Biol. Bull. 213: 122-134. (October 2007)
© 2007 Marine Biological Laboratory
Central Nervous System of Chaetoderma japonicum (Caudofoveata, Aplacophora): Implications for Diversified Ganglionic Plans in Early Molluscan Evolution
Shuichi Shigeno1,*,
Takenori Sasaki2 and
Gerhard Haszprunar3
1 Department of Neurobiology, Pharmacology and Physiology, The University of Chicago, 947 E 58th St., Chicago, Illinois 60637
2 The University Museum, The University of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo 113-0033, Japan
3 Zoologische Staatssammlung München, Münchhausenstrasse 21, 81247 München, Germany
* To whom correspondence should be addressed. E-mail: sshigeno{at}uchicago.edu
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Abstract
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The organization of the central nervous system of an "aplacophoran" mollusc, Chaetoderma japonicum, is described as a means to understand a primitive condition in highly diversified molluscan animals. This histological and immunocytochemical study revealed that C. japonicum still retains a conservative molluscan tetra-neural plan similar to those of neomenioids, polyplacophorans, and tryblidiids. However, the ventral and lateral nerve cords of C. japonicum are obviously ganglionated to various degrees, and the cerebral cord-like ganglia display a lobular structure. The putative chemosensory networks are developed, being composed of sensory cells of the oral shield, eight precerebral ganglia, and eight neuropil compartments that form distinct masses of neurites. In the cerebral cord-like ganglia, three anterior, posterior, and dorsal lobes are distinguished with well-fasciculated tracts in their neuropils. Most neuronal somata are uniform in size, and no small globuli-like cell clusters are found; however, localized serotonin-like immunoreactivity and acetylated tubulin-containing tracts suggest the presence of functional subdivisions. These complicated morphological features may be adaptive structures related to the specialized foraminiferan food in muddy bottoms. Based on a comparative scheme in basal molluscan groups, we characterize an independent evolutionary process for the unique characters of the central nervous systems of chaetoderms.
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Introduction
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Caudofoveata (Chaetodermomorpha) consists of small vermiform molluscs with shining aragonite spicules embedded in a chitinous cuticle (see Fig. 1A). They are one of the most primitive extant molluscan groups to have several plesiomorphic characters (Thiele, 1902; Hyman, 1967; Salvini-Plawen, 1972, 1980, 1985, 1991, 2006; Haszprunar, 1987), but highly specialized apomorphic characters are also known (Scheltema, 1988, 1993, 1996; Scheltema et al., 1994; Haszprunar, 2000). Caudofoveata is thought to be derived from a creeping neomenioid-like ancestor (Haszprunar, 2000) or from flat-shaped stem molluscs (Scheltema, 1993; Caron et al., 2006; but see Butterfield, 2006). The organization of the nervous system displays a complex lobular form (Wirén, 1892) compared to those of other molluscs such as polyplacophorans and patellogastropods (see Bullock and Horridge, 1965).
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Figure 1. Diagram showing a generalized structure for body plans (above) and the central nervous system (below) in Chaetoderma japonicum. In all drawings, anterior is toward the top. The central nervous system is composed of the cerebral, ventral, and lateral nerve cords with small serially repeated ganglia (blue). The major nerve tracts are shown by simple lines (red). Some internal organs and nerves are omitted for simplification.
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Caudofoveates generally burrow in muddy bottom and feed mostly on foraminiferans (Salvini-Plawen, 1981, 1988). No visual and static (equilibrium) organs are developed, and instead the chemosensory organs (e.g., oral shield posterior or around the mouth) and the anterior regions of the cerebral ganglia are elaborated as precerebral ganglia (Hoffman, 1949). Thus, Caudofoveata is well suited for exploration of how chemosensory-motor networks have evolved in basal molluscan lineages. However, the neural network structure, including inter- and intra-lobe connectivity, remains largely uncertain. This paper reports the first neuroanatomical results using immunocytochemical neural markers for a species traditionally classified as an "aplacophoran," and offers insights into the evolution of the nervous system of vermiform shell-less molluscs.
