studied together with cell differentiation. For analyses of cellular processes during development, it is helpful to use species that have the least number of cells, because small numbers of cells allow observation of events that involve every cell. Ascidians provide us with excellent models for cell-by-cell analyses during embryogenesis because of the small number of cells in their embryos and larvae.

Ascidians are members of the subphylum urochordates, which is a group of the phylum chordates along with cephalochordates and vertebrates (Satoh, 2003). As the phylogenetic position suggests, ascidians possess the basic chordate body plan (Fig. 1A-C). The larvae of most ascidians are tadpoles, which have a notochord in the tail (the name "urochordate" was derived from this feature) and a dorsally located and hollowed neural tube (Fig. 1A). The tadpole body of many ascidian larvae with a low number of cells is extraordinarily simple compared to those of vertebrates. Typically, the fates of most cells are restricted to a single type at the 64- to 110-cell stage (Fig. 1D). Gastrulation starts as early as the 110-cell stage, neurulation is initiated in embryos with about 500 cells, and the tadpole larva is composed of approximately 3000 cells.

In several ascidians such as Halocynthia roretzi and Ciona intestinalis, cell lineages during embryogenesis have been described in detail (Fig. 1E: Kumano and Nishida. 2007). The cell lineages give us a set of basic information about the cellular events of embryogenesis--namely, when and how many times cells are divided and the timing of the cells' differentiation. Because the mechanisms of differentiation of cells have been well documented in reviews (e.g., Nishida, 2005; Lemaire, 2009), in this article we review the recent studies of ascidians that focus on the behaviors of cells during embryogenesis.

Unequal Cleavage of the Posterior Blastomere and Formation of the Germ Cells

Ascidian embryos show unequal cleavage as early as the 8-cell stage (Fig. 2A). The cleavage plane of the third cell division is somewhat closer to the animal pole. This results in smaller animal cells and bigger vegetal cells. A pair of vegetal blastomeres named B4.1 are the cells forming the posterior part of the body. The B4.1 pair and its daughter cells perform unequal cleavages during the three successive cleavages (Fig. 2B). The daughter cells located at the posterior side become smaller than their anterior sisters. The anteriorly located daughter cells cleave equally in the following cleavages, while the posterior daughter cells undergo further unequal cleavages, suggesting that the machinery for the unequal cleavages is inherited by the posterior daughter cells. Indeed, embryos that have gotten rid of the posterior cytoplasm show equal cleavage (Nishida, 1994), suggesting that the posterior cytoplasm is responsible for the unequal cleavage.

The centrosome-attracting body (CAB) is the structure that attracts the nuclei to the posterior pole (Fig. 2C; Hibino et al., 1998; Nishikata et al., 1999). Clusters of the microtubules are observed between the CAB and the nuclei, and the microtubule clusters are thought to anchor the nuclei and spindle to the posterior pole to restrict their position. The CAB also functions as the localizing center of maternal mRNAs (Fig. 2C). Several tens of mRNAs are known to be localized at the CAB (Yamada, 2006; Prodon et al., 2007). The proteins encoded by the localized mRNAs function in the determination of cell differentiation, spindle attraction, and localization of mRNAs (Yoshida et al., 1996; Nishida and Sawada, 2001; Nakamura et al., 2005; Negishi et al., 2007). The localized mRNA includes vasa mRNA, which is a well-known marker of the primordial germ cells (Schup-bach and Wieschaus, 1986; Fujimura and Takamura, 2000). Indeed, the CAB region contains an electron-dense matrix resembling the germ plasm (Iseto and Nishida, 1999), and the posterior daughter cells express VASA protein and are thought to differentiate into germ cells (Shirae-Kurabayashi et al., 2006). The formation of the germ cells at the poste-riormost region of embryos is also observed in protostomes, suggesting that the manner of germ cell formation is conserved among protostomes and deuterostomes.

