Skip to main content

Peripheral sensory neurons govern development of the nervous system in bivalve larvae


Recent findings regarding early lophotrochozoan development have altered the conventional model of neurogenesis and revealed that peripheral sensory elements play a key role in the initial organization of the larval nervous system. Here, we describe the main neurogenetic events in bivalve mollusks in comparison with other Lophotrochozoa, emphasizing a novel role for early neurons in establishing larval nervous systems and speculating about the morphogenetic function of the apical organ. We demonstrate that during bivalve development, peripheral sensory neurons utilizing various transmitters differentiate before the apical organ emerges. The first neurons and their neurites serve as a scaffold for the development of the nervous system. During veliger stage, cerebral, pleural, and visceral ganglia form along the lateral (visceral) nerve cords in anterior-to-posterior axis. The pedal ganglia and corresponding ventral (pedal) nerve cords develop much later, after larval settlement and metamorphosis. Pharmacological abolishment of the serotonin gradient within the larval body disrupts the navigation of “pioneer” axons resulting in malformation of the whole nervous system architecture. Comparative morphological data on neurogenetic events in bivalve mollusks shed new light on the origin of the nervous system, mechanisms of early axon navigation, and sequence of the tetraneurous nervous system formation. Furthermore, this information improves our understanding of the basic nervous system architecture in larval Bivalvia and Mollusca.

Unsolved questions on lophotrochozoans neurodevelopment

Lophotrochozoa, a morphologically varied group of animals, include worm-like forms, shelled animals, and soft-bodied slugs, which makes searching for common anatomical and morphological features in adults daunting and unsuccessful to date. However, during lophotrochozoan early larval development, ancestral morphological and molecular patterns are present with formation of the nervous system being among the most well-conserved and prominent features of this development. Therefore, development of the nervous system has long attracted the attention of comparative biologists for testing evolutionary and phylogenetic hypotheses.

Although for more than 100 years, researchers have attempted to understand the details and general patterns of this development utilizing available methodology [1,2,3,4,5,6], some major questions remain concerning the following points: (1) homologous characters that have been reduced or strongly modified in adults by adaptation (for example, simplification of the organization of the nervous system); (2) phylogenetic relationships among lophotrochozoans and their kinship with other groups of animals (Ecdysozoa and Deuterostomia); and (3) the potential body plan (bauplan) of the last common ancestor for particular groups. Answering these questions requires a variety of methodological approaches, including morphological/physiological and molecular methods.

Bivalves are seldom used in studies of lophotrochozoan neurodevelopment. These animals, with sedentary or free-living lifestyles as adults, develop from free-swimming larvae that hatch as trochophores, develop through the veliger stage, undergo metamorphosis, and then settle [1,2,3,4, 6].

Adult and larval neuroanatomy: correction of morphological misinterpretations

Previous investigations on bivalves have concluded that the central nervous system (CNS) of adults consists of paired cerebropleural and visceral ganglia, the latter typically fused and connected to the cerebropleural ganglia by longitudinal lateral cords [6,7,8,9,10,11,12] (Fig. 1a). These neurostructures innervate most internal organs. Fused pedal ganglia belong to CNS and innervate the foot, and are connected to the cerebropleural ganglia via the ventral (pedal) cords (Fig. 1a) [4, 13]. Accordingly, the nervous system of adult bivalves is tetraneural, consisting of two prominent pairs of longitudinal neurite bundles (cords) ganglionated to various extents [4, 6,7,8,9,10,11,12,13].

Fig. 1

Architecture of the nervous system of adult bivalves (a) and larvae before and after metamorphosis (bd). a Tetraneurous nervous system (ventral + pedal cords) of the adult bivalve. b Nervous system of bivalve larvae. c The tetraneurous bivalve nervous system after settlement/metamorphosis. d Nervous system of a mactrid juvenile Mactromeris polynyma from plankton as visualized by whole-mount immunostaining for FMRFamide (green) and acetylated alpha-tubulin (red) (the arrows indicate the lateral nerve cord and ventral (pedal) nervous cords). CPG cerebropleural ganglion, PNC pedal nervous cord, PG pedal ganglion, LNC lateral nerve cord, VG visceral ganglion, AO/CG apical organ/cerebral ganglion, PlG pleural ganglion

Research on the developmental neurogenesis of bivalves is limited and interpretation of larval morphology often contradictory [3, 4, 6, 14, 15]. Findings published to date do not allow for an exact phylogenetic placement of bivalves within Mollusca or the evolutionary origin of the last common ancestor of bivalves and the other molluskan taxa to be determined. A number of evolutionary hypotheses concerning the origin of bivalves and their relationship to potential sister groups within the mollusks (Testaria, Conchifera, Diasoma), as well as the relationship of mollusks to other lophotrochozoan taxa (the annelid-mollusk and mollusk-entoproct hypotheses) have been proposed [4, 16,17,18,19,20]. However, available developmental data provide no clear support for any of these hypotheses. Moreover, the neurogenetic mechanisms by which the ganglia develop along the nerve cords or organize into a two-ring tetraneural structure remain unclear.

In particular, the variability of early neurogenetic events in bivalve species [3, 14, 15, 21] and the lack of information on the late stages of development are the major barriers at present to fully understand neurogenesis in mollusk larvae. Nevertheless, studies have shown that bivalve larvae have a ganglionic morphology of the nervous system and paired ganglia (cerebral and visceral) lie along the lateral (visceral) nerve cord in veliger larvae of mytilids [3, 22], ostreids [14, 22,23,24], and mactrids [25] Additional paired ganglia, identified as pedal ganglia sitting on lateral nerve cords in these same larvae, are very probably the same as the pleural ganglia detected earlier in larvae of Ostrea edulis, Crassostrea virginica, and Mytilus galloprovincialis [1, 23, 26].

