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Keywords:

  • ascidian culture;
  • budding;
  • development;
  • histology;
  • regeneration;
  • take-over;
  • sexual reproduction;
  • staging method;
  • stem cells

Abstract

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. BOTRYLLUS SCHLOSSERI: A VALUABLE EXPERIMENTAL MODEL
  5. BUD DEVELOPMENT
  6. BUD DEVELOPMENT AND COLONIAL BLASTOGENETIC CYCLE
  7. BUD DEVELOPMENT: THE CONTRIBUTION OF HISTOLOGY
  8. BUD DEVELOPMENT: RELATIONSHIPS WITH SEXUAL REPRODUCTION
  9. PERSPECTIVES AND CONCLUSIONS
  10. Acknowledgements
  11. REFERENCES

Botryllus schlosseri, a cosmopolitan colonial ascidian reared in the laboratory for more than 50 years, reproduces both sexually and asexually and is used as a model organism for studying a variety of biological problems. Colonies are formed of numerous, genetically identical individuals (zooids) and undergo cyclical generation changes in which the adult zooids die and are replaced by their maturing buds. Because the progression of the colonial life cycle is intimately correlated with blastogenesis, a shared staging method of bud development is required to compare data coming from different laboratories. With the present review, we aim (1) to introduce B. schlosseri as a valuable chordate model to study various biological problems and, especially, sexual and asexual development; (2) to offer a detailed description of bud development up to adulthood and the attainment of sexual maturity; (3) to re-examine Sabbadin's (1955) staging method and re-propose it as a simple tool for in vivo recognition of the main morphogenetic events and recurrent changes in the blastogenetic cycle, as it refers to the developmental stages of buds and adults. Developmental Dynamics 236:335–352, 2007. © 2006 Wiley-Liss, Inc.


INTRODUCTION

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. BOTRYLLUS SCHLOSSERI: A VALUABLE EXPERIMENTAL MODEL
  5. BUD DEVELOPMENT
  6. BUD DEVELOPMENT AND COLONIAL BLASTOGENETIC CYCLE
  7. BUD DEVELOPMENT: THE CONTRIBUTION OF HISTOLOGY
  8. BUD DEVELOPMENT: RELATIONSHIPS WITH SEXUAL REPRODUCTION
  9. PERSPECTIVES AND CONCLUSIONS
  10. Acknowledgements
  11. REFERENCES

Chordata constitute the major Deuterostome phylum. They show primary bilateral symmetry and share at least four anatomical characteristics: (1) the presence of a permanent or temporary notochord in the form of an elastic dorsal rod, which prevents shortening of the body when longitudinal muscles contract; (2) a hollow dorsal nerve cord, modified to some extent at the front end; (3) a ventral gut, which anteriorly forms a pharynx provided with gill slits or pharyngeal pouches, and a ventral glandular structure (endostyle/thyroid gland) able to fix iodine; and (4) a muscular tail (post-anal part of body). Most chordates belong to the subphylum vertebrata (47,000 species), whereas invertebrate Chordata, representing approximately 6% of the species of the phylum, are collectively named Protochordata or Prochordates: they include Tunicata (Urochordata) and Cephalochordata. Unlike vertebrates, which are widely distributed on land and in fresh and sea waters, and are active predators or grazers, Protochordates are filter-feeding marine animals, most of which have a sedentary lifestyle.

Tunicates owe their name to the test, or tunic, the peculiar tissue in which the adult body is embedded. They are also called Urochordata, as the notochord is present only in the tail of larvae. The pharynx is well developed in adults and generally occupies most of the body volume. Tunicates include solitary and colonial, benthic and pelagic forms.

Most tunicate species are represented by ascidians or sea squirts, sessile animals widespread all over the world, mainly in shallow tropical and temperate waters. Approximately 3,000 species have been reported so far, both solitary and colonial.

In recent years, solitary ascidian species (Ciona intestinalis, Ciona savignyi, Halocynthia roretzi) have emerged as model organisms for studying the molecular control of embryogenesis and differentiation of specific cell lines (Nishida,2002a,b; Oda-Ishii et al.,2005; Passamaneck and Di Gregorio,2005; Satoh and Levine,2005; Dufour et al.,2006), and their genome has been partially or fully sequenced (Dehal et al.,2002; Yokobori et al.,2003). Although less well studied at the molecular level, compound ascidians have the advantage that, in the same organism and at various levels (morphological, biochemical, molecular), several developmental pathways (embryogenesis, blastogenesis, and regeneration) leading to adult, filter-feeding zooids may be compared. From this viewpoint, it is interesting to note that, among chordates, great ability for regeneration and asexual reproduction has been maintained only in tunicates.

The stolidobranch colonial ascidian Botryllus schlosseri has emerged as a model species for the study of asexual reproduction (Sabbadin,1958,1975; Rinkevich,2002; Laird et al.,2005a; Manni and Burighel,2006), as it is a cosmopolitan organism, easy to find and rear in the laboratory, and able to reproduce in captivity, both sexually and asexually. Embryogenesis is rapid, and larvae for dispersal of new colonies can be obtained within a week from fertilization. In addition, it is already known as a valid model organism for studying natural apoptosis and clearance of effete cells (Burighel and Schiavinato,1984; Lauzon et al.,1992,1993,2002; Cima et al.,2003), allorecognition (Sabbadin,1962,1982; Sabbadin and Astorri,1988; Ballarin et al.,1995,2002a; Rinkevich et al.,1998a; Cima et al.,2004a; De Tomaso et al.,2005; Nyholm et al.,2006), immunobiology (Ballarin et al.,1994,2002b; Ballarin and Burighel,2006), and regeneration (Berrill,1951; Zaniolo and Trentin,1987; Laird et al.,2005a).

B. schlosseri was introduced as a laboratory animal more than 50 years ago by Prof. Armando Sabbadin, at the University of Padova (Italy) who, for the first time, set up conditions for the permanent culture of this species, which had only occasionally been reared before, mainly due to interest in its budding (Metschnikow,1869; Della Valle,1882; Oka,1892; Pizon,1893; Hjort,1893; Berrill,1941a,b; Watterson,1945). Since then, many papers on B. schlosseri have been published, especially in the field of developmental biology and immunobiology, thanks to the possibility of collecting larvae and letting them metamorphose in laboratory aquaria in controlled conditions.

In this species, budding occurs continuously within a colony, in an orderly and synchronized manner, so that cyclical changes of adult generations occurs. Thus, the interval of time from one generation change to the next one can be defined as a blastogenetic cycle. During this period, various asexual developmental stages can be identified. However, a uniform method of staging both bud development and blastogenetic cycle is lacking, hindering proper comparison of data from different laboratories. In the present study, we present B. schlosseri as a valuable chordate model for studying, in the same organism, sexual and asexual development and regeneration. A detailed description of bud development up to adulthood and the attainment of sexual maturity is presented and Sabbadin's (1955a) staging method for bud and colony development is re-examined and re-proposed as a simple tool for in vivo recognition of the main morphogenetic events and recurrent changes in the blastogenetic cycle.

Botryllus schlosseri Colonies

An adult colony of Botryllus schlosseri (Fig. 1) is formed of (1) several filter-feeding blastozooids, grouped in star-shaped systems with a central common cloacal siphon, into which individual atrial siphons open; (2) buds (primary buds) on zooids; and (3) budlets (secondary buds) on buds. Zooids, buds, and budlets are embedded in a common, gelatinous tunic, and share a circulatory system represented by a network of vessels of epidermal origin, branching out from the zooids and buds, crossing the tunic, and connected to a marginal vessel, which runs along the contour of the colony (Brunetti and Burighel,1969). Sausage-like blind ends, known as ampullae, depart from these vessels toward the tunic surface and store blood cells. One side of the colonies (the leading end) advances on the substrate and, here, ampullae are particularly elongated and bear epithelial columnar “pad cells” in their tips, which help colony adhesion to the substrate and are involved in allorecognition between contacting colonies (Rinkevich et al.,1998a; Cima et al.,2006a).

