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date: 17 August 2018

Neurobiology of Reproduction in Mollusks: Mechanisms and Evolution

Abstract and Keywords

Ultimately, the outcome of successful reproduction—fertilization of eggs and production of surviving offspring—is relevant for how these processes evolve. However, a thorough understanding of the underlying, proximate mechanism is essential for interpreting evolutionary outcomes properly. Comparing neuroendocrine processes across different species, with different sexual systems, is one way of uncovering similarities and differences in regulation of their reproductive processes. Here, we provide an overview of the neuroendocrine control of reproductive processes in mollusks. In doing so, we also illustrate that it is relevant to consider the sexual system when addressing the neurobiology of reproduction. For example, our overview indicates that hermaphroditic mollusks seem to regulate their male and female reproduction via largely nonoverlapping neurobiological wiring and neuroendocrine substances, whereas this is not necessarily the case for separate-sexed mollusks. Clearly, this has implications for the available degrees of freedom within sexual systems in terms of evolutionary pathways.

Keywords: cephalopods, copulation, gastropods, mating, neuroendocrinology, regulation, reproductive organs, sexual systems

A Brief History of Neurobiology in Molluscs and Their Sexual Systems

The field of neurobiology as it stands now, with its many subdisciplines, would not be where it is now if it were not for mollusks. This animal group includes cephalopods (derived from the Greek words for “head-feet”), gastropods (Gr. for “stomach-foot”), and bivalves (Gr. for “two valves”). A number of these molluskan species have essentially stood at the basis of the development of the physiological side of neurobiology. After all, the Nobel Prize–winning insights from the axon potential theory developed by Hodgkin and Huxley were largely based on experiments with the giant axon of the veined squid (Loligo forbesii: Hodgkin et al., 1952; Hodgkin & Huxley, 1952a, 1952b, 1952c, 1952d). The size and accessibility of this axon allowed them to measure currents that were, at the time, impossible in other experimental systems. Since then, lots has happened and species like the great pond snail, Lymnaea stagnalis, and the sea hare, Aplysia californica, have championed an impressive body of literature spanning the full range of neurobiology (reviewed, e.g., in Chase, 2002).

Besides their historical importance for this field of research, the group of the Mollusca is also of importance because it comprises the second most speciose group of animals (after arthropods) and contains a large variety of animal forms ranging from slugs and snails, via bivalves, to cephalopods. They include many marine, freshwater, and terrestrial species. Among these are also the largest known invertebrate species, the giant squid (Architeuthis dux, over 12 m long) and the colossal squid Mesonychoteuthis hamiltoni (10 m long). For comparison, the smallest shelled mollusks reach maximum shell lengths that stay below 1 mm. But besides the large variety in sizes, this taxonomic group also comprises all possible forms of sexual systems (i.e., how the organism expresses whether it is male, female, or both at the same time or in sequence), including separate sexes, asexuality, parthenogenesis, selfing, and sequential or simultaneous hermaphroditism (e.g., Heller, 1993; Auld & Jarne, 2016; Koene, 2016), and fertilization occurs externally or internally (e.g., Michiels, 1998; Anthes, 2010).

Considering the mode of sexual system is clearly highly relevant when addressing the neurobiology of reproduction. The complexity of the neuroendocrine system regulating reproductive processes is clearly different between, for example, separate-sexed and hermaphroditic species (because the latter have both genders at the same time or in sequence over their lifetime). Regulation of the male and female system will need to be coordinated by largely nonoverlapping neurobiological wiring and neuroendocrine substances in simultaneous hermaphrodites, whereas this is not necessary in separate-sexed species (Fig. 1). In other words, for simultaneous hermaphrodites there is a lot of potential overlap in the regulatory systems, resulting in excitatory and inhibitory crosstalk between the male and female processes in order to avoid the simultaneous expression of conflicting reproductive processes. For separate-sexed mollusks, although timing in neuroendocrinological regulation of sexual processes may differ, such overlap between mating cues and signals, detection of these by receptors, the subsequent processing of this information, and the resulting motor output leading to behavior is largely absent (Fig. 1). Whether this leads to evolutionary constraints in either of the modes of sexual system is still largely unclear. By making a start with comparing the neurobiological regulation of several key processes, we point out a number of processes where this may be the case and hope that this will stimulate further comparative work in the future. We also identify some commonalities between the mechanisms underlying the different forms of sexual systems.

Neurobiology of Reproduction in MollusksMechanisms and EvolutionClick to view larger

Figure 1 A schematic overview of how male and female reproductive processes need to be regulated in separate-sexed and simultaneously hermaphroditic mollusks. As can be seen, for simultaneous hermaphrodites there is a lot of potential overlap in the regulatory systems, as indicated by the excitatory (triangles) and inhibitory (circles) cross-talk between the male and female processes. Clearly, this is needed to avoid conflicting reproductive processes to be executed at the same time. As illustrated, such overlap between mating cues and signals, detection of these by receptors, the subsequent processing of this information, and the resulting motor output leading to behavior do not occur when the sexes are separated, meaning that regulatory processes (and the involved messengers) can be similar in male and female mollusks.

Reproductive Processes and Their Neurobiology

Ultimately, the outcome of successful reproduction—fertilization of eggs and the production of surviving offspring—is relevant for how these processes evolve. However, if one wants to interpret these evolutionary outcomes properly, a thorough understanding of the underlying, proximate mechanism is essential. For example, different animal species may seem to arrive at the same evolutionary outcome that is relevant for sexual selection, but they may achieve this via very different mechanisms. Hence, while at the ultimate level one might conclude that these adaptations are the same, at the proximate level one would have to conclude that very different evolutionary trajectories were taken (that happen to achieve the same end result, i.e., convergence). Hence, comparison of neuroendocrine processes across different species, with different sexual systems, can uncover similarities and differences between the way in which they have been selected (by evolution) to regulate their reproductive processes. Therefore, in the following, we will describe some of the essential reproductive processes of mollusks, including what is known about their neuroendocrine regulation, in order to identify similarities and differences. In doing so, we start each section with separate-sexed cephalopods followed by hermaphroditic gastropods.

Cephalopods: Reproductive Behavior and Its Neuroendocrine Control

Reproduction is one of the most studied biological aspects of cephalopods. In this class of short-lived mollusks (they generally reach an age of 1 to 2 years only), the sexes are separate and there are no simultaneous or sequential hermaphrodites. In other words, there are no currently known cephalopod species that, repectively, have both sexes at the same time or change from one to the other sex during their lifetime (both of which do occur in other mollusks; see later discussion). Although one case of intersexuality or pseudohermaphroditism has been reported, in the squid Ancistrocheirus lesueurii (Hoving et al., 2006), the authors pointed out that one likely explanation would be that this was caused by exposure to pollutant in the form of endrocrine disruptors. In most cephalopod species, males and females are rather alike in form, but there are exceptions: many loliginids and the secondarily pelagic octopod Argonauta show extreme sexual dimorphism, with minute males (Hanlon & Messenger, 1996).

Sexual maturity does differ between the sexes in cephalopods. Males are reproductively capable for the greater part of their life cycle. Females, which can mature later in life, can receive and store sperm for the majority of their lives, so that fertilization and egg laying are temporally independent of mating. Most cephalopods produce eggs only once, with death occurring soon afterward (Boyle, 1983, 1987). The immediate causes of this apparently universal mortality are not clearly understood, but the sequence of physical changes brought about by the optic gland hormone seems to be irreversible. Some direct evidence for this hypothesis is available from experiments in which the optic glands were excised from mature octopuses, whose gonads subsequently regressed, while feeding and growth resumed (Wodinsky, 1977; Tait, 1986).

However, such semelparous behavior is not universal in cephalopods: Octopus chierchiae is iteroparous (Rodaniche, 1984) and some oceanic squids such as Sthenoteuthis oualanimsis (Harman et al., 1989; Nigmatullin & Laptikhovsky, 1994) and the deep sea cirrate octopods Opisthoteuthis spp. (Villanueva, 1992) produce series of mature eggs throughout most of their life cycle. Estimates of individual fecundity vary widely between species, and there is a trade-off between egg size and fecundity (Boyle & Rodhouse, 2005). Compared with other marine invertebrates, the eggs of cephalopods are large, well-protected, and produced in relatively low numbers (ranging from a few dozen to some hundreds of thousands; Boyle & Rodhouse, 2005), depending on the size of the animal as well as the species’ spawning pattern.

The anatomy of the cephalopod reproductive system is rather similar across orders. The essential physiological processes of the onset and progress of reproductive maturity in females are the meiotic maturation of the oocyte, the production and sequestration of yolk in the oocyte (vitellogenesis), the development of organs for the formation of protective individual egg coats (oviducal glands), and the encapsulation of the spawned egg mass (nidamental glands). In most species of cuttlefish, squid, and octopus, the enlargement of oviducal and nidamental glands marks the beginning of breeding competence and reproduction. Such processes seem to be controlled by neurohormones and hormones (see Di Cristo, 2013, and references). In males, sexual maturity is reached when mature spermatozoa are packaged into complex spermatophores stored in the spermatophoric (Needham’s) sac. Also these aspects seem to rely on hormonal control (Wells & Wells, 1959; D’Aniello et al., 1996).

Fertilization of eggs can be achieved once female individuals have been inseminated (there are no known reports of self-fertilization occurring). To achieve fertilization, during mating, spermatophores are transferred from the male to the female using an arm that is specially modified for use in the mating process (the hectocotylus) (Hanlon & Messenger, 1996). There may be competition for mates, but multiple mating is common. Egg masses are attached to the rocks (in most octopuses, loliginid squid, and cuttlefish) or released into the water column in fragile gelatinous masses (in most squid families) (Hanlon & Messenger, 1996).

The basis of our understanding of the neuroendocrine control of cephalopod sexual maturation is based on Wells and Wells’s (1959) findings using Octopus vulgaris (Fig. 2A). All their studies on the function and action of the optic gland and its hormone are milestones in the study of reproductive endocrinology in cephalopods (Wells, 1978). Subsequent work contributed by adding biochemical information (O’Dor & Wells, 1973, 1975; Wells et al., 1975; Wells & Wells, 1975), but it has unfortunately not yet resulted in the purification of the gonadotropic hormone from the optic gland.

Neurobiology of Reproduction in MollusksMechanisms and EvolutionClick to view larger

Figure 2 Different models of the nervous control of Octopus reproduction. (A) Model proposed by Wells and Wells (1959). (B) Model proposed by Di Cosmo and Di Cristo (1998). (C) Model proposed by Di Cristo (2013). Black arrowheads represent inhibitory control; white arrowheads represent excitatory control; arrows represent unknown relationships. Dashed lines indicate supposed pathways.

More recent studies suggest that the olfactory lobe, as well as the subpedunculate lobe, is involved in the control of the activity of the optic gland (Di Cosmo & Di Cristo, 1998). These data led to the neuroanatomical evidence that the olfactory lobe might influence the optic gland by releasing a positive factor (gonadotropin-releasing hormone, GnRH) to counterbalance the inhibitory effect of the tetrapeptide FMRFamide from the subpedunculate lobe on the glandular cells. This model, consisting of two centers controlling the activity of the optic gland, constitutes a simple feedback loop to switch optic gland activity on and off (Fig. 2B). This “two centers” model is, however, incomplete and leaves many questions unanswered. According to a revised model, it is now hypothesed that the olfactory lobe neurons (with their neuroproducts) may function as regulators of two apparently mutually excluding, or at least barely overlapping, processes: growth and reproduction (Fig. 2C; Di Cristo, 2013). As in many organisms, production of eggs in Octopus, as well as cephalopods in general, is a very energy-demanding physiological process that normally involves a shift of metabolic energy from somatic growth to reproductive maturation. The nervous system (and in turn gonadotropic glands) generally tempers the fertility of individuals to match nutritional availability. Some neuropeptides that are present in the olfactory lobe (NPY, galanin; Suzuki et al., 2000, 2002) may provide information about the state of the amount of stored energy that is available to the animal. In addition, in one of the last studies on the optic gland performed by Wells (O’Dor & Wells, 1978), it was demonstrated for Octopus that an optic gland hormone (and, hence, the nervous control of this gland) suppresses protein synthesis in the muscles. This suppression is associated with an increase in the concentration of free amino acids circulating in the blood. In females, these events coincide with a rapid growth of the ovary and its ducts, and a loss of weight elsewhere. That study also established that this effect could also be dependent on an unknown hormone originating from the (female) gonads. Hence, according to this evidence, the production of yolk-rich eggs (vitellogenesis) by female cephalopods, which requires the conversion of energy from food into the large amount of required yolk, is dependent on the optic gland hormone (O’Dor & Wells, 1973).

Egg production is also the terminal event in the life cycle in cephalopods; the animals breed once and then die. In females, egg laying is preceded by a period during which the ovary enlarges considerably, followed by a period during which the female broods the eggs. By the time the eggs hatch, the female appears notably emaciated and soon afterward dies. The death of female octopuses is perhaps not very surprising; the animals starve and at the same time produce great quantities of yolk. Interestingly, male octopuses also die around the same time as their mating partners; they too cease to feed during the last few weeks of their lives, and, like females, they show degenerative changes that range from a failure to control skin color and texture to the development of skin lesions that fail to heal (van Heukelem, 1973).

The situation in males suggests that the terminal condition of females is not simply due to starvation and the demands of a developing ovary. In fact, this semelparous behavior, as well as the ensuing death, also seems to be controlled by the optic gland hormone. Wodinsky (1977) demonstrated in Octopus hummelincki that removal of the optic glands from females that had laid and were brooding eggs induced a complete change of behavior. These animals abandoned their eggs, resumed feeding and growth, and lived for considerably longer than mature animals retaining their glands. A resumption of growth is in agreement with the role of an optic gland hormone in shifting energy balance in cephalopods (O’Dor & Wells, 1978). All the evidence points to activation of the optic glands being an irreversible process, resulting in postreproductive animals continuing to waste away until they die.

According to all these data, some hints about the functional role of optic gland hormone can be deduced. It is undoubtedly a gonadotrophic hormone (from the Greek word τροϕή, “food”). The early steps of gametogenesis, follicular cell division, and vitellogenesis strictly depend on it. All these stages point to gonad growth; although it is still unknown whether this effect is direct or indirect, it is interesting to note that progesterone causes the same effect (Abate et al., 2000; Di Cosmo et al., 2001; Cuomo et al., 2005; Di Cristo et al., 2010). Moreover, while there is some evidence that progesterone is involved in the meiotic maturation of oocytes, nothing is known about the meiotic maturation of spermatocytes, which does not depend on optic gland hormone (Wells, 1978).

In the new scheme of nervous control of female reproduction in Octopus (Di Cristo, 2013), it is proposed that during the early stages of life (Di Cosmo et al., 2001), NPY neurons “perceive” the energy demands. They could then affect feeding behavior and, at the same time, shut down optic gland activity almost completely, in coordination with the subpedunculate lobe, either directly or by negatively modulating the activity of GnRH neurons. This would shift energy to body growth rather than egg production. Subsequently, when internal signals of satiation indicate that a sufficient level of stored energy has been reached and can be converted into yolk, these inputs could inhibit NPY neurons. This would release the inhibition of those neurons activating the optic gland (GnRH; galanin) and stop the activity of the subpedunculate lobe. Under the influence of the rising optic gland hormone titer, energy flow will be shifted to reproduction and vitellogenesis, and gonadal maturation could then be initiated.

Such a process could also be affected by sex steroids, as well as their receptors, whose levels fluctuate during the life cycle (Di Cosmo et al., 2001). The role of an estrogen receptor in Octopus in the olfactory lobe is still unclear (De Lisa et al., 2012; Keay et al., 2006, see earlier). What is known is that some of the classical “vertebrate” steroidogenic enzymes are not present in mollusks (Markov et al., 2009), but enzymatic reactions on cholesterol-derived steroidal backbones are likely to happen, at least in Octopus and Sepia. This might indicate that either the real ligand of this receptor is still unknown or, as suggested elsewhere, it could work as a sensor, binding endogenous (or exogenous) hydrophobic molecules present in the diet (Markov & Laudet, 2011). If the latter hypothesis were confirmed, the pathway mediated by this receptor would be influenced by the “metabolic milieu.” The foregoing hypothetical scenario implies that there is an unknown factor that signals the “state of satiety.” Interestingly, oxytocin, whose homolog has been demonstrated to be present in cephalopods (cephalotocin: Reich, 1992; Takuwa-Kuroda et al., 2003), might function as a “satiety hormone” (as it does in rats; Arletti et al., 1990). Its release is then hypothesized to positively depend on the optic gland hormone titer.

The most intriguing aspect of this topic concerns the optic gland hormone, whose role is not yet well defined. It is not known whether it is just a trophic hormone, which induces gonad ripening and vitellogenesis, as gastropods dorsal bodies do (Roubos et al., 1980; see later), or whether it is also involved in meiotic maturation and egg laying. Wells and Wells (1959) reported that only three out of sixty-nine females with lesions that activate the optic gland laid eggs. This suggestion of the existence of an optic gland “trophic” hormone obviously poses the question about the true nature of the hormone that is responsible for gametogenesis and egg laying in Octopus. Screening of the Octopus vulgaris transcriptomes (Moroz et al., 2011; Zhang et al., 2012), as well as the Octopus bimaculoides genome (Albertin et al., 2015), has revealed the presence of an egg laying-like peptide (ELH), but its localization and function are still unknown.

In a sense, the hypothesis that such a cephalopod ELH could function as the gonadotropin in cephalopods (two gonadotropins hypothesis) would provide possible answers about the control of sexual maturation in male cephalopods. In fact, this neurohormone could explain why Octopus males, as well as other cephalopod males, produce mature spermatophores long before optic gland enlargement (O’Dor & Wells, 1978), where the ELH is expected to be produced. Moreover, this would imply that the continued production of spermatozoa in young males probably does not impose a high-energy demand on the individual. Hence, the trophic optic gland hormone is not needed for sperm maturation (meiosis), production, and donation, indirectly confirming the role of cephalopods ELH in regulating gametogenesis and gamete release.