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Materials and Methods
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Specimens
Three specimens of Chaetoderma japonicum Heath, 1911, were collected from muddy sediments dredged from off Shizuoka Prefecture, Japan (R/V Tansei-Maru, cruise KT-02-5, station EN2-3, from 34°32.620'N, 138°03.110'E to 34°32.440'N, 138°03.231'E, 240–260 m deep, on 26 May 2002) near the type locality (off Oigawa River, Shizuoka Prefecture, Albatross Station 3721, 207–250 fathoms: Heath, 1911: p. 9 and p. 67). All of the specimens used in this study were identified as C. japonicum on the basis of external characters such as (1) the thick head region sharply demarcated from the trunk, (2) the trunk covered with vertically aligned spicules in the anterior half and posteriorly directed spicules in the posterior half, and (3) long flaring spines on the tail regions (for original description see Heath, 1911: pp. 67–68, pl. 3, figs. 7–8). Although Heath (1911) originally described the species as C. japonica, C. japonicum is correct, because the gender of the generic name is neuter, not feminine. All specimens were fixed on board with 4% paraformaldehyde in phosphate buffered saline (PBS) at 4 °C (overnight), and then transferred to 70% ethanol for long-term storage at –30 °C. One individual was examined for wholemount immunohistochemistry and others for immunohistochemical studies as described below. Two series of histological sections (sagittal and cross sections) were deposited at The University Museum, The University of Tokyo (registration number: UMUT RM29448-29449).
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Figure 7. Summary diagrams for the cerebral cord of Chaetoderma japonicum. (Left) Four precerebral ganglia (pc1–4) and three lobes (anterior, dorsal, and posterior) are shown with other structures in the head region such as the radular sac (rad), buccal ganglia (buc), and oral shield (os). (Middle) Major nerve pathways (dorsal root, dr, and ventral root, vr) from the oral shield (os) to precerebral ganglia with neuropil compartments (np1–np4). The dark brown areas indicate densely acetylated tubulin-stained neuropil. Some connective and commissural tracts are also shown in the anterior and posterior lobes, with connections to the interbuccal commissure (ic), the buccal nerve (bn), and the ventral and lateral nerve cords (vn+ln). Dark region indicates dense tubulin-positive regions. (Right) Large cells (black circles) are distributed in the three regions (lc1–3) in cell body layers of the anterior, posterior, and dorsal lobes. Serotonin-containing neurites are notably accumulated in the posterior lobe and the buccal ganglia, as shown in dark gray against the pale color of the anterior lobe and the no color of the precerebral ganglia.
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Histology and immunocytochemistry
Wholemounts and sections were immunostained using a standard protocol (Shigeno and Yamamoto, 2002) with some modification. Sharp scissors were used to cut transverse fragments of the head, trunk, and tail regions of specimens in 70% ethanol. For wholemount immunocytochemistry (see Fig. 3), some regions of the surrounding spicules in trunk regions were carefully removed. Specimens stored in the cold were rehydrated in graded series of ethanol, washed well in PBST (PBS with 1% Tween 20), and treated with proteinase K (5 µg/ml in PBS) for 5 min at 37 °C to facilitate penetration of antibodies. The specimens were washed in PBST and blocked in 10% bovine serum albumin (BSA) in PBST for 1 h at room temperature. A mouse anti-acetylated 
-tubulin antibody (Sigma, T6793, 1:3000 dilution in PBST and 1% BSA) and an Alexa Fluor 488-conjugated anti-mouse IgG antibody (Molecular Probes, 1:400 in PBST) were used as the primary and secondary antibodies, respectively. Stained samples were further dissected along the sagittal axis and then mounted in 10% glycerol in PBST. They were examined using an LSM-510 confocal laser scanning microscope (Carl Zeiss) or fluorescent BX62 microscope (Olympus).