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Figure 1. A: Colony of Botryllus schlosseri (dorsal view). B: Histological section of a colony showing coexistence of sexual (embryos and testes in adult, oocytes in bud) and asexual reproduction (adults and bud). Arrow, atrial siphon primordium in primary bud; arrowhead, dorsal lamina in adult. Hematoxylin and eosin stain. C–H: Development of secondary bud, from stage 1 to 6 (arrow in C–H), and primary bud at stages 7 (C) and 8 (D,E,G). Stage numbers are in brackets. Buds at stage 8 in G and F bear three and two secondary buds, respectively: note that anterior, right budlets are more developed than left/posterior ones and represent reference for colony stage (right side of bud is on left in figure). All secondary buds are in lateral view, except bud stage 6 (H), which is in frontal view; buds at stages 7 and 8 are in ventral view. I: Sketch showing changes in axis orientation during bud development. Left: bud passage from stages 3 to 4; tilt of anteroposterior axis with respect to the parent; anteroposterior bud elongation; dorsoventral axis remains parallel to parent per lateral axis. Right: bud passage from stages 6 to 7; bud axes coincide with parent axes for bud rotation along the longitudinal axis. J: Bud at stages 4, 5, 6, and 7, viewed from left side (modified from Burighel et al.,1998). Scale bar = 650 μm in A, 200 μm in B, 100 μm in C–H.

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Each colony is a clone, deriving from the metamorphosis of one tadpole larva, 1.5 mm in length, bearing a photolith for light and gravity perception, three adhesive papillae, and eight ampullae on the anterior of the trunk branching from a large lacuna ventral to the endostyle (Brunetti and Burighel,1969; Boyd et al.,1990; Manni et al.,1999; Sorrentino et al.,2000). After hatching, the larvae swim freely, before adhering to a suitable substrate with their papillae and undergoing metamorphosis. The ampullae, embedded in a thin tunic layer, extend radially on the substrate and remain connected to the ventral lacuna by means of thin stalks. Within 12–24 hr of adhesion, the stalks anastomize to form a single arch-shaped vessel, which becomes the marginal vessel of the young zooids (Brunetti and Burighel,1969). Within 36–48 hr, the larvae metamorphose into fully functional oozooids, approximately 0.5 mm in length. Each oozooid has four long transverse protostigmata on each side of the branchial basket, a stomach with longitudinal plications, and a palleal bud on the right side. After a week, at 19°C, the oozooids are resorbed in the course of a process called take-over and are replaced by their buds, which develop into mature zooids (blastozooids of the first asexual generation). The first blastozooid usually produces two palleal buds, one on each side (second blastogenetic generation), which replace the first blastozooid at the next take-over.

Bilateral budding characterizes healthy colonies, the right side of the zooid having a higher blastogenetic potential than the left one (Sabbadin,1955a,1956,1958); the budlets, especially those on the right side, can split into two at early developmental stages (Fig. 1G). Commonly, budlets on the left side and the posterior budlet on the right side are less well developed than the anterior right budlet (which is the reference for the colony stage); however, all the surviving budlets reach the same developmental stage at the beginning of take-over. Because adult zooids are cyclically resorbed and replaced by growing buds, a healthy colony can grow in size up to hundreds or thousands of zooids and buds, kept synchronized in their development by the common vascular system. In the field, sexual reproduction usually starts when water temperature exceeds 10°C (spring), and large colonies tend to die after the production of huge quantities of eggs (Brunetti,1974; Grosberg,1988). Overwintering occurs in the form of small juvenile colonies. Colony life spans seem to be genetically controlled (Rinkevich et al.,1992).

BOTRYLLUS SCHLOSSERI: A VALUABLE EXPERIMENTAL MODEL

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. BOTRYLLUS SCHLOSSERI: A VALUABLE EXPERIMENTAL MODEL
  5. BUD DEVELOPMENT
  6. BUD DEVELOPMENT AND COLONIAL BLASTOGENETIC CYCLE
  7. BUD DEVELOPMENT: THE CONTRIBUTION OF HISTOLOGY
  8. BUD DEVELOPMENT: RELATIONSHIPS WITH SEXUAL REPRODUCTION
  9. PERSPECTIVES AND CONCLUSIONS
  10. Acknowledgements
  11. REFERENCES

B. schlosseri offers several advantages as an experimental organism: (1) it is easily found in the field; (2) it can grow and reproduce in laboratory conditions; (3) colonies are embedded in a soft, transparent tunic, which allows zooids and buds to be observed under the microscope; (4) large colonies can be cut into clonal fragments able to reproduce; (5) through controlled crosses, it is possible to select colonies carrying defined genes for allorecognition or pigmentation; and (6) colonies undergo recurrent generation changes, with diffuse cell death in zooid tissues. Taking advantage of these features, blastogenesis has been studied by means of several kinds of experimental approach. For example, thanks to the tunic features and bidimensional arrangements of zooids, it is possible not only to select colonies at specific developmental stages, but also to operate on them with a thin, tungsten needle (e.g., to remove zooids of different generations from a system, to damage them, isolate them from other individuals or from the vascular net, graft buds on to the tunic, and so on). This kind of experimental procedure was first introduced by Sabbadin in 1956 and then adopted by many researchers to study bud regeneration (Watkins,1958; Zaniolo et al.,1976; Majone,1977), regulation of programmed cell death and cell removal during take-over and relationships between the resorption of old zooids and bud growth (Sabbadin,1956; Lauzon et al.,2002), establishment of bud axes during development (Sabbadin,1960; Sabbadin et al.,1975), and competition between different generations in the colony (Sabbadin,1958). The differential ability of zooids to survive and resume the colonial life cycle in varying experimental conditions has revealed that colonies have a great regulative potential, which manifests itself in differing ways, for example, with anticipation of regression of filtering adults, whose components can be re-used by growing buds, and variations in zooid growth rate, duration of zooid life span, and budding intensity (Sabbadin,1958).

Another useful characteristic of B. schlosseri is colony subcloning: fragments of a limited number of systems can detach naturally or be separated experimentally from the parent colony. When small subclones (one to two systems) from different colonies are juxtaposed and left to adhere to a supporting glass slide, their leading ends facing each other, allorecognition can be more easily observed and studied: this is typically manifested as colony specificity, which occurs whenever two colonies come into contact through their peripheral vascular ampullae, either in the form of fusion of tunics and anastomosis of vascular systems or nonfusion (Sabbadin,1962,1982). In the latter case, a rejection reaction is typically observed and shares many features with vertebrate inflammation, such as chemotaxis, blood cell infiltration and degranulation, and cytotoxicity at the “points of rejection” along the contact border (Rinkevich,1992; Ballarin et al.,1998; Cima et al.,2006a). The possibility of separating colonies differing by one allele after their fusion has revealed the alteration of their original fusibility for some time (Sabbadin and Astorri,1988), thus confirming the role of blood in allorecognition, previously demonstrated in Botryllus primigenus (Taneda,1985). Using the allorecognition assay reported above, Oka and Watanabe (1957,1960) showed that, in Botryllus primigenus, allorecognition is controlled by a single polymorphic gene locus with many codominant alleles, and Sabbadin (1962,1982) demonstrated that the same holds true for B. schlosseri. The locus was named FuHC by Weissmann et al. (1990): fusion occurs when at least one allele is shared by the contacting colonies. The high polymorphism at the allorecognition locus is devoted to limiting fusion to genetically related colonies to prevent somatic or germ cell parasitism in the resulting chimeric colonies (Stoner et al.,1999; Laird et al.,2005b; De Tomaso,2006; Nyholm et al.,2006).

In B. schlosseri, transplantation of single buds and zooids can also be successfully performed. Buds transplanted from one colony into the tunic of another colony can be vascularized and complete their development only if donor and host colonies are fusible (Sabbadin,1982). Adult zooids transplanted into another colony are resorbed at take-over, independently of type of transplant (isograft or allograft) (Rinkevich et al.,1998b).

Through selected crosses, Sabbadin (1959) and Sabbadin and Graziani (1967a) were able to demonstrate that the presence/absence of orange, blue, and reddish pigmented cells in colonies is controlled by three independent Mendelian loci, and two other loci control the distribution of nephrocytes around and between the siphons. The diverse combinations of these characters give rise to a variety of color morphs differing in both geographical and ecological distribution (Sabbadin and Graziani,1967b; Sabbadin,1978). The pigmentation pattern is a useful experimental marker, as it allows easy identification of colonies by simply ascertaining their color (Sabbadin,1971; Sabbadin and Zaniolo,1979; Laird et al.,2005b).