The primordial germ cells of ascidians are located in the tail at the larval stage (Shirae-Kurabayashi et al., 2006). The tail is regressed during metamorphosis, and most of the tail cells are lost from the juvenile body (Chambon et al., 2002). An exception is the primordial germ cells; these cells survive after tail regression and migrate into the gonadal rudiments (Okada and Yamamoto, 1999; Shirae-Kurabayashi et al., 2006).

Confocal Microscopy

Confocal microscopy is a powerful technique for observing cell morphology in detail (see Fig. 3C). For example, confocal microscopy provided a detailed cellular description of the middle tailbud embryos of Ciona intestinalis. Accurate numbers of the cells in each tissue were determined, and novel cells whose presence was previously not known were discovered (Nakamura et al., 2012). Two databases of the confocal images of embryos at each developmental stage have been constructed for this ascidian (Tassy et al., 2006; Hotta et al., 2007a; Nakamura et al., 2012): Aniseed ( and FABA (Hotta et al., 2007b). These databases provide us with a standard for the time course of the developmental events. In addition, researchers can now perform "in silico" observations of the embryos to look for novel phenomena without the labor of retrieving the confocal images themselves.

An outstanding achievement of the confocal microscopy studies is the quantification of the cell volume and superficial area based on the retrieved cell images. For example, the difference in the cell volume between sister cells suggests the occurrence of the unequal cleavages (Tassy et al., 2006). The calculation of the contact areas between cells is crucial for understanding the induction during cell fate decisions (Tassy et al., 2006). At the 32-cell stage, some of the animal blastomeres differentiate into the anterior neural cells, while others have an epidermal cell fate. The specification of the anterior neural cells requires an inductive signal from the A-line blastomeres (A-line blastomeres are the cells located at the anterior half of the vegetal hemisphere, derived from A4.1 cells at the 8-cell stage; Figs. 1D and 2A) situated at the vegetal hemisphere (Bertrand et al., 2003). All of the animal cells have the potential to differentiate into anterior neural cells (Hudson and Lemaire, 2001), and many of them have contact with the inducing vegetal cells. However, only some of the animal cells undergo the anterior neural fate. The calculation of the the area of contact between the animal cells and A-line vegetal cells has explained why the restricted animal cells differentiate into the anterior new-al cells; the animal cells fated to be the neural cells have a larger area of contact between the A-line cells compared to the epidermally fated cells. Therefore, the strength of the inductive signals may be determined by the mass of the contact surface, and the recipient cells receiving a signal that surpasses the threshold may have an anterior neural fate. This means that the differentiation of the cells is not accomplished by a simple all-or-none mechanism. Future studies must address (a) the quantity of the signals necessary for converting the cells into a specific type, (b) how the signals that surpass the threshold are maintained or strengthened in the recipient cells, and (c) how the effect of the signals below the threshold is erased.

Cellular and Molecular Mechanisms of Gastrulation

Gastrulation is the first major morphogenetic movement during embryogenesis for forming the digestive tube. In ascidians, gastrulation starts as early as the 110-cell stage. The endodermal cells and mesodermal cells located in the vegetal half of embryos invaginate into the animal region to form the gut. During gastrulation, the shapes of the moving cells are changed dramatically. The shape change is thought to create the force to bend the tissue inward toward the body to accomplish gastrulation. Sherrard et al., (2010) used confocal microscopy to measure the shape change of the endodermal cells during gastrulation to make a model for calculating the force exerted upon the cells. Their study divided the onset of gastrulation into two steps. In Step 1, the apical surface of the endoderm that covers the major part of the vegetal half before gastrulation shrinks, while the whole vegetal hemisphere flattens. In Step 2, the endodermal cells shorten their apical-basal axis by extending their basal plane, and the vegetal cell plate invaginates toward the animal pole.