Optic Gland

The optic glands are small, rounded bodies that lie on the optic tracts of cephalopods (Fig. 3). In young O. vulgaris, they are pale yellow in color, becoming swollen and orange as the animal matures. Owen (1832) also indicated them in a drawing of the brain of Sepia, and they are illustrated and labeled as “glandula ottica” in O. aldrovandi by Delle Chiaie (1828). Subsequent descriptions in the literature nearly all reported them as nervous tissue, but Boycott and Young (1956) had no doubt that these bodies were endocrine glands. Their fine structure has been described by Bjorkman (1963), Nishioka et al. (1970), and Froesch (1974).

The glands are heavily vascularized (Young, 1971). Apart from those associated with blood vessels, there are two types of cell in the gland: large stellate cells, with massive (10–15-µm diameter) rounded nuclei, called chief cells. These are responsible for the production of the optic gland hormone since they are the ones that change in size and appearance as the gland matures. A smaller group of supporting cells, probably fibrocytes that are responsible for the production of a connective tissue framework for the gland, is dispersed among chief cells (Boycott & Young, 1956; Wells & Wells, 1959). The cytoplasm of the large stellate cells contains many mitochondria, an extensive Golgi apparatus, and large numbers of ribosomes. The endoplasmic reticulum (in the resting gland) consists of a sparse system of fine tubules. The fine structure of the supporting cells is similar but with fewer mitochondria and ribosomes, and they resemble glial cells (Bjorkman, 1963). The cytoarchitecture of the gland suggests that it is of nervous origin.

These glands are innervated by the subpedunculate lobe at the back of the supraoesophageal brain through the optic gland nerve. Removal of this lobe is followed by degeneration of at least some of the nerves, which enter the gland from a bundle running along the optic stalk (Wells & Wells, 1959; Froesch, 1974). Ultrastructural analyses of the optic glands (Bjorkman, 1963) revealed that, in the immature optic gland, there are two types of synapses, axoaxonal (among fibers of optic gland nerve) and axoglandular (contacts on glandular chief cells), whereas there is only one type of synapse, the axoglandular, in the mature gland (Froesch, 1974). It was proposed that the axoaxonal synapses might inhibit the axoglandular synapses, according to the widespread pattern of presynaptic inhibition (Froesch, 1974). This idea was supported by the absence of axoaxonal synapses in the gland of adult animals, which is supposed to be active only in the absence of inhibition. Note that no trace of the optic glands is visible in either the embryonic stages or the planktonic larvae. They appear in young animals and seem to develop from nerve cells close to the olfactory lobe (Bonichon, 1967).

In a study of the neuropeptidergic innervation of the optic gland of Sepia, Le Gall et al. (1988) found that the neuropeptide Phe-Met-Arg-Phe-NH2 (FMRFamide) is the substance present in the optic gland nerve that apparently inhibits optic gland activity. Later it was also demonstrated that in Octopus vulgaris, the optic glands are innervated by FMRFamide immunoreactive fibers originating from neurons in both the subpedunculate and the olfactory lobes (Di Cosmo & Di Cristo, 1998).

The Function of the Optic Glands

In 1956, Boycott and Young noted in O. vulgaris that the optic glands were enlarged in a proportion of the animals that they were using for experiments on brain function in learning. Animals in which the optic tracts had been sectioned had large glands and grossly enlarged gonads. Similar effects were observed after destruction of the subvertical lobes. By now, the optic glands are recognized as the endocrine organs that control the maturation of the reproductive system. Wells and Wells (1959) hypothesized that optic glands secrete a gonadotropic hormone, and they clarified the relationship between the gland and the central nervous system (CNS). They experimentally demonstrated that either cutting the optic gland nerve or making a surgical lesion in the subpedunculate lobe resulted in an enlargement of the gland and a subsequent hypertrophy of the gonads (Fig. 3).

A similar (but weaker) hypertrophy of both optic gland and gonads is produced by cutting the optic nerves or removing the optic lobes (Fig. 3). If immature animals are blinded by lesioning of the optic nerves, the optic glands enlarge and the octopuses mature precociously. This observation led Defretin and Richard (1967) to test the effect of reducing the day length on the state of the glands in Sepia. They kept cuttlefish in tanks in the dark for 22 out of every 24 hours. A control group remained in continuous light. The stellate cells in the glands of animals maintained in the short day length treatment enlarged greatly, with a massive increase in endoplasmic reticulum and Golgi apparatus. Budding from the Golgi apparatus were large numbers of dense vesicles 100 nm in diameter, which were absent in the resting glands and presumably contain the glandular secretion to be shed into the bloodstream.

Wells and Wells (1959) then proposed that the CNS exerts an inhibitory control on the optic gland via the optic gland nerve and that photoperiod also plays a crucial role in controlling gonad maturation; that is, darkness activates it. Moreover, the gonad remains small, whatever the operation, if the optic glands are removed. Since the effects of blinding and more central brain lesions were not additive, it was postulated that control of the condition of the glands always ran via the subpedunculate lobe (see Fig. 2B). In this chain of events, excess of light triggers subpedunculate lobe inhibition of optic gland activity. The last part of this chain is excitatory and hormonal, affecting the gonad. Subsequent work has, in general, confirmed these hypotheses. Removal of the subpedunculate lobe is always followed by enlargement of the optic glands. If the operation is unilateral, only the gland on that side enlarged. Activation of one or both of the optic glands always produced enlargement of the gonad. The effect is most spectacular in females, where the ovary may increase in size from about 1/500th to as much as 1/5th of the body weight within a month of the operation. Even in males, these activating lesions induce weight increase of the testis, particularly in small undividuals.

Removal of the optic glands from males that are already producing sperm is followed by a decrease in the weight of the testis and the eventual cessation of sperm and spermatophore production; the effect is most marked in the largest males. The optic glands in males weighing less than 1,000 g are generally small and not obviously secreting. It has been hypothesized that even glands that are not visibly enlarged must be producing some secretion which seems sufficient to stimulate sperm production in males but insufficient to stimulate yolk production in the ovary (Wells & Wells, 1972).

Whereas the relation between subpedunculate lobe, optic gland, and gonad seems nearly resolved, the link between day length and secretion of the gonadotropin is less certain (Wells, 1978). When Sepia is kept under short day length conditions, it matures precociously (Defretin & Richard, 1967; Richard, 1967), and particularly some wavelengths of light seem to have an inhibitory effect on gonad development. However, attempts to replicate the cuttlefish experiments using Octopus have yielded only equivocal results: Only a few of the animals appear to respond (Buckley, 1977). The effect of blinding the animals, moreover, might be unconnected with the detection of day length, since other operations that damage structures in the eye’s orbit, or the implantation of foreign bodies into the orbit can also result in optic gland enlargement, albeit much more slowly than optic nerve lesion (Wells, 1978). A possible explanation for this discrepancy could come from physiological studies indicating that neuropeptides, mainly FMRFamide and dopamine, can modulate retina adaptation in female Octopus either to light or to darkness, according to the sexual maturity of the animal (Di Cristo et al., 2003). This would imply that the brain control of optic gland secretion involves assessment of photoperiod, chemical stimulus, and modulation of photoreceptor activity.

To confirm the endocrine importance, activated optic glands have been implanted into the system of blood sinuses behind the eyes of recipient animals from the same species. Once these glands attract an arterial blood supply, they begin to secrete. A month later the gonads of recipients that would otherwise be immature are noticeably enlarged; neither the sex of the donor animal nor the condition of the glands (active or inactive) when implanted makes any difference for the final result (Wells & Wells, 1975). It has furthermore been demonstrated that the implant could be species and genera independent, the optic gland of different species (or genera) being active and inductive in the recipients, indicating evolutionary conservation. Surprisingly, implanting activated optic glands from decapods into octopods has no effect, even though there is reason to believe that the hormone produced by the optic glands of decapods is very similar to that found in Octopus (Wells & Wells, 1975). The effect of the optic gland on gonad development and male and female gametogenesis can also be demonstrated in vitro (Durchon & Richard, 1967; Richard, 1970; Wells, 1978).

More recent work (Di Cristo et al., 2010) has revealed that there are two different patterns of yolk protein in the eggs of Octopus. These mirror the two main periods characterizing the reproductive cycle of the female of O. vulgaris: the nonvitellogenic period and the vitellogenic period. It seems that progesterone plays a role in Octopus vitellogenesis (Di Cristo et al., 2010) by inducing proliferation of follicle cells (the site of synthesis of yolk protein; see later). This suggests that this steroid, together with the optic gland hormone, may be involved in vitellogenesis in Octopus (Abate et al., 2000; Di Cristo et al., 2010).

When octopus oocytes from the vitellogenic phase were used in in vitro experiments, O’Dor and Wells (1973) showed that the ovary is the site of synthesis of yolk proteins. Injected, labeled amino acid rapidly disappeared from the blood and began to accumulate in the ovary. Uptake and synthesis stopped when the optic glands were removed. However, these experiments did not show which part of the ovary is responsible. But experiments on oocytes and follicle cells in cuttlefish (Lankester, 1875; Yung Ko Ching, 1930) indicated the follicle cells as the likely site of yolk secretion. These light microscope studies have been followed by investigations at the electron microscope level in Octopus (Wells et al., 1975; Buckley, 1977) and confirmed this hypothesis.

The follicle cells are packed with rough endoplasmic reticulum and active Golgi apparatus. Electron-dense material, originating from the Golgi apparatus, is exported to the oocyte down finger-like processes. Follicle cells from an animal deprived of its optic glands 2 or 3 days previously have ceased to produce dense granules (Wells et al., 1975). In contrast to this, the oocyte itself has few organelles and shows no effect of hormone deprivation. Its main phase of synthetic activity comes earlier in development, in the period of carbohydrate and lipid accumulation, which precedes the full development of the follicle cells wrapping. This stage is independent of the optic gland hormone (Buckley, 1977). It seems that the hormone peaks twice during oocyte development. In early oocyte development (which corresponds roughly to the juvenile stage), it is required for oogonia division. This is followed by a period during which the ovary will grow, apparently at a normal rate, in the absence of the optic glands. The glands become essential again only in the last phases of ovarian maturation, when the animal begins to accumulate the massive quantities of vitellogenin (Wells, 1978). Interestingly, sex steroids, namely progesterone, seem to be required in both circumstances (Cuomo et al., 2005; Di Cristo et al., 2010). This opens a new window in the endocrine control of cephalopod gonad development (at least in females); the combined action of the unknown optic gland hormone, which seems to act as a trophic hormone, and the sex steroids that are locally produced (Di Cosmo et al., 1998, 2001, 2002; Di Cristo et al., 2010; Di Cristo, 2013). Nothing is known about the presumptive feedback action of steroids on the brain, however, but this is something worth exploring in the future.

The optic gland hormone exerts also an enlarging effect on the oviducts and oviducal glands, as well as the apparatus responsible for packaging the sperm into spermatophores. The effect is greatest in the smallest animals (Froesch & Marthy, 1975; Wells, 1960). Castration does not appear to alter the condition of the ducts in males or females nor to affect hectocotylus regeneration (Callan, 1940; Wells & Wells, 1972), which suggests that neither male nor female gonads produce hormone(s) involved in this process. Nevertheless, although no physiological evidence has been reported, an endocrine role has been proposed for the cephalopod gonads (D’Aniello et al., 1996; Di Cosmo et al., 1998, 2001, 2002; Tosti et al., 2001) based on the presence of “vertebrate” sex steroids, steroidogenesis, and steroids-binding receptors.

Finally, although few behavioral studies have been done on the effects of reproductive hormones in cephalopods, Wells and Wells (1972) demonstrated that either castration or surgery affecting the condition of optic glands in mature octopus males did not interfere with the reaction of males to females placed in their tanks. Octopuses with their testis and male ducts removed approached females, inserted the hectocotylus, and passed imaginary spermatophores down the groove in the hectocotylus into the mantle of the female; even the entire set of stereotyped movements during the mating were not impaired by that surgery (Wells & Wells, 1972). The only operation that prevented correct behavior was removal of the tip of the hectocotylus; evidently this indicates that a particular signal from the tip is required to successfully locate an oviduct in the mantle and initiate spermatophore transfer. This is also supported by the finding that males will copulate with females having the distal ends of the oviducts removed, but they do not pass spermatophores (Wells, 1978).

The Neurosecretory System of the Vena Cava

The neurosecretory system of the vena cava (NSV) is localized in the ventral part of the posterior subesophageal mass of cephalopods, particularly in the ventral median vasomotor lobe. It is undoubtedly a neurosecretory/neurohaemal system. The endings of its many neurons penetrate the walls of the vena cava and are filled with a variety of large vesicles (Martin, 1968). From the vena cava, secreted products can reach visceral organs located in the mantle cavity. The peripheral NSV system was described by Alexandrowicz (1964, 1965), and the neurosecretory cells reported from the visceral lobe by Bonichon (1967) and Martin (1968) are almost certainly parts of the same system. The neurosecretory cells are present and active in animals of all ages and at all times of year (Laubier-Bonichon, 1973).

To uncover its neuroendocrine role, several researchers used extracts of the peripheral part of the NSV system and were able to show that these had marked effects upon the heartbeat of Octopus and Eledone (Bianchi, 1969; Berry & Cottrell, 1970; Bianchi & De Prisco, 1971). This effect is probably brought about by FMRFamide, which is a potent cardioaccelerator (Di Cosmo & Di Cristo, 1998). But apart from its role in cardioregulation, the importance of the NSV system in the reproduction of cephalopods comes from the discovery that it contains two members of the vasopressin/oxytocin superfamily: cephalotocin and octopressin (Reich, 1992; Takuwa-Kuroda et al., 2003), two neuropeptides that have been shown to be released into the vena cava to reach target organs in Octopus. These peptides have also been found in S. officinalis (Henry et al., 2013).

Cephalopods seem to be the only invertebrates with two members of the oxytocin/vasopressin superfamily of peptides, and receptors for these peptides have also been identified and cloned (Kawada et al., 2004; Kanda et al., 2005). Interestingly, octopressin evoked contractions of the smooth muscles of the oviduct, while cephalotocin had no effect on this tissue. Octopressin mRNA is also expressed in the buccal lobes, which control feeding behavior (Young, 1971). We should recall here that in Sepia, other peptides released from eggs and the female reproductive ducts are able to control the contraction of oviducts (Zatylny et al., 2000a,b; Bernay et al., 2004, 2006b).

The Control of Optic Gland Activity: cephGnRH and Other Neuropeptides

To date, the control on the onset and maintenance of sexual maturity in cephalopods has been linked to several neuropeptides that directly (or indirectly) control the activity of the optic gland. Le Gall et al. (1988) first demonstrated in Sepia officinalis that the tetrapeptide FMRFamide is the molecule present in the subpedunculate lobe as well as in the optic gland nerve, hypothesizing its inhibitory role on the release of optic gland hormone. The same neuropeptide was later also found in the same lobes in Octopus vulgaris (Di Cosmo & Di Cristo, 1998).

The olfactory lobe has also been demonstrated to be part of the circuit controlling activity of the optic gland. In the neurons of the posterior olfactory lobule, a specific isoform of GnRH was first localized (Di Cosmo & Di Cristo, 1998) and later characterized (Iwakoshi et al., 2002). This cephalopod GnRH dodecapeptide (cephGnRH) is a member of the wide family of GnRH neuropeptides, which in vertebrates are essentially involved in the activation of gonadotropin release from the pituitary gland (Morgan & Millar, 2004). The ancient origin of this peptide was suggested by the recent description of orthologous GnRH in many invertebrates, including nonvertebrate chordates (Tsai & Zhang, 2008; Roch et al., 2011). In mollusks, GnRH has been sequenced in three cephalopods (Iwakoshi et al., 2002; Di Cristo et al., 2009; Onitsuka et al., 2009), two bivalves (Bigot et al., 2012; Treen et al., 2012), and three gastropods (Tsai & Zhang, 2008; Zhang et al., 2008; De Lisa et al., 2013). The presence of a peptide with the structural features of both GnRH and insect adipokinetic hormone in the nematode Caenorhabditis elegans (termed GnRH-AKH) (Lindemans et al., 2009) suggests the presence of a GnRH peptide superfamily (Roch et al., 2011).

In contrast to the sequence elucidation of GnRHs, only scarce functional data are available in mollusks (Osada & Treen, 2013). Many studies have been performed primarily using vertebrate GnRH peptide isoforms (Goldberg et al., 1993; Pazos & Mathieu, 1999; Young et al., 1999; Zhang et al., 2000; Gorbman et al., 2003; Nakamura et al., 2007), complicating data interpretation. To date, it is still an open question whether GnRH affects reproduction in cephalopods. Some evidence favors such an hypothesis, but some comparative studies seem to exclude a direct role of GnRHs in reproduction in mollusks. Physiological studies based on homologous GnRH administration have been conducted on few species. In O. vulgaris, octGnRH stimulates oviduct contraction and gonadal steroidogenesis, suggesting a role in the reproductive process (Iwakoshi-Ukena et al., 2004; Kanda et al., 2006). In Aplysia, however, homologous GnRH has little effect on the activation of egg production, although it modulates activity of diverse central neurons and inhibits after-discharge in bag cells (Tsai et al., 2010; Sun & Tsai, 2011; see later for explanation of bag cells). Finally, putative scallop GnRH-like peptide stimulated spermatogonial cell division in cultured scallop testis (Treen et al., 2012). Interestingly, in C. elegans, although homologous peptide administration was not performed, RNA interference that disrupted the production of GnRH-AKH prohormone resulted in delayed egg laying (Lindemans et al., 2009).

This mismatch between structure and function in GnRH in invertebrates has no simple explanation, and we should probably ask why reproduction seems to be affected by GnRH only in some species. One possible line of inquiry that could help in addressing this problem would be to study the presence, expression, pharmacology, and physiology of GnRH receptors in invertebrates. In Octopus, a GnRH receptor has been identified and characterized (Kanda et al., 2006). This receptor is distributed throughout nervous and peripheral tissues, and interestingly in the olfactory lobe, optic gland, as well as in the gonads. It responds specifically to its native peptide (octGnRH) to activate the classical GnRH-induced signal transduction pathway; this results in steroidogenesis in reproductive tissues during bioassays (Kanda et al., 2006). However, without data on the Octopus gonadotropin, it is impossible to define the role of GnRH in such a function.