Both blood and epithelial cells can be cultured in vitro in appropriate conditions, representing an interesting tool in studying cell differentiation (Sala,1973; Rinkevich and Rabinowitz,1993,1994; Rabinowitz and Rinkevich,2003). Hemocytes can also easily be obtained by puncturing blood vessels in the tunic with a fine tungsten needle to prepare short-term primary cultures. Using this technique, morphological, ultrastructural, cytochemical, immunocytochemical, and cyto-enzymological analyses have been performed to clarify hemocyte differentiation pathways and mutual relationships (Sabbadin,1955b; Milanesi and Burighel,1978; Burighel et al.,1983; Schlumpberger et al.,1984a,b; Ballarin et al.,1993; Ballarin and Cima,2005; Lapidot and Rinkevich,2005), their role in immunobiology (Ballarin et al.,1994,2002b; Ballarin and Burighel,2006; Cima et al.,2006a), and alterations in functionality when exposed to environmental pollutants (Cima et al.,1995; Kamer and Rinkevich,2002; Cima and Ballarin,2004).

In addition, the peripheral vascular ampullae can easily be reached by a capillary mounted on a microinjector and are suitable sites for microinjection of various substances (tracers, inhibitors) and blood cells. Hemocytes can be injected into other colonies, either isogenic or allogenic, to study their effects on the host colony (Sabbadin and Ballarin,1990; Laird et al.,2005b). Moreover, colonies can bear the injection of several microliters of solution, which is rapidly dispersed throughout the colony's vascular net. This property has been used to study the ability of cells to take up particles from the bloodstream (Manni et al.,1993).

BUD DEVELOPMENT

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. BOTRYLLUS SCHLOSSERI: A VALUABLE EXPERIMENTAL MODEL
  5. BUD DEVELOPMENT
  6. BUD DEVELOPMENT AND COLONIAL BLASTOGENETIC CYCLE
  7. BUD DEVELOPMENT: THE CONTRIBUTION OF HISTOLOGY
  8. BUD DEVELOPMENT: RELATIONSHIPS WITH SEXUAL REPRODUCTION
  9. PERSPECTIVES AND CONCLUSIONS
  10. Acknowledgements
  11. REFERENCES

Blastogenesis in Botryllus was initially described by several 19th century researchers (Metschnikow,1869; Della Valle,1882; Oka,1892; Hjort,1893; Pizon,1893) and in the 20th century by Ärnbäck-Christie-Linde (1923), Berrill (1941a,b), Watterson (1945), Sabbadin (1955a), Milkman (1967), and Izzard (1973). A bud primordium appears as a thickening of the peribranchial wall and the overlying epidermis, which then arches progressively to form a double vesicle. The inner vesicle folds to form branchial and atrial chambers, gut, and the nervous system; the outer one gives rise to the zooid epidermis.

A staging method to describe bud morphogenesis, based on in vivo and histological observations, was first introduced by Berrill (1941a), who identified 11 stages. It was then modified by Sabbadin (1955a; Table 1; Figs. 1C–H, 2), who combined Berrill's stages 1 and 2 (owing to difficulties in identifying Berrill's stage 1 in current laboratory practice, under the binocular microscope, and in the absence of a parallel histological control), and called the disc-shaped thickening of the atrial wall, representing the bud primordium, stage 1. He also subdivided Berrill's stage 3 into two stages, for a better description of the early blastogenetic development of budlets up to the double vesicle stage. A modification of Sabbadin's staging was proposed by Izzard (1973), on the basis of detailed study of the appearance of anteroposterior polarity and bilateral asymmetry during blastozooid development. The same information was added by Sabbadin to his initial staging proposal (Sabbadin et al.,1975), with the introduction of intermediate stages (1+, initial arching of bud primordium; 2+, skewing of the hemisphere from its lateral orientation toward the anterior end of the parent; 3+, elongation and expansion of the inner vesicle along the anteroposterior axis of the parent) following stages 1, 2, and 3, respectively. Figure 1I shows two steps of bud development involving changes in axis orientation: the appearance of morphological polarity and bilateral asymmetry (from stages 3 to 4, according to Sabbadin) and later rotation of the bud along the anteroposterior axis (occurring during take-over, from stages 6 to 7, according to Sabbadin). The three staging methods reported above are compared in Table 1. Although they look similar, Berrill's method is mainly based on histological observations, and time is required to prepare sections for stage identification, particularly at the beginning of bud development. Similarly, Izzard's method is complicated by its focus on axis development. Sabbadin's method has the advantage of simplicity, because it requires only a good stereomicroscope for recognition of bud stages. In addition, unlike the methods proposed by Berrill and modified by Izzard, which are not widely applied, Sabbadin's method is currently in use and the majority of histological data on B. schlosseri blastogenesis are based on it.

Table 1. Comparison Among Staging Methods of Zooid Development of Botryllus schlosseri
Blastogenetic stages according to:
 Berrill,1941aSabbadin,1955aaIzzard,1973
  • a

    Modified according to Sabbadin et al.,1975.

Secondary bud1. Initial disc of peribranchial cuboidal cells accompanied by epidermis.1. Thickening disc on parent atrial wall. 1+, initial arching of bud primordium.1. Bud primordium: first appearance to maximal size.
 2. Maximal disc of columnar cells.  
  2. Disc expanded and arched in hemisphere. 2+, skewing of hemisphere toward anterior end of parent.2. Formation of hemisphere by arching of primordium.
 3. Arching and closure of disc to form a vesicle.3. Double vesicle stage: inner vesicle originated by closure of disc, outer vesicle from parent epidermis. 3+, bud elongation according to anteroposterior axis, which diverges anteriorly from that of parent.3. Skewing of hemisphere and vesicle formation. Primary circulation established.
   4. Expansion of atrial vesicle and development of bilateral asymmetry.
   5. Evagination of gut rudiment.
 4. Appearance of atrial folds and gut primordium.4. Appearance of atrial folds (peribranchial chamber rudiments) and gut primordium.6. Formation of atrial folds and pericardial rudiment.
 5. Branchial and peribranchial chambers recognizable; appearance of heart and neural complex rudiment.5. Branchial and peribranchial chambers and gut recognizable.7. Elaboration of organ rudiments: intestine and pyloric caecum arise from gut rudiment; neural complex and siphons form; heart infolds.
  6. Heart recognizable. 
Primary bud6. Bud rudiment.7. Appearance of atrial wall thickening, representing bud rudiments of following generation.8. Appearance of bud primordium.
 7. Stigmata primordium.  
 8. Heart beat.8. Heart beating. 81, heart beating slowly; 82, heart beating at normal rhythm.9. Heart beating. 91, heart beating slowly; 92, heart beating at normal rhythm.
Adult9. Stigmata and siphons active.9. Functional maturity. 91, oral siphon aperture; 92, atrial siphon aperture; 93, common cloacal siphon aperture.10. Functional maturity: perforation of siphons.
 10. Gonad maturation.10. Gonad maturation.11. Gonad maturation.
 11. Dissolution.11. Resorption. 111, siphons retraction and closure; 112, general shrinkage of zooids; 113 further contraction of zooids and branchial dissolution; 114, heart beat stops.12. Dissolution.
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Figure 2. Bud development from stages 1 to stage 7. Note in bud stage 7, budlet of new generation at stage 1. Ventral view.

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BUD DEVELOPMENT AND COLONIAL BLASTOGENETIC CYCLE

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. BOTRYLLUS SCHLOSSERI: A VALUABLE EXPERIMENTAL MODEL
  5. BUD DEVELOPMENT
  6. BUD DEVELOPMENT AND COLONIAL BLASTOGENETIC CYCLE
  7. BUD DEVELOPMENT: THE CONTRIBUTION OF HISTOLOGY
  8. BUD DEVELOPMENT: RELATIONSHIPS WITH SEXUAL REPRODUCTION
  9. PERSPECTIVES AND CONCLUSIONS
  10. Acknowledgements
  11. REFERENCES

As stated above, three blastogenetic generations are usually found in a colony: adults (filtering zooids), their palleal (primary) buds, and budlets (secondary buds) on buds (Fig. 3). The development of buds and zooids is highly synchronized, so that when adults are resorbed, their place is taken by primary buds, which open their siphons 24–36 hr after the beginning of take-over, while secondary buds become primary buds and give rise to a new budlet generation (Sabbadin et al.,1975). Old zooids contract and undergo massive, diffuse apoptosis of their tissues (in parallel with some necrosis in the digestive tube), and are gradually resorbed by either wandering professional or fixed occasional phagocytes (Burighel and Schiavinato,1984; Lauzon et al.,1993; Cima et al.,2003). The colonies, which cannot feed until the new adult generation opens its siphons, rely on recycling of components of dying zooids, which are used for growth of the developing buds (Sabbadin,1956; Lauzon et al.,2002). The blastogenetic cycle starts with the opening of the siphons of new adult zooids and ends with take-over, when the next blastogenetic generation reaches functional maturity. This time interval, in which buds and budlets gradually grow, takes 1 week at 19°C.