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Figure 3. Anti-acetylated -tubulin antibody wholemount staining, showing the structure of the nervous system in Chaetoderma japonicum. (A) The anterior end of the dorsal head region. The tubulin-containing neurons are scattered in the apical part (ap), and neurons are arranged in an orderly fashion at the posterior part (arrow). (B) The ventral and lateral nerve cords with knob-like structures (arrowheads) at the trunk; lateral view. Some nerves run obliquely against the radial axis (arrows). (C) An enlarged view of the ventral and lateral nerve cords at the trunk, showing the positions of nerve projections (arrowheads). One thick muscle fiber (m) runs between these cords. (D) A lateral view of the trunk with fine longitudinal nerve tracts (lon). (E) Radial nerves project from the lateral nerve cord from the ventral to the dorsal side. Some branches run to the anterior region (arrows). Arrowheads indicate the repeated roots of radial nerves, and asterisks are major branching points. (F) The ventral and lateral nerve cords connect to the suprarectal commissure (sc) at the tail region. (G) An enlarged view for the radial nerves and peripheral neurons in the body wall of the head region. Monopolar neurites toward the dorsal (arrow) or ventral (arrowheads) side. (H) An enlarged surface view of panel G, showing the scattered distribution of various cell types. Small (arrowheads) and large (arrows) cells are seen. (I) An enlarged view of the surface region of the lateral surface of the trunk. Similar ovoid cells are distributed at the base of aragonite spicules (arrows) with small cells (arrowheads). lon, longitudinal nerve tracts; m, muscle fibers; see Figure 1 for other abbreviations. Scale bars: A, G–I, 50 µm; B–F, 100 µm.
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For immunohistochemistry, paraffin-embedded samples were cut at 8-µm thickness except for one specimen cut at 10 µm (Fig. 5A–E). The sections were deparaffinized with xylene, rehydrated, and blocked with 10% BSA in PBST for 1 h at room temperature. They were transferred into primary antibody and kept overnight at 4°C, then washed and kept in secondary antibody for 3 h at room temperature. The post-stained sections were placed for about 30 min in 4'-6-diamidino-2-phenylindole (DAPI) (Sigma, 0.1 µg/ml in PBST) and rhodamine-conjugated concanavalin A (ConA) (Vector Laboratories, 1:1000 in PBST) for one-step staining of nuclei and cell membranes, respectively. The samples were observed using the same microscopes as described above.
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Figure 5. Structures of the cerebral cord, precerebral ganglia, and attached nerves. (A) Sagittal section showing the relationship between the cerebral cord and tubulin-containing cells at the oral shield. (B) Nerve bundles from the oral shield to precerebral ganglia. (C) Distinct regions of sensory cilia (a' and b') project and fasciculate into nerve bundles (a and b, respectively). Arrowhead indicates possible direct connection from cilia. (D) Some nerve bundles from sensory cilia run to the cerebral cord (arrowheads). Concanavalin A-positive fine neurites in the central part of precerebral ganglia do not display strong tubulin immunoreactivity (arrow). (E) Fasciculate nerve tracts from precerebral ganglia run into the neuropil of the cerebral cord (arrowheads). (F) Serotonin-like immunoreactive cells and neurites are abundant at the posterior and dorsal lobes of the cerebral cord. Small (arrow) and large (arrowhead) cells are distinguished as serotonin-positive cells. Anti-acetylated -tubulin antibody staining (Tube) with DAPI and/or Concanavalin A (ConA) (A–E). Anti-serotonin (5-HT) antibody staining (F). al, anterior lobe; dl, dorsal lobe; dn, dorsal root of nerves from the oral shield; pl, posterior lobe; see Figure 1 for other abbreviations. Scale bars: A, 100 µm; B, F, 50 µm; C–E, 20 µm.
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For serotonin labeling (Figs. 2B, 4G, H, 5F), the sections were incubated in a rabbit anti-serotonin antibody (Sigma, S5545, 1:500 in PBST) and then in an alkaline-phosphatase (AP)-conjugated rabbit IgG antibody (Molecular Probes, 1:400 in PBST). To visualize AP, a standard protocol was used (Agata et al., 1998).
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Figure 2. (A) A fixed female specimen of Chaetoderma japonicum. Anterior is toward the top and ventral is left. Asterisk is an artifactual slit. (B) The para-sagittal section (a lateral part of mouth opening, mo) shows the head region of C. japonicum and the relationship between the cerebral cord and the ventral nerve cord. Anti-serotonin antibody staining. The dotted line indicates the posterior end of two large ganglia (vn1 and vn2). See Figure 1 for abbreviations. Scale bars: A, 1 mm; B, 100 µm.