Besides GnRH and FMRFamide, the olfactory lobe also contains several other (neuro)peptides. Immunoreactive-like signals for APGWamide (Di Cristo et al., 2005; Di Cristo, 2013), galanin (Suzuki et al., 2000), neuropeptide Y (NPY; Suzuki et al., 2002), and corticotropin-releasing factor (CRF; Suzuki et al., 2003) have been reported in O. vulgaris. Some of these results are exclusively based upon immunological data, so there is a need for strong molecular data supporting these findings. However, some evidence is already available; for example, an APGWamide mRNA has been reported to be present in the transcriptome of Octopus CNS (Di Cristo, 2013), and one nonapeptide, referred to as peptide tyrosine phenylalanine (PYF), has been isolated from the CNS of the squid Loligo vulgaris (Smart et al., 1992). This peptide shows high homology with the C-terminal end of the other molluskan NPYs (seven out of nine residues are identical) and could be a processed form of a genuine Loligo NPY neuropeptide.

Galanin (and galanin-like peptides), NPY, and CRF, whose presence has been hypothesized in the olfactory lobe, are known to play important roles in the balance between metabolism and reproduction, not only in vertebrates (Crown et al., 2007) but also in invertebrates (see de Jong-Brink et al., 1999, 2001, for the role of NPY in invertebrates). In vertebrates, (1) neuropeptides, including galanin-like peptide (GALP) and NPY, all reside in the hypothalamic area involved in the regulation of metabolism and reproduction; (2) neurons producing these peptides are targets of metabolic hormones, such as leptin and insulin; and (3) these neuropeptides either directly or indirectly affect feeding and metabolism, as well as the secretion of GnRH and gonadotropins.

Steroids in Cephalopods

Sex steroids are key molecules in the endocrine mechanisms of vertebrates (Bentley, 2001). In many mollusks, “vertebrate-like sex steroids” were essentially believed to act as “endocrine disruptors” (Lafont & Mathieu, 2007). However, the discovery that these animals synthesize steroids and show tissue expression of steroid receptors has resulted in a reconsideration of the role that these molecules may play in invertebrate reproduction (Kohler et al., 2007).

In cephalopods, steroids deserve a special mention. The presence of steroids in Octopus (but also in Sepia) has been widely reported (D’Aniello et al., 1996; Di Cosmo et al., 1998, 2001; Tosti et al., 2001; Di Cosmo et al., 2002; Cuomo et al., 2005; Di Cristo et al., 2008, 2010; De Lisa et al., 2012). These papers cover the biochemistry, physiology, and pharmacology of vertebrate-like sex steroids in this species and establish three key points: (1) Octopus and Sepia are capable of synthesizing steroids in both gonads and the CNS; (2) these steroids interact with specific receptors localized in the gonads, reproductive tracts, and specific CNS regions; and (3) these steroids can affect the physiology of the gametes of Octopus.

One of the key aspects concerning steroids in cephalopods is the presence of putative steroidogenic enzymes in tissues of these animals. Some canonical enzymatic reactions on “cholesterol-derived backbones” are likely to occur in the CNS and gonads of cephalopods. However, structural evidence on the presence of the (presumptive) cephalopod steroids is lacking to date. The absence of studies on the structural characterization of steroids in cephalopods could also explain some different results on steroid receptors in Octopus. For example, an orthologous gene of the estrogen receptor has been cloned in O. vulgaris (octER; Keay et al., 2006) and localized in the olfactory lobe (De Lisa et al., 2012). While this receptor seems to be able to bind estradiol pending conformational changes (De Lisa et al., 2012), previous findings reported that the same receptor was constitutively active as transcription factor and unable to bind estradiol (Keay et al., 2006). Interestingly, stimulation of Octopus with estradiol increases the octGnRH transcript, as well as the octER transcript, in the olfactory lobes (De Lisa et al., 2012). This result links the expression of GnRH in the olfactory lobe to the presence of estrogens, whose level fluctuates throughout the entire life cycle of, at least, the female of Octopus (Di Cosmo et al., 2001), peaking just before egg laying. It is noteworthy that the level of octER gene expression in the olfactory lobe also fluctuates.

Male Processes: Sperm Maturation, Spermatophore Transfer, and Spermatophoric Reaction

Cephalopod sperm are produced in the median single testicle, and mature sperm are released into the vas deferens (and following spermatophoric glands), where they are packaged in spermatophores and stored in the spermatophoric (Needham’s) sac (Mann et al., 1970). The mature sperm often have a species-specific morphology, and detailed descriptions are available for several squid, cuttlefish, and octopus species (Franzen, 1955, 1956, 1967; Richard, 1971; Maxwell, 1974, 1975; Fields & Thompson, 1976; Healy, 1989, 1990, 1993). The mature spermatophore is a coiled mass of sperm and an ejaculatory apparatus wrapped in a roteic and elastic tunic. Inside, a transparent viscous fluid, the spermatophoric plasma, surrounds the sperm rope. In front of the sperm rope lies the gelatinous rod of the ejaculatory apparatus, separated from the sperm rope by a region filled with an amber-colored syrupy cement (Mann et al., 1970).

During mating, spermatophores are pumped through the distal portion of the vas deferens to the single exiting duct. This region of the vas deferens is referred to as the penis, but it does not act as intromittent organ. Spermatophores are transferred to the female by the hectocotylized arm. The structure and position of the hectocotylus vary greatly between species. Many oceanic squids have no hectocotylus, and it is absent from Vampyrotheuhis and the finned octopus Cirrotheuthis (Murata et al., 1982; Arnold, 1984). Some sepioids and teuthoids have two arms hectocotylized. In cuttlefishes and some squids, the hectocotylus is evident only by the reduction or absence of suckers on the arm. In octopuses, the whole arm is usually modified by a groove along its ventral surface, terminating near the arm’s tip, which is modified and often spoon-like. During copulation, the hectocotylized arm acquires the spermatophore (either by reaching in with the arm tip, as in squids and cuttlefishes; or by extension of the penis to the base of the hectocotylized arm, as in octopods) and transfers the spermatophore to the female in a variety of ways (Hanlon & Messenger, 1996). In decapods, donated spermatophores are stored in seminal receptacles, which are quite distant from the female’s oviducts; generally they are situated just below the mouth or in a ring around the mouth (Drew, 1911; Ikeda et al., 1993). In some squids, males attach the spermatophore onto a pad located on the inner wall of the mantle (Hanlon & Messenger, 1996). In O. vulgaris spermatophores are typically 2–3 cm long and about 0.5 mm across at their blunt, sperm rope end. They are tubular and turgid. There is a posterior or “male-oriented” part that is opaque white, filled with compacted sperm. The other, anterior, region is more translucent; it ends in a thin thread, which is carried back along the body of the tube. This “female-oriented” region is an invaginated folded tube, the “ejaculatory apparatus,” the anterior end of which forms a cap (Wells, 1978). These features vary considerably from one species to the next (Marchand, 1913).

During mating in Octopus, the spermatophore, held in the muscular penis, is carried down to the groove in the hectocotylized arm. The arm tip is then inserted into the mantle cavity of the female to reach the opening of the distal oviduct. After transfer to the female or release into seawater, the spermatophore swells and bursts, ejaculating the contained sperm by the spermatophoric reaction (Mann, 1970). The uptake of seawater at the hind end of the spermatophore provokes this “spermatophoric reaction.” The sperm rope moves away from the blunt end of the capsule, pushing the ejaculatory apparatus, until the tunic breaks and the ejaculatory apparatus is extruded. The latter evaginates slowly, pushed forward by the advancing sperm rope and sperm plasma, but suddenly there is an explosive increase in the rate of evagination and the whole apparatus and the front part of the outer tunic of the spermatophore swells to form a bladder, into which the rest of the sperm rope passes. The driving force for extrusion of the sperm bladder appears to be dilution of the spermatophoric plasma by seawater coupled with the considerable elasticity of the outer tunic of the spermatophore (Mann et al., 1970). The subsequent fate of the sperm bladder apparently differs from one genus to the next. In O. vulgaris, it bursts, liberating sperm into the oviduct, which carries them toward the oviducal glands by peristalsis. Here, they accumulate in the spermathecal sections of the oviducal glands, where they are presumably stored until the eggs are ready (Belonoschkin, 1929a,b; Wells, 1960).

Female Processes: Oocyte Maturation, Fertilization, Egg Deposition, and Parental Care

Depending on the cephalopod species considered, average female fecundities vary from a few hundred eggs (e.g., sepiolid squids), to several thousands (e.g., sepiid cuttlefish), to hundreds of thousands (e.g., merobenthic octopods like O. vulgaris or O. cyanea), or around 1 million, as in the pelagic octopod Ocythoe tuberculata (Salman & Akalin, 2012). The main processes of the onset and progress of reproductive maturity in the female are egg maturation (oogenesis and vitellogenesis) and the development of organs for the formation of protective individual egg coats (oviducal glands) and the encapsulation of the spawned egg mass (nidamental glands). Oogenesis, the production of eggs, is described for representative types such as Sepia (Richard, 1971; Dhainaut & Richard, 1976), Loligo (Selman & Arnold, 1977; Knipe & Beeman, 1978; Selman & Wallace, 1978), several ommastrephid squid species (Takahashi, 1978; Lipinski, 1979; Schuldt, 1979), and Octopus (Bolognari et al., 1976; Di Cosmo et al., 2001).

There are very limited data about the meiotic maturation of oocytes in cephalopods. In O. vulgaris, L-type Ca2+ currents play a role in oocyte growth and cytoplasmic maturation, and possibly in preparing the plasma membrane for the interaction with the spermatozoon (Cuomo et al., 2005). A large germinal vesicle is characteristic of the immature oocytes (100–300-µm diameter), while in subsequent stages of growth (up to 1,000-µm diameter) the nucleus is no longer visible and the metaphase spindle appears. This suggests that in O. vulgaris, oocytes are arrested in the first meiotic prophase up to the early-vitellogenic stage and resume meiosis at this stage up to a second block, presumably in metaphase I. Progesterone probably functions as hormonal stimulus for the first prophase–metaphase meiotic transition (Cuomo et al., 2005).

Small oocytes first arise from a germinal epithelium and are present from a very early stage (Fig. 3). Three phases of oocyte development are recognized along oocyte maturation (see Bottke, 1974, for the loliginid Alloteuthis; Boyle & Chevis, 1992, for Eledone cirrhosa; Di Cosmo et al., 2001, for O. vulgaris). In the first stage (immature-previtellogenic phase), oocytes are less than 1 mm in length, and each is attached to the connective tissue of the ovary by a short stalk (Fig. 3). In the second stage (early vitellogenic phase), there is a migration of flattened follicles from the stalk in a single layer over the surface of the oocyte. At this point the oocyte increases in volume, while the proliferation of the follicular cell layer increases at a greater rate, forming a deeply enfolded double layer well supplied with blood vessels, giving the egg a reticulated appearance (Fig. 3).

Neurobiology of Reproduction in MollusksMechanisms and EvolutionClick to view larger

Figure 3 Diagrammatic drawing of Octopus central nervous system as seen from above (modified from Di Cosmo et al., 2004). Abbreviations: g.opt., optic gland; n.ol., olfactory nerve; n.opt., optic nerves; ol., olfactory lobe; opt., optic lobe; subpd., subpedunculate lobe; tr.opt., optic tract.

In the third stage (full to late vitellogenic phase), the follicular cell layer acquires columnar characteristics and secretes proteinaceous yolk into the lumen of the oocyte. The eggs, consisting of an oocyte and follicular cells complex, undergo a rapid increase in volume. When the accumulation of yolk is complete, the follicle cell layer degenerates and the chorion is formed. The formation of the oocyte/follicular cell complex seems to be the key to the rapid growth of the cephalopod ovary during sexual maturation, since it seems that this complex accelerates the process of vitellogenesis. The extraordinary rate of development of the follicular cell layer is achieved by high rates of follicular cell division, with many nuclei becoming polyploid at this time (Boyle & Chevis, 1991). This follicular cell division and the subsequent huge release of vitellogenin into the ooplasm seem to be controlled by vertebrate-like steroids (Abate et al., 2000; Di Cristo et al., 2008). Progesterone induces vitellogenin synthesis in previtellogenic ovaries of O. vulgaris and is responsible for follicle cell proliferation (Di Cristo et al., 2008). The morphological changes in the oviduct and oviducal gland observed throughout the reproductive cycle in O. vulgaris mirror progesterone and estradiol fluctuations, suggesting that both steroids might work in synergy to sustain the growth and differentiation of the female reproductive system in this species (Di Cosmo et al., 2001). Moreover, the localization of both progesterone and 17β-estradiol receptors in the nuclei of the follicle cells in the ovary of O. vulgaris sustains the theory that there is a role for these steroids in the regulation of yolk synthesis (Di Cosmo et al., 1998, 2002). Finally, the presence of steroidogenic enzymes in the follicle cells, namely the 3β-hydroxysteroid dehydrogenase (3β-HSD), supports the concept of gonadal synthesis of steroids in Octopus.

At full maturity, the chorion is fully formed and encloses the egg, which detaches from its formative epithelial stalk. The size of eggs growing from the same string of germinal epithelium is not constant, but varies along the length of the string, and these differences are a further index of the stage of maturation of the animal (Boyle & Chevis, 1992; Di Cosmo et al., 2001). These mature eggs are temporarily accumulated in the proximal oviduct (Boyle & Rodhouse, 2005).

Upon ovulation, mature eggs pass from the proximal oviduct into the fertilization chamber of the oviducal gland. Here, spermatozoa are immobilized in the mucosa of the spermathecae (Hanlon & Messenger, 1996; Di Cosmo et al., 2001). In O. vulgaris, it has been shown that stimulation of spermatozoa, collected from the spermatophores in the male reproductive tract, with progesterone or the Ca2+-ionophore induces an acrosome-like reaction (Tosti et al., 2001). Supporting this hypothesis, spermatozoa stored in the spermathecae of the female oviducal gland do not show any acrosomal region (Tosti et al., 2001). Moreover, when the mature eggs enter into the fertilization chamber, they release a chemo-attractant factor, named sperm-attracting peptide (Octo-SAP), that induces the mobilization of spermatozoa toward its source (which is followed by Ca2+ mobilization and membrane protein phosphorylation; De Lisa et al., 2013). These events facilitate egg fertilization, although the molecular mechanisms underlying this event are far from being fully characterized. However, in S. officinalis, various sperm attractant factors have been purified (Zatylny et al., 2002). These chemotactic molecules were characterized as being small peptides synthesized in the embedded oocytes during vitellogenesis, released in the external media during egg laying, and able to facilitate fertilization by increasing the chance of gamete collision.

After fertilization and during spawning, in Octopus, eggs pass through a large section of the oviduct, the oviducal gland, in the distal portion. The glandular region of the oviducal gland provides secretion for the protection of the eggs and also stores the sperm that accumulated after mating (Froesch & Marthy, 1975). Finally, in decapods, the nidamental glands secrete the mass of jelly material, or fragile gelatinous globes, that surround all the eggs. In cuttlefish, the whole process of egg laying seems to be controlled by a network of peptides; about 10 neuropeptide/peptide families have been found to meet the criteria for displaying a regulatory activity during spawning (Zatylny-Gaudin et al., 2016): They could trigger egg laying by integrating environmental stimuli across a neurosensory network, synchronize oocyte transport and egg-capsule secretion, and induce male attraction and mate aggregation (waterborne sex pheromones).

In cuttlefishes, external fertilization occurs within the arm bundle. Sperm are released from the seminal receptacle on the buccal region or in the mantle cavity as eggs are extruded from the oviducts. Sepia eggs are often exceptionally large (up to 2 cm diameter; Corner & Moore, 1980; Arnold, 1984). These eggs, usually blackened with ink to obscure the developing embryo and perhaps reduce predation, are attached to plants or stones. Males in some species accompany females during the several hours needed for egg laying (e.g., Corner & Moore, 1980; Boletzky, 1983); this form of temporary mate guarding is thought to be important because of sperm competition. Females usually die shortly after spawning, although spawning can extend over several weeks or even months in the laboratory (Boletzky, 1983, 1987, 1988). No form of egg care has ever been reported.

In squids, fertilization is also external, but there are two mating styles: Eggs can be fertilized either as the capsules exit the oviduct (when spermatophores have been placed in the mantle cavity) or amid the arms (when spermatozoa are released from the seminal receptacle near the mouth). Then eggs are deposited either as large communal masses on the substrate or small groups laid on rocks, seagrasses, or corals. The behavioral cues initiating egg laying are not understood yet. Female loliginid squids such as D. pealei or D. opalescens are spent after producing large numbers of eggs. Parental care of eggs or young is unknown among squids.

In octopods, fertilization is internal, occurring in the oviducal gland (where sperm are stored) as eggs pass down the oviduct. Females lay their eggs (many thousands of eggs; 1 × 3 mm) in strings, attaching them to the rocks. The operation seems to be preceded by an attempt to clean off the area by continually running over it and plucking with the suckers (Arakawa, 1962). Suckers manipulate laid eggs, pressing them onto the rock and one another, sticking them together by the stalks, so that they hang freely in bunches from a common stem. Eggs are deposited by two methods: either cemented individually to a hard substrate in the den; or individual eggs are intertwined with other eggs in clusters of many tens, and then cemented to a substrate. Females guard and care for the eggs, and there is a vital behavioral and physiological change associated with this phase of the life cycle. Basic metabolic changes occur in O. vulgaris females that approach full maturity, with somatic growth stopping (O’Dor & Wells, 1978), and the ovary becoming full of ripe eggs; in some octopods the other internal organs become squeezed in the mantle cavity. Feeding behavior becomes erratic. After egg laying, the female never leaves the den, does not feed, and continuously cleans and aerates the eggs while she broods them, which can be for as long as 1–3 months, perhaps a quarter of her life span. Declining physiological conditions in spawning females are often recognizable by the production of more or less disorganized egg masses (Boletzky, 1987). Females invariably become emaciated and die soon after hatching occurs. Males also die at about the same age, due to the same physical degradation as females: hormonally induced muscle catabolism, high amino acid levels in the blood, and consequent high metabolic rate (O’Dor & Wells, 1978; see earlier). This semelparous behavior, as well as the ensuing death, also seems to be controlled by the optic gland hormone (Wodinsky, 1977).