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Figure 3. A: Detail of whole-mount sexually mature blastozooids. Inset: enlargement of primary bud carrying a secondary bud with ovary rudiment. Hematoxylin stain; ventral view. B: Explanatory drawing of an adult zooid, with primary bud and secondary bud (ventral view). Scale bar = 250 μm.

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It is therefore useful to describe the cycle in such a way as to obtain direct information on the developmental stages of zooids, buds, and budlets, which, as already stated, are closely related. A first staging proposal for the blastogenetic cycle was that of Watanabe (1953) for the Japanese species Botryllus primigenus: he distinguished four recurrent phases (A to D), the latter corresponding to take-over (Table 2). Colonies were reared in the bay facing the Shimoda Marine Biological Station and, due to the high water temperature during the study period, blastogenesis was particularly rapid and the blastogenetic cycle was completed within a period of 4 days. Therefore, the four phases corresponded to the daily developmental condition of zooids and buds. These recurrent phases were applied to B. schlosseri as a staging method by researchers in the United States and Israel (Lauzon et al.,1996,2002; Laird and Weissman,2004; Rabinowitz and Rinkevich,2004; Voskoboynik et al.,2004; Rosner et al.,2006). Despite its apparent simplicity, this method gives poor information on the developmental stages of the three related blastogenetic generations (for instance, phase A includes primary buds in Sabbadin's stages 6 and 7 and phase B includes secondary buds in Sabbadin's stages 2 and 3) and cannot document the rapid developmental changes of buds and irregularities in the synchronization of the three blastogenetic generations, which frequently occur when colonies suffer from stress or are experimentally manipulated, so that its use is limited to studies (e.g., take-over) in which detailed knowledge of bud developmental stages is not necessary. More recently, the need for a more accurate description of bud development prompted Lauzon et al. (1996,2002) to modify Watanabe's proposal by introducing some intermediate stages (A1, A2, B1, B2, C1, C2, and D; see Table 2 with Watanabe's and Sabbadin's stages).

Table 2. Comparison Among Staging Methods Colonial Cycle of Botryllus schlosseria
Colonial developmental stages according to:
Sabbadin,1955aWatanabe,1953bLauzon et al.,2002
  • a

    Sexual reproduction (stage 10 according to Sabbadin,1955a, and Berrill,1941a) is not considered.

  • b

    Watanabe (1953) adopted Berrill's (1941) method for Botryllus primigenus. Berrill's stages are in brackets.

9/7/1A (1-6-9)A-1. Secondary bud evaginates from lateral wall of primary bud.
  Onset of a new cycle: oral and excurrent siphons of zooid open.
9/8/2B (2/3-8-9)A-2. Skewing of secondary bud toward anterior hemisphere of parent zooid.
  B-1. Heartbeat begins in primary bud.
9/8/3 B-2. Secondary bud forms a closed double-layered vesicle.
9/8/4C (4-8-9)C-1. Organogenesis begins in secondary bud.
9/8/5 C-2. Primary subdivisions are completed in secondary bud.
  Pigment cells accumulate in outer epithelium of primary bud.
9/8/6  
11/8/6D (5-9-11)D. Onset of take-over: shutdown of oral and excurrent siphons; dorsal movement of buds and polarized contraction of zooid along its anteroposterior axis: programmed cell death: massive apoptosis of cell corpses by blood-derived phagocytes; cessation of heart beat in zooid.

Because the developmental stage of a colony is univocally defined by the developmental stages of the three blastogenetic generations, a better alternative is to the express stages of colony development by a formula of three numbers separated by slashes (e.g., 9/8/3), as defined by Sabbadin (1955a). Each number refers to the development of the coexisting generations in the colony, that is, the first to adult filtering zooids, the intermediate to primary buds, and the last to secondary buds, respectively. In this way, we know the developmental condition of zooids, buds, and budlets exactly, and the information related to each blastogenetic stage is not lost. A similar “combination of stages,” with reversed order of numbers, was used by Watanabe (1953) in his description of the “asexual reproduction phases” of B. primigenus.

In optimal conditions, the cycle starts with 9/6/0 (a brief interval following the take-over, during which colonies contain newly formed adults and their buds, without budlet generations) and continues with 9/7/1 and 9/8/2-5 until 9-11/8/6, when take-over occurs again. Intermediate or mid-cycle stages, as defined by Lauzon et al. (1992), correspond to stages 9/8/2 to 9/8/4 (Table 2).

Stage 10 refers to sexually mature zooids and is not considered in our presentation of B. schlosseri blastogenesis, as the sole difference with respect to stage 9 is related specifically to gonad and gamete differentiation. Therefore, as far as blastogenetic development is concerned, stages 9 and 10 are equivalent.

BUD DEVELOPMENT: THE CONTRIBUTION OF HISTOLOGY

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. BOTRYLLUS SCHLOSSERI: A VALUABLE EXPERIMENTAL MODEL
  5. BUD DEVELOPMENT
  6. BUD DEVELOPMENT AND COLONIAL BLASTOGENETIC CYCLE
  7. BUD DEVELOPMENT: THE CONTRIBUTION OF HISTOLOGY
  8. BUD DEVELOPMENT: RELATIONSHIPS WITH SEXUAL REPRODUCTION
  9. PERSPECTIVES AND CONCLUSIONS
  10. Acknowledgements
  11. REFERENCES

The fine anatomy of Botryllus schlosseri zooids has been studied since the second half of the past century, when electron microscopy began to be available in addition to classic histological procedures. These studies were devoted either to the development of particular organs, such as the digestive system (Burighel,1970), circulatory system (Brunetti and Burighel,1969; Burighel and Brunetti,1971), blood (Milanesi and Burighel,1978; Burighel et al.,1983), heart (Nunzi et al.,1979), branchial basket (Casagrande et al.,1993), and nervous system (Burighel et al.,1997; Zaniolo et al.,2002), or to better comprehension of biological processes (Tables 3, 4), such as sexual reproduction (Sabbadin and Zaniolo,1979; Zaniolo et al.,1987; Manni et al.,1993,1994) and take-over (Burighel and Schiavinato,1984; Lauzon et al.,1993; see also Table 3 for references). Most of these studies describe bud development according to Sabbadin's (1955a) staging. Today, they represent a solid basis for interpretation of results obtained with a variety of techniques, including immunohistochemistry and molecular biology.

Table 3. Main Developmental Events During Zooid Development
Zooid stageNeural complex and motor nervous systemSiphonsPharynxGutVascular apparatus and heart
  • aZooid stages according to Sabbadin (1955a). Gonad and embryonic development not considered (see Table 5).

  • b

    See Table 4 for further details.