The shrinkage of the apical surface of the endoderm in Step 1 requires RhoA-mediated phosphorylation of myosin. In contrast, the shortening of the apical-basal axis of the endoderm in Step 2 is largely independent of the RhoA pathway. Gastrulation was simulated by estimating the force at the cell membrane within a cell and between cells. Changing the force parameter can change the estimated embryonic structure during gastrulation. The parameter sets that fit the actual morphological changes support the two-step mechanism of gastrulation. The simplicity of ascidian embryos is an advantage for such simulation analyses, because quite a few parameters are necessary for constructing the simulation.


The notochord of ascidians, which is present at the center of the tail (Fig. 3A), is extraordinarily simple compared to those of vertebrates. The notochord of most ascidians invariably consists of 40 cells (Satoh 1994). Their cell division is finished by the neurula stage (Miyamoto and Crowther, 1985; Munro and Odell, 2002a; Jiang et al., 2005). This simple notochord enables us to observe events of notochord formation at single-cell resolution. Here we discuss recently obtained knowledge about the cellular events associated with formation of the notochord, namely cell division control, morphogenetic movement, and vacuole formation.

The invariant number of the notochord cells suggests that their cell division number is strictly regulated. At the 64-cell stage, four A-line blastomeres are restricted to the notochord cell fate. These cells divide three times after the 64-cell stage, and then they enter the quiescent stage. Likewise, two B-line blastomeres (B-line blastomeres are the cells located in the posterior half of the vegetal hemisphere, and they are derived from B4.1 cells at the 8-cell stage; Figs. 1D and 2A) at the 110-cell stage are destined to differentiate into notochord cells. The notochord cells further divide twice, and their cell division then ceases. The A-line and B-line notochord cells are respectively called primary and secondary notochord cells. The 32 primary and 8 secondary notochord cells are located at the anterior and the posterior side of the notochord, respectively (Nishida, 1987).

The fixed cell division number of notochord cells is thought to be determined in accordance with their differentiation (Fujikawa et al., 2011). Differentiation of ascidian notochord cells is induced by fibroblast growth factor (FGF) signaling and the key transcription factor gene brachyury (Yasuo and Satoh, 1994, 1998; Nakatani et al., 1996; Corbo et al., 1997). Expression of brachyury is induced by FGF signaling from the endodermal cells adjacent to the notochord precursor cells (Nakatani et al., 1996; Nakatani and Nishida, 1997; Imai et al., 2002; Yasuo and Hudson, 2007). Disrupting FGF or brachyury functions results in the conversion of the notochord-fated cells into neural cells, whereas the artificial treatment of neural cells with FGF or the forced expression of brachyury in the neural cells results in the cells becoming notochord cells (Yasuo and Satoh, 1998; Kumano and Nishida, 2007).

In Ciona, the number of times a cell divides is strictly determined by the cell's fate. Neural cells that are treated with FGF and those in which brachyury is overexpressed divide three times after the 64-cell stage, and then they do not divide further. In contrast, notochord cells that are artificially converted into neural cells divide more than three times (usually five times) after the 64-cell stage. Therefore, FGF and brachyury may express the downstream cell-cycle regulators to control the cell division number of notochord cells. Notochord cells that have finished their division are thought to be in the G1/GO phase (Fig. 3C; Ogura et al., 2011). However, the timing of when they come into the G1/GO phase has not been studied in detail.

How do FGF and brachyury determine the cell cycle number? In Halocynthia roretzi and Ciona intestinalis, brachyury starts to be expressed at about the 64-cell stage (Yasuo and Satoh, 1994; Corbo et al., 1997), after which the cells divide three more times. A simple model is that the total time required for the downstream gene of Brachyury to restrict cell division corresponds to the time for the three rounds of cell-cycle progression. Transcription of the downstream genes and translation and maturation of the proteins is a plausible explanation for the lag. However, this hypothesis does not fully explain the control of the number of cell divisions by FGF/brachyury, because inducing the precocious expression of brachyury did not change the total number of notochord cell divisions (Fujikawa et al., 2011). It is thus thought that there is an additional mechanism determining the cell division number, which is turned on by FGF/brachyury.