Accordingly, although morphology varies, the nervous system of bivalve veligers consists of three paired ganglia (cerebral, pleural, and visceral) connected by the lateral cord (Fig. 1b). However, larval neuromorphology also differs from the adult nervous system with respect to the degree of fusion of the cerebral and pleural ganglia, indicating that this ganglionic fusion occurs during late development. As published previously [13, 27], cerebral and pleural ganglia develop independently in some mollusk larvae and only fuse in the adults (e.g., in Dreissena, Mactra, and Arca pernoides. Location and timing of the development of paired pedal ganglia remain unknown. Researchers schematically depict pedal ganglia and their connections with cerebral ganglia, emphasizing that in larvae, paired pedal (ventral) and lateral (visceral) nerve cords together build a tetra-cord structure [4, 6, 20]. Nonetheless, no detailed description of the development of the pedal nervous system has yet appeared.

Here, we first review the findings on later developmental stages (after the settlement and metamorphosis of pediveligers), when pedal ganglia with their cerebropleuro-pedal connections (pedal cords) arise (Fig. 1c). Immunostaining of larvae (caught in plankton) during the late stages of development revealed that the pedal ganglia develop during late ontogeny only (Fig. 1d) and subsequently fuse later as shown for other bivalves [13]. Altogether, the early set of paired ganglia, including cerebral, pleural, and visceral, is more fully developed in the veliger stages, while the ventral part of the nervous system (paired pedal ganglia) differentiate later on (Fig. 1c, d), in association with the foot formation and functions. Thereafter, we consider the underlying cellular mechanisms in more detail, including how the various parts of the nervous system are formed during early development of bivalve mollusks.

Early events in bivalve neurodevelopment

It is generally accepted that the first neurons to appear in larvae are sensory cells, responsible for converting external stimuli from the environment and appropriately regulating larval behavior and development in Lophotrochozoa [28,29,30], Echinodermata [31, 32], and Cephalochordata [33]. Most commonly, the apical organ (AO) is the first sensory structure to appear. This organ consists of neurons that employ several different transmitters to regulate the rate of development and locomotion of larvae as well as settlement and metamorphosis [30, 34,35,36].

However, some evidence indicates that simultaneously with or even before the neurotransmitter expression in AO cells, solitary sensory cells appear on the periphery of the body of several trochozoan larvae. The function of these pioneer sensory neurons remains unknown. One hypothesis is that they help to provide scaffolding for the definitive nervous system [14, 21, 37,38,39].

Recent studies on the bivalve mollusk Crassostrea gigas confirm the early appearance of peripheral sensory neurons [14] and, together with the experimental findings on Mytilus trossulus, larvae presented here indicate the morphogenetic significance of these cells in connection with formation of the nervous system.

In greater detail, two groups of sensory cells can be detected on the dorsal and ventral sides of the trochophore larvae of the oyster C. gigas prior to the emergence of the AO. The neurites of the FMRFamide-immunoreactive (FMRF-ir) neurons in the dorsal sensory group (dorsal sensory center, DSC) extend towards the neurons of the ventral sensory group (ventral sensory center, VSC) (Fig. 2a). As neurogenesis proceeds, these long axons of the dorsal neurons terminate near the ventral cells and these interconnections are organized into paired dorso-ventral cords (Fig. 2b). However, there is no direct connection to the differentiating cells of the AO located between the dorso-ventral cords [14].

Fig. 2

(Schematic drawings adapted by Olga Kharchenko from Yurchenko et al. [14])

Neurogenesis of the oyster Crassostrea gigas: FMRF-immunoreactive nervous system. a Early neurons that appear in the dorsal (DSC) and ventral (VSC) peripheral sensory centers, before appearance of transmitters in the apical organ (AO). b, c They provide morphogenetic scaffolding for the anlagen of the lateral nerve cords (LNC) during oyster development. vn ventral neurons, pn posterior neurons

At later stages, the AO and neurons located posteriorly extend their processes to the ventral cells and along the dorso-ventral cord. Thus, in the veliger of C. gigas, the anlagen of the lateral nerve cords have already been formed (Fig. 2c). Interestingly, the early dorsal and ventral sensory centers are transient, and their cells and processes cease to express immunoreactivity and probably disappear during the veliger stage. However, the questions whether the cells of the ventral sensory centers are precursors of the pleural ganglia and the posterior cells are precursors of the visceral ganglia remain unanswered.

We speculate that early peripheral sensory cells provide scaffolding for the differentiating nervous system, marking the location for subsequent cell migration and organization of the major central ganglia and nerve cords. The exact sources of neuronal progenitors during lophotrochozoan development are largely unknown. Some data indicate that cells in cerebral placodes give rise to neurons within the cerebral ganglia [40, 41]. Several mechanisms for the differentiation of ganglionic neurons have been proposed, involving local nerve-dependent signals [42], migration of neuronal precursors from outside into ganglia anlagen [43], and/or the glial or sensory epithelium (unpublished data). However, none of these hypotheses has experimental support to date.

The transmitters employed by the primary sensory neurons

At present, we have no explanation why serotonin (5-HT) and FMRFamide-related peptides (FMRF-RP) are the primary transmitters expressed by the first sensory neurons that appear during bivalve development. FMRFamide and related peptides, in addition to 5-HT, influence multiple physiological processes, including heart rate, blood pressure, motility of the gut, feeding, and locomotory behavior in both adult and larval invertebrates [44,45,46,47]. The peripheral sensory neurons in the larvae of different lophotrochozoan groups contain FMRF-RP (gastropod mollusks [48,49,50]) or 5-HT and FMRF-RP in different neurons (annelids [51] and bivalve mollusks [3, 14]) or in one and the same neuron (polyplacophoran mollusk [52] and annelids [53]).