1-2     
3    Bud peduncle invaginated into two peduncular vessels; bud equatorial sinus formed
4Dorsal thickening of inner vesicle Appearance of atrial folds (peribranchial chamber rudiments)Median-posterior evagination of inner vesicleHeart primordium as compact mass; equatorial sinus in tilted plane
5Evagination of dorsal tube from posterior branchial chamber; first delamination of neuroblast from dorsal tube Branchial and peribranchial chambers recognizableStomach rudiment; initial formation of perivisceral leafletsHeart, in form of vesicle, enlarges and elongates; early evagination of radial vessel
6Fusion of dorsal tube with anterior branchial chamber; closure of posterior apertureOral and atrial siphon primordia recognizable in section as thickenings of branchial and atrial epitheliaEndostyle evident; expansions of chambers; peribranchial chambers posterodorsally fused in atrial chamberIntestine rudiment; pyloric gland and caecum rudimentHeart raphe formation (pericardium and myocardium separate); radial vessel formation; main blood sinuses and lacunae present
7Cerebral ganglion and neural gland recognizable; first appearance of motor nerves: pericoronal and subendostylar nerves; feeble gut and heart innervation Stigmatal primordia in form of peribranchial thickenings; dorsal lamina evident; primordium of three longitudinal vesselsDistinct pyloric gland and caecum rudiments; gland in form of duct and first ampullae; initial stomach foldsHeart myocardium invaginates deeply; flattening of pericardial and myocardial cells; radial vessel connected to marginal vessel
8/2Neural gland begins to separate from dorsal organ; visceral nerve recognizable; transverse interstigmatic nerves presentBrachial and atrial epithelia fused with epidermis Anal aperture; initial gastric foldsHeart beats slowly (81); heart beats at normal rhythm (82); in this and successive stages, progressive development of blood sinuses
8/3Cortex and medulla recognizable in cerebral ganglion; neural gland cytodifferentiation; nerves in gastric folds; longitudinal interstigmatic nerves presentRudiment of oral tentaclesStigmata perforationPerivisceral leaflets completely formed; 8 definitive gastric folds 
8/4Extended oral siphon innervation; nerve ring around each stigma Anteroposterior enlargements of stigmata and cytodifferentiation  
8/5Dorsal organ entirely isolated from neural gland  In this and successive stages, progressive cytodifferentiation of gut cells 
8/6Nerve number reduction    
9/7/1Definitive innervation (pericloacal nerve and nerves to dorsal cloacal lip present)Oral siphon aperture (91); atrial siphon aperture (92); common cloacal siphon aperture (93)  Establishment of adult blood circulation
9/8/2-9/8/6     
11/8/6b Siphon retraction (111)Shrinkage and disgregation (112)Shrinkage and disgregationLast heart beat (113); heart beat stops (114)
ReferencesBurighel et al.,1998; Zaniolo et al.,2002Burighel,1970; Burighel and Brunetti,1971; Burighel et al.,1998Burighel and Schiavinato,1984; Casagrande et al.,1993; Manni et al.,2002; Lauzon et al.,1993Burighel and Schiavinato,1984; Burighel,1970; Burighel and Milanesi,1975; Lauzon et al.,1993Nunzi et al.,1979; Burighel and Brunetti,1971; Mukai et al.,1978
Table 4. Main Events in Adult Zooid Generation During the Take-overa
Take-over stageMain events
111Unresponsiveness of the oral siphons to mechanical stimuli; end of filtration; closure of oral and then cloacal siphons.
112General shrinkage of zooids; contraction of the branchial basket and agglutination of branchial cilia.
113Further contraction of zooids; heart still beating.
114Heart stops; remnants of zooids in the form of dark vesicles in the centre of each system.

In reviewing bud development, we follow Sabbadin's staging method and indicate secondary bud stages with a single number (from 1 to 6); primary bud stages with a combination of two numbers, the first one (underlined) referring to primary buds and the second to secondary buds (7/1, 8/2, 8/3, 8/4, 8/5, 8/6); and adult stages with a combination of three numbers, as reported above, indicating filtering zooids (underlined), primary, and secondary buds, respectively (9/7/1, 9/8/2, 9/8/3, 9/8/4, 9/8/5, 9/8/6, 11/8/6). The advantage of this staging method, which considers younger generations is particularly evident in the analysis of stage 8, because this stage lasts a considerable number of days (approximately 5 days at 19°C), with respect to the duration of the blastogenetic cycle. In this period, secondary buds form the rudiments of the main organs, passing through stages 2 to 6, whereas important morphogenetic events occur day by day in the primary bud, which progressively differentiates toward the adult condition. Thus, subdivision of stage 8 into five steps (8/2–6), with reference to secondary bud development, allows us to follow in detail the organogenetic processes occurring in the primary bud. In the same way, it is useful to define the adult stage (9) by referring to bud and budlet stages by a triad of numbers. Indeed, adults undergo morphogenetic processes only at the beginning (when they open their siphons) and end (when they undergo take-over) of filtering activity. As stated below, reference to a triad also gives useful information on the development of embryos brooding in the parental peribranchial chambers, other than the specific phase of the adult life span.

The next section describes the development of zooids, which involves three blastogenetic cycles and lasts approximately 20 days at 18°–19° C (Sabbadin,1955a,1960; Fig. 4).

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Figure 4. Histology of secondary bud at stage 6 (column A1–G1, toluidine blue), primary bud at stage 8/2 (column A2–G2, toluidine blue), adult in mid-cycle (column A3–G3, hematoxylin and eosin stain), and adult in take-over (column A4–G4, hematoxylin stain). Note apoptotic nuclei (dark spots) of cells during take-over (column A4–G4). Lines A and B: transverse sections at the level of the branchial chamber (A) and gut (B); line C: neural complex; line D: stigmata (in D2: arrowheads, longitudinal branchial vessels; arrow: stigma primordium); line E: gut (in E1, esophagus is open in presumptive atrial chamber, because this is not yet separated from branchial chamber); line F: heart (in F2: arrow, heart raphe); line G: oral (G1, G3–4) and atrial (G2) siphon (in G2: arrowheads, tunic covering atrial epithelium; in G3: arrowheads, oral tentacles). A1, A2 and B1 modified from Burighel et al. (1998). Scale bars = 100 μm in A1–B1, 25 μm in C1–G1, 100 μm in A2–B2, 50 μm in C2–G2, 200 μm in A3–B3 and A4–B4, 100 μm in C3–G3 and C4–G4.

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Secondary Bud: Stages 1–6

The early bud appears as a small disc-like thickening of the peribranchial wall, accompanied by overlying epidermis (stage 1, Figs. 1C, 2, 3A); the bud arches symmetrically (stage 1+), becomes a hemisphere (stage 2, Figs. 1D, 2), and then skews toward the anterior end of the parent (stage 2+). Next, the inner layer of the hemisphere folds into a sealed vesicle enclosed by an epidermal vesicle (double vesicle stage, or stage 3, Figs. 1E,I, 2). The two epithelial layers and the connective tissue between them form the mantle; during bud development, the heart, gonads, blood sinuses, muscles, neural complex, and nerves form and locate in it. While the inner vesicle loses its connections with the peribranchial wall, the outer one remains associated with the parent epidermis through a hollow peduncle (or stalk), through which blood arriving from the parent flows. The bud then elongates along an anteroposterior axis (stage 3+), which diverges slightly anteriorly from that of the parent; the dorsal area of the bud faces the parent (Fig. 1I). The bud now is ready to begin organogenesis, which involves the inner vesicle and the mantle; the outer vesicle constitutes the bud's epidermis.

Branchial and peribranchial chambers.

In the inner vesicle, two parallel invaginations of the prospective ventral side grow dorsally (stage 4, Figs. 1F,I,J, 2), to divide the inner vesicle into a central branchial chamber (whose ventral region contains the prospective endostyle) flanked by two peribranchial chambers (stage 5, Figs. 1G,J, 2). The latter contact each other dorsally and posteriorly, fusing into the atrial chamber (stage 6, Figs. 1H,J, 2, 4A1–G1). Now the oral and atrial siphons are recognizable as thickenings of the branchial and atrial epithelia and the overlying epidermis (Fig. 4G1). During these movements, the peribranchial epithelium accompanies gut growth, forming the perivisceral leaflet and defining a series of perivisceral blood lacunae.

Neural complex.

The region from which the neural complex (cerebral ganglion and neural gland) develops is represented by a median dorsal thickening of the inner vesicle (stage 4). Some cells proliferate from it and migrate as presumptive neural cells into the space between the epidermis and the underlying roof of the inner vesicle. The thickening grows as a tubular evagination (dorsal tube; stage 5, Figs. 1G,J, 2), elongates anteriorly, opens into the prospective prebranchial zone, and loses its original posterior communication (stage 6, Fig. 4A1–C1). It represents the neural gland rudiment. During these changes, the process of delamination of neuroblasts from its wall continues; they migrate lateroventrally to the tube (stage 6), where they aggregate (stage 7/1) to form the cerebral ganglion rudiment.

Gut.

A medioposterior evagination of the inner vesicle represents the rudiment of the gut (stage 4). It grows as a rounded evagination (stomach rudiment; stage 5, Figs. 1G,J, 2) while its base narrows progressively and the esophagus develops. Two evaginations of the stomach wall grow backward, flanking the left side of the branchial basket (stage 6): the smaller one represents the pyloric caecum and pyloric gland rudiment; the larger one is the intestine rudiment (Fig. 4E1). The latter grows dorsally and posteriorly and remains blind until stage 8/2, when the anal aperture forms.

Heart and blood vessels.

The heart primordium appears in the form of a compact mass located in the mantle (stage 4), ventrally, on the right side of the branchial rudiment; it then develops a cavity and elongates into a tubular structure (stage 5, Figs. 1G,J, 2). Its wall (Fig. 4F1), facing the endostyle, invaginates along a dorsolateral line, forming an open myocardial fold. The two lips of the invagination separate the external pericardium from the internal myocardium. The primordium of the radial vessel appears as an evagination of the ventral epidermis of the bud (stage 5); it later connects to and fuses with a similar evagination coming from the marginal vessel of the tunic (stage 7/1; Fig. 1H).