The invariant number of cells is not a unique feature of the notochord (Fig. 3C). The number of larval muscle cells is invariant among individuals, but the number varies slightly among species (Nishida and Satoh, 1985; Nishida, 1987; Satoh, 1994). Curiously, the muscle and notochord cells show similar behavior with respect to cell division. First, both muscle and notochord cells stop dividing by the larval stage. Second. the primary muscle and notochord cells divide three times after the 64-cell stage (Nishida, 1987). Third, numbers of larval muscle and notochord cells are invariant among individuals. In contrast to the similarities, the mechanisms of differentiation of the muscle and notochord are quite different in relation to the maternal factors and the inductive signal. Muscle cell fate is determined by the maternal factor macho-1, whereas notochord cell fate is induced by the surrounding endoderm cells (Nakatani et al., 1996; Nishida and Sawada. 2001). When the muscle cells receive the inductive signal from the endoderm. their fate is changed into that of mesenchymal cells (Kobayashi et al., 2003). Mesenchymal cells divide more than three times after the 64-cell stage. As mentioned above, the notochord cells are converted into neural cells when they lose induction from the endoderm. The neural cells divide more than the notochord cells do. Therefore, the inductive signal from the endoderm mediated by the same FGF gene results in completely different responses by the recipient cells, namely, increasing the cell division number of the muscle and mesenchyme cells while restricting the number of the notochord and neural cells. Such different responses can be explained by the different combinations of transcription factor genes expressed in the cells receiving the inductive signal (Nishida, 2005; Lemaire, 2009). Transcription factor genes can modify the downstream cascade of the FGF signaling, and the differences may lead to different consequences for the recipient cells.

Although larval notochord and muscle cease their cell divisions, the total number of cells in larvae increases during the period after hatching (Yamada and Nishida, 1999), suggesting that some tissues continue to undergo cell division. Observations of cell-cycle progression in the larval body with a fluorescent cell-cycle indicator (Sakaue-Sawano et al., 2008) suggested that most cells in the tail are in the G1/GO phases, whereas many cells in the trunk are in the S/G2/M phases (Ogura et al., 2011). The tissues showing an invariant cell number as represented by the notochord and muscle are the major tissues constituting the tail. The tail is lost during metamorphosis, and thus these cells do not contribute to the adult body (Fig. 1B). Programmed cell death occurs in the larval tail and is thought to be necessary for the initiation of metamorphosis (Chambon et al., 2002, 2007; Nakayama-Ishimura et al., 2009). In contrast, larval mesenchyme and endoderm cells do not stop dividing, and thus these cells are the cause of the increase in the larval cell number. These trunk tissues of most solitary ascidians arc in a premature state in the larval body (Cloney 1982), and they remain after metamorphosis to form the adult organs. Therefore, there should be a substantial difference in cell-cycle control between the tissues lost during metamorphosis and tissues for the adult body. The epidermal cells show intermediate behavior with respect to cell division during embryogenesis. After neural tube closure in C. intestinalis, the epidermal cells divide once, and their further cell division is temporally arrested (Ogura et al., 2011). The epidermal cell cycle at the trunk is reactivated during the larval stage by reentering the S-phase (Nakayama et al., 2005), while the epidermal cells at the tail remain in the G1/GO phase (Ogura et al., 2011), suggesting that the epidermal cell-cycle control is accomplished in a region-specific manner.

The notochord of Ciona intestinalis and C. savignyi is a tubular structure. The 40 notochord cells align to form a single row (Fig. 3A). The notochord cells finish their cell division at the end of the gastrula stage, forming a 4 X 10 sheet in a bilaterally symmetric manner (Fig. 3B). Following gastrulation, the notochord plate invaginates about the axial midline to make a cylindrical rod during tailbud formation, and then the notochord cells are moved toward the midline to form a single row by an event known as convergent extension (Fig. 3C). The convergent extension of notochord cells is dependent on actin filaments, and the movement is regulated by molecules similar to those involved in the formation of the planar cell polarity (PCP) of epithelial cells, such as Prickle (Tree et al., 2002; Wallingford et al., 2002a; Sasakura et al., 2003; Jiang et al., 2005).