Neurons with different immunoreactivity (FMRF-RP and 5-HT) innervating several larval organs may perform common functions, e.g., affecting the beating of cilia or innervating muscles [22, 54,55,56,57,58]. Thus, both may be involved in the regulation of locomotion, distribution, and defensive behavior. In addition, the FMRFamide peptide family modulates ligand-gated channels that transport Na+ and associated water, rendering larvae resistant to osmotic pressure [29, 59,60,61,62]. FMRFamide, expressed in the early peripheral neurons of bivalve and gastropod larvae [3, 14, 21, 48], has been suggested to modulate their behavior in response to environmental changes (for example, in salinity, temperature, and pH) in their ecological niche, where appropriate movement in the water column determines successful survival. However, more experiments on the role of early sensory neurons in larval development are clearly required.

Neurites of sensory neurons play important roles in morphogenetic events

Neurogenesis in several groups of Lophotrochozoa has been found to begin with the emergence of peripheral neurons. Importantly, the locations of these early larval neurons vary in different species. In the bivalve mollusk Mytilus trossulus, they appear in the episphere and their processes organize into two nerve bundles that run in parallel along the ventral side of the larvae [3] (Fig. 3a). In the polyplacophora Ischnochiton hakodadensis, the bodies of these early cells are located in the episphere and their long axons extend underneath the neuropil of the AO and thereafter caudally in two bundles on the ventral side [48] (Fig. 3a).

Fig. 3

Schematic depiction of the location of early sensory elements in representative members of different lophotrochozoan groups. The long axons of early peripheral neurons (gray color) are arranged similarly in two parallel nerve bundles along the ventral side of the larvae, where the anlagen of the lateral nerve cords will later form. a Larvae of the mussel Mytilus trossulus and the chiton Ischnochiton hakodadensis; b Larva of the oyster Crassostrea gigas; c Larva of the gastropods Tritonia diomedea, Aplysia californica, and Lymnaea stagnalis; d Larva of the annelids, Phyllodoce maculata and Platynereis dumerilii

Recently, we demonstrated that in the trochophore larvae of C. gigas, early FMRF-ir peripheral cells differentiate in the hyposphere and send their long axons from the dorsal to the ventral side [14] (Fig. 3b). In the gastropod mollusks Tritonia diomedea, Aplysia californica, and Lymnaea stagnalis, peripheral cells are located caudally in the hyposphere and their processes run anteriorly along the curvature of the larval body towards the AO and then turn ventrally [21, 37, 50, 63, 64] (Fig. 3c). In the polychaetes Phyllodoce maculata and Platynereis dumerilii, the solitary early cells are located in the most caudal region of the hyposphere, their long axon bifurcating close to the cell body with two long processes turning to the ventral side and running anteriorly [51, 53] (Fig. 3d).

All early peripheral cells bear cilia and are considered to be sensory [21]. The prominent features of the early sensory peripheral cells in the larval body are their long axons. Independent of the location of the perikaryon, episphere or hyposphere, dorsally or ventrally, the emerging axons extend towards and follow the ventral side in two parallel longitudinal lateral paired cords. Thus, the pioneer processes ultimately delicate a position or configuration for future developed lateral nerve cords, which form by several pathways (dorso-ventral way or anterior–posterior way, or posterior–anterior way). In general, longitudinal nerve cords are a common feature for non-lophotrochozoan and lophotrochozoan protostomes, evolutionarily ancient and can be considered to be an ancestral character of Bilateria.

The role of serotonin in the navigation of the axons of early neurons

During development, the navigation and guidance of dendritic and axonal extensions and connections between neurons and non-neuronal targets are strictly regulated by a multitude of growth and transcriptional factors, adhesion molecules, and target receptors [65, 66]. Among the well-studied factors, 5-HT has been found to be a navigational cue during neural development in vertebrates and invertebrates [67, 68].

In the gastropod mollusk Helisoma, 5-HT and dopamine inhibit the elongation of neurites of specific identified neurons [69,70,71], while the initial phase of the process, 5-HT can also inhibit neurite growth [71, 72].

During early larval development, the primary source of 5-HT is the apical organ (AO), where specific neurons secrete transmitter via their varicose neuropil [73]. Apical restriction of secretion in this manner results in a gradient of 5-HT along the anterior–posterior axis of the larval body and this may help to navigate the axons of peripheral sensory neurons. Clearly, application of 5-hydroxytryptophan (5-HTP), the precursor of 5-HT, to trochophores of the annelid Platynereis dumerilii increases levels of 5-HT within AO neurons specifically and influences the rate of larval development without altering the structure of the major nerve elements. In contrast, application of 5-HT itself, increasing the concentration of this transmitter throughout the larval body and thereby abolishing the gradient, causes irreversible structural damage to the larval nervous system and, ultimately, larval death [21].

We have performed similar pharmacological manipulations during the development of the mussel Mytilus trossulus as described for Platynereis [21]. The experimental data presented in Fig. 4 demonstrate normal neurodevelopment (Fig. 4a–a2) and larval nerve elements malformed by application of 5-HT (10−6 M to early trochophores, up to the veliger stage) (Fig. 4b–b2). Numerous nerve processes were located chaotically in experimental larvae, reminiscent of neuronal sprouting. No compact structures, such as neurite bundles characteristic of the corresponding stage of normal development, were detected. These findings are consistent with previous hypothesis that 5-HT produced by AO neurons and released from the AO compact neuropil is involved in navigation of the long axons of early peripheral sensory cells. Further investigations to gain a deeper understanding of the action of 5-HT in axonal navigation and its consequence for nervous system formation in lophotrochozoan larvae are required.