Primary Bud: Stages 7/1–8/6

The passage from stages 6 to 7/1 occurs during take-over, when the bud is ready to produce a new generation of budlets and becomes a primary bud. In the meantime, it rotates along its longitudinal axis, becoming oriented like its parent, so that buds and parents lie in parallel (Fig. 1I). The primary bud completes its morphogenesis and undergoes cytodifferentiation. At stage 8/6, the buds are ready to substitute adult zooids, which are undergoing take-over. In this way, the “generation change” is again set in motion and is completed when the old generation has been fully resorbed. Concurrently with take-over, buds arrange themselves into new systems: their atrial siphons elongate until they join (stage 9), thus forming the common cloacal chamber of each zooid system.

Branchial and peribranchial chambers.

The primordia of the stigmata appear (stage 7/1) on the peribranchial epithelium in the form of small thickenings disposed in parallel, dorsoventrally oriented rows. The peribranchial epithelium invaginates against the branchial epithelium (stage 8/2; Fig. 4A2,D2), and its wall becomes steadily thinner, preparing for fusion and perforation, which occur in the next stage (stage 8/3). Then (stage 8/4), the stigmata enlarge, assuming an elliptical shape, while the flanking cells arrange themselves in seven rows; rudimentary cilia begin to form (stage 8/5) and progressively elongate and arrange themselves into a single row per cell.

The primordia of the three pairs of longitudinal branchial vessels become recognizable as thickened folds of the branchial epithelium (stage 7/1). In the siphon primordia, the epidermis fuses with the underlying branchial and atrial epithelia; the latter are involved in the production of a thin layer of tunic toward the chamber lumina (stage 8/2, Fig. 4G2). The rudiments of oral tentacles appear as short evaginations (stage 8/3).

Neural complex and peripheral nervous system.

In the early primary bud (stage 7/1, Fig. 1J), the cerebral ganglion is identifiable as a distinct cell mass, ventral to the neural gland rudiment. It progressively enlarges and is soon (stage 8/2) organized into a cortex of neuronal somata and a medulla of loosely packed neuritic processes (Fig. 4C2). The neural gland rudiment begins to separate from the dorsal organ (corresponding to the dorsal strand typical of other ascidians; stage 8/2) and becomes involved in cytodifferentiation (stage 8/3): it differentiates into an anterior aperture (ciliated duct), endowed with ciliated cells, and a large central lumen defined by epithelial cells that progressively flatten.

First nerves become recognizable precociously (stage 7/1) at the base of the rudiment of the oral siphon, around the stomach, intestinal primordium, and heart. The peripheral innervation pattern follows blood sinus formation between epithelia: along them, nerves find their preferred pathways to the developing organs. The nerve net becomes more complex as blastogenesis continues and many nerves appear: the visceral nerve and transverse interstigmatic nerves (stage 8/2), nerves along the gastric folds and longitudinal interstigmatic sinuses (stage 8/3), many thin nerves in the gut and oral siphon, and a complete nerve ring around each stigma (stage 8/4). Maximal complexity is reached at approximately stage 8/5; subsequently (stage 8/6), the number of nerves decreases dramatically to the nerve pattern of adults, composed of anterior, posterior, and visceral nerves (the main nerves of the blastozooid) and four or five thin nerves per side in the cerebral ganglion.

Gut.

The pyloric gland rudiment now bifurcates into two branches (stage 7/1): one representing the rudiment of the pyloric caecum and the other the rudiment of the gland, which progressively differentiates into a net of tubules, ending in blind ampullae, encrusting the mid-intestinal epithelium. The stomach wall begins to fold into eight longitudinal folds, which progressively extend to their full length. The intestinal end and overlying perivisceral epithelium perforate, forming the anal aperture (stage 8/2); progressively, three intestinal regions (preglandular, glandular, rectal) become visible (stage 8/5). During the final steps, the primary bud completes its differentiation, and all the cell types of the digestive tract are recognizable.

Heart and blood vessels.

The heart assumes the form of a double-walled, tubular structure (stage 7/1, Figs. 1C,J, 2): the invaginating myocardium penetrates deeper into the pericardial cavity, and the two lips of the invagination approximate each other to define the luminal cavity. Both pericardial and luminal cavities undergo progressive enlargement. The heart (Fig. 4F2) begins its activity at stage 8/2: initially, the heart beat is slow (stage 81, see Table 1), but gradually reaches its normal rhythm (stage 82). It is coordinated by the heart of the adult generation, because it reverses the direction of its contractions soon after cardiac reversion has taken place in the adult.

As the heart begins its activity, the deeply located circulatory system, pertaining to the pharynx and digestive tract, begins to be defined, being distinguished from that of the mantle, which remains superficial. New blood lacunae gradually form and organize the typical adult circulatory tree. At take-over, the young zooids remain connected to their dissolving parents by means of a vessel, which belongs to the interzooid circulatory tree. Owing to this change, the deeply located circulation is also modified.

Filter-Feeding Adult Stage

Many aspects of adult anatomy have been described in detail by several authors using light and electron microscopy; references regarding B. schlosseri can be found in Burighel and Cloney (1997) and Mackie and Burighel (2005). Here, we concentrate on the main morphogenetic processes occurring in adults, i.e., the beginning of filtering activity (stage 9/7/1) and resorption of adults (stage 11/8/6).

The oral siphons open first (91, see Table 1; Fig. A3–G3), followed shortly after by the atrial siphons (92); next, the blastozooids modify their disposition and meet each other to form the common cloacal siphon of the system (93): the dorsoposterior regions of the blastozooids grow backward and rise to form the cloacal lips which, all together, define the common cloacal chamber. Water flows into the branchial chamber by branchial ciliary activity, and both respiration and feeding begin. Young blastozooids display definitive innervation and circulatory patterns. In intermediate stages of the blastogenetic cycle (except for male gonads, which continue to develop), the blastozooids are not involved in morphogenesis or important processes of tissue renewal, as also shown by scarce mitotic and very low apoptotic activities (Tiozzo et al.,2006).

The resorption of adult zooids represents the final process in the blastogenetic cycle. It is known as take-over, and at least four steps can be distinguished (Tables 1, 4).

During the shrinkage of adult zooids, the epidermis and peribranchial epithelium contract and play a special role in compressing the underlying organs: a wave of contraction moves anteroposteriorly along the body (Burighel and Schiavinato,1984; Lauzon et al.,1992). The pharynx and peribranchial cavities collapse, the branchial walls disintegrate, and circulating phagocytes massively infiltrate senescent tissues and rapidly ingest effete cells (Burighel and Schiavinato,1984; Cima et al.,2003). The gut epithelium retains its apparent continuity longer; it begins contracting immediately after the branchial sac; stomach and intestine display a combination of necrotic and apoptotic changes (Burighel and Schiavinato,1984; Lauzon et al.,1993). In later stages, degenerating organs undergoing resorption are immersed in a single blood lacuna. The heart continues to beat during disintegration of the digestive organs; the direction of its beats is affected by the hearts of daughter zooids, which have reached adulthood. Within 24–36 hr, the zooids and their organs are completely resorbed.

The cells of senescent adults die mainly by apoptosis (Lauzon et al.,1993; Cima et al.,2003): they are rapidly ingested by professional, circulating phagocytes that massively infiltrate zooid tissues, and sometimes by occasional phagocytes represented by neighboring epithelial cells (Burighel and Schiavinato,1984; Tiozzo et al.,2006). Phagocytes recognize a variety of “eat-me” signals, such as phosphatidylserine and oxidized lipids, on the surface of cell corpses; CD36, a component of the vitronectin receptor, is also involved in this recognition (Cima et al.,2003; Voskoboynik et al.,2004).

During take-over, the colony is rejuvenated: the tissues of old zooids die and are resorbed, while buds grow to adulthood, sustained by the recycling of the material engulfed and digested by phagocytes through the common circulation (Sabbadin,1958; Lauzon et al.,2002). Blocking phagocyte ingestion of senescent cells leads to an interruption of zooid degeneration (Voskoboynik et al.,2004), suggesting that removal signals on apoptotic cells, clearance of cell corpses by phagocytes, and completion of the blastogenetic cycle are closely linked. Bud maturation is also related to and conditioned by the regressing zooids: when, in conditions of stress, take-over occurs earlier, maturation of primary buds is anticipated as the result of the sudden availability of great quantities of nutritive material deriving from the clearance of senescent cells and tissues from the old zooids (Sabbadin,1956,1958). In this case, colonies frequently have only two generations (9/3–5), as the buds cannot yet yield new budlets (Sabbadin,1958). Conversely, extirpation of all buds except one from adult zooids significantly increases the life span of the adult generation (stage 9) and of its buds (stage 8), as well as the size of the buds and their blastogenetic potential, so that colonies with up to five generations can be obtained (Sabbadin,1958); zooids can reach giant sizes when derived from a single bud left in a colony (Lauzon et al.,2002). This finding indicates that sustaining bud growth is an arduous task for zooids, which cannot support the development of their buds to adulthood, except through their death and dissolution and recycling of their tissue components (Sabbadin,1956,1958).