Indeed, the mutants of Prickle of the ascidian Ciona savignyi show imperfect convergent extension of the notochord cells, and the abnormality results in the short tail phenotype (Jiang et al., 2005). The Prickle-mutated notochord cells are capable of lamellipodia formation, suggesting that the mutant cells are motile. Therefore, the abnormal notochord formation in Prickle mutants may be associated with incorrect movements of notochord cells. The Prickle-deficient cells show the abnormality in their medial-lateral and anterior-posterior polarities, suggesting that the PCP pathway acts in the polarization of the notochord cells, which is probably crucial for proper movement of the cells. According to this function, the proteins composing the PCP pathway, namely Prickle and Dishevelled, are colocalized at the cell membrane of notochord cells in the asymmetric manner along the medial-lateral axis during convergent extension (Jiang et al., 2005). The asymmetry is suggested to provide the cue for the orientation of the movement of notochord cells.

The morphogenetic movement of cells requires a positional cue from surrounding tissues to determine the direction of their movement by setting up the position of the migrating cells' filopodia or lamellipodia. In addition, cells exhibit boundary capture (Wallingford et al, 2002b) in which cells anchoring to a tissue boundary express protrusions away from the boundary and exert forces pulling the neighboring cells toward the boundary. Because the notochord is surrounded by the muscle, endoderm, and neural ectoderm, these tissues are thought to affect the orientation of notochord cell movement during convergent extension. Munro and Odell (2002a, b) performed a tissue isolation experiment with the ascidian Boltenia villosa, in which the notochord cell-mass was isolated with one or several surrounding tissues to see whether the convergent extension of notochord cells occurred normally in the cultured tissues. It was concluded that although the notochord requires contact with another tissue for proper tube formation, it does not require any specific tissue for morphogenetic movement. As mentioned above, notochord cells have an asymmetric feature along the medial-lateral axis. Indeed. Prickle and Dishevelled proteins are absent from the cell membrane adjacent to the surrounding muscle cells at the lateral side (Jiang et al., 2005), suggesting that contact with the surrounding tissues may function in suppressing the accumulation of these proteins. It is likely that the notochord cells require contact with the surrounding cells at the lateral side to identify their medial-lateral axis; however, the formation of this axis may not need a specific signal from the surrounding tissues, as was suggested by the tissue-culturing experiments (Munro and Odell, 2002a).

Signaling molecules that control the convergent extension of notochord cells have been characterized (Sasakura and Makabe, 2001; Niwano et al., 2009; Shi et al., 2009). In Halocynthia roretzi, Wnt5 is expressed in the notochord cells (Sasakura et al., 1998). Both the overexpression of Wnt5 and disruption of its function by morpholino oligonucleotides result in defects to the convergent extension of notochord cells (Niwano et al., 2009). Tissue-specific expression and disruption of Wnt5 have shown that Wnt5 is required in the notochord cells themselves, although this gene is expressed in several tissues including muscle and epidermis (Sasakura et al., 1998). This is consistent with the results in Bottenia villosa (Munro and Odell, 2002a) where the notochord does not need signaling from any specific tissue to support its morphogenesis.

The notochord of Ciona intestinalis expresses a gene encoding FGF receptor (FGFR) during convergent extension, suggesting the involvement of FGF signaling for notochord morphogenesis (Shi et al., 2009). Indeed, disrupting FGFR by its dominant negative form results in failure in lamellipodia formation and convergent extension of notochord cells. A form of Ciona FGF, FGF3, serves as a ligand required for convergent extension of the notochord. Curiously, FGF3 is strongly expressed in the most ventral side of the neural tube, which is thought to be homologous to the vertebrate floor plate. This expression pattern of FGF3 suggests that the notochord of C. intestinalis needs signaling from the neural tube for proper morphogenesis, which is in contrast to the previously mentioned results in Boltenia. An interesting question is whether this contrast reflects differences between the different species or whether other tissues can compensate for the function of the neural tube in explants of Boltenia.