Gastropods: Reproductive Behavior and Its Neuroendocrine Control

In contrast to the cephalopods, where the male and female reproductive processes seem largely regulated by the same neuroendocrine systems, gastropods tend to have clearly separated systems. The main reason for this seems to be that they are hermaphroditic, meaning that they can perform the male and female role and do not necessarily perform this all at the same time (Fig. 1). For example, sperm transfer and sperm receipt can occur at the same time, but egg laying is uncoupled in time (occurring only when the conditions are right). Because the sexual roles are regulated separately, we will here describe them one by one.

Male Reproductive Behavior and Its Neuroendocrine Control

In most species that have been investigated in this respect, the main centers for the control of male reproductive behavior are situated on the right side of the brain, reflecting the fact that the male organs are mostly located on the right side of the body and come out via the male gonopore just behind the tentacle (e.g, Koene et al., 2000; Fig. 4). However, it should be noted that this is based on species that have a dextral development (and generally a right-winding shell, but see Aplysia) and their male gonopore on the right side. Obviously, the whole central nervous morphology is mirrored in sinistral species, that is, with a left winding shell and their male gonopore on the left side of their body (e.g., Davison et al., 2009).

Neurobiology of Reproduction in MollusksMechanisms and EvolutionClick to view larger

Figure 4 Schematic drawings for comparison of the central nervous system and reproductive morphology of the three main orders of gastropods mentioned in the text. (A) Central nervous systems are redrawn from Chase (2002) and include the known neuronal and neuroendocrine cell cluster involved in male (blue) and female (red/pink) neuroendocrine regulation. AL, anterior lobe; BCC, bag cell cluster; Bu, buccal ganglion; CDC, caudo-dorsal cluster; Ce, cerebral ganglion; DB, dorsal bodies; Me, mesocerebrum; Pe, pedal ganglion; PeI, pedal I-cluster; VL, ventral lobe. (B) Reproductive morphology based on the three main model species described. The drawing from L. stagnalis was adapted from Koene (2010; see also Jarne et al., 2010), and the morphologies are drawn in the same style to ease comparison. a, atrial gland; ad, allosperm duct; ag, albumen gland; bc, bursa copulatrix; bt, bursa tract; btd, bursa tract diverticulum; c/fp, carrefour/fertilization pouch; dmg, digitiform mucus gland; ds, dart sac; ep, epiphallus; fgp, female genital pore; fl, flagellum; fp/sp, fertilization pouch/spermathecal; ga, genital atrium; gp, genital pore; hd, hermaphroditic duct; m, muciparous gland; mu, mucus gland; mgp, male gental pore; o, oothecal gland; od, oviduct; ot, ovotestis; p, penis; pco, pars contorta; pe, preputium; pg, prostate gland; pp, preputium; ps, penis sheath; sd, sperm duct; sv, seminal vesicles (in opisthobranchs this is also referred to as ampulla, a); u, uterus; v, vaginal duct; vd, vas deferens; wd, winding gland. (C) Illustration of the three main model species used and mentioned in the text: the sea hare, Aplysia californica; the great pond snail, Lymnaea stagnalis; and the brown garden snail, Cornu aspersum (formerly known as Helix aspersa). The positions of the genital pores are also indicated in the drawing of each species, to place the morphological drawings into perspective.

The main clusters of neurons involved in the regulation of male behavior of the dextral pond snail L. stagnalis are the anterior lobes (especially on the right side), right ventral lobe, right pedal Ib cluster, and some more dispersed cells in the right parietal and pleural ganglia (De Boer et al., 1996; El Filali et al., 2015). Neurons in all these clusters project into the penial nerve, which is the only nerve that innervates the male copulatory organs. Backfills of the penial nerve of Helisoma trivolvis reveal a very similar pattern (Young et al., 1999). Depending on the species, the exact terminology of the involved clusters varies, but it seems to be relatively conserved across gastropods (Koene et al., 2000). For example, the equivalent of the anterior lobe of hermaphroditic freshwater snails is called the mesocerebrum in Stylommatophora (land snails) and the H-cluster in Opisthobrachia (sea slugs; Koene et al., 2000). Likewise, each of these groups also has a clear cluster of cells that are located in the right pedal ganglion that is part of the network. Because the involved neurons and their endocrinology, as well as how this results in male behavior, has been investigated in most detail in L. stagnalis, we will briefly describe this process in the following. Wherever possible, we will include information about other species of gastropods (see also Fig. 4).

The bilateral anterior lobes are located in the cerebral ganglia, and they usually show a striking right-left asymmetry. In dextral species, the right lobe is bigger (e.g., Koene et al., 2000; Fig. 4A) while in sinistral species the left one (e.g., Biomphalaria glabrata: Lever et al., 1965; L. stagnalis: Davison et al., 2009) is bigger; as mentioned earlier, this corresponds to the side where the male organs can be everted from the body via the male gonopore. There are many different neuropeptides that have been reported to be present in the right anterior lobe, but APGWamide seems to play a key role in the execution of the male copulatory behavior (De Lange & Van Minnen, 1998; Koene et al., 2000; Koene, 2010) and is expressed in virtually all right anterior lobe neurons as well as in some on the left. The APGW transcript consists of 10 copies of APGWamide and one C-terminally located Anterior Lobe Peptide (CALP: Li et al., 1992; Smit et al., 1992) and is coexpressed in anterior lobe neurons with either Conopressin, the Lymnaea form of the neuropeptide tyrosine (LyNPY) or Pedal Peptide (Croll & Van Minnen, 1992; Smit et al., 1992; De Lange et al., 1997). APGWamide is also found in the freshwater snail Bulinus truncatus (De Lange & Van Minnen, 1998) and in male Pygmy squid, Idiosepius pygmaeus (Sirinupong et al., 2011); however, it may have a more general role in reproduction in bivalves since in the sea scallop Placopecten magellanicus it was detected in both sexes (Smith et al., 1997).

The second important cluster, the ventral lobe, is very prominent in the right cerebral ganglion of a dextral basommatophoran species and also contains numerous substances. The most prominent one is FMRFamide, which is a commonly occurring peptide that is here coexpressed with a number of the other peptides encoded on the same exon (Schot & Boer, 1982; Bright et al., 1993; Santama et al., 1993, 1995; Van Golen et al., 1995). Furthermore, APGWamide, Conopressin, LyNPY, and Pedal Peptide are all singly expressed in right ventral lobe neurons of L. stagnalis (Croll & Van Minnen, 1992; De Lange et al., 1997). In the Stylommatophora and Opisthobranchia, no clear parallel structure seems to have been identified (Fig. 4A). However, recent work showing that substances like FMRFamide, Conopressin, and NPY are expressed in the CNS of Theba pisana, a Stylommatophoran land snail, encourages looking for a neuron cluster where their expression overlaps as it does in the ventral lobe (Adamson et al., 2015).

The pedal Ib cluster protrudes clearly from the right pedal ganglion of L. stagnalis. There is abundant evidence that these cells are serotonergic (Elekes et al., 1988; Croll & Chiasson, 1989; Kemenes et al., 1989; Hetherington et al., 1994). As mentioned earlier, seemingly equivalent clusters can also be found in Stylommatophora (Helix aspersa; Eberhardt & Wabnitz, 1979; Li & Chase, 1995) and Opisthobranchia (Aplysia californica; Rock et al., 1977) (Fig. 4A). In addition to these clusters, at least for L. stagnalis, a number of neurons that are dispersed in the right parietal and pleural ganglia have been shown to be involved in the male network and contain a number of different neuropeptides (El Filali et al., 2015). Not much more is known about these neurons, but some send branches into both the penial and intestinal nerve (El Filali et al., 2015), which may imply a dual function that warrants further attention. Because of their dispersed nature, identifying similar neurons in Stylommatophora and Opisthobranchia is currently not possible but might be done based on their peptide content.

How these CNS regions control the male organs can, in part, be deduced from the projection of their axon fibers (see Fig. 4B for an overview of the reproductive organs). In L. stagnalis the penial complex, which is composed of the preputium (penis-carrying organ), penis, and retractor muscles, is innervated by peptidergic neurons of the right anterior lobe, ventral lobe, pedal Ib cluster, and dispersed neurons from the right pleural and parietal ganglia (e.g., Kemenes et al., 1989; Croll et al., 1991; Li et al., 1992). Peptides like Conopressin and APGW colocalize in neurons innervating the penial complex and vas deferens (Van Kesteren et al., 1992; Van Golen et al., 1995) and fibers containing FMRF and its coexpressed peptides are found in the penial complex (Bright et al., 1993; Santama et al., 1993, 1995; Van Golen et al., 1995). In addition, axons from the anterior lobe have also been found to innervate the pedal Ib cluster directly (Croll et al., 1991; Croll & Van Minnen, 1992).

The male behavior of gastropods is typically composed of a series of courtship events that can include shell mounting, circling, positioning, preputial eversion, and probing, and it typically ends in full penis eversion and intromission after which sperm transfer takes place. A distinction can be made in the way in which sperm is transferred. This can occur via unilateral copulation, where one individual acts in the male role and the other in the female role (after which roles can, but need not, swap; Koene & Ter Maat, 2005; Anthes et al., 2006). In such cases, one can typically also encounter so-called mating chains or mating rings, in which animals copulate in both roles but with different partners. In a chain the individual in front only acts as a female, the individual in the back only as a male, and each individual in between acts as male to the partner directly in front of it and female to the partner directly in its back. When the frontmost individual also inseminates the backmost, a circle is formed. Copulation can also be reciprocal (or bilateral), meaning that both individuals donate and receive sperm during a single mating interaction, via simultaneous intromission of the copulatory organs (e.g., Anthes et al., 2010; Lodi & Koene, 2016a). Note that in a few normally unilateral mating species, it seems possible to mate simultaneously reciprocal (reviewed in Jordaens et al., 2007). In addition, sperm can be transferred in an ejaculate, composed of sperm and accessory gland products, or in a spermatophore, in which case the ejaculate is packaged in a more or less elaborate casing (Zizzari et al., 2014).

The neuroendocrinology aspects of the male behavioral sequence have, again, been worked out in most detail in L. stagnalis and will be reviewed in the following, largely based on the behavioral descriptions of Van Duivenboden and Ter Maat (1985) and De Boer and coworkers (1996) and the review by De Lange et al. (1998a). Not much is known about the regulation of the behavioral components circling and positioning on the shell. During circling, which lasts on average 5 minutes, the sperm donor performs counterclockwise locomotion along the windings of the partner’s shell, first toward its tip and then toward its margin. When the margin is reached, which may take several attempts, the sperm donor spends on average 17 minutes to position itself and to start eversion of the preputium. This partially everted preputium becomes visible as a white bulge behind the right tentacle (and, evidently, the left tentacle in sinistral species). This eversion requires the relaxation of the circular muscles surrounding the male gonopore as well as the preputium retractor muscle bands (De Boer et al., 2010). Many of the aforementioned neuropeptides have been found around the male gonopore and in the preputial muscles, via immunocytochemistry, and affect contractions of the preputium retractor muscle bands in vitro. Importantly, injection of APGW into the haemolymph causes penial eversion in vivo (De Boer et al., 1997). Although in Biomphalaria glabrata, application of APGW into the water did not cause eversion, FMRF did (Muschamp & Fong, 2001; Fong et al., 2005). In addition, the presence of selective serotonin-uptake inhibitors, like prozac, and serotonin-antagonists in the water caused eversion (Muschamp & Fong, 2001; Fong et al., 2005), corroborating that the classic transmitter serotonin itself is involved and may prevent eversion. These peptides probably act in concert with the existing hydrostatic pressure in the head sinus and/or increased pressure in the arteria penis (Bekius, 1972; see also De Boer et al., 2010). De Lange et al., (1998a) speculated that longitudinal and circular muscles near the gonopore could contribute to full eversion, and that these muscular contractions and relaxations might be controlled by the motor neurons of the ventral lobe and dispersed pleural and parietal neurons.

The latter is supported by the presence of substances affecting these muscles (El Filali et al., 2015) and would allow for the very fine-tuned movements that are essential for successful intromission of the penis (De Lange et al., 1998b). This is attempted during probing, when the fully everted preputium makes movements under the lip of the partner’s shell in search of its female gonopore (De Boer et al., 1996). This presumably requires a sensory mechanism ensuring the correct position of the preputium prior to eversion and intromission of the penis. The most obvious candidates that could serve this function are the neurons that are found at the distal tip of the preputium. Several of these cells seem to be sensory neurons that contain Conopressin, with a dendrite extending though the preputial epithelium and an axon in the penis nerve (respectively, Zijlstra, 1972; De Lange et al., 1998a), while others form varicosities on local musculature and contain APGW, Pedal Peptide, and/or LIP (De Lange et al., 1998a). The latter suggests that these neurons may also directly control the minor positional adjustments that are necessary for penis intromission (De Lange et al., 1998a) and may even be involved in the release of the ejaculate.

The end of these courtship behaviors, and the start of copulation, is marked by the intromission of the penis into the female gonopore. The position of nerve fibers containing different neuropeptides in the longitudinal and circular muscle layers of the penis suggests that alternate contraction of these muscle layers could result in the eversion of the penis (De Lange et al., 1998a). Like the preputium, the everted penis is probably also given support by blood pressure (in the arteria penis muscularis) (L. stagnalis: Bekius, 1972; Biomphalaria glabrata: De Jong-Brink, 1969). Once intromission is achieved, the ejaculate is transferred into the vaginal duct of the recipient in around 35 minutes. Ripe spermatozoa, produced in the ovotestis (De Jong-Brink et al., 1977; Rigby, 1982), are released from the seminal vesicles. These are also innervated, and Geoffroy et al. (2005) speculated that this innervation provides the necessary nervous control for the correct timing of the release of spermatozoa, which is critical for effective sperm transfer and would allow for control over the exact amount.

Once released, the spermatozoa get transported via the sperm duct toward the prostate gland, where seminal fluid components (SPFs, often also referred to as accessory gland products or ACPs) are added (Fig. 4B). The ejaculate is then transported to the penis via the vas deferens. The latter organ then shows rhythmic muscle contractions that travel over its whole length and presumably start in the so-called pacemaker area near the prostate gland (unpublished data in De Lange et al., 1998a). This peristalsis has been proposed to be caused by the antagonizing effects of APGW and Conopressin on the vas deferens. This idea is supported by the resemblance of Conopressin with Vasopressin/Oxcytocin, which are involved in ejaculation in vertebrates (Van Kesteren et al., 1996). Additionally, the pacemaker area seems essential for peristalsis and its neurons contain LyNPY and receive input from the CNS via penial nerve branch 1 (np1; and the contraction frequency can be changed via electrical stimulation of this nerve: unpublished data in De Lange et al., 1998a). Further down the vas deferens its muscles are innervated by axons containing APGW, LIP, or serotonin, but not Conopressin. The prostate gland cells are surrounded by small muscle fibers, which are innervated by fibers containing APGW (Croll & Van Minnen, 1992) and DEILSR (the latter only in the area of the sperm duct; De Lange et al., 1998a), suggesting that the release of their products can also be fine-tuned depending on the circumstances or partner’s quality.

Once ejaculate transfer has finished, the penis and preputium are retracted. The retraction of the penis is most likely regulated by relaxation of the aforementioned preputium muscles and a decrease in the hydrostatic pressure. For the much larger preputium, retraction seems to require the preputium retractor muscles, because when this set of muscles is cut, these snails are incapable of retracting the organ (De Boer et al., 2010). The preputium retractor muscles are affected by many of the identified substances, mostly the neuropeptides mentioned earlier, and their effects agree with the presence of nerve fibers containing these substances. Thus, many substances are involved in the contraction and relaxation of the preputium retractor muscles, which ultimately results in the eversion and retraction of the male copulatory organ. Evidently, some of these substances may also be involved in the fine coordination of preputial movement during courtship as well as copulation.

Female Reproductive Behavior and Its Neuroendocrine Control

The main neuroendocrine center for the control of female reproductive behavior in the Basommatophora, mostly based on work in L. stagnalis, is the bilateral Caudo-Dorsal Cell cluster (CDC) situated in the cerebral ganglia, and its neurohaemal area in the cerebral commissure (Fig. 4A). Opisthobranchs possess a bilateral cluster with a very similar function, called the bag cell cluster, but a very different position as they are found in the rostral end of the abdominal ganglion of the CNS (e.g., Kupfermann, 1967; Li et al., 1999; Fig. 4A). Cells that are morphologically similar to CDCs of L. stagnalis have been found in Stagnicola palustris, Radix ovata, Bulinus truncatus, Planorbis planorbis, Planorbis vortex, Planorbarius corneus, and Biomphalaria glabrata (Boer et al., 1977; Roubos & Van der Ven, 1987). The following will first briefly describe these neuron clusters and their endocrinology, followed by their involvement in the production of female behavior.

In the Basommatophora, the importance of the CDCs was first revealed by cauterizing these cells, which completely abolished egg laying (Geraerts & Bohlken, 1976). These cells project into the neurohaemal area of the cerebral commissure, are electrically coupled, and send axons through the cerebral commissure to the contralateral cell cluster (Joosse, 1964; Wendelaar-Bonga, 1971; Roubos, 1976; De Vlieger et al., 1980). As a result, these cells show a synchronous, long-lasting discharge both in vitro and in vivo (respectively, De Vlieger et al., 1980; Ter Maat et al., 1986). That these CDCs are neurosecretory and that their products induce egg laying was first shown by injecting extracts from the cerebral commissure (e.g., Geraerts & Bohlken, 1976). This also works in different lymnaeids, although cross-species tests show that there is family specificity (comparing Lymnaea, Bulinus, Biomphalaria: Dogterom & Van Loenhout, 1983). A very elegant in vitro experiment confirmed that the CDCs release hormones into the blood during their bursting activity: the CDCs induced to discharge in one CNS preparation caused the CDCs in a second preparation in the same saline bath to start their long-lasting discharge (Ter Maat et al., 1988).