Fig. 4

Effect of serotonin on the growth of FMRFamide-immunoreactive processes during development of the mollusk Mytilus trossulus. Mussel trochophores (32 hpf) were cultured in filtered seawater in the presence of 5-HT (10−6 M) or without any additives (the control). Larvae were collected at the veliger stage (60 hpf) and subjected to whole-mount immunostaining for FMRFamide (green) and acetylated alpha-tubulin (blue) as described earlier [14]. ab2 right lateral view of untreated (a, a2) and 5-HT-treated (b, b2) larvae. a1, b1 Ventral view of untreated and 5-HT-treated larvae. Note the enhanced sprouting and absence of compact ventral neurite bundles in the 5-HT-treated larvae. CG cerebral ganglion, LNC lateral nerve cord, PlG pleural ganglion

Involvement of neurons that secrete acetylcholine, serotonin, and FMRFamide in the development of bivalve mollusks

Immunolabeling is widely utilized to visualize differentiation in specific nerve elements of various taxa, but, unfortunately, only a limited number of commercial antibodies are applicable for invertebrates. Among these, antibodies against 5-HT and FMRF-RP are most commonly used. In addition, antibodies directed towards acetylated alpha-tubulin provide a marker for neuronal processes and some neuronal cell bodies in annelids and mollusks (personal data, [74,75,76]).

Histochemical reactions and immunochemical labeling against enzymes involved in biochemical degradation have been employed in attempts to visualize acetylcholine-containing (ACh-containing) regions as well as individual cells and nerve bundles; but these approaches in many cases have produced inconsistent results which were hard to interpret. In our recent study on neurogenesis in C. gigas, we showed that an antibody against the vesicular acetylcholine transporter (VAChT) is a reliable marker of ACh-containing elements [14]. With this antibody, we could demonstrate that at the trochophore stage, VAChT-positive reaction visualizes neurons within the AO as well as posterior neurons (Fig. 5). At the veliger stage, VAChT-ir neurons are present in cerebral and pleural ganglia and VAChT-ir fibers are found along the lateral nerve cords [14]. In addition, immunolabeling and whole-mount in situ hybridization for choline acetyltransferase were shown to identify ACh-containing elements in the central nervous system and peripheral neurons of gastropods, bivalves, and annelids [14, 26, 77,78,79,80,81].

Fig. 5

(Schematic drawings adapted by Olga Kharchenko from Yurchenko et al. [14])

Arrangement of neurons expressing different transmitters at the veliger stage of the bivalve mollusk Crassostrea gigas. Only the right side of the body is shown. FMRFamide-immunoreactive fibers first grow dorso-ventrally and then turn posteriorly. 5-HT-ir fibers grow in anterior-to-posterior direction from the apical organ (AO), while VAChT-ir grows in the posterior-to-anterior direction towards the AO. Altogether, these processes form a compact neurite bundle along the ventral side of the larval body. The right and left bundles run in parallel on the respective sides of the larval body, organizing rudiments of the paired lateral nerve cords. AO apical organ, dn dorsal neurons, vn ventral neurons, pn posterior neurons


In summary, neurogenesis in bivalve mollusks begins with the emergence of peripheral FMRF-ir cells with their growing processes and apical 5-HT-ir sensory cells. Together, these early elements delineate the main structure of the nervous system, including the apical/cerebral ganglia and paired nerve cords. VAChT-ir cells later appear in the periphery and their processes follow the pathway already established by early “pioneer” neurons (Fig. 4). Both early 5-HT-ir and FMRF-ir fibers grow in an anterior-to-posterior direction, from the AO to the ventral portion of the larvae, while the processes of VAChT-ir cells extend in the opposite direction sending axons towards the AO and along the ventral part of the larvae. In bivalve mollusks, FMRF-ir and VAChT-ir neurites participate in the formation of nerve cords to a greater extent than 5-HT-ir neurites. At the same time, a gradient of 5-HT probably plays an important role in the navigation of FMRF-ir axons and, finally, in formation of the lateral nerve cords.

The second neuronal cord, i.e., the pedal (ventral) cord with associated pedal ganglia, appears late in bivalve development, after pediveliger settling. Further research on bivalve neurogenesis, with a particular focus on VAChT-ir structures, is required to determine the cellular compositions of the AO and the central and peripheral nervous systems of veligers in detail.

Future perspectives

At present, many major questions and gaps remain regarding the cellular and molecular mechanisms of neurogenesis in bivalve mollusks:

  1. 1.

    Are some of the early neurons a prerequisite for the main ganglia of the definitive nervous system in mollusks?

  2. 2.

    When do ganglia fuse and what is the underlying mechanism and in response to what stimulus it starts?

  3. 3.

    Why does the pedal nervous system appear later in the development of bivalve mollusks and how is its development connected to that of the foot? What happens to the pedal ganglia in mollusks where the foot is resorbed (cementing species)?

  4. 4.

    Do mollusks have a peripheral (autonomous) nervous system?

  5. 5.

    How and where (placodes, neuroectoderm, and ectoderm) do neural precursors form and neuroblasts migrate?

  6. 6.

    What characters of the early development are common for various taxa and thus may reflect the neuroarchitecture of the last common ancestor?

Availability of data and materials

Not applicable.


  1. 1.

    Erdmann W. Nr. 5. Ober die Entwicklung und die Anatomie der ansatzreifen Larve von Ostrea edulis mit Bemerkungen iiber die Lebensgeschichte der Auster. Untersuchungen iiber die Lebensgeschichte der Auster; 1934. p. 1–25.

  2. 2.

    Nielsen C. Trochophora larvae: cell lineages, ciliary bands, and body regions. 1. Annelida and Mollusca. J Exp Zool B Mol Dev Evol. 2004;302:35–68.