Following convergent extension, 40 coin-shaped notochord cells (Fig. 3C) keep their asymmetric character along the anterior-posterior axes (Jiang et al., 2005). For example, their nuclei are located at the posterior side. Therefore, maintenance of the axes is thought to be necessary to form the mature notochord. Formation of the lumen of the notochord shows a strong relationship with the cellular axes. After forming a single row, the notochord cells secrete extracellular matrices at the junction between two notochord cells, forming luminal pockets (Fig. 3D). Enlargement of the extracellular lumens changes the shape of the notochord cells to form a bi-concave shape along the anterior-posterior axis. Subsequently, the luminal pockets tilt, and the notochord cells become triangular in the medial section. This change in cell shape allows the previously separated lumens to fuse. Eventually the notochord cells become flattened and assume the morphology of endothelial cells. The cells surround the central lumen and then form a single and straight notochord tube. The tilting and fusion of the luminal pockets is not achieved by a simple expansion of the lumens, but rather requires a morphogenetic movement of the notochord cells that is dependent on the actin filaments. Indeed, treatment of the embryos with an actin-depolymerizing agent prevents both the tilting and the fusion of the luminal pockets, while enlarging the lumens (Dong et al., 2009). Therefore, notochord cells move themselves in a highly regulated manner to form lumens between cells, which is thought to be a good model for tubulogenesis (Denker and Jiang, 2012).

Coordination Between Cell Division and Nlorphogenetie Movement

During early embryogenesis, blastomeres cleave rapidly to increase the cell number in a restricted period. The rapid cell cycles during embryogenesis include a very short or no-gap (01 and 02) phases. Because the cells constituting the mature body usually have long gap phases, the cells must change their manner of cell-cycle progression to include the long gap phases as embryogenesis proceeds (Budirahardja and Gonczy, 2009). How this change is accomplished is an important issue in our understanding of the cell-cycle regulation during development. In several organisms, the insertion of the long gap phases during development is known to be correlated with morphogenetic movements. This phenomenon is thought to be required for coordination between cell division and movement (Duncan and Su, 2004; Copp and Greene, 2010), because cells cannot perform these two events simultaneously; the cells must arrest one of the events during the occurrence of the other.

In Ciona intestinalis. coordination between cell division and morphogenetic movement is observed in the epidermal cells during neural tube closure (Ogura et al., 2011). The central nervous system of chordates is formed at the dorsal ectodermal region. At the initial stage, the central nervous system of chordates is a flattened plate called the neural plate. The neural plate invaginates beneath the epidermal layer to form the neural tube. During neural tube formation, the epidermis adjacent to the neural plate migrates toward the midline; this movement is thought to provide the force to bend the neural plate into a tubular structure. In C. intestinalis, the epidermal cells divide 11 times from fertilization to the larval stage. Their 10th division, neural tube closure, and the 11th division occur in that order. The cells that finished the 10th division start their 11th cell cycle. The 11th cycle contains a long G2 phase compared to the previous cycles, and the cells migrate toward the midline. After migration, the cells finish the G2 phase and enter the M phase to complete the 11 th division. The necessity of the long interphase at the llth cycle has been shown by causing a precocious 11th division by shortening the 02 phase by overexpressing cdc25, the cell-cycle regulator facilitating the G2--M transition. The 02 phase is not necessarily the phase for movement of the epidermis, because elongation of the S phase instead of the 02 phase does not disrupt the movement. Therefore, the long interphase during the 11 th cell cycle gives the cells the time required to move into place.