BUD DEVELOPMENT: RELATIONSHIPS WITH SEXUAL REPRODUCTION

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. BOTRYLLUS SCHLOSSERI: A VALUABLE EXPERIMENTAL MODEL
  5. BUD DEVELOPMENT
  6. BUD DEVELOPMENT AND COLONIAL BLASTOGENETIC CYCLE
  7. BUD DEVELOPMENT: THE CONTRIBUTION OF HISTOLOGY
  8. BUD DEVELOPMENT: RELATIONSHIPS WITH SEXUAL REPRODUCTION
  9. PERSPECTIVES AND CONCLUSIONS
  10. Acknowledgements
  11. REFERENCES

Sexual differentiation and the production of zooids in a colony depend on three main factors: (1) the time at which the colony reaches sexual maturity, (2) bilateral asymmetry, and (3) input of germ cells from other zooids conveyed by the bloodstream.

The gonads do not form in the oozooids, nor in first-generation blastozooids. When the gonads appear, their maturation may require several generations, passing through the phase of incipient sexuality, with the appearance of undifferentiated gonad blastema in secondary buds, to a phase with abortive testes and oocytes not undergoing vitellogenesis, followed by the phase of male and then female sexual maturity (Sabbadin,1960; Sabbadin and Zaniolo,1979).

Colonial bilateral gonadal primordia appear in the secondary bud at stage 3 as clumps of undifferentiated cells on either side of the inner vesicle (Table 5; Sabbadin and Zaniolo,1979): their lateral portion becomes an ovary. Oocytes appear in secondary buds (Fig. 4A1), ripen (only one or a few on either side) in primary buds (Fig. 1B), and ovulate when the primary buds are about to pass to the adult stage (10/7/1) (Milkman,1967). Fertilization occurs just after siphon opening, and the larvae hatch when adults are almost at the end of their life cycle.

Table 5. Relationship Between Main Events of Sexual Reproduction and Zooid Developmental Stagesa
Zooid stage34-567/1-8/38/48/5-8/69-10/7/110/8/210/8/310/8/4-511/8/6
  • a

    Data according to Sabbadin (1955a). Embryonic stages from Manni et al. (1999).

EventAppearance of gonad rudiment Testis recognizable as solid mass; beginning of vitellogenesis Beginning of spermatogenesis Ovulation, fertilization, beginning of embryo developmentEarly tail bud embryo stageMost of sperm release; mid-tail embryo stageLate tail bud embryo stageLarval hatching

The medial portion of the gonad primordium differentiates into a plurilobular testis, in the form of a coherent structure, which reaches maturity in adults (Fig. 1B) and discharges most of the sperm 1–2 days after ovulation, thus avoiding selfing (Milkman,1967; Sabbadin,1971; Sabbadin and Zaniolo,1979; Stewart-Savage et al.,2001).

Gonadogenetic power, like blastogenetic power, varies in zooids according to the dextral or sinistral, anterior or posterior origin of their buds, but it is significantly higher on the left side with regard to the presence, size, and measurability of testes, and the numbers of eggs and developing embryos (Sabbadin,1955a; Sabbadin and Zaniolo,1979).

During its growth, the oocyte is surrounded by three cell envelopes: test cells, and inner and outer follicle cells, all deriving from primary follicle cells (Manni et al.,1993,1994). Test cells are separated from follicle cells by a fibrous layer, the vitelline coat, or chorion. At ovulation, the egg hatches from the outer follicle cells, which may persist for some time as a kind of corpus luteum. The inner follicle cells participate in forming the placental cup, together with the parental peribranchial and oviductal epithelia, and the embryo is held in the peribranchial chamber (Fig. 1B; Zaniolo et al.,1987).

At 18–19°C, embryonic development lasts approximately 5 days. Sexual and asexual cycles coincide: the stage of development and the day of hatching of embryos in a colony are closely related to the colonial blastogenetic stage (Table 5). The embryo reaches the early tail bud, mid-tail bud, and late tail bud stages when the colony is at stages 10/8/2, 10/8/3, and 10/8/4–5, respectively (Manni et al.,1999).

Larvae, which are visible inside the parental body, can be removed with a thin needle and collected separately. If the colony begins take-over too early, before the larvae hatch, the latter remain in the tunic and are lost (Sabbadin,1955a,1958).

Colonies are physiological units in a dynamic, balanced situation based on competition between zooids of the same and coexisting generations: if environmental conditions worsen, sexual reproduction is arrested, the budding rate is reduced to one bud per zooid, and the colony stops growing. Conversely, in good environmental conditions, as many as four budlets per bud can develop, two or three oocytes per bud can enter vitellogenesis, and more than one embryo can be brooded by adults (Sabbadin,1958).

It has long been known that oocytes and male elements can be captured by the bloodstream within the tunic vessels and conveyed to other zooids. Their recycling through successive generations has been regarded as the necessary condition for their maturation (Mukai and Watanabe,1976; Sabbadin and Zaniolo,1979).

Stem Cells

Several pieces of evidence suggest that B. schlosseri possesses circulatory pluripotent stem cells, which originate in the embryonic mesenchyme (Sala,1973) and which are the progenitors of the blood cell line (Sabbadin,1955b) and germ line (Izzard,1968, Sabbadin and Zaniolo,1979). The high potential of circulatory hemoblasts has been experimentally proven in colonies deprived of all their zooids (adults and buds) and reduced only to the peripheral tunic with colonial circulatory vessels containing blood moved by the contractile ampullae. In these conditions, hemoblasts adhere to vessel walls and give rise to budlets (vascular budding), which mature into functional zooids, thus restoring the colony to its original phenotype (Sabbadin et al.,1975).

In an elegant experiment, Sabbadin and Zaniolo (1979) demonstrated that germ stem cells can be exchanged between colonies carrying the opposite genotypes AAbb and aaBB, relative to two Mendelian loci controlling pigmentation. They were separated after a few days of fusion and double-crossed, as males and females, to colonies of the double recessive genotypes aabb. Offspring, phenotypically Ab from aaBB × aabb crosses and aB from AAbb × aabb crosses, were obtained, arising from germ cells received from the partner during the fusion period. Heterochtonous offspring could still be collected at the 14th and 15th blastogenetic generations after interruption of fusion; in some cases, all the offspring were heterochthonous.

The above and other recent experiments (Stoner et al.,1999; Laird et al.,2005b; De Tomaso,2006) showed that stem cells for the somatic and/or germ line can be exchanged between fused colonies. Fusion confers three major benefits upon the chimeric colony (De Tomaso,2006): (1) increase in size, with consequent selective advantage in substrate competition; (2) earlier achievement of sexual maturity (which occurs only over a minimum size); (3) increased genetic diversity. However, the histocompatibility gene limits fusion to kin colonies, to avoid somatic or germ cell parasitism.

Laird et al. (2005b) identified a population of multipotent stem cells from B. schlosseri colonies, which were able to give rise to both somatic and germ cells. The authors succeeded in inducing somatic and germ cell parasitism by transplanting these cells between colonies, showing that injected cells can give rise to either soma or germ. This finding may be interpreted in the light of the presence of either separate stem cells for the somatic and germ lines, or pluripotent stem cells that differentiate according to microenvironmental signals received from the niche in which they land after transplantation.

PERSPECTIVES AND CONCLUSIONS

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. BOTRYLLUS SCHLOSSERI: A VALUABLE EXPERIMENTAL MODEL
  5. BUD DEVELOPMENT
  6. BUD DEVELOPMENT AND COLONIAL BLASTOGENETIC CYCLE
  7. BUD DEVELOPMENT: THE CONTRIBUTION OF HISTOLOGY
  8. BUD DEVELOPMENT: RELATIONSHIPS WITH SEXUAL REPRODUCTION
  9. PERSPECTIVES AND CONCLUSIONS
  10. Acknowledgements
  11. REFERENCES

While all animal phyla contain representatives that undergo sexual reproduction by means of fertilization followed by embryogenesis, the ability to reproduce asexually appears to be confined to relatively few taxa: of these, tunicates are the closest relatives to vertebrates. B. schlosseri offers the unique advantage of exhibiting cyclical, natural budding, as well as induced regeneration of some tissues, and is thus a species that can open up interesting future research perspectives.