The latter experiment convincingly demonstrated that the initiation of the bursting activity is chemically mediated, and as it turns out that several peptides are involved and released into the blood (e.g., Geraerts et al., 1983, 1985; Geraerts & Hogenes, 1985). The different peptides that have so far been identified are mostly encoded on the same gene. Two of the three identified copies of the gene, called CDCH-I and -II, code for at least 11 peptides (Hermann et al., 1997) and are both expressed in the CDCs (Van Minnen et al., 1989). Moreover, CDCH-I is found in neurons throughout the female tract except the albumen gland (Van Minnen et al., 1989). CDCH-I and -II are also found in nerve fibers in the prostate gland, whereas CDCH-I-positive secretions are found in the secretory cells of the sperm duct and semen and only CDCH-II-positive fibers in the preputium (Van Minnen et al., 1989). The main peptides that have been identified in L. stagnalis are Amino Terminal Peptide (Nt), Calfluxin (CaFl), Carboxyl Terminal Peptide (CTP), CDCH, α-, β1-, β2-, β3- CDCPeptide, γ-peptide, δ-peptide, ε-peptide (Geraerts et al., 1983, 1985; Vreugdenhil et al., 1985; Li et al., 1994), and two additional peptides (Jiménez et al., 2004). α-CDCP has also been found in Biomphalaria glabrata (Van Minnen et al., 1992). The key peptide for egg laying seems to be CDCH, which is therefore also often referred to as Egg Laying Hormone (ELH), because the injection of this hormone alone triggers egg laying (Ter Maat et al., 1987) whereas some of the other involved peptides seem to be more involved in coordinating the egg-laying process once it has started, as described later.

The bag cells, which have been investigated in great detail in Aplysia californica, show similar synchronous coupling (e.g., Dargaei et al., 2014; Fig. 4A). They also synthesize an egg-laying hormone (ELH) that is encoded on a precursor protein that also contains multiple other peptides, a-, b- and e-bag cell peptides (BCPs) and acidic peptide (AP) (e.g., Kupfermann, 1967; Li et al., 1999), many of which have distinct physiological and behavioral roles with ELH setting everything in motion (Conn & Kaczmarek, 1989). As shown by Ferguson et al., (1989), in vivo stimulation of bag cells in Aplysia californica can elicit full egg-laying behavior. ELH has also been identified in a number of other species, including Opishtobranchs like Aplysia dactylomela (Cummins et al., 2010), but also bivalves like the Akoya pearl oyster Pinctata fucata, the Pacific oyster Crassostrea gigas (Matsumoto et al., 2013; Stewart et al., 2014), the Owl limpet Lottia gigantea (Veenstra, 2010), and very recently also the land snail Theba pisana (Stewart et al., 2016b). The latter finding will now allow for the identification of the CNS regulating egg laying in land snails, which is so far unknown (Fig. 4A).

The dorsal bodies (DB) comprise a second bilateral endocrine organ that is important for female development and oogenesis (Fig. 4A). Besides in L. stagnalis, dorsal bodies have also been found in Planorbarius corneus, Biomphalaria glabrata, Ancylus flaviatilis, and Bulinus truncatus (Lever et al., 1965; Boer et al., 1968, 1977). No clear equivalent structure seems to exist in the other main gastropod groups, although the cephalopod optic gland seems to serve a very similar function (see earlier). The surgical removal of the dorsal bodies results in no egg production in juveniles and low egg production in adults. Moreover, vitellogenesis is blocked in the ovotestis, whereas spermatogenesis seems to continue normally (Geraerts & Joosse, 1975). The fact that the female function can be recovered by implantation of dorsal bodies indicates that a hormonal factor, referred to as Dorsal Body Hormone, is involved (Geraerts & Joosse, 1975), but its molecular identity remains elusive. This hormone is probably produced in the single type of endocrine cells that the organ is composed of (Joosse, 1964; Wijdenes et al., 1983). This dorsal body hormone also stimulates growth and development of female accessory sex organs, while it does not affect male accessory organs (Geraerts & Joosse, 1975; Geraerts & Algera, 1976). No similar structure seems to have been described in either Opisthobranchs or Stylommatphora (Fig. 4A).

Most Basommatophora lay their eggs in egg masses that they fix to the substrate. A number of the behavioral components described later are found in several basommatophoran species (e.g., L. stagnalis: Ter Maat et al., 1987; Bulinus octoploides: Rudolph & White, 1979; Ancylus flaviatilis: Bondesen, 1950). The following descriptions of egg-laying behavior and its underlying mechanisms are largely based on detailed behavioral descriptions (Goldschmeding et al., 1983; Ferguson et al., 1993) integrated with knowledge about the involved substances in L. stagnalis (Hermann et al., 1997).

The aforementioned CDC discharge triggers the start of egg laying, which starts behaviorally with a so-called resting phase, and releases all peptides encoded on the CDCH gene via the neurohaemal area into the animal’s haemolymph (Ferguson et al., 1993; also Jansen & Ter Maat, 1985). The resting phase, which lasts on average 40 minutes, is characterized by the animal stopping locomotion, holding its shell still and slightly pulled forward over the tentacles and no rasping of the substrate. Locomotion most likely stops because CDCH inhibits the right pedal motor neurons (RPeN). The excitation of the CDCs, which also receive external input, is enhanced by the local auto-excitatory action of α-CDCP (previously called CDCA) and CDCH (Ter Maat et al., 1988). This positive feedback makes both CDCH and α-CDCP, but not the β-CDCPs, necessary for the CDC discharge, thus generating an all-or-none response (Ter Maat et al., 1988; Brussaard et al., 1990). CDCH also initiates the actual ovulation, that is, the release of ripe eggs from the ovotestis, which is followed by their fertilization and packaging by the female accessory glands (details about the female organs involved in the different stages of egg packaging can be found elsewhere (Plesch et al., 1971; Wijsman, 1989; Jarne et al., 2010; Koene, 2010).

The passing of eggs through the female tract is sensed by ciliated cells that send nervous signals to the CNS via the intestinal nerve. This signal triggers the release of β3-CDCP and α-CDCP, from other cells than the CDCs, at which point the animal enters the turning phase. This phase, which lasts for around 60 minutes, is characterized by the animal starting locomotion again, turning its shell back and forth by 90° and rasping the substrate. Locomotion is probably initiated by the increased activity of RPeN motor neurons induced by CDCH in combination with β3-CDCP, and possibly also via direct axonal projections from the CDCs (Hermann et al., 1997). β3- and α-CDCP also excite other motor neurons that are involved in shell movements and buccal rasping (Hermann et al., 1997). Some of these motor neurons project into the inferior cervical nerves that control the columellar muscle, and bilateral lesioning of these nerves eliminates shell turning during egg laying (Hermann et al., 1994). Buccal rasping is performed to clean the substrate for proper attachment of the egg mass (Ter Maat et al., 1989). This process continues as long as the passage of eggs through the female tract is registered, which explains the clear correlation between the duration of the turning phase, as well as the following oviposition, with the size of the egg mass (Ter Maat et al., 1987, 1989; Ferguson et al., 1993). In turn, the number of eggs in an egg mass depends on the time since last oviposition, indicating that ripe oocytes accumulate in the ovotestis (Ter Maat et al., 1983; see also Koene et al., 2006) and explaining why egg mass size is independent of the injected dose of CDCH (Dogterom et al., 1984).

Oviposition starts roughly 2 to 3 hours after the CDC discharge and is the moment when the egg mass emerges from the female gonopore and is fixed to the substrate. This phase usually lasts about 10 minutes during which the animal continues rasping, stops shell turning, and hardly locomotes. When the egg mass has been deposited, the animal crawls along the mass, without rasping or shell turning, before leaving it behind; this is referred to as the inspection phase and can last up to 30 minutes but may also be much shorter.

Adult L. stagnalis can lay egg masses at a frequency of at least one mass per week, under laboratory conditions, and such masses contain on average between 50 and 150 eggs, depending on the individual’s body size (Koene et al., 2007). This species seems to have a preference for laying eggs during the day (Ter Maat et al., 2012), but other species have been found to predominantly lay eggs during the night (Planorbarius corneus and Helisoma trivolvis: Cole, 1925). The number of eggs in an egg mass and the laying frequency also vary widely for other species (Bondesen, 1950; Dogterom & Van Loenhout, 1983). L. stagnalis that are older than 300 days usually show a decline in egg production (Janse et al., 1990; see also Hermann et al., 2009). Other factors that inhibit egg laying are starvation and dirty water (Ter Maat et al., 1982; Dogterom et al., 1984). As a consequence of the latter, egg laying can be triggered by a transfer from dirty to clean water. Although this is known as the clean water stimulus, the effect is actually caused by a combination of clean water, clean surface, and higher oxygen (Ter Maat et al., 1983, 2012).

Spiking activity of the CDCs is not affected by dirty water, but during starvation they become hyperpolarized, which elevates the threshold for egg laying (Ter Maat et al., 1982). Dopaminergic neuronal input from most nerves tested can also arrest CDC spiking (Ter Maat, 1979; Ter Maat & Lodder, 1980), and one important nerve in this respect is the intestinal nerve, which transmits information from the female reproductive tract, via the pleuroparietal connective, to the cerebral ganglia (Ferguson et al., 1993). The RPeNs also receive input from this intestinal nerve (Hermann et al., 1997), and lesioning this nerve results in decreased rasping and shell turning during egg laying (Ferguson et al., 1993); however, it does not eliminates the effects of α-CDCP and β3-CDCP (stimulation of shell movements and rasping rate; inhibition of locomotion; Hermann et al., 1997). Once dischared, the CDCs have a refractory period during which they are inhibited for up to 4 hours after egg laying, after which they return to their normal resting state (Kits, 1980). This inactivity likely correlates with the depletion of secretory products in the commissure after egg laying. These products are replenished in the hours following egg laying with the reserve pool of product that is present in the cell bodies of the CDCs (Jiménez et al., 2004).

The electrical properties and secretory products of the bag cells of Aplysia californica show striking similarities with those of the CDCs. For one, they are electrically coupled and show a similar all-or-none discharge, which results in the release of the egg-laying hormones that initiate ovulation and egg-laying behavior; subsequently these cells also go into a refractory period (reviewed in Conn & Kaczmarek, 1989). Also in this species, the egg-laying behavior follows a rather fixed sequence of events and the following description is based on previous detailed descriptions (Strumwasser et al., 1980; Cobbs & Pinsker, 1982a,b; Blankenship et al., 1983; Conn & Kaczmarek, 1989). Before laying eggs, animals cease eating and often locomote to a vertical surface. Other signs are puckering mouth movements, swelling of the common genital groove, and cessation of locomotion. They also start head-weaving movements and twitch their rhinophores (see later) more frequently. Head weaving signified the movement of the head back and forth in a very stereotyped fashion, which, in combination with tucking movements, allows the animal to deposit a compact egg mass that is properly attached to the substrate (which is aided by a sticky mucus).

Aplysiid egg laying is a seasonal activity and an increase in seawater temperature enhanced it (Pinsker & Parsons, 1985). Another natural stimulus that may lead to the initiation of egg laying is related to pheromones released by other conspecifics: Aplysiids seem to aggregate for mating and egg laying, which is mediated by a number of pheromones, including one called attractin, resulting in synchronized laying (Audesirk, 1977; Painter et al., 2003, 2004; Cummins et al., 2004; see also later). A single egg mass may contain well over 100 million eggs (MacGinitie, 1934; Susswein et al., 1984), and under laboratory conditions this can amount to 478 million eggs over roughly 4 months (MacGinitie, 1934).

Precopulatory Detection of Mating Cues

One of the prominent factors that affect mate choice is mating history of both the donor and recipient. For example, while from the male perspective it will be beneficial if the individual is able to fertilize many sperm recipients (Bateman, 1948; Anthes et al., 2010), there is generally some limit to the number of inseminations that an animal is willing to perform. Part of this is due to the (substantial) costs involved in the production and transfer of ejaculates (Dewsbury, 1982; Hoffer et al., 2010; Franklin et al., 2012). As a consequence, sperm donors should choose to inseminate mates with a high reproductive potential. This can, for instance, be achieved by preferentially mating with virgin partners, as a sperm donor can then potentially monopolize the reproductive resources of its mate. Also, avoiding partners that have recently mated would lead to avoiding sperm competition (i.e., inseminating individuals that already have allosperm from other partners stored in their storage organ; Jarne et al., 2010; Koene, 2010). Other factors that can be relevant for the assessment of the quality of potential mates include body size and shape, morphology, parasite infection, disease resistance, genetic relatedness, mating history, mate identity, and population origin (e.g., Nakadera & Koene, 2013). In the following we will explore some of these factors as well as which proximate mechanisms possibly lie at the basis of such mating decisions and which cues (such as water-borne chemicals, substances in mucus trails, or tactile information) may be involved.

Cephalopods—Reproductive Strategies: Motivation, Mate Choice, and Mating History

Like most organisms, cephalopods use several strategies to reach maximal reproductive fitness. Their very different lifestyles and body forms could explain why the mating systems among these animals are quite different (Hanlon & Messenger, 1996). They have evolved a high adaptive flexibility (Rocha et al., 2001). For example, the reproductive options of coleoids, which are usually short-lived, extend from massive and simultaneous spawning at the end of the animal’s life to continuous spawning during long portions of their life spans. As a consequence, several different classifications have been suggested to describe reproductive patterns in cephalopods (Rocha et al., 2001; Boyle & Rodhouse, 2005). It is obvious, however, that both the frequent semelparous and the rare iteroparous behaviors of cephalopods (Boyle, 1983, 1987; Boyle & Rodhouse, 2005; Franklin et al., 2012) imply a high-energy demand on females to produce eggs.

In these promiscuous mollusks, the act of mating may also be an energy-demanding process. For example, after mating both male and female individuals of the dumpling squid, Euprymna tasmanica, have been observed to display reduced swimming endurance (Franklin et al., 2012). Such a locomotion impairment would be ecologically disadvantageous for squids since it could compromise future mating and successful reproduction (Franklin et al., 2012).

At least two different mating systems can be distinguished: (1) in some cuttlefishes and squids there is elaborate agonistic behavior, extensive courtship, and brief copulation, but no protection of the eggs; (2) in octopuses there is little, if any, agonistic or courtship behavior, yet copulation is relatively lengthy, and females care for the eggs (Hanlon & Messenger, 1996). Some of these behavioral aspects of reproduction are controlled by hormones/paracrine substances (Wodinsky, 1977; Cummins et al., 2011; Enault et al., 2012).

Mate choice is one important aspect for maximizing reproductive fitness, although many questions about mate choice remain open for decapods and octopods. What is known is that visual display behavior of decapods can be interpreted as a warning signal for other, competing males and/or as an “ornament” to attract females. Such visual display is apparently absent in octopods, and this difference in reproductive behavior with decapods remains a key question in cephalopods.

In general, decapods use body patterns (deriving from the chromatophores and other structural “elements” in the skin: Hanlon & Messenger, 1996) to communicate intra- and intersexually. Such chromatic displays, coupled with specific arm movements, can also be indicative of the sex of the animal; that is, during fighting, courting, and mating, decapods display their sex. For example, in S. officinalis, agonistic contests are characterized by an Intense Zebra Display from the males (see Hanlon & Messenger, 1988, 1996, for references) with the arms either stretched forward or arched toward other males. The visual display can lead to intense fighting, culminating in the winning male approaching the female and copulating with her.

In squids, visual communication also plays a very important role in the intrasexual as well as intersexual relationship during mating. Displays are made in the shoal, where courting parties (one female to four to five males) are formed. There is often a large male that approaches the female, trying to drive away other males. Such intrasexual and intersexual messages are transmitted by specific body patterns and, surprisingly, females can use these to attract males other than the one close to her.

Fighting and competitive aggression among decapod males is a means by which males gain access to preferred females for mating. Males of cuttlefishes and squids generally approach females in small groups (cuttlefishes, two to three males for female) or large groups (courting parties in squids). This approach has been observed at the moment of mating in Sepia, and it may be present in Sepioteuthis or Doryteuthis (Hanlon & Messenger, 1996). Generally, in cuttlefishes, the male–female pair mates in a “head-to-head” position. Interestingly, courtship and mating in S. officinalis can be highly tactile; a male will hover over and alongside the female, continually drawing his arms softly over her mantle, head, and arms (Hanlon & Messenger, 1996). Unfortunately, there are as yet no clues as to how this tactile aspect of male courtship affects the receptivity of the female. During mating, the male will pass spermatophores that are stored mainly in the seminal receptacle below the mouth.

In squids, males engage in agonistic contests among themselves until a winner emerges who keeps away all the others from his partner. As a pair becomes isolated, mating begins. A head-to-head as well as a male-parallel position is used: Spermatophores are passed to the female and stuck on the mantle, around the head, or in the seminal receptacle, according to the species (Hanlon & Messenger, 1996).

Females synthesize a protein named Loligo β-microseminoprotein (Loligo β-MSP) in their reproductive exocrine glands and embed the protein in the outer tunic of egg capsules (Cummins et al., 2011). This protein is able to immediately and dramatically change the behavior of male squid from calm swimming and schooling to extreme fighting, even in the absence of females. Males are attracted to the eggs visually, but upon touching them and contacting Loligo β-MSP, they immediately escalate into intense physical fighting with any nearby males. Such evidence highlights the role of chemical communication during reproduction in cephalopods.

In contrast, octopuses are solitary animals that show little agonistic or courtship behavior before mating. In O. vulgaris, no specific display seems to exist that would distinguish between the octopus sexes (Wells, 1978), so cohabitation is not present (Hanlon & Messenger, 1996). O. cyanea (Wells & Wells, 1972) seems to show some striking males-discouraging displays. Packard (1961) suggested that O. vulgaris males show the larger suckers at the base of their second and third pair of arms as a signal of “maleness.” However, there is no definitive evidence for this, and it can only be supposed either that there are subtle visual cues that we have not detected so far (Wells & Wells, 1972), or that sex recognition is chemotactile in this species.