The elongated interphase in the epidermal cell cycle during neurulation is the time for remodeling of the actin filament (Ogura et al., 2011). Prior to neural tube closure, F-actin is accumulated at the medial plane of the epidermal cells adjacent to the neural plate. This accumulation depends on Rho and Rho kinase (ROCK), because it is abolished by pharmacological inhibition of ROCK. When the prolonged G2 phase at the 11th cell cycle is inhibited by overexpressing cdc25, F-actin does not accumulate. This failure in F-actin accumulation may occur because shortening the G2 phase results in the F-actin being used to form the cell division apparatus for the precocious cell division. Therefore, the long G2 phase acts as a buffer that gives the epidermal cells time to use F-actin for morphogenesis.

It is likely that there are many events in ascidian embryogenesis that require cell-cycle controls in addition to the epidermal cells described above. It has been suggested that the endodermal cells during gastrulation and neural cells during neural tube formation show extended cell cycles (Nicol and Meinertzhagen 1988a, b; Hotta et al., 2007). Determining the duration of each cell-cycle phase in these cells is important because this knowledge will identify the phase(s) at which the cell cycles are controlled to achieve the morphogenetic movements. The length of the interphases during embryogenesis is controlled at the S phase as well as at the G2 phase (McClelancl et al., 2009). Both of these cell-cycle phases should be the focus of future studies.

Control of the epidermal cell cycle during vertebrate neural tube closure is not well understood, and future studies may reveal conservation between ascidians and vertebrates. In contrast, the regulation of the cell cycle of the neural cells during neural tube closure has been documented in vertebrates. Studies using mouse mutant strains have suggested that some neuroepithelial cells begin to exit the cell cycle and start differentiation at about the stage of neural tube closure, and a number of genes have been identified that are essential for the balance between proliferation and differentiation (reviewed by Copp and Greene, 2010). The coordination between the proliferation and differentiation of neural plate cells seems to be essential for proper neural tube closure. Elongation of the cell cycle of the neural cells during neurulation has also been reported in ascidians (Nicol and Meinertzhagen, 1988a. b). suggesting that the importance of cell-cycle control in the neural cells is shared among chordates for forming the dorsal neural tube.

In addition to cell-cycle control, programmed cell death during neural tube closure is well known in vertebrates. Massive programmed cell death, mainly apoptosis, occurs during the neural tube closure of vertebrate embryos, especially at the neural ridges and midline before and after fusion (Weil et al., 1997: Copp et al., 2003). Moreover, chicken embryos treated with a pan-caspase inhibitor exhibit failure in neural tube closure (Weil et al., 1997), suggesting an important role of apoptosis in neural tube closure. A 2011 study in mouse exploited a FRET-based fluorescent reporter to observe caspase activation in vivo (Yamaguchi et al., 2011). Simultaneous imaging of the apoptosis and morphogenesis has revealed that inhibiting apoptosis caused the delay in the progression of cranial neural tube closure and increased the frequency of exencephaly.

In ascidians, the occurrence of programmed cell death during neural tube closure has not been observed. Because ascidian embryos consist of much smaller numbers of cells compared to vertebrates, the death of a cell may have more impact on ascidian individuals than on vertebrates. The occurrence of programmed cell death during neural tube closure in ascidians must be examined carefully to observe the conservation or divergence of the mechanisms of neurulation among chordates.

Conclusions and Future Perspectives

As discussed in the preceding sections, ascidians are excellent models for studying the cellular events in the formation of the chordate body. The features of the development of ascidians can be summarized by the term "simplicity." The cell fate decision, cell division, and morphogenetic movement of all cells constituting the ascidian larval body can be described at the single-cell level. This feature is extraordinarily important for observing cellular events in live embryos.