    Article  Google Scholar 

  3. 3.

    Voronezhskaya EE, Nezlin LP, Odintsova NA, Plummer JT, Croll RP. Neuronal development in larval mussel Mytilus trossulus (Mollusca: Bivalvia). Zoomorphology. 2008;127:97–110.

    Article  Google Scholar 

  4. 4.

    Wanninger A. Evolutionary developmental biology of invertebrates 2: lophotrochozoa spiralia. Wien: Springer; 2015.

    Google Scholar 

  5. 5.

    Martín-Durán JM, Pang K, Børve A, Lê HS, Furu A, Cannon JT, Jondelius U, Hejnol A. Convergent evolution of bilaterian nerve cords. Nature. 2018;553:45–50.

    Article  Google Scholar 

  6. 6.

    Schmidt-Rhaesa A, Harzsch S, Purschke G. Structure and evolution of invertebrate nervous systems. Oxford: Oxford University Press; 2016.

    Google Scholar 

  7. 7.

    De Biasi S, Vitellaro-Zuccarello L. Distribution of 5HT-immunoreactivity in the pedal ganglion of Mytilus galloprovincialis. Cell Tissue Res. 1987;249:111–6.

    Article  Google Scholar 

  8. 8.

    Vitellaro-Zuccarello L, De Biasi S, Bernardi P, Oggioní A. Distribution of serotonin-, gamma-aminobutyric acid- and substance P-like immunoreactivity in the central and peripheral nervous system of Mytilus galloprovincialis. Tissue Cell. 1991;23:261–70.

    CAS  Article  Google Scholar 

  9. 9.

    Too CKL, Croll RP. Detection of FMRFamide-like immunoreactivities in the sea scallop Placopecten magellanicus by immunohistochemistry and Western blot analysis. Cell Tissue Res. 1995;281:295–304.

    CAS  Article  Google Scholar 

  10. 10.

    Henry M, Benlinmame N, Belhsen O, Jule Y, Mathieu M. Immunohistochemical localization of FMRFamide-containing neurons and nerve fibers in the ganglia and the gonad wall of the scallop, Pecten maximus (L). Neuropeptides. 1995;28:79–84.

    CAS  Article  Google Scholar 

  11. 11.

    Siniscalchi A, Cavallini S, Sonetti D, Sbrenna G, Capuano S, Barbin L, Turolla E, Rossi R. Serotonergic neurotransmission in the bivalve Venus verrucosa (Veneridae): a neurochemical and immunohistochemical study of the visceral ganglion and gonads. Mar Biol. 2004;144:1205–12.

    CAS  Article  Google Scholar 

  12. 12.

    Meechonkit P, Kovitvadhi U, Chatchavalvanich K, Sretarugsa P, Weerachatyanukul W. Localization of serotonin in neuronal ganglia of the freshwater pearl mussel, Hyriopsis (Hyriopsis) bialata. J Mollus Stud. 2010;76:267–74.

    Article  Google Scholar 

  13. 13.

    Bullock TH, Horridge GA. Structure and function in the nervous systems of invertebrates. San Francisco: WH Freeman; 1965.

    Google Scholar 

  14. 14.

    Yurchenko OV, Skiteva OI, Voronezhskaya EE, Dyachuk VA. Nervous system development in the Pacific oyster, Crassostrea gigas (Mollusca: Bivalvia). Front Zool. 2018;15:10.

    Article  Google Scholar 

  15. 15.

    Pavlicek A, Schwaha T, Wanninger A. Towards a ground pattern reconstruction of bivalve nervous systems: neurogenesis in the zebra mussel Dreissena polymorpha. Org Divers Evol. 2018;18:101–14.

    Article  Google Scholar 

  16. 16.

    Salvini-Plawen LV, Steiner G. The testaria concept (polyplacophora + conchifera) updated. J Nat Hist. 2014;48:2751–72.

    Article  Google Scholar 

  17. 17.

    Friedrich S, Wanninger A, Brückner M, Haszprunar G. Neurogenesis in the mossy chiton, Mopalia muscosa (Gould) (polyplacophora): evidence against molluscan metamerism. J Morphol. 2002;253:109–17.

    Article  Google Scholar 

  18. 18.

    Haszprunar G, Wanninger A. Molluscs. Curr Biol. 2012;22:R510–4.

    CAS  Article  Google Scholar 

  19. 19.

    Kocot KM, Cannon JT, Todt C, Citarella MR, Kohn AB, Meyer A, Santos SR, Schander C, Moroz LL, Lieb B, Halanych KM. Phylogenomics reveals deep molluscan relationships. Nature. 2011;477:452–6.

    CAS  Article  Google Scholar 

  20. 20.

    Wanninger A. Shaping the things to come: ontogeny of lophotrochozoan neuromuscular systems and the tetraneuralia concept. Biol Bull. 2009;216:293–306.

    Article  Google Scholar 

  21. 21.

    Nezlin LP, Voronezhskaya EE. Early peripheral sensory neurons in the development of trochozoan animals. Russ J Dev Biol. 2017;48:130–43.

    Article  Google Scholar 

  22. 22.

    Dyachuk V, Wanninger A, Voronezhskaya EE. Innervation of bivalve larval catch muscles by serotonergic and FMRFamidergic neurons. Acta Biol Hung. 2012;63:221–9.

    CAS  Article  Google Scholar 

  23. 23.

    Ellis I, Kempf SC. Characterization of the central nervous system and various peripheral innervations during larval development of the oyster Crassostrea virginica. Invertebr Biol. 2011;130:236–50.

    Article  Google Scholar 

  24. 24.