Male octopuses fight, and when the smaller cannot escape, he is killed. A female typically submits to the demands of the males. Each male carries about 50 spermatophores, each 2–3 cm long, that they place individually at the base of the hectocotylized arm with an “arched” posture followed by an explosive “pumping” action that somehow sends the spermatophore down the arm and into the oviduct. Competition among males for female is common in Octopus. Several observations report of two to three, or even six octopi mating with one female (Wood, 1963; Voight, 1991; but see Hanlon & Messenger, 1996).

Given the promiscuity of cephalopods, familiarity as well as mating history may influence reproductive behavior. In decapods, females that have not mated recently show a more receptive behavior (Schnell et al., 2015), but only when they mate with unfamiliar males. Mating behavior in males is unaffected by positive or negative female receptiveness, but it is strongly inhibited by familiarity with females, preferring unfamiliar matings (Schnell et al., 2015). Moveover, mating history can be detected albeit in a not-yet-specified way (possibly chemical, see Squires et al., 2013), as evidenced by increased mating duration in pygmy octopus (Cigliano 1995).

Chemosenses

The Olfactory Lobe and the Olfactory Organ in Cephalopods

In cephalopods, the olfactory lobe lies on the optic tract, close to both the peduncle lobe and the optic gland, and is subdivided into three lobules (anterior, middle, and posterior in O. vulgaris) (Messenger, 1967; Young, 1971). From the anatomy, one could infer that the olfactory lobe receives and processes distant chemosensory information (olfaction sense) (Young, 1971; Hanlon & Messenger, 1996). In fact, the name of this lobe comes from its connection, through the olfactory nerve, to the olfactory organ, a chemoreceptor organ (Woodhams & Messenger, 1974; Polese et al., 2016). However, the function of the olfactory organ, at least in Octopus, is still far from clear, although a role of olfaction in this mollusk’s reproduction has been hypothesized (Polese et al., 2015).

Knowledge about the olfactory organ in cephalopods comes from studies on Nautilus (Basil et al., 2000; Ruth et al., 2002) and decapods (Lucero et al., 1992, 1995, 2000; Piper & Lucero, 1999; Mobley et al., 2007; Mobley et al., 2008a,b; Villanueva & Norman, 2008). In squid, the olfactory organ is a sensory epithelium made of ciliated supporting cells and sensorial bipolar neurons. Each receptor neuron is connected to the olfactory lobe and other areas of the CNS (Messenger, 1967, 1979). Relatively recently, Walderon et al., (2011) investigated the role of olfaction in the distance chemoreception of conspecifics in Octopus bimaculoides, but almost nothing is known about the mechanisms, functions, and modulation of the olfactory organ in octopods.

It should be emphasized that in Octopus, which uses its arms for detecting food, there are millions of chemoreceptors on the suckers (Graziadei, 1964). The role of olfactory organs and olfactory lobes is therefore presumably related to distance chemical reception (olfaction), rather than chemotactile perception via the suckers (taste by touch; Wells, 1962; see also Hanlon & Messenger, 1996). The presence in this lobe, particularly in the posterior lobule, of peptidergic neurons that innervate optic gland cells (see earlier), has led to a critical revision of the role played by the olfactory lobe in reproduction (Di Cosmo & Di Cristo, 1998). These neurons send their axons to the chief cells of the optic gland (Di Cosmo & Di Cristo, 1998; Suzuki et al., 2000, 2002; Iwakoshi et al., 2002; Iwakoshi-Ukena et al., 2004; Di Cristo et al., 2005), possibly mediating the control exerted by the olfactory system on optic gland activity.

Apparently, some of the GnRH (and other neuropeptidergic) neurons in the olfactory lobe also send their axons to the mucosa of the olfactory organ (Polese et al., 2015). According to these data, signals from the chemical world outside the animal are relayed to the optic gland, modulating its activity, through the two-way connection between olfactory organ and lobe. If this is anatomically correct, we cannot yet formulate any definitive theory about the role that both chemical stimuli and GnRH could have on the functioning of the optic gland and its hormone. So far, there is no evidence of any molecule perceived by the olfactory organ that could trigger sexual activity, nor has a clear “receptive” role in reproduction been described for the olfactory organ. According to Wells and Wells (1972), chemotactile stimuli could have some importance, suggesting a role of suckers in reproductive behavior (at least in Octopus).

Incidentally, another neuropeptide has been found in the posterior olfactory lobule: the tetrapeptide Ala-Pro-Gly-Trp-NH2 (APGWamide; Di Cristo et al., 2005; Di Cristo, 2013). APGWamide is known to play a key role in male sexual behavior in gastropods (De Lange, Joosse, & Van Minnen, 1998; De Lange & Van Minnen, 1998, see earlier). In the Octopus CNS, APGWamide-immunoreactive cell bodies are confined, other than olfactory lobe, to the inferior frontal system. This system of lobes is involved in the reception and analysis of chemosensorial information coming from suckers in Octopus (Young, 1971). Interestingly, and in connection to this, research on Loligo demonstrated that visual and chemotactile input evokes intramale competition for mates (Buresch et al., 2004). In octopuses too, female touch may alter sexual behavior of males, by making them very active. This observation of mating behavior in octopuses suggests that contact may play a part in chemical-mediated sex recognition (Hanlon & Messenger, 1996). This “taste by touch” strategy, common in many behaviors of Octopus, fits well with the identification of APGWamide neurons in the inferior frontal system in both male and female Octopus. This system, which is exclusively present in octopods, forms a functional unit correlated with chemotactile information from arms and their use (Young, 1971). It could be suggested that “chemotactile approach” might be fundamental during mating. Supporting this idea is the evidence that the only operation that prevents normal male sexual behavior in octopus, which comprises the transfer of spermatophores by using the third right arm, the hectocotylus, is the removal of the tip from such an arm; evidently, some signal from the tip indicates when it is in the right position (Wells & Wells, 1972). Irrespective of whether this “probing” behavior comprises exclusively tactile information or also chemical cue, the presence of APGWamide in the inferior frontal system suggests that this neuropeptide is involved in this behavior. Moreover, the parallel seems clear with probing in Lymnaea, where APGWamide is also known to play a role in preputium positioning, and may relay sensory information about the preputium’s tip to the CNS (De Lange et al., 1998b).

Gastropods: Motivation, Mate Choice, and Mating History

Mating History and Mate Choice

Male reproductive costs have been shown to be roughly equal to the female costs in L. stagnalis by experimental elimination of the male function (De Visser et al., 1994; Hoffer et al., 2010). So, given these costs, do sperm donors choose their mates strategically? In the freshwater snail L. stagnalis, male-acting individuals do not seem to discriminate between virgins and mated individuals, the likelihood of mating with either is the same (Koene et al., 2008), but they do transfer more sperm once mating takes place (Loose & Koene, 2008; see next section on postcopulatory sexual selection). If inseminations are energetically costly, an individual that has already inseminated a partner would be predicted to prefer to inseminate a novel partner. This effect has become known as the Coolidge effect and can be tested by offering individuals the choice between familiar and novel partners (Wilson et al., 1963; Koene & Ter Maat, 2007). When this was done in L. stagnalis, this revealed that this species prefers a novel over a familiar partner (Koene & Ter Maat, 2007). Especially when ejaculates are expensive to produce and transfer, it is beneficial for the male role to choose a novel partner, because the donor can expect to fertilize more eggs in that novel partner than in the partner it has already given sperm to (and invested an ejaculate in). Such a decision to allocate sperm optimally over many mating partners, rather than to give everything to only one, is predicted to lead to a higher male reproductive success in the long run, to help avoid potential fertilization incompatibilities and effects of inbreeding (if some of the partners happen to be related to the sperm donor) as well as local sperm competition (among sperm from the same donor; e.g., Schärer, 2009).

The Coolidge effect found in L. stagnalis seems to depend on chemical cues, given that this preference was no longer observed when the mating trials were performed in a clean aquarium. Whether this is then due to the mucus trail or to specific chemicals released by the recipients remains to be investigated (Koene & Ter Maat, 2007). Interestingly, the Coolidge effect was also tested in the freshwater snail Biomphalaria glabrata and the sea slug Chelidonura sandrana but found for neither (respectively, Häderer et al., 2009; Werminghausen et al., 2013). One explanation that has been invoked for both species is that, given their promiscuity, they do not obtain sufficient benefits from discriminating between different mating partners. While this is partially supported by the higher mating rates of this species, which is 4–13 times within 12 hours (Vernon & Taylor, 1996), we will see later that B. glabrata does distinguish mating partners based on other traits. It would now be interesting to see whether partner novelty (and also virginity) affects mate choice in other gastropods than the three that have been tested so far (also the opisthobranch sea hare Aplysia fasciata, for which the effect was proposed but never experimentally tested; Ziv et al., 1989), since this could give an indication of the importance of such cues for mate choice (as well as sperm competition). For instance, the Coolidge effect may lie at the basis of why snails tend to perform more inseminations when they are in larger groups, a situation in which they encounter many novel partners (e.g., Koene & Ter Maat, 2007).

Motivation to Mate

There are, of course, many environmental factors that influence whether mating will take place or not. One can think of the influence of temperature, food availability, parasites, and the like (e.g., Nakadera & Koene, 2013). But even when all these factors are optimal, one can still observe that a period of sexual isolation will influence the likelihood of mating. Such increased eagerness to mate after sexual isolation seems to be a common phenomenon in a range of simultaneously hermaphroditic gastropods (e.g., Aplysia fasciata: Ziv et al., 1989; Helix aspersa: Adamo & Chase, 1990). There are two main reasons for this that we will explore here: either it is profitable to mate (in one of the sexual roles) or the mating history has temporarily motivated the individual.

The profitability to mate has been best investigated, and it indicates that one of the main factors causing variation in motivation to mate is sexual isolation. This is often used experimentally to increase the likelihood of mating (Koene & Ter Maat, 2005). For example, for the description of the copulation strategy of Stagnicola elodes and Biomphalaria glabrata, sexually isolated individuals were used (respectively, Rudolph, 1979a; Vernon & Taylor, 1996). Given that these animals are hermaphroditic, but generally mate unilaterally, it is important to consider which role drives this decision to mate.

This has been investigated in detail in L. stagnalis, which has also been shown to become more motivated after sexual isolation. More important, the motivation to mate in the male role seems decisive. For example, previous work has shown that individuals that have been sexually isolated for a longer time than their partner will act as sperm donors (Van Duivenboden & Ter Maat, 1985). When both individuals are motivated to donate sperm, role alternation will take place so that both individuals of the mating pair get to donate (and receive) sperm (Koene & Ter Maat, 2005). The one that inseminates first is generally referred to as the primary donor, and the other as the secondary donor (e.g., Nakadera et al., 2014a). To ensure the chance to also inseminate, the secondary donor may even display a typical mating posture in which it already holds onto the shell of the partner, ready to mount, well before this one has finished inseminating (Koene & Ter Maat, 2005). However, if more than one partner is around, that is, if the pair is not confined to a small mating arena, but rather in a group, the secondary donor need not inseminate its primary donor, but is just as likely to choose a different partner to donate sperm to (e.g., Koene & Ter Maat, 2007). In other words, this species mating behavior is based on unconditional unilateral mating and reciprocity is not conditional for initiating a mating (Koene & Ter Maat, 2005).

So how does the individual “decide” about when to be male or, in other words, what is the underlying physiological mechanism? One of the main factors determining this decision seems to be the availability of seminal fluid, which is produced in the prostate gland. After sexual isolation, the animal has had time to fully replenish the seminal fluids that have been transferred in previous matings, which can be measured in terms of gland weight (de Boer et al., 1997). Electrophysiological experiments have demonstrated that increases in gland size are detected by the CNS via a small branch of the penial nerve (np1; De Boer et al., 1997). This nerve branch feeds into the different regions within the CNS that are known to be involved in male mating behavior (reviewed in Jarne et al., 2010; Koene, 2010). That this information is crucial for the execution of the male role has been demonstrated by lesioning this nerve, which results in complete elimination of the male function, that is, feminization (Van Duivenboden et al., 1985; De Visser et al., 1994; Koene et al., 2009a; Hoffer et al., 2010).

A study on Physa heterostropha pomilia also shows that isolated individuals tend to mate more often as males than mated individuals (Wethington & Dillon, 1996). Moreover, role alternations were predominantly seen between isolated individuals (Wethington & Dillon, 1996; Koene & Ter Maat, 2005). That this may be a general pattern is further supported by reports that role alternation is reportedly very rare in “spontaneous” copulations, that is, copulations between nonisolated snails (Bulinus globosus: Rudolph, 1979b; L. stagnalis: Van Duivenboden & Ter Maat, 1988; Physa heterostropha pomilia: Wethington & Dillon, 1996). What now remains to be demonstrated for these species is whether the underlying physiological mechanism is the same as in L. stagnalis.

Besides motivation to perform the male role, the female-acting individual may also show sexual drive, or receptivity, depending on circumstances. For example, the ability to self-fertilize is common within this group, yet the tendency to favor outcrossing over selfing seems to vary widely between species and families (Escobar et al., 2011). L. stagnalis, for instance, has a strong preference for outcrossing (Cain, 1956; Knott et al., 2003; Koene et al., 2009). As mentioned earlier, most freshwater pulmonates can mate in the male and female role, but within one copulation each individual only performs one sexual role. Generally, it is assumed that these snails are usually receptive as females, because they seem rather inactive when copulating in this role (Van Duivenboden & Ter Maat, 1985). However, behavioral work on several freshwater snails has shown that these hermaphrodites can display behaviors that discourage the mating partner from inseminating or even dislodge the sperm donor from the shell. For example, in P. acuta this is done by vigorous shaking of the shell and biting in response to a conspecific mounting the individuals shell (DeWitt, 1996). In Bulinus globosus, the mounted individual twists its body (and hence its shell) in such a way that the prospective sperm donor cannot reach the female gonopore for insemination (Rudolph & Bailey, 1985). Similar insemination avoidance behaviors may also be present in L. stagnalis (as indicated by Van Duivenboden & Ter Maat, 1988), but this remains to be fully confirmed.

There are several convincing studies that show that such behaviors are indeed used in mate choice. One example comes from P. acuta, where the use of insemination avoidance behaviors has been shown to depend on the genetic background of the mating partners (Facon et al., 2006). In that experiment, the female acting (mounted) snails showed a higher frequency of shell swinging and biting of the partner’s preputium (phallus) when these partners were related (siblings; Facon et al., 2006). Given the high inbreeding depression that this species shows, such insemination avoidance behavior may help in preventing inbreeding, via earlier termination of unwanted mating attempts. In contrast, a closely related species from the same genus, P. gyrina, has been reported to show more intense avoidance behavior when mated by partners from other populations (McCarthy, 2004). These findings, taken together, may indicate that these species prefer to mate with sympatric individuals as long as they are not highly related (i.e., siblings). Such female behaviors remain to be explored for other species.

Detection of Mate Choice Cues via Mucus Trails and Pheromones

So, a crucial question emerges from this: How are traits and reproductive states of potential mating partners detected by sperm donors? Obviously, given that these animals have rather limited visual and acoustic senses, much of this is detected via chemical information (e.g., Chase, 2002; Cummins & Wyeth, 2014). However, even though several studies have demonstrated that pheromones and/or mucus trails are important for such mate choice decisions (e.g., Ng et al., 2013), the mechanistic link to how the animals detect these cues has remained very underexposed. In the following, we will explore what is known and what remains to be investigated in order to obtain a clearer picture of the involved physiology.

Nearly all gastropods release mucus, a sticky slime-like substance. They do this ventrally, via their foot, for which they are often equipped with mucous glands that are embedded in the foot (sometimes called the pedal gland). This mucus is used for gliding over surfaces and for adhering to these same surfaces. Via the dorsal surface of their skin they also produce mucus that prevents desiccation and dilutes or washes off irritants (Chase, 2002). Moreover, chemical or tactile irritants that discourage predators can be present and are produced in epidermal gland cells. For example, in L. stagnalis, 11 different epidermal gland types can be distinguished in the skin. Three types of gland cells are found in numerous places in the snail’s skin, while other types are specific to the foot, lip, or mantle regions (Zijlstra, 1972).

The mucus is composed of water in combination with proteins, carbohydrates, and lipids (Zijlstra, 1972; Davies & Hawkins, 1998). The major structural components of molluskan mucus are glycosaminoglycans and proteoglycans (proteins that are highly glycosylated). These proteoglycans are thought to provide stability to other proteins present in the mucus and are therefore less easily degraded. In the mucus trail of L. stagnalis at least three types of high molecular weight glycoconjugates, which are mucin-like, have been identified and have the ability to form gels (Ballance et al., 2004).

In addition to the general constituents that make the mucus sticky, it also contains specific pheromones and other types of chemicals that can provide information about the animal. In aquatic environments, such substances can often dissolve and/or diffuse into the water and can thus travel over long distances. In such cases, such pheromonal information can lead to aggregations, not only for mate searching purposes but also by way of food sharing and group defence (Kuanpradit et al., 2012). Several of such pheromones, the best known of these is a protein pheromone called attractin, have been shown to have such functions in the Opisthobranch sea hare Aplysia californica and its congeners (and is released from the egg cordon; e.g., Painter et al., 2003, 2004; Cummins et al., 2004).

A similar pheromone to one of the components of the Aplysia blend, called temptin, seems also to be produced by the hermaphroditic freshwater snail B. glabrata (Hathaway et al., 2010), as well as the broadcast spawning separate-sexed abalone Haliotis discus (Kuanpradit et al., 2012), for which it has been shown that females are attracted to pheromones released by the male prior to spawning (Nhan et al., 2010). This suggests that some of these pheromones are highly conserved, even across mating systems and habitat types. This is also supported by the fact that L. stagnalis is also attracted to the Aplysia pheromone attractin (Painter et al., 2004). However, whether these pheromones play a similar role as the one proposed in Aplysia—aggregation for egg laying and mating—remains to be convincingly shown in any species.