The recent studies introduced in the preceding sections clearly suggest that morphogenetic movement, cell cycle, and cell division are two key cellular events for understanding ascidian embryogenesis. Morphogenetic movements include gastrulation, convergent extension of notochord cells, and neurulation, and these morphogenetic movements share two features: the actin filament has important roles and the small GTPase Rho mediates F-actin formation. Future analyses of these mechanisms will highlight the core cascade and components of the actin regulation shared in these cells. In contrast, the cell cycle is regulated in different fashions among tissues, and therefore it is an interesting question how such specific mechanisms in cell-cycle control are coupled with the cell-type specification.

Here we emphasize that ascidian embryos and vertebrates share common characteristics that represent the chordate body plan. The notochord and dorsal neural tube are the features specific to chordates (Satoh, 2003; Lemaire, 2011), and the mechanisms that drive the formation of these structures can only be studied by analyzing chordates. It has been suggested that the ascidians possess neural crest and placodes (Jeffery et al., 2004; Mazet and Shimeld. 2005; Kourakis and Smith, 2007; Abitua et al., 2012; Sasakura et al., 2012). The molecular mechanisms conserved among ascidians and vertebrates that conduct the formation of these structures should be the keys for constructing the chordate-specific features. These key mechanisms may have been inherited from the ancestor of chordates. An important question for zoology, i.e., "how the chordates evolved" can be answered by studies of the molecular mechanisms of ascidians and comparisons of the mechanisms among chordates.

There is an issue that must be addressed for a better understanding of the mechanisms of cellular events during ascidian embryogenesis. In contrast to the detailed description of the cellular events, the molecular mechanisms underlying the cellular processes are insufficiently understood. For the cell division of notochord cells, it has been shown that the key transcription factor Brachyury controls the number of cell divisions. Identification of the downstream gene(s) of Brachyury that directly cause arrest of the cell cycle is needed. The three events of morphogenesis--namely the movement of the notochord cells during the convergent extension, the invagination of the endodermal cells during gastrulation, and the movement of the epidermal cells during neural tube closure--are achieved by actin filaments. The regulative cascades of actin filament regulators are only partially known. Characterization of the whole cascade is necessary for understanding how F-actin is formed in an asymmetric manner in the cells and how the asymmetry is associated with the polarity of the cells. The regulative cascades of actin polymerization have been extensively studied in Ciona intestinalis with respect to the heart formation during a later developmental stage (Christiaen et al., 2008). The Christiaen study will be the milestone in the understanding of actin regulation during embryogenesis. The coordination between the cell cycle and morphogenetic movement is not fully described in morphogenetic movement other than the epidermal cells during neural tube closure. Future studies will address the detailed mechanisms of the cellular events of ascidians, and these mechanisms will be compared to those of other multicellular organisms to identify conservation of the mechanisms.


We thank the members of the Shimoda Marine Research Center at the University of Tsukuba for their kind cooperation during our study. We also thank the National Biore-source Project, MEXT, Dr. Nobuo Yamaguchi, Dr. Kunifumi Tagawa, and Dr. Shigeki Fujiwara and his colleagues for providing us with Ciona adults. We are grateful to Dr. Koji Hotta for his kind provision of the confocal image. We thank Nicholas Treen for critical reading of this manuscript. This study was supported by Grants-in-Aid for Scientific Research from the Japan Society for the Promotion of Science (JSPS) and the Ministry of Education, Culture, Sports, Science and Technology (MEXT), Japan (21112004, 23681039, 23657140 to Y. S. and 23.127 to Y. 0.). Y. S. was supported by the Toray Science and Technology Grant.

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Received 9 January 2013; accepted 31 May 2013.

* To whom correspondence should he addressed. E-mail:

(E) The cell lineage of the notochord cells from the 8-cell stage until the 64-cell stage according to Nishida (1987), Cell lineage analysis in ascidian embryos by intracellular injection of a tracer enzyme. III. Up to the tissue restriction stage. Reprinted with permission from Developmental Biology 131: 526-541.


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Author:Ogura, Yosuke; Sasakura, Yasunori
Publication:The Biological Bulletin
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Geographic Code:9JAPA
Date:Aug 1, 2013
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