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Figure 4. (A) Horizontal section of the ventral and lateral nerve cords at the trunk. (B) Enlargement of a part of the nerve cords and neuropils. (C, D) The anterior end of the lateral nerve cords at the head region, showing the ganglionic structure (ln1 and ln2) (see Figure 1 for the position). (E) Horizontal section of the tail part showing the suprarectal commissure with a nerve to the ctenidium (ct) (arrowhead). (F) A horizontal section of the tail region showing a distinct neuropil in the suprarectal ganglion (closed dotted line). A large cell is indicated by the arrowhead. (G) An enlarged view of the ventral nerve cord with nerves from the ganglion (vn1) at the base of the pedal pit (arrowheads). (H) Ganglion 2 (vn2) and repeated ganglion-like ventral nerve cord. A densely serotonin-stained region is present in the neuropil of ganglion 2 (arrowhead). Hematoxylin staining (A–F) and serotonin immunostaining (G and H). m, muscle fiber; see Figure 1 for other abbreviations. Scale bars: A, E, F, 100 µm; B–D, G, H, 50 µm.
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A series of serial sections (Fig. 6) were incubated with a mouse anti-acetylated -tubulin primary antibody and an anti-mouse IgG secondary antibody conjugated to peroxidase as described previously (Shigeno and Yamamoto, 2002). After 3,3'-diaminobenzidine (DAB) reaction, they were stained with Mayer's hematoxylin solution (WAKO) and mounted as permanent slides.
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Figure 6. (A–E) Oblique horizontal sections showing acetylated -tubulin-positive neuropil compartments (np 1–4), fasciculated tracts (black arrowheads), larger nuclei (white arrowheads), and three lobes in the cerebral cord (al, pl, and dl) of Chaetoderma japonicum. The cerebral cord is surrounded by an outer membrane (arrows). The medial dotted line indicates a midline in A–C. (F) A dorsal view of the entire cerebral cord, reconstructed from oblique horizontal sections. The most dorsal section is outlined (star), and anterior is toward the top. (G) The dense tubulin-containing region of the neuropil compartment (np1) and another neuropil region in the anterior lobe (al). (H) Large nuclei (white arrowheads) and small scattered neuropils (asterisks) in the anterior and dorsa lobes. (I) In the anterior and posterior lobes (black arrowheads), fasciculated tracts connect a commissural pathway that is possibly connected to the ventral and lateral nerve cords (con). Some large nuclei are distributed (white arrowheads). Anti-acetylated -tubulin (Tub), and hematoxylin (Hem) double-staining (A-H, and H-I). See Figure 7 for summaries and abbreviations. Scale bars: A–E, G–I, 100 µm.
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Image processing
Panfocus images (Fig. 6A-H, except for F) were automatically prepared by stacking a series of three or four digitally captured optical sections, using the Nikon All-in-focus DM-255i program. Three-dimensional reconstruction of the cerebral cord-like ganglia was performed with scanned photographs of serial hematoxylin-stained sections (8-µm thickness) that were processed using Scion Image, ver. Beta 4.03, software according to the manufacture's protocol (http://www.scioncorp.com/ [accessed 6/09/2005]). Additional processing of images for contrast, brightness, and color balance was made as needed with Adobe Photoshop CS2 (Adobe Systems Inc., USA) or Paintshop Pro 7 (Jasc Software, USA). Schematic diagrams were created with Paintshop Pro 7 or Adobe Illustrator CS2 (Adobe Systems Inc., USA).
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Results
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The body of C. japonicum is externally divided into three parts: the cerebral region (the head region), the trunk, and the tail region (Figs. 1, 2A) (for terminology and general description, see Salvini-Plawen, 1985; Scheltema et al., 1994). The cerebral part includes a protrusible head-end with a circular oral shield (head shield or buccal shield), its mouth opening, and a pedal pit. It contains the cerebral cord-like ganglia, buccal ganglia, a purely muscular buccal apparatus, a radular sac, and salivary glands. The body wall consists of longitudinal, circular, and oblique body wall musculatures used in locomotion, but a foot sole is lacking. The trunk is cylindrical and is the widest part of the body. It contains the midgut, the digestive gland, and the dorsally situated gonad. The posterior tail region includes the single fused gonopericardial duct, the intestine, and the most posteriorly situated knob with the hemocoel, gametoduct, rectum, and a pair of bipectinate ctenidia in the bell-like mantle cavity.