    Ellis I, Kempf SC. The presence and location of small cardioactive-like peptides in larvae of Crassostrea virginica. Invertebr Biol. 2011;130:251–63.

    Article  Google Scholar 

  25. 25.

    Kreiling JA, Jessen-Eller K, Miller J, Seegal RF, Reinisch CL. Early development of the serotonergic and dopaminergic nervous system in Spisula solidissima (surf clam) larvae. Comp Biochem Physiol A Mol Integr Physiol. 2001;130:341–51.

    CAS  Article  Google Scholar 

  26. 26.

    Raineri M. Is a mollusc an evolved bent metatrochophore? A histochemical investigation of neurogenesis in Mytilus (Mollusca: Bivalvia). J Mar Biol Assoc UK. 1995;75:571–92.

    Article  Google Scholar 

  27. 27.

    Heath H. The anatomy of the pelecypod family Arcidae. Trans Am Philos Soc. 1941;31:287–319.

    Article  Google Scholar 

  28. 28.

    Glebov K, Voronezhskaya EE, Khabarova M, Ivashkin E, Nezlin LP, Ponimaskin EG. Mechanisms underlying dual effects of serotonin during development of Helisoma trivolvis (Mollusca). BMC Dev Biol. 2014;14:14.

    Article  Google Scholar 

  29. 29.

    Vekhova E, Ivashkin E, Yurchenko O, Chaban A, Dyachuk V, Khabarova M, et al. Modulation of Mytilus trossulus (Bivalvia: Mollusca) larval survival and growth in culture. Acta Biol Hung. 2012;63:230–4.

    Article  Google Scholar 

  30. 30.

    Voronezhskaya EE. Apical sensory neurones mediate developmental retardation induced by conspecific environmental stimuli in freshwater pulmonate snails. Development. 2004;131:3671–80.

    CAS  Article  Google Scholar 

  31. 31.

    Yoshihiro M, Keiko W, Chieko O, Akemi K, Baba SA. Regulation of ciliary movement in sea urchin embryos: dopamine and 5-HT change the swimming behaviour. Comp Biochem Physiol Part C Comp Pharmacol. 1992;101:251–4.

    Article  Google Scholar 

  32. 32.

    Soliman S. Pharmacological control of ciliary activity in the young sea urchin larva. Effects of monoaminergic agents. Comp Biochem Physiol Part C Comp Pharmacol. 1983;76:181–91.

    CAS  Article  Google Scholar 

  33. 33.

    Kaltenbach SL, Yu J-K, Holland ND. The origin and migration of the earliest-developing sensory neurons in the peripheral nervous system of amphioxus. Evol Dev. 2009;11:142–51.

    Article  Google Scholar 

  34. 34.

    Trapido-Rosenthal HG, Morse DE. l-α, ω-Diamino acids facilitate GABA induction of larval metamorphosis in a gastropod mollusc (Haliotis rufescens). J Comp Physiol B. 1985;155:403–14.

    CAS  Article  Google Scholar 

  35. 35.

    Hadfield M, Meleshkevitch E, Boudko D. The apical sensory organ of a gastropod veliger is a receptor for settlement cues. Biol Bull. 2000;198:67–76.

    CAS  Article  Google Scholar 

  36. 36.

    Urrutia PM, Okamoto K, Fusetani N. Acetylcholine and serotonin induce larval metamorphosis of the Japanese short-neck clam Ruditapes philippinarum. J Shellfish Res. 2004;23:93–101.

    Google Scholar 

  37. 37.

    Croll RP, Voronezhskaya EE. Early elements in gastropod neurogenesis. Dev Biol. 1996;173:344–7.

    CAS  Article  Google Scholar 

  38. 38.

    Voronezhskaya EE, Ivashkin EG. Pioneer neurons: a basis or limiting factor of lophotrochozoa nervous system diversity? Russ J Dev Biol. 2010;41:337–46.

    Article  Google Scholar 

  39. 39.

    Voronezhskaya EE, Croll RP. Mollusca: gastropoda. In: Schmidt-Rhaesa A, Harzsch S, Purschke G, editors. Structure and evolution of invertebrate nervous systems. Oxford: Oxford University Press; 2015. p. 196–221.

    Google Scholar 

  40. 40.

    Raven CP. Morphogenesis: the analysis of molluscan development. Oxford: Pergamon Press; 1966.

    Google Scholar 

  41. 41.

    Jacob MH. Neurogenesis in Aplysia californica resembles nervous system formation in vertebrates. J Neurosci. 1984;4:1225–39.

    CAS  Article  Google Scholar 

  42. 42.

    Schacher S, Kandel ER, Woolley R. Development of neurons in the abdominal ganglion of Aplysia californica: I. Axosomatic synaptic contacts. Dev Biol. 1979;71:163–75.

    CAS  Article  Google Scholar 

  43. 43.

    McAllister LB, Scheller RH, Kandel ER, Axel R. In situ hybridization to study the origin and fate of identified neurons. Science. 1983;222:800–8.

    CAS  Article  Google Scholar 

  44. 44.

    Dale N, Kandel ER. Facilitatory and inhibitory transmitters modulate spontaneous transmitter release at cultured Aplysia sensorimotor synapses. J Physiol. 1990;421:203–22.

    CAS  Article  Google Scholar 

  45. 45.

    Fong PP, Warner M. Serotonin-induced parturition in the fingernail clam Sphaerium (Musculium) transversum (Say). J Exp Zool. 1995;272:163–6.

    CAS  Article  Google Scholar 

  46. 46.

    Conzelmann M, Offenburger S-L, Asadulina A, Keller T, Munch TA, Jekely G. Neuropeptides regulate swimming depth of Platynereis larvae. Proc Natl Acad Sci USA. 2011;108:E1174–83.