Such pheromones that are released into the water column may serve the more general purpose of locating potential mating partners (or suitable habitat or egg-laying sites). But mucus trails can also contain pheromonal information that works at the closer range for finer-scale location of conspecifics that have left the trail. This can often result in so-called trail following, where individuals follow the mucus trail (laid by themselves or others). Several reviews indicate that such trail following can serve many adaptive purposes, which include homing, prey location, and mate finding (e.g., Ng et al., 2013). Obviously, the latter is of relevance here. Such research has, for example, shown that sexually motivated Biomphalaria glabrata are more likely to display trail following than nonmotivated individuals (Townsend, 1974). And both Biomphalaria glabrata and Physa acuta seem to even be able to detect the direction in which the trail was laid (Wells & Buckley, 1972; Townsend, 1974), possibly due to a chemical gradient in the trail (Bousfield et al., 1980).

Chemosenses: Osphradium and Cephalic Sensory Organs

In the Gastropoda, the osphradium is a peripheral sensory organ that has mechano-, osmo-, and chemosensory properties (Chase, 2002; Cummins & Wyeth, 2014). This organ is widespread within the Mollusca and is generally located in the mantle cavity (e.g., Haszprunar, 1987). The osphradium of aquatic pulmonate snails is found just anterior to the pneumostome, the animal’s lung opening (Chase, 2002; Cummins & Wyeth, 2014). The general morphological layout of the osphradium is similar within all aquatic pulmonates that have been investigated. Among these are the blood fluke planorb, Biomphalaria glabrata (e.g., Michelson, 1960); the great rams horn, Planorbarius corneus (e.g., Benjamin & Peat, 1971); and the great pond snail, L. stagnalis (e.g., Wedemeyer & Schild, 1995).

In L. stagnalis, again the species in which this organ has been studied most extensively, the organ is described as having a Y- or λ-shaped epithelial canal that has a connection to the outside via a small pore. Although the exact shape of this epithelial canal may differ between species, they all conform to the following morphological description. From the osphradial pore, the epithelial canal leads inward and its epithelium can be divided into three regions (e.g., Townsend, 1973; Nezlin et al., 1994; Wedemeyer & Schild, 1995). In the region closest to the opening of the canal, the epithelium is ciliated, and subepithelial receptor neurons are present. The inner, sometimes bifurcated, part of the canal is covered with secretory cells. The middle, where the canal bifurcates in L. stagnalis, consists of both ciliated cells and numerous sensory cell dendrites that terminate at the surface (Nezlin, 1995; Kamardin et al., 1998). The latter is referred to as the sensory epithelium (Wedemeyer & Schild, 1995).

The osphradial canal is surrounded by a layer of muscle cells and an osphradial ganglion that contains the subepithelial receptor neurons, the sensory neurons, and other ganglion cells. The few studies that have been performed on these osphradial neurons indicate that several regulatory substances are present (γ-amino-butyric acid/GABA, FMRFamide, serotonin/5HT, methionine- and leucine-enkephalin; Nezlin et al., 1994; Nezlin & Voronezhskaya, 1997). Besides sensory neurons, serotonin-containing motor neurons are present (Benjamin & Peat, 1971; Nezlin et al., 1994) that have been proposed to control the contractions of the osphradial muscle cells to force the water surrounding the animal in and out of the epithelial canal (Nezlin et al., 1994).

The osphradial nerve leads into the right internal parietal nerve that connects this ganglion to the parietal ganglion of the CNS (Crisp, 1973; Wedemeyer & Schild, 1995; Nezlin, 1997; Kamardin et al., 1998). The osphradial nerve runs very superficially under the skin, just behind the female gonopore, and is therefore relatively accessible in an anaesthetized snail. Hence, experimentally lesioning of this nerve has been instrumental in answering several questions about the function of the osphradium. For example, osphradectomy (i.e., the denervation or removal of the osphradium) has revealed that the organ is involved in detecting changes in respiratory gas concentrations as well as in chemoreception of organic compounds (Wedemeyer & Schild, 1995). This is also corroborated by neurophysiological experiments demonstrating that the osphradial receptor neuronss detect changes in O2, CO2, NaCl, KCl, and amino acids as well as more complex odour molecules (Kamardin, 1995; Wedemeyer & Schild, 1995). However, whether food cues are also detected via the osphradium remains unclear, especially because studies using an osphradectomy approach in Biomphalaria glabrata have come up with mixed results (e.g., Michelson, 1960; Townsend, 1973). Of course, freshwater pulmonates do locate food successfully but may use different chemosenses (Bovbjerg, 1968). The fact that amino acids are detected by the osphradium suggests that food detection may play a role as well (at least in L. stagnalis: Wedemeyer & Schild, 1995). More relevant for the topic at hand, the osphradium has been shown to pick up chemosensory information from injured conspecifics (Dalesman et al., 2006), low calcium availability (Dalesman et al., 2011), and predators (in the form of kairomones; Dalesman & Lukowiak, 2011). Moreover, such cues have been shown to affect respiration and behavioral responses to food sources and predator presence (Dalesman et al., 2011), and they can even interfere with long-term memory formation that is thought to be related to stress (e.g., Karnik et al., 2012).

Besides detecting environmental cues important for survival, detected olfactory information seems to extend to nonaquatic compounds that are known to stimulate olfactory neurons in nonaquatic animals (Wedemeyer & Schild, 1995). All this does indicate that the osphradium is capable of mediating information needed for homeostatic responses as well as real olfactory responses involved in distance chemoreception (Wedemeyer & Schild, 1995; Kamardin et al., 1998; Cummins & Wyeth, 2014). Whether this extends to detection of information that is relevant for reproductive decisions is less clear (Wedemeyer & Schild, 1995). There are two suggestions indicating that this may be the case. For marine pulmonate gastropods, of the genus Siphonaria, it has been suggested that the osphradium could play a role in homing (Kamardin, 1983, 1988). In addition, Townsend (1973) speculated about the osphradium’s potential role in pheromone detection in Biomphalaria glabrata.

The ospradium does seem to be involved in the decision to lay eggs, which is partly regulated by the presence of clean water (Ter Maat et al., 1988). It has been shown that spiking activity in the osphradial nerve increases when clean water comes through the epithelial canal, while this activity decreases with dirty water (containing urea; Kamardin et al., 2001). Since neurons in the osphradial ganglion project all the way into the cerebral ganglia, and there are even two bilateral neurons with projections into the osphradial nerve (Nezlin, 1995), suggests that it is possible that information about water quality is relayed to the caudo-dorsal cells (CDCs). This latter bilateral cluster of neurosecretory cells, which is responsible for releasing the egg-laying hormone and associated peptides that set ovulation and egg laying in motion, is known to become active after transfer from dirty to clean water (see later, reviewed in Ter Maat et al., 1988; Koene, 2010). Interestingly, osphradectomy increased, rather than reduced, egg laying, suggesting that this organ generally has a tonic inhibitory effect on egg laying under the circumstances this was tested (Nezlin, 1997). Nevertheless, this does reveal that information coming from the osphradium about water quality, which correlates with (un)favorable egg-laying conditions, is used for this reproductive decision (Nezlin, 1997; Kamardin et al., 2001).

Albeit admittedly rather speculative, there may also be a connection to the regulation of male mating behavior. The central neuronal network that innervates the penial complex includes many peptidergic neurons that are found clustered in the anterior and ventral lobe of the right cerebral ganglion, the I-cluster of the right pedal ganglion cluster, and dispersed cells in the right pleural and parietal ganglia (Smit et al., 1992; De Boer et al., 1997; El Filali et al., 2003, 2015). Interestingly, the two right parietal neurons that are backfilled via the osphradial nerve seem to be close to the B-cells, where the aforementioned parietal dispersed cells are also found (Kamardin, 1995; El Filali et al., 2003, submitted). Some of these latter neurons do have projections in both the penis nerve and the right internal parietal nerve. Whether these connect to the osphradium, or whether the neurons near this B-cluster integrate chemical information from the osphradium into the network regulating male behavior, is unknown, but worth investigating.

Besides the osphradium, much of the perception of the external world in relation to feeding, homing, aggregation, mating, escape, and avoidance, happens via the cephalic sensory organs (e.g., Chase, 2002; Wyeth & Croll, 2011). When referring to the cephalic sensory organs, one simply means the lips and tentacles. These organs are involved in most of the chemosensory and mechanosensory reception of gastropods, meaning that they are also used in both contact and distance chemoreception (Chase, 2002).

Basommatophora have only one set of tentacles with an eye spot at the base of each tentacle. This is in contrast to Stylommatophora, which have two sets of tentacles, the inferior and superior (or cephalic) ones; the latter bearing the eye spots and the chemosensory receptors at their tips. In the Basommatophora, species generally have chemosensory receptors all over the tentacles, but there are exceptions where a specific patch of chemosensory receptors is present at the base of each tentacle (e.g., Biomphalaria and Helisoma trivolvis: Emery, 1992).

The pair of triangular tentacles of L. stagnalis is covered with sensory endings of various types and also contains free nerve endings (Nezlin, 1995; Zaitseva, 1999). Their chemoreceptors respond to food items, their extracts, and amino acids (Zaitseva, 1994; Bovbjerg, 1968), while they do not react to sucrose (Nakamura et al., 1999a,b). Furthermore, experimental work has shown that it is possible that the chemoreceptors in the tentacles are peripheral neurons that either have central projections themselves or have peripheral synapses onto other peripheral neurons that project centrally (Wyeth & Croll, 2011; Cummins & Wyeth, 2014). Investigations into the central projection, which occurs via the left and right tentacle nerves, reveal that many neurons in nearly all ganglia of the CNS are innervated (Planorbarius corneus: Zaitseva, 1999; L. stagnalis: Zaitseva & Bocharova, 1981; Nakamura et al., 1999a,b). This clearly indicates that such information is used extensively by many biological functions of the animal (Wyeth & Croll, 2011).

The lip of L. stagnalis is innervated by the median and superior lip nerves that originate from both cerebral ganglia (Nakamura et al., 1999a,b). The superior lip nerve mainly innervates the mouth and the dorsal side of the head, while the median lip nerve mainly innervates the lip (Nakamura et al., 1999a,b). Afferent fibers from the lip and mouth area have been shown to not only terminate in the cerebral ganglia but also in the pleural, parietal, and pedal ganglia (reviewed for L. stagnalis in Cummins & Wyeth, 2014; Planorbarius corneus: Zaitseva, 1999).

The chemosensory neurons in the lips detect sugar, and much of the work on the control of feeding behavior has used this stimulus to unravel the mechanisms underlying feeding (e.g., Kemenes et al., 1986; Nakamura et al., 1999a,b; Ito et al., 2013). Such sugar stimulation seems to be mainly registered via the superior but not the median lip nerves (Nakamura et al., 1999a,b). The median lip nerves may therefore be more involved in mediating signals from mechanoreceptors that are used in the search for food and may be important in detecting contact pheromones.

Four types of peripheral sensory cells could be identified in the lips and tentacles. These include bipolar catecholaminergic, histaminergic, and nitrergic cells with dendrites that penetrate the sensory epithelia. The fourth cell type is unipolar nitrergic cells (Wyeth & Croll, 2011; Cummins & Wyeth, 2014). Information from these sensory cells reaches the CNS via the bilateral tentacle and lip nerves. And this information reaches many ganglia of the CNS, as evidenced by the extensively branched networks that can be revealed by backfilling these nerves. The fact that the further complexity of the peripheral sensory neuroanatomy has only recently been revealed indicates that there is still lots of scope for discoveries in this area of research, making it very difficult to integrate what is known into a comprehensive picture (e.g., Wyeth & Croll, 2011). Finally, it should be noted that much of this evidence is based on work looking at the control of feeding, using relevant stimuli for this, leaving the field of pheromonal detection via the cephalic sensory organs wide open, as well as how such information might reach the central neurons involved in reproductive processes.

Opisthobranchs have two pairs of tentacles. The posterior pair of tentacles is called rhinophores, literally meaning bearers of a nose. The anterior tentacles are called oral tentacles because they constitute the lateral margins of the oral veil. The epithelia of all these tentacles are specialized for chemoreception. Abolition of the rhinophores in Aplysia blocks the facilitation of mating, feeding, and respiratory pumping that is caused by the presence of pheromones. The tip of the rhinophores contains a specialized primary afferent organ which captures the odorant pheromones (Levy et al., 1997; Cummins & Degnan, 2010). This indicates that these rhinophores are involved in sensing pheromones and other chemicals.

Cephalopods: Postcopulatory (Neuro)physiological Processes

There is some evidence to suggest that male cephalopods may mate completely randomly and that females are either promiscuous or practice “simultaneous polyandry.” A stable pair almost never occurs in octopods or in decapods, where a temporary pair may be formed (sometimes “joined” by a sneaker male; Hanlon & Messenger, 1996). Clearly, repeated mating by females before laying eggs provides opportunity for sperm competition (Smith, 1984).

Sperm competition was defined originally by Parker (1970) as the competition within a single female between the sperm from two or more males for the fertilization of the ova. The ability to produce large quantities of sperm, sperm packaging in multiple spermatophores, sperm storage by females, the physical nature of the oviduct and spermathecae, the existence of “polygamous” mating systems, and nonsynchronous sexual maturation are all strong hints that sperm competition indeed exists in cephalopods, although proper evidence has only started to emerge recently (Hanlon et al., 1997; Squires et al., 2014, 2015).

In decapods, there is some evidence that the female can choose the spermatophores that are stuck around the head or in the mantle. Moreover, their seminal receptacle seems to be too small to contain millions of sperm. It is possible that the sperm from the most recent mating displace those from earlier matings. The male can also displace sperm from the female spermathecae, as revealed by the enlargement of the tip of the hectocotyli at the end of mating in Euprymna tasmanica (Squires et al., 2013). Similarly, in octopods, sperm competition is not unlikely. Sperm may be stored in the spermathecae of the oviducal gland, even months before the fertilization. Also in this order it has been suggested that the tip of the hectocotylus, which is spatula-like, could be inserted into the oviduct and reach spermathecae, removing old sperm (or sperm from another males) before depositing new sperm. Similar adaptations for sperm displacement from the female spermathecae, such as the presence of scoops, plates, or suckers on the hectocotyli that seem to function to break open competitors’ spermatophores, are found in other cephalopod species (Hanlon et al., 1999; Naud et al., 2005; Naud & Havenhand, 2006; Squires et al., 2013).

Once fertilization has been achieved in cephalopods, eggs should be protected by several layers of proteinaceous material and then laid and attached to rocks or stones (Hanlon & Messenger, 1996). Endocrine control on oviduct contraction could be performed through the release of neurohormones/neuropeptides from neurons of the visceral lobe into the vena cava to reach reproductive organs (Berry & Cottrell, 1970; Kanda et al., 2005). Alternatively, the fusiform ganglion of the peripheral nervous system might directly control muscle contraction of the reproductive ducts (in Octopus; Young, 1971). Peptides from the oxytocin/vasopressin family (Takuwa-Kuroda et al., 2003; Zatylny-Gaudin et al., 2016) seem to play some role in contraction of the female accessory gland and oviducts. FMRFamide and GnRH might also be involved in these physiological processes (Di Cristo et al., 2002; Iwakoshi-Ukena et al., 2004). APGWamide has, moreover, been shown to inhibit the contraction of the oviducts in Sepia officinalis (Henry et al., 1997). In Octopus vulgaris, APGWamide immunoreactivity is present in the visceral lobe as well as the visceral nerve (Di Cristo et al., 2005). This tetrapeptide is also present in the glandular cells of the oviducal gland, likely acting on the oviduct in an exocrine manner (Di Cristo et al., 2005). The only evidence that there may be male accessory gland substances, transferred along with the sperm in the spermatophore that influence these female processes comes from Sepia officinalis. In that species, APGWamide has been reported to be present in the case of the spermatophore (Bernay et al., 2006a) and to be involved in oocyte transport (Henry et al., 1997). In combination, these findings might hint at a possible allohormonal function for APGWamide.

Gastropods: Postcopulatory (Neuro)physiological Processes

Seminal fluid components, which are often referred to as accessory gland proteins, have been shown to play a major role in postcopulatory sexual selection processes in many animals (e.g., Arnqvist & Rowe, 2005; Perry et al., 2013). The presence and use of accessory gland proteins that are added to the sperm before transfer have only been investigated in one hermaphroditic freshwater snail so far. The prostate gland of L. stagnalis has been shown to produce a number of proteins and peptides that are transferred to the partner (Koene et al., 2010); these are added to the sperm when it passes through the gland’s lumen (Jarne et al., 2010). The receipt of one of these, referred to as ovipostatin or LyAcp10, induces a delay in egg laying of recipients (Koene et al., 2010). This delay seems to result in a higher investment per egg and has been proposed to enhance storage duration and/or fertilization chances of the donated sperm (Hoffer et al., 2012). The latter might occur because the delay in egg laying, which is relatively frequent in this species, might enable the donated sperm to reach the sperm storage site before a new set of eggs is fertilized and an egg mass is laid.

Very recent work has revealed that two of the identified proteins affect the male function of the recipient, something that is unique to hermaphrodites. These novel proteins cause a snail to transfer half the amount of sperm to its next partner. This sperm-number-reduction effect has been demonstrated both after natural insemination and artificial injection (via the female gonopore) with the isolated protein. Moreover, it has been shown that this reduction in sperm numbers is relevant for paternity success, because such snails achieved less paternity with their recipient (Nakadera et al., 2014a). The finding demonstrates that these hermaphrodites directly influence their partner’s male investment (called a cross-sex effect; Anthes et al., 2010, see later), while one normally expects males to affect female physiology. This novel insight profoundly affects the way in which one thinks about postcopulatory sexual selection in general. Clearly, this now also needs to be further investigated and verified in other hermaphroditic snails as well as other (molluskan) species where males can influence other males.

As becomes evident from the preceding, there are numerous processes that are being coordinated during and after the receipt of an ejaculate. This is a field of research that has only emerged relatively recently, especially when considering research on hermaphrodites, so some of the following will be rather speculative. Nevertheless, because a lot is known about the physiological regulation of many of the male and female reproductive processes that are involved, some tentative links can be made.