In this head region, there is a larger neural mass at the dorsal side (Fig. 2B) that is difficult to observe in wholemount specimens (Fig. 3A). The neural mass has been called a brain (Hoffman, 1949) or cerebral ganglia (Salvini-Plawen, 1972, 1978, 1985), but here we call it the "cerebral cord-like ganglia" or simply the "cerebral cord" due to its morphological condition without obvious distinction between left and right ganglia and neuropils (see Discussion and Fig. 8 for a summary). In the ventral head region, the cerebral cord of C. japonicum connects to the ventral nerve cord through the cerebro-buccal connectives (Fig. 1, 2B). A pair of buccal ganglia is located at the posterior lateral position of the buccal mass (Fig. 2B).
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Figure 8. Generalized cerebral ganglia or cords in the context of early molluscan evolution. Neuropils, a mass of neurites enclosed by layers of cell bodies, are represented with dotted lines. Cord-like cerebral ganglia in Chaetoderma are shown with eight large precerebral ganglia, distinct lobes, and neuropil compartments. Phylogenetic relationships of some molluscan groups are still controversial but are depicted here in an evolutionary tree modified from Caron et al. (2006), (1) Stem-groups of Mollusca only known as fossils; ladder-like nervous system and anteriorly accumulated neurons might be present, as described in recent acoelomorphs (Reuter et al., 2001; Raikova et al., 2004). (2) Cerebral ganglia or cord-like ganglia as an anterior part of the archetypical molluscan tetraneural plan (Salvini-Plawen, 1972; Haszprunar, 1992). (3) Tubiform ‘Aplacophora’: (3a) Neomeniomorpha, medially centralized cord-like ganglia with six precerebral ganglia in Syngenoherpia (Salvini-Plawen, 1972) or a centralized pair of cerebral ganglia in Rupertomenia (Schwabl, 1955; see also Scheltema et al., 1994); (3b) Prochaetodermomorpha; an obviously distinct pair of cerebral ganglia with a long cerebral commissure in Prochaetoderma (Salvini-Plawen, 1972); (3c) Chaetodermomorpha; fusion to the one cerebral cord-like ganglion with many neuropil compartments in Chaetoderma (Hoffman, 1949; Salvini-Plawen, 1972; and the present study). (4–6) Some crown groups of Mollusca. (4) Polyplacophora; cerebral cord-like ganglia (cerebro-buccal ring) in the chiton Lepidochitona, with a simple neuropil (Gantner, 1989; see also Eernisse and Reynolds, 1994; Voronezhskaya et al., 2003; Wanninger and Haszprunar, 2003, for other species). (5) Tryblidiida (Monoplacophora); a pair of, but indistinct, swellings are considered as cerebral ganglia in Vema (Wingstrand, 1985; Haszprunar and Schaefer, 1997). (6) Other crown groups such as Gastropoda; a pair of distinct cerebral ganglia and simple neuropils in the archaegastropod Patella, or multiple lobes, neuropil compartments, fasciculated fiber tracts in many derived gastropods (Bullock and Horridge, 1965; Chase, 2002, for references).
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At the trunk, a pair of ventral and lateral nerve cords run from the cerebral cord to the end of the posterior part of the tail (Fig. 1), and they are detected by acetylated -tubulin (Fig. 3B–F) and serotonin-like immunoreactivity as a mass of long connectives of nerves (Fig. 2B). The radial nerves appear at a certain interval from the ventral and lateral nerve cords (Fig. 3C, E). They generally run from ventral to dorsal with a branching pattern (Fig. 3E), whereas some radial nerves run obliquely from ventral to dorsoanterior (Fig. 3B). A fine tubulin-positive longitudinal connective is present in the lateral region of the trunk (Fig. 3D, E). Wholemount immunocytochemistry revealed many tubulin-containing neural cells along the radial nerves at the head and trunk region (Fig. 3G). The cells usually have a single neurite, and their cell bodies are about 10–20 µm in diameter. Distal to the radial nerves, tubulin-positive cells are scattered together with cells of various sizes in the head region (Fig. 3H). The small cells are about 8 µm in diameter (the nuclei, about 5 µm) and the large cells are about 15 µm in diameter (the nuclei, 8 µm) with short bipolar or multipolar neurites (Fig. 3H). This condition is somewhat different at the surface of the trunk, where the cells are similar in size with ovoid cell bodies (Fig. 3I; cell body about 10 µm in diameter and nuclei about 8 µm).