    CAS  Article  Google Scholar 

  47. 47.

    Kerbl A, Martín-Durán JM, Worsaae K, Hejnol A. Molecular regionalization in the compact brain of the meiofaunal annelid Dinophilus gyrociliatus (Dinophilidae). Evodevo. 2016;7:20.

    Article  Google Scholar 

  48. 48.

    Croll RP, Voronezhskaya EE. Early FMRFamide-like immunoreactive cells in gastropod neurogenesis. Acta Biol Hung. 1995;46:295–303.

    CAS  PubMed  Google Scholar 

  49. 49.

    Voronezhskaya EE, Elekes K. Transient and sustained expression of FMRFamide-like immunoreactivity in the developing nervous system of Lymnaea stagnalis (Mollusca, Pulmonata). Cell Mol Neurobiol. 1996;16:661–76.

    CAS  Article  Google Scholar 

  50. 50.

    Dickinson A, Croll R, Voronezhskaya E. Development of embryonic cells containing serotonin, catecholamines, and FMRFamide-related peptides in Aplysia californica. Biol Bull. 2000;199:305–15.

    CAS  Article  Google Scholar 

  51. 51.

    Voronezhskaya EE, Tsitrin EB, Nezlin LP. Neuronal development in larval polychaete Phyllodoce maculata (Phyllodocidae). J Comp Neurol. 2003;455:299–309.

    Article  Google Scholar 

  52. 52.

    Voronezhskaya EE, Tyurin SA, Nezlin LP. Neuronal development in larval chiton Ischnochiton hakodadensis (Mollusca: Polyplacophora). J Comp Neurol. 2002;444:25–38.

    Article  Google Scholar 

  53. 53.

    Starunov VV, Voronezhskaya EE, Nezlin LP. Development of the nervous system in Platynereis dumerilii (Nereididae, Annelida). Front Zool. 2017;14:27.

    Article  Google Scholar 

  54. 54.

    Beiras R, Widdows J. Effect of the neurotransmitters dopamine, serotonin and norepinephrine on the ciliary activity of mussel (Mytilus edulis) larvae. Mar Biol. 1995;122:597–603.

    CAS  Article  Google Scholar 

  55. 55.

    Braubach OR, Dickinson AJG, Evans CCE, Croll RP. Neural control of the velum in larvae of the gastropod, Ilyanassa obsoleta. J Exp Biol. 2006;209:4676–89.

    Article  Google Scholar 

  56. 56.

    Penniman JR, Doll MK, Pires A. Neural correlates of settlement in veliger larvae of the gastropod, Crepidula fornicata. Invertebr Biol. 2013;132:14–26.

    Article  Google Scholar 

  57. 57.

    Christopher K, Chang J, Goldberg J. Stimulation of cilia beat frequency by serotonin is mediated by a Ca2+ influx in ciliated cells of Helisoma trivolvis embryos. J Exp Biol. 1996;199:1105–13.

    CAS  PubMed  Google Scholar 

  58. 58.

    Dyachuk V, Odintsova N. Development of the larval muscle system in the mussel Mytilus trossulus (Mollusca, Bivalvia). Dev Growth Differ. 2009;51:69–79.

    CAS  Article  Google Scholar 

  59. 59.

    Perry SJ, Straub VA, Schofield MG, Burke JF, Benjamin PR. Neuronal expression of an FMRFamide-gated Na+ channel and its modulation by acid pH. J Neurosci. 2001;21:5559–67.

    CAS  Article  Google Scholar 

  60. 60.

    López-Vera E, Aguilar MB, Heimer de la Cotera EP. FMRFamide and related peptides in the phylum Mollusca. Peptides. 2008;29:310–7.

    Article  Google Scholar 

  61. 61.

    Chaban A, Voronezhskaya E. Involvement of transient larval neurons in osmoregulation and neurogenesis in the freshwater snails, Lymnaea stagnalis and Helisoma trivolvis. Acta Biol Hung. 2008;59:123–6.

    Article  Google Scholar 

  62. 62.

    Battonyai I, Voronezhskaya EE, Obukhova A, Horváth R, Nezlin LP, Elekes K. Neuronal development in the larvae of the invasive biofouler Dreissena polymorpha (Mollusca: Bivalvia), with special attention to sensory elements and swimming behavior. Biol Bull. 2018;234:192–206.

    CAS  Article  Google Scholar 

  63. 63.

    Voronezhskaya EE, Elekes K. Expression of FMRFamide gene encoded peptides by identified neurons in embryos and juveniles of the pulmonate snail Lymnaea stagnalis. Cell Tissue Res. 2003;314:297–313.

    CAS  Article  Google Scholar 

  64. 64.

    Marois R, Carew TJ. Ontogeny of serotonergic neurons in Aplysia californica. J Comp Neurol. 1997;386:477–90.

    CAS  Article  Google Scholar 

  65. 65.

    Waites CL, Craig AM, Garner CC. Mechanisms of vertebrate synaptogenesis. Annu Rev Neurosci. 2005;28:251–74.

    CAS  Article  Google Scholar 

  66. 66.

    Wen Z, Zheng JQ. Directional guidance of nerve growth cones. Curr Opin Neurobiol. 2006;16:52–8.

    CAS  Article  Google Scholar 

  67. 67.

    Daubert EA, Condron BG. Serotonin: a regulator of neuronal morphology and circuitry. Trends Neurosci. 2010;33:424–34.

    CAS  Article  Google Scholar 

  68. 68.

    Xing L, Son J-H, Stevenson TJ, Lillesaar C, Bally-Cuif L, Dahl T, et al. A serotonin circuit acts as an environmental sensor to mediate midline axon crossing through EphrinB2. J Neurosci. 2015;35:14794–808.