Allorecognition

B. schlosseri, with its capacity for allorecognition of contacting colonies, represents a reference species for studying nonself recognition and the evolution of adaptive vertebrate immunity, strongly supporting the rise of comparative immunology (Oka and Watanabe,1960; Sabbadin,1962). The initial hypothesis of the presence of a histocompatibility locus in this species (Sabbadin,1962; Scofield,1982a) has recently been confirmed by the isolation and characterization of a protein, which represents the first molecule expressed by a nonvertebrate histocompatibility gene described so far (De Tomaso et al.,2005), as well as a putative receptor involved in allorecognition (Nyholm et al.,2006).

Histocompatibility and Self-Sterility

Interestingly, the highly polymorphic locus for histocompatibility probably also controls self-sterility in Botryllus primigenus, in such a way that homologous colonies, when crossed, produce no offspring (Oka and Watanabe,1960). In this respect, the B. schlosseri populations from the Lagoon of Venice are different, because no hindrance exists in experimental self-fertilization between homologous colonies (Sabbadin,1971,1982). However, B. schlosseri populations from Woods Hole and Monterey appear to be more similar to the B. primigenus model, rather than to the Venetian population of B. schlosseri (Scofield et al.,1982a,b). Thus, an extension of studies on the relationships between fertilization, inbreeding depression, and histocompatibility may add information, for a more exhaustive picture about the role of kin in the evolution of a population.

Stem Cells

B. schlosseri is an excellent model for the study of toti- and pluripotency of adult stem cells, which may be compared with embryonic stem cells thanks to their potential to form entire individuals, as in the case of vascular budding (Sabbadin et al.,1975). As evidenced by the experiments of Laird et al. (2005a), the possibility of isolating single stem cells and introducing them into the circulation of fusible host colonies is a very useful way of following their differentiation pathways, commitment before transplantation, and the influence of various landing niches for final differentiation. This approach allows the formation of chimeras and the recognition of heterologous components, avoiding the need to fuse colonies and collect descendants, and is thus very suitable for studying the basis of cell differentiation in tunicates. From another point of view, it allows testing of the parasitic potential of cells injected into the host colony, and contributes further to the discussion on possible mechanisms which, in heterogeneous populations, favor the development of histocompatibility as a method for enhancing kin recognition and increasing their reproductive success (Weissman,2000).

Apoptosis

As previously stated, the natural occurrence of massive, cyclical cell death, causing the resorption of entire individuals, renders B. schlosseri an interesting model organism for the study of apoptosis, its genetic control (two genes whose expression changes during take-over were described by Lauzon et al.,1996), and the role played by phagocytes in recognizing and clearing senescent cells and regulating tissue resorption (Voskoboynik et al.,2004). This finding is important because, despite the high number of in vitro model systems, mainly selected cell lines, there is an increasing need to find reliable models for in vivo studies of the biological role of apoptosis in the organism as a whole.

Morphogenesis and Chordate Evolution

For its simplicity and the homology of several structures with vertebrates, B. schlosseri can be used to study various aspects of morphogenesis and evolution in chordates. For example, the universal developmental mechanism of fusion and perforation of epithelia, difficult to study in vertebrates due to body complexity, has been studied in B. schlosseri during the formation of the branchial fissures, which occurs as a pluri-repetitive module at each blastogenetic generation (Manni et al.,2002). This or other aspects of organogenesis and differentiation may be studied with this ascidian. In addition, study may be facilitated and also extended by the possibility of culturing cells obtained by tissue dissociation (Rinkevich and Rabinowitz,1994; Rabinowitz and Rinkevich,2003).

Developmental Mechanisms

In this animal, both asexual and sexual forms of reproduction form zooids with comparable morphology. This finding has aroused the interest of many authors, thanks to the possibility of comparing two different developmental pathways (i.e., determinative development of zygotes and regulative development of buds) to verify whether steps in embryogenesis are repeated during blastogenesis, or if the latter is a completely new form of development (Manni and Burighel,2006). This kind of research has opened a window on scarcely explored horizons of developmental biology. Although the availability of B. schlosseri gene sequences is still limited, with respect to solitary species of the genera Ciona and Halocynthia, efforts to compare the two types of development in terms of gene expression are now in progress (Laird et al.,2005a: Tiozzo et al.,2005). For example, a novel gene, Athena, specifically involved in B. schlosseri blastogenesis, has recently been identified and studied by means of RNAi and antisense morpholinos, also introducing the use of molecular techniques in analyzing knockdown phenotypes (Laird et al.,2005a). It would be of interest to extend comparisons between embryogenesis and blastogenesis to explore other aspects, such as the molecular mechanisms regulating body pattern formation during budding. For example, the expression of Hox genes could show if, during the vegetative formation of zooids, a specific developmental phase comparable with the phylotypic stage of the embryo can be identified. Analysis of other genes could also tell us whether specific territories with potentials comparable with embryonic germ layers can be defined in the bud.

Body Axis Establishment

Many biological problems that arose when using the experimental approaches developed in the 1950s and 1960s (budectomy, zoodectomy, isolation of buds, grafting of buds; Sabbadin,1956,1958), have only occasionally been re-examined with more modern techniques (Lauzon et al.,2002). For example, in special experimental conditions (Sabbadin,1956; Sabbadin et al.,1975), ablation of zooids causes buds to develop reverse bilateral asymmetry (situs inversus viscerum and cordis), with zooids with their digestive tract located on the right and the heart on the left; this reversion also influences blastogenetic and gonadogenetic powers, more developed on the left and right, respectively. Because the reversed bilateral asymmetry of a zooid is transferred to its buds, but not genetically inherited by its embryos, it offers the possibility of studying how genetic and epigenetic factors influence body axis formation.

Ecological Implications

Lastly, B. schlosseri has also recently been used as an experimental model to monitor the risk caused by the presence of various biocidal compounds in the environment (Cima et al.,2004b) as well as a reference species for studying the disturbing action of xenobiotics on the macrofouling biocenosis of hard substrates, where it is often a dominant species (Cima et al.,2006b). This finding highlights the versatility of the species and its future possible applications also in the ecological field.

The multiplicity of research directions offered by B. schlosseri and the possibility of using modern molecular techniques to face fundamental questions on developmental biology now require a staging method for development of this species. This method ideally should be easy and applicable to living colonies; referred to the three blastogenic generations, codifying precise morphogenetic information on each zooid generation; and shared by the scientific community. In our opinion, the staging method introduced by Sabbadin (1955a) combines simplicity with abundance of information on bud morphogenesis and colonial blastogenetic cycle, and we propose it as a reference method for future study on this species.

Acknowledgements

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. BOTRYLLUS SCHLOSSERI: A VALUABLE EXPERIMENTAL MODEL
  5. BUD DEVELOPMENT
  6. BUD DEVELOPMENT AND COLONIAL BLASTOGENETIC CYCLE
  7. BUD DEVELOPMENT: THE CONTRIBUTION OF HISTOLOGY
  8. BUD DEVELOPMENT: RELATIONSHIPS WITH SEXUAL REPRODUCTION
  9. PERSPECTIVES AND CONCLUSIONS
  10. Acknowledgements
  11. REFERENCES

The authors thank Mr. Marcello Del Favero for his enthusiasm and skill in rearing from many years Botryllus schlosseri in the laboratory of the Department of Biology at the University of Padova. P.B. and L.B. were funded by the Ministero della Università e Ricerca Scientifica e Tecnologica. The authors dedicate this paper to Armando Sabbadin, professor emeritus at the University of Padova, who, more than 50 years ago, introduced Botryllus schlosseri as a model organism in the laboratories of the Department of Biology (University of Padova). They thank him for having passed on to them his great passion for ascidian biology and for his scientific rigor.

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  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. BOTRYLLUS SCHLOSSERI: A VALUABLE EXPERIMENTAL MODEL
  5. BUD DEVELOPMENT
  6. BUD DEVELOPMENT AND COLONIAL BLASTOGENETIC CYCLE
  7. BUD DEVELOPMENT: THE CONTRIBUTION OF HISTOLOGY
  8. BUD DEVELOPMENT: RELATIONSHIPS WITH SEXUAL REPRODUCTION
  9. PERSPECTIVES AND CONCLUSIONS
  10. Acknowledgements
  11. REFERENCES
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