Notable features in the ventral and lateral nerve cords are some large, distinct ganglionic masses with ovoid neuropils in the head region (Fig. 1), whereas serially repeated ganglion-like swellings are present from the posterior part of the head-to-tail region (Fig. 4A, B). In the head region, at least two distinct ganglia are identified in both of the ventral and lateral nerve cords (Figs. 2B, 4C, D). In the tail region, the ventral and lateral nerve cords finally form a continuous loop as the suprarectal ganglion, from which the nerves for ctenidia and posterior organs are derived (Fig. 4E). No ganglionic swellings are seen in the suprarectal ganglion, but a round distinct neuropil is identified in this ganglion (Fig. 4F). In addition to these histological features of the nerve cords, serotonin-like immunohistochemistry revealed two notable features: some nerves from the ganglionic mass (vn1) of the ventral nerve cord that project to the posterior region of the pedal pit (Fig. 4G), and densely labeled serotonin-positive fibers in the neuropils of the ganglia (vn2) (Fig. 4H).
Other nerves projecting from the cerebral cord are of two types. These are nerves to the oral shield, which possibly innervate sensory cilia (Fig. 5A), and the cerebro-buccal connectives, which innervate the buccal and radular sacs through the buccal ganglia (not shown). The tubulin-positive sensory cilia are distributed uniformly, and the nerves merge to form thick fasciculated bundles (Fig. 5B, C). The nature of the sensory cells and positions of these neuronal cell bodies are still largely unclear in this study, but it seems that these sensory nerves directly connect to the precerebral ganglia (Fig. 5B) and the cerebral cord (Fig. 5D). Tubulin immunoreactivity is evident in long thick tracts, and ConA-positive fine fibers also exist (Fig. 5D). The fasciculated tracts in the cerebral cord connect directly to the precerebral ganglia and possibly to sensory cells of the oral shield (Fig. 5E).
A histological study based on oblique horizontal sections reveals that the cerebral cord is lobed and is a single bilaterally symmetrical structure (see Fig. 6A–E for a series and 6F for a three-dimensional view) with the notable fine organization (Fig. 6G–I, and summary in Fig. 7). Eight large precerebral ganglia are detected in total. Each ganglion is distinct, with a thin membranous cover (Fig. 6A, B, arrows). The internal division of the cerebral cord is largely uncertain so far (Salvini-Plawen, 1972), but we identified three regions in the cerebral cord of C. japonicum: the anterior, posterior, and dorsal lobes (Fig. 6C–E). In the anterior lobe at the base of each precerebral ganglion, eight distinct and densely tubulin-positive compartments are evident (e.g., Fig. 6G). All projections from the precerebral ganglia and some nerves from the head-part converge into these neuropils. We name these distinct regions the first to fourth neuropil compartments of the anterior lobe (np1–4). Tracts from the neuropil compartments connect to a thick commissure that extends to the ventral and lateral nerve cords and cerebro-buccal connectives. The posterior lobe contains a large neuropil, and serotonin-like immunoreactivity is intense in the neurites and cells in this lobe (Fig. 5F). The dorsal lobe is located in the dorsal region of the anterior and posterior lobes, which are characterized by small neuropils (asterisks in Fig. 6H) and dense serotonin-like immunoreactivity (Fig. 5F). Some fasciculated tracts enter into this lobe from the anterior lobe (Fig. 6I). The large round nuclei are broadly distributed throughout the lobes (e.g., Fig. 6H, I). The large nuclei are about 16 µm in diameter compared with the 8 µm of cells in the precerebral ganglia and cerebral cords. It is likely that the large nuclei are particularly abundant at the cell body layers of the anterior and dorsal lobes.
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Discussion
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On the basis of earlier studies (e.g., Wirén, 1892; Heath, 1904, 1905; Hoffman, 1949), Salvini-Plawen (1972, 1978, 1985) has presented a synthesis of the neuroanatomy of the "Aplacophora." As seen in a proposed hypothetical ancestral mollusc (Salvini-Plawen, 1972, 1985, 1991, 2006; Haszprunar, 1992), the nervous system of Caudofoveata is primitively tetraneurous, consisting of ventral (pedal) and lateral nerve cords extending from the cerebral ganglia (Fig. 1). Solenogastres has a simpler neural organization in the head region, being regarded as primitive (Scheltema, 1993, 1996), although opposing views have been offered (Salvini-Plawen, 1985; Salvini-Plawen and Steiner, 1996).