    CAS  Article  Google Scholar 

  69. 69.

    Haydon PG, McCobb DP, Kater SB. The regulation of neurite outgrowth, growth cone motility, and electrical synaptogenesis by serotonin. J Neurobiol. 1987;18:197–215.

    CAS  Article  Google Scholar 

  70. 70.

    McCobb DP, Haydon PG, Kater SB. Dopamine and serotonin inhibition of neurite elongation of different identified neurons. J Neurosci Res. 1988;19:19–26.

    CAS  Article  Google Scholar 

  71. 71.

    Diefenbach TJ, Sloley BD, Goldberg JI. Neurite branch development of an identified serotonergic neuron from embryonic Helisoma: evidence for autoregulation by serotonin. Dev Biol. 1995;167:282–93.

    CAS  Article  Google Scholar 

  72. 72.

    Koert CE, Spencer GE, van Minnen J, Li KW, Geraerts WP, Syed NI, et al. Functional implications of neurotransmitter expression during axonal regeneration: serotonin, but not peptides, auto-regulate axon growth of an identified central neuron. J Neurosci. 2001;21:5597–606.

    CAS  Article  Google Scholar 

  73. 73.

    Lacalli TC. Structure and organization of the nervous system in the trochophore larva of Spirobranchus. Philos Trans R Soc Lond B Biol Sci. 1984;306:79–135.

    Article  Google Scholar 

  74. 74.

    Shigeno S, Parnaik R, Albertin CB, Ragsdale CW. Evidence for a cordal, not ganglionic, pattern of cephalopod brain neurogenesis. Zool Lett. 2015;1:26.

    Article  Google Scholar 

  75. 75.

    Fischer AHL, Henrich T, Arendt D. The normal development of Platynereis dumerilii (Nereididae, Annelida). Front Zool. 2010;7:31.

    Article  Google Scholar 

  76. 76.

    Kempf SC, Page LR. Anti-tubulin labeling reveals ampullary neuron ciliary bundles in opisthobranch larvae and a new putative neural structure associated with the apical ganglion. Biol Bull. 2005;208:169–82.

    Article  Google Scholar 

  77. 77.

    Raineri M, Ospovat M. The initial development of gangliar rudiments in a posterior position in Mytilus galloprovincialis (Mollusca: Bivalvia). J Mar Biol Assoc UK. 1994;74:73.

    Article  Google Scholar 

  78. 78.

    Denes AS, Jékely G, Steinmetz PR, Raible F, Snyman H, Prud’homme B, Ferrier DEK, Balavoine G, Arendt D. Molecular architecture of annelid nerve cord supports common origin of nervous system centralization in Bilateria. Cell. 2007;129:277–88.

    CAS  Article  Google Scholar 

  79. 79.

    Simionato E, Kerner P, Dray N, Le Gouar M, Ledent V, Arendt D, Vervoort M. atonal- and achaete-scute-related genes in the annelid Platynereis dumerilii: insights into the evolution of neural basic-Helix-Loop-Helix genes. BMC Evol Biol. 2008;8:170.

    Article  Google Scholar 

  80. 80.

    Vergara HM, Bertucci PY, Hantz P, Tosches MA, Achim K, Vopalensky P, Arendt D. Whole-organism cellular gene-expression atlas reveals conserved cell types in the ventral nerve cord of Platynereis dumerilii. Proc Natl Acad Sci USA. 2017;114:5878–85.

    CAS  Article  Google Scholar 

  81. 81.

    D’Este L, Casini A, Kimura S, Bellier JP, Ito E, Kimura H, Renda TG. Immunohistochemical demonstration of cholinergic structures in central ganglia of the slug (Limax maximus, Limax valentianus). Neurochem Int. 2011;58:605–11.

    Article  Google Scholar 

Download references


The authors wish to thank Igor Adameyko (Karolinska Institutet), Roger Croll (Dalhouse University), and Alexey Chernyshev (NSCMB FEB RAS) for their valuable comments on this manuscript. The work was conducted using equipment of the Far East Center for Microscopy (NSCMB FEB RAS) and equipment of the Core Centrum of Institute of Developmental Biology RAS. We are also grateful to the staff of the Vostok Biological Station (NSCMB FEB RAS) for their generous assistance and to Olga Kharchenko for the art drawings. The work of EEV was conducted in the frame of IDB RAS Government basic research program, No. 0088-2019-0005. This work was financially supported by the Government of the Russian Federation through the ITMO Fellowship and Professorship Program.


This research was funded by RFBR grants # 18-34-20064 (immunostaining), # 18-04-01213 (axonal navigation experiments), and # 19-04-00792 (plankton sampling).

Author information




EEV and VAD planned the study, interpreted the results, and wrote the manuscript. OVY participated in data acquisition. AVS collected the plankton and NKK identified the bivalve species. All authors read and approved the final manuscript.

Corresponding author

Correspondence to Vyacheslav A. Dyachuk.

Ethics declarations

Ethics approval and consent to participate

Not applicable.

Consent for publication

All of the authors have consented to publication of the manuscript.

Competing interests

The authors declare that they have no competing interests.

Additional information

Publisher's Note

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

Rights and permissions

Open Access This article is distributed under the terms of the Creative Commons Attribution 4.0 International License (, which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver ( applies to the data made available in this article, unless otherwise stated.

Reprints and Permissions

About this article

Verify currency and authenticity via CrossMark

Cite this article

Yurchenko, O.V., Savelieva, A.V., Kolotuchina, N.K. et al. Peripheral sensory neurons govern development of the nervous system in bivalve larvae. EvoDevo 10, 22 (2019).

Download citation


  • Mollusk
  • Neurotransmitters
  • Nerve cords
  • Ganglia
  • Tetraneuralia