One might assume that the many serially repeated swellings of the ventral and lateral nerve cords in the head region and trunk are an artifact caused by shrinkage of the body, because the longitudinal tracts (probably a mass of axonal bundles) are also repeatedly curved (e.g., Fig. 4D). However, this phenomenon has been described in various Caudofoveata species and also in relaxed specimens (G. Haszprunar, pers. obs.). Although the patterns of neuromuscular junctions are still unclear, these serial neural structures do not seem to correspond with serial body wall muscles or with any other structure of the body. Thus, this feature cannot be interpreted as segmentation or metamerism.
The anterior region of the ventral and lateral nerve cords of C. japonicum has an obvious ganglionated structure with distinct morphological units of different ganglionic size (vn1 and vn2 in Fig. 2B; ln1 and ln2 in Fig. 4C). Interestingly, the posterior end of these ganglia is restricted at the anterior ventro-lateral head region. The precise innervation patterns on the lateral nerve cords were not elucidated in this study, but some serotonin-containing nerves from the ganglia 1 (vn1) of the ventral nerve cords were observed in the posterior tissues of the pedal pit (Fig. 4G). This suggests that ganglion-like features may be related to a specialized sensory or motor function of the posterior part of the pedal pit such as muscle regulation or sensory detection of food in the substrate. Similar ganglionic features have also been described in the ventral and lateral nerve cords of other species in Caudofoveata and Solenogastres (Neomeniomorpha) (Salvini-Plawen, 1972, 1978). To address homologies between the ganglia of Aplacophora and other molluscan groups, more comprehensive ontogenetic and comparative studies are needed, since adult caudofoveates entirely lack a pedal sole and it is uncertain whether the posterior part of the head region of chaetoderms may correspond to the pedal region of other molluscan groups.
Each of eight distinct precerebral ganglia seems to contain a population of primary or secondary neurons connected from the sensory cells of the oral shield (Fig. 7). At the base of the precerebral ganglia, distinct neuropil compartments exist as a dense acetylated -tubulin-positive region with many fasciculated tracts toward the center of the cerebral cords. Distinct precerebral ganglia at the anterior head region have been reported in some species of Aplacophora (Fig. 8), where precerebral parts supply some nerves to the so-called atrial sense organ, a mechano-chemoreceptive area in a position comparable to that of the oral shield in the Caudofoveata (Salvini-Plawen, 1978, 1985; Haszprunar, 1986; Scheltema et al., 1994). To determine whether the precerebral ganglia of Solenogastres are homologous to those of Chaetoderma or Caudofoveata in general requires immunocytochemical studies in Solenogastres comparable to those presented here. In contrast, no such anterior complex centers have been reported in polyplacophorans and conchiferans such as tryblidiids (monoplacophorans) and basal gastropods (Fig. 8; and see also Bullock and Horridge, 1965; Eernisse and Reynolds, 1994; Haszprunar and Schaefer, 1997; Chase, 2002). Only derived or "advanced" gastropods and cephalopods show an even higher degree of specialization of the cerebral part of their nervous systems, which include elaborated chemosensory networks with numerous interneurons, fasciculated tracts, and many distinct neuropils (see Young, 1971; Budelmann, 1995; Chase, 2002). Thus, the anterior region of the cerebral cord of C. japonicum, with its distinct neuropil compartments, is among the most highly differentiated nervous systems in the basal molluscs studied to date (Fig. 8).
The anterior and posterior subdivisions of the cerebral cord-like ganglia were first described in Chaetoderma nitidulum (Hoffmann, 1949). In addition to their subdivisions, we distinguished one additional subdivision as the dorsal lobe in the cerebral cord of C. japonicum. Such a tripartite pattern has never been described in other molluscs, Solenogastres, Polyplacophora, and Tryblidiida (Plate, 1898; Lemche and Wingstrand, 1959; Bullock and Horridge, 1965; Salvini-Plawen, 1972, 1978, 1985; Wingstrand, 1985; Eernisse and Reynolds, 1994; Haszprunar and Schaefer, 1997). The functional roles of the subdivisions are unclear owing to the lack of cellular-level neurotracing studies in chaetoderms. However, on the basis of neural bundle patterns (Fig. 7), we suggest that the anterior, posterior, and dorsal lobes have been specialized as higher order sensory centers to integrate mechano- and chemosensory information from the oral shield and whole head regions, or as moto-sensory associative centers related to the development of both precerebral ganglia and ganglionated ventral and lateral nerve cords. Comparison of these subdivisions with the cerebral ganglia of gastropods also suggests the following: (i) Olfactory or mechano-chemosensory centers are basically situated in the antero-lateral parts of the cerebral ganglia, and second- or higher order chemosensory interneurons develop in the anterior bases of the ganglia (Van Mol, 1967; Sonetti et al., 1982; Chase and Tolloczko, 1989, 1993; Chase 2000). (ii) The basal or ventral regions of the cerebral ganglia usually include many motor neurons with tight connectivity to other motor centers such as pedal and pleural parts (Benjamin and Ings, 1972; Hernádi et al., 1984; Balaban and Chase, 1990; Ierusalimsky et al., 1994). (iii) The anterior dorsal region of the cerebral ganglia tends to differentiate into association or integration centers, as seen in the procerebrum of pulmonates (Hanstroem, 1928; Bargmann, 1930; Van Mol, 1967; Chase and Tolloczko, 1989; and see Chase, 2002, for a recent review). The presence of many small and distinct neuropils in C. japonicum indicates that the dorsal lobe may include a pool of associative interneurons.
Intense serotonin-like immunoreactivity is present in the nervous system of C. japonicum (Figs. 2B, 4G, H, 5F). It is not clear whether the antibody used in our study detects serotonin itself; however, the labeling patterns for C. japonicum are similar and partially comparable to those of other molluscs. In the nervous system of chitons, serotonergic cells are uniformly distributed, but staining is particularly dense in the pedal cords, with some cells in the cerebral cord (Friedrich et al., 2002; Voronezhkaya et al., 2003; see also Moroz et al., 1994). In a scaphopod (Wanninger and Haszprunar, 2003) and in bivalves (Matsutani and Nomura, 1986; Croll et al., 1995), the labeled cells are also uniformly localized in pedal, cerebral, and other peripheral ganglia. In gastropod embryos and adults, a similar expression pattern is restricted to the anterior, medio-dorsal parts of cerebral ganglia, as seen in the vetigastropod Haliotis (Barlow and Truman, 1992; Hinman et al., 2003), the caenogastropod Crepidula (Dickinson et al., 1999), and in many heterobranch species, e.g., Helisoma (Goldberg and Kater, 1989), Lymnaea (Croll and Chiasson, 1989), Aplysia (Dickinson et al., 2000), and Clione (Slatternly et al., 1995), and even in cephalopods, e.g., Idiosepius and Octopus (Shigeno et al., unpubl. data). The labeled cells in C. japonicum are located in the ventral nerve cords (or pedal components), as seen in most of the above-mentioned molluscs. Interestingly, most cells in the precerebral ganglia and the anterior lobe did not exhibit intense immunoreactivity, whereas serotonergic cells exist in the dorsal and posterior lobes of the cerebral ganglia. These comparative data indicate that the serotonergic cells have already been uniformly present in an early molluscan nervous system, and they tend to localize in independent lineages of Caudofoveata, Bivalvia, Gastropoda, and Cephalopoda during molluscan evolution. Further comparative studies are required to identify the homological patterns of serotonergic cells in various molluscan lineages.
Additionally, some derived gastropods such as pulmonates have distinctive chemosensory interneurons called "small cells" or "globuli cells," which are characterized by having round and chromatin-rich cell bodies organized into clusters located at the anterio-dorsal regions of the cerebral ganglia (see Bullock and Horridge, 1965; Chase, 2002, for many references). Such small cells are not found in C. japonicum, indicating that this species has less elaborate sensory processing abilities than those of derived gastropods.
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Acknowledgments
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We thank Luitfried von Salvini-Plawen (University of Vienna) for reading the manuscript and for important suggestions. We thank also Charles Derby (Georgia State University) for revision of this manuscript. For materials, we are indebted to the crew of R/V Tansei-Maru and researchers on board cruise KT-02-5 for their kind assistance in obtaining the material. This research was supported in part by a grant from the Japanese Society for Promotion of Science (no. 15340175, 18770063) to T.S.
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Footnotes
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Received 19 January 2007; accepted 12 June 2007.
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Abstract
Introduction