The identified accessory gland proteins, which have been shown to immediately affect postcopulatory processes, are the best place to start (see earlier; Koene et al., 2010; Nakadera et al., 2014a). For example, for the delay in egg laying that ovipostatin (LyAcp10) causes, one prediction could be that this directly, or indirectly, affects the excitability of the main neuroendocrine center that controls egg laying: the bilateral caudo-dorsal cell (CDC) cluster in the cerebral ganglia. While the CDC system has been described in detail in L. stagnalis (e.g., Ter Maat et al., 1983), it is also present in other freshwater species (see earlier), as is consistent with the presence of an ovipostatin-like substance in other species (Adema et al., in press).

There are different ways in which accessory gland proteins, like ovipostatin, could influence egg laying. First, this could be achieved by the protein being taken up into the blood of the recipient and directly targeting the activity of the CDCs. Secondly, peripheral receptors within the female reproductive tract may relay information relevant to egg laying to the CDCs. Ovipostatin could then either be a protein that serves as a cue telling the female system to delay egg laying because the received ejaculate needs to be processed, or it could act as a manipulative agent “hijacking” receptors that are normally used for the regulation of egg laying. Clearly, to find out, it will be necessary to identify the receptor on which ovipostatin acts as well as its site of action.

As with the female system, the accessory gland proteins (LyAcp5 and 8b) that have been shown to influence the male function of the recipient could also act via different routes. For example, given the decrease in sperm numbers, the release of ripe sperm from the seminal vesicles may be targeted. Alternatively, the transport of the ejaculate could be slowed down. This transport is controlled by the peristalsis of the vas deferens, which pumps the ejaculate across during copulation. The transport process has been proposed to be caused by the antagonizing effects of the neuropeptides APGWamide and conopressin (Van Golen et al., 1995). But there is a whole legion of other neuropeptides that is involved in the regulation of the male function, so it remains to be investigated how accessory gland proteins bring about their immediate sperm-number-reducing effect (Koene, 2010; Nakadera et al., 2014b).

Finally, although this remains to be investigated for Lymnaea stagnalis, accessory gland proteins may also target other processes in the recipients, such as allosperm storage and digestion, the recipient’s male or female behavior during mating interactions, as well as sex allocation. The latter, which is defined as the allocation toward the male and female reproductive function (Charnov, 1982; Schärer, 2009), could potentially also explain some of the effect that accessory gland proteins have in L. stagnalis. Although one generally measures effects on reproductive output in a specific sex function, it should not be forgotten that these are simultaneous hermaphrodites that perform both sexual functions. Therefore, assuming a fixed reproductive budget, a decrease in one sex function may actually result in an increase in the other (Charnov, 1979; Schärer, 2009; Koene, 2016). Hence, rather than targeting specific male or female processes, these accessory gland proteins could also be targeting the sex allocation decision of the recipient. In other words, by reducing the amount of sperm transferred, less new sperm will need to be produced and these leftover resources can be put into egg laying (either by increasing the total number of eggs produced or by increasing the quality and/or size of eggs; see also Schärer, 2014). The latter has been suggested to be the case in L. stagnalis, where repeated receipt of ejaculates increases the investment per egg (Hoffer et al., 2012). This is also in agreement with previous work showing that this species is very flexible in its resource investment (Van Duivenboden et al., 1985; Koene et al., 2009a). But while it has been shown that the elimination of the male function, by lesioning the aforementioned np1 nerve, leads to double the amount of eggs laid (De Visser et al., 1994; Koene et al., 2009; Hoffer et al., 2010), it remains unknown how such allocation decisions are executed by the CNS (Koene, 2010).

In land snails, research related to the transfer of accessory gland products has mainly been done in the brown garden snail, C. aspersum. This species exchanges sperm packages (also called spermatophores) during a single mating interaction. Accessory gland products are transferred prior to this exchange, during courtship, via so-called love dart shooting. This rather odd behavior implies the stabbing of a love dart (expelled via the genital pore by the forceful eversion of the dart sac) through the body wall of the mating partner. This action results in the love dart injecting digitiform gland mucus, which the dart is coated with into the partner’s haemolymph. The mucus causes conformational changes to the part of the female reproductive system that receives the spermatophore, resulting in altered spermatophore uptake and delayed sperm digestion (Koene & Chase, 1998a), thereby increasing sperm storage (Rogers & Chase, 2001) and ultimately paternity of the successful dart user (Landolfa et al., 2001; Rogers & Chase, 2002; Chase & Blanchard, 2006).

The dart’s accessory gland component that induces one of the muscular contractions in the spermatophore receiving organ was recently identified. The active peptide turns out to resemble buccalin, a known modulator of which different forms are used to change muscle contractions in freshwater and marine mollusks (Stewart et al., 2016a). This particular form, found in both C. aspersum and Theba pisana, is named love-dart allohormone (Stewart et al., 2016a) and supports that the effects of dart mucus are evolutionarily conserved within a number of dart-bearing species (Kimura et al., 2014; Lodi & Koene, 2016a,b). In the context of neuroendocrine regulation, one can at least speculate that the accessory gland products injected by the love dart may directly act on the (peripheral and/or) CNS, given that they circulate in the recipient’s haemolymph, but this remains to be demonstrated.

At least one neuropeptide, APGWamide, has been shown to be present as a male accessory gland product in the ejaculate of the bivalve Crassostrea gigas and, as mentioned earlier, in the spematophore case of the cephalopod Sepia officinalis (Bernay et al., 2006a; where it is directly involved in oocyte transport: Henry et al., 1997; see earlier). In Crassostrea gigas, APGWamide has been shown to modulate adductor muscle contractions that ensure the release of oocytes in the external medium. At the same time, unlike in Basommatophora (see earlier), APGWamide seems to have no effect on contractions of the penis and proximal vas deferens (Bernay et al., 2006b). As concluded by the authors of the latter study, at least in the bivalve the effect of APGWamide on the adductor muscle (oocyte release) is probably achieved via a pheromonal pathway (Bernay et al., 2006b). Given the difference in fertilization mode, external versus internal, the APGwamide on the spermatophore would then most likely function as an allohormone (Koene & Ter Maat, 2001).

Conclusions and Future Perspectives

The nervous system exerts a key role on the regulation of sexual maturity and sexual behavior in animals. Although this is true for all the animals we discussed in this chapter, they show different aspects of reproductive physiology that complicate the neurobiology of their reproduction. In fact, many gastropods are (simultaneous or sequential) hermaphroditic while cephalopods are gonochoric (separate sexed). This implies that in gastropods, different areas of the CNS control either male or female reproduction. Lymnaea provides the best example of how there is seemingly no overlap among neuropeptides and neurons controlling male and female reproductive behavior (see also Koene, 2010). Whether there are such neuronal and molecular sexual difference in cephalopods still remains to be determined. So far, the male and female cephalopod CNS (brain) shows no clear dimorphic anatomy, although the timing in the nervous control of sexual maturation is clearly sexually dimorphic. The two gonadotropin hypothesis, that is, the presence of the optic gland hormone as well as of the cephELH, could partially explain this asynchronous sexual maturation. At the same time, for hermaphroditic gastropods, it should be noted that a clear neuronal or neuroendocrine “switch” that prevents male and female behaviors from being executed at the same time remains to be uncovered (see Koene, 2010).

Another fundamental difference lies in reproductive strategies: iteropary in gastropods, semelpary in (almost all) cephalopods. This difference may have severe implication on how the neuroendocrine regulation of reproduction has been able to evolve in these different animal groups. For example, the energy demands are very different between the two classes: Cephalopods lay hundred of thousands of yolk-rich eggs once at the end of their life. At that life stage, energy from food should then be converted efficiently into substrate for vitellogenesis. The optic glands seem to be the main driver behind this, as they are involved in the somatic development of gonads and accessory organs. Interestingly, the dorsal bodies seem to have a very similar function in gastropods, making the possible homology between these two systems an interesting topic for future investigation.

Although both the optic gland and the dorsal body hormones are involved in the process of vitellogenesis and induce the enlargement of gonad and accessory reproductive organs, in female cephalopods the semelparous strategy, including the necessity of vitellogenesis, implies the mandatory action of optic gland hormone upon the putative egg-laying hormone action. The different reproductive strategy of gastropods does not affect the different action of the hormones (DBH and CDCH): The neuronal circuitry involved in growth inhibits egg laying in L. stagnalis. Yet there is an additional similarity between the two reproductive systems, which is based on the influence of light. This pond snail detects via ocular as well as nonocular photoreceptors (Ter Maat et al., 2012), and day length also seems to play a key role in the energetic regulation of egg laying. For example, long-day conditions cause L. stagnalis to channel most of its acquired resources into the production of eggs over the summer. Only under shorter day conditions do they also store some of their energy for later use (Ter Maat et al., 2007). These parallels, despite the different ecological strategies, seem fruitful avenues to investigate at the morphological, physiological, and molecular levels.

What also emerges from the preceding is that, although some cephalopods do use visual cues for sex and mate recognition, much of what happens during mating interactions seems to be governed by olfactory and tactile cues. Clearly, most gastropods have very limited visual capabilities, so they seem to largely rely on pheromonal cues for finding mates and tactile cues for successful sperm transfer. Interestingly, there seems to be a clear similarity between the use of sensory information by the cephalopod hectocotylus and the molluskan preputium/copulatory organ. Hence, an interesting topic for further investigation would be to see if similar, evolutionarily conserved, neuroendocrine substances and sensory neurons are involved in these processes. Such a comparison could be further amplified by also including sequential hermaphrodites, since the male and female system are clearly (and temporally) separated in such gastropods. In addition, this would allow for investigating how the neuroendocrine regulation changes when such species change sex during their lifetime. One obvious place to start, although far from a neurobiological model organism, would be the Owl limpet Lottia gigantea, mainly because the genome of this sequential hermaphrodite has been fully sequenced (e.g., Veenstra, 2010; although using the genome of the sequentially hermaphroditic bivalve Crassostrea gigas will also be interesting in this respect).

The aforementioned would also aid in identifying pheromones and male accessory gland products as well as their receptors. On the one hand, identification of pheromones and their sites of action (receptors) would provide a better fundamental understanding of how mollusks choose their mating partners and use chemical information in order to make mating decisions. On the other hand, such knowledge might eventually lead to the development of methods that can help to contain mollusks in places where they have become a pest and/or cause trouble as intermediate hosts for parasites. Likewise, a better understanding of the mechanism via which male accessory gland proteins exert their effects on reproduction (especially egg laying) could lead to using this knowledge for either preventing or improving reproduction, the latter clearly also being valuable for species that are bred for consumption.

The modes of action of male accessory gland products, albeit proteins or peptides, are also of interest in the evolutionary context of sexual selection. Especially when considering that there may be limits to how a sperm donor (male) may be able to affect processes in a sperm recipient (female; see also Koene, 2016), it is now important to come to a better understanding of how ACPs evolve and make use of (or “hijack”) the sperm recipient’s reproductive neuroendocrine system. That this is of importance becomes especially clear from reproductive interactions during which ACPs are transferred via injection devices that seem to have solely evolved for this purpose (Zizzari et al., 2014). Examples of this are the love darts of lands snails (mentioned earlier) but also the stylets of some sea slugs that seem to inject accessory gland products into the head of the partner, close to the CNS. For this latter species, belonging to the genus Siphopteron, it has been suggested that the injected substance, of unknown identity, may target the CNS directly (Anthes & Michiels, 2007; Lange et al., 2014).

As explained earlier, similar effects may also be achieved by ACPs that are added to the sperm. These clearly also deserve further attention in future research; especially given that the current knowledge is largely based on very few gastropods, it will be interesting to compare this to the role of ACPs in bivalves and cephalopods. In contrast, morphological adaptations for sperm displacement from the female spermathecae, such as the presence of scoops, plates, or suckers on the hectocotyli that seem to function to break open competitors’ spermatophores, have not been reported in gastropods, whereas these are found in more cephalopod species than the ones mentioned earlier in this chapter (e.g., Hanlon et al., 1999; Squires et al., 2013; Naud et al., 2005; Naud & Havenhand, 2006). Hence, investigating whether such adaptations exist in gastropods will be worthwhile, given that many of the copulatory organs seem morphologically elaborated enough, and neurophysiologically sufficiently agile, to also perform such functions.

As becomes clear from the aforementioned studies, many specific questions still remain to be answered in the study of reproduction of mollusks. In some cases, like cephalopods, serious technical difficulties hinder proper progress in our understanding of the undoubtedly complex control of reproduction in these “advanced invertebrates” (Wells, 1978). The housing and maintenance of cephalopods in captivity present many problems, as do molecular manipulation (see Fiorito et al., 2014). New genomic and transcriptomic approaches (Moroz et al., 2011; Albertin et al., 2012, 2015) may open new windows to understanding some key steps of reproduction in these animals. Among some of the issues that clearly require investigation are the identification of the optic gland hormone, the possible role of the egg-laying hormone, the hormonal role of the gonads, the role of olfaction and chemotactile stimuli, and sex recognition. Although the practical limitations are fewer in gastropods, where experimental manipulation is generally easier, this also holds. Like in many fields of research, also here we expect that the development of genetic tools for molluskan model species—such as RNAi, transgenics, and CRISPR genome editing—in combination with more classical approaches, will provide new opportunities to address the aforementioned questions.

References

Abate, L., Bertolucci, E., Conti, M., Di Cosmo, A., Di Cristo, C., Mettivier, G., … Russo, P. (2000). Quantitative dynamic imaging of biological processes with solid state radiation detectors. IEE Transactions on Nuclear Science, 47, 1907–1910.Find this resource:

Adamo, S. A., & Chase, R. (1990). Dissociation of sexual arousal and sexual proclivity in the garden snail Helix aspersa. Behavioral and Neural Biology, 54, 115–130.Find this resource:

Adamson, K. J., Wang, T., Zhao, M., Bell, F., Kuballa, A. V., Storey, K. B., & Cummins, S. F. (2015). Molecular insights into land snail neuropeptides through transcriptome and comparative gene analysis. BMC Genomics, 16, 308.Find this resource:

Adema, C., Hillier, L. Jones, C., Loker, E., Knight, M., Minx, P., … Wilson, R. (2017). Whole genome analysis of a schistosomiasis-transmitting freshwater snail. Nature Communications, in press.Find this resource:

Albertin, C. B., Bonnaud, L., Brown, C. T., Crookes-Goodson, W. J., da Fonseca, R. R., Di Cristo, C., … Ragsdale, C. W. (2012). Cephalopod genomics: A plan of strategies and organization. Standards in Genomic Sciences, 7, 175–188.Find this resource:

Albertin, C. B., Simakov, O., Mitros, T., Yan Wang, Z., Pungor, J. R., Edsinger-Gonzales, E., … Rokhsar, D. S. (2015). The octopus genome and the evolution of cephalopod neural and morphological novelties. Nature, 524, 220–224.Find this resource:

Alexandrowicz, J. S. (1964). The neurosecretory system of the vena cava in Cephalopoda, I. Eledone cirrosa. Journal of the Marine Biology Association of the United Kingdom, 44, 111–132.Find this resource:

Alexandrowicz, J. S. (1965). The neurosecretory system of the vena cava in Cephalopoda, II. Sepia officinalis and Octopus vulgaris. Journal of the Marine Biology Association of the United Kingdom, 45, 209–228.Find this resource:

Anthes, N. (2010). Mate choice and reproductive conflict in simultaneous hermaphrodites. In P. M. Kappeler (Ed.), Animal behaviour: Evolution and mechanisms (pp. 329–357). Berlin, Germany: Springer Verlag.Find this resource:

Anthes, N., David, P., Auld, J. R., Hoffer, J. N. A., Jarne, P., Koene, J. M., … Schärer, L. (2010). Bateman gradients in hermaphrodites: An extended approach to quantify sexual selection. American Naturalist, 176, 249–263.Find this resource:

Anthes, N., & Michiels, N. K. (2007). Precopulatory stabbing, hypodermic injections and unilateral copulations in a hermaphroditic sea slug. Biology Letters, 3, 121–124.Find this resource:

Anthes, N., Putz, A., & Michiels, N. K. (2006). Hermaphrodite sex role preferences: The role of partner body size, mating history and female fitness in the sea slug Chelidonura sandrana. Behavioral Ecology and Sociology, 60, 359–367.Find this resource:

Arakawa, K. Y. (1962). An ecological account of the breeding behaviour of Octopus luteus. Venus, 22, 176–180.Find this resource:

Arletti, R., Benelli, A., & Bertolini, A. (1990). Oxytocin inhibits food and fluid intake in rats. Physiology and Behavior, 48, 825–830.Find this resource:

Arnold, J. M. (1984). Cephalopods. In A. S. Tompa, N. H. Verdonk, & J. A. M. van den Biggelar (Eds.), The Mollusca, Vol. 7. Reproduction (pp. 419–454). New York, NY: Academic Press.Find this resource:

Arnqvist, G., & Rowe, L. (2005). Sexual conflict. Princeton, NJ: Princeton University Press.Find this resource:

Audesirk, T. E. (1977). Chemoreception in Aplysia californica III. Evidence for pheromones influencing reproductive behaviour. Behavioral Biology, 20, 235–243.Find this resource:

Auld, J. R., & Jarne, P. (2016). Sex and recombination in snails. In R. M. Kliman (Ed.), Encyclopedia of evolutionary biology (Vol. 4, pp. 49–60). Oxford, England: Academic Press.Find this resource:

Ballance, S., Howard, M., White, K. N., McCrohan, C. R. Thornton, D. J., & Sheehan, J. K. (2004). Partial characterisation of high-molecular weight glycoconjugates in the trail mucus of the freshwater pond snail Lymnaea stagnalis. Comparative Biochemistry and Physiology Part B: Biochemistry, 137, 475–486.Find this resource:

Basil, J. A., Hanlon, R. T., Sheikh, S. I., & Atema, J. (2000). Three-dimensional odor tracking by Nautilus pompilius. Journal of Experimental Biology, 203(9), 1409–1414.Find this resource:

Bateman, A. J. (1948). Intra-sexual selection in Drosophila. Heredity, 2, 349–368.Find this resource:

Bekius, R. (1972). The circulatory system of Lymnaea stagnalis (L.). Netherlands Journal of Zoology, 22, 1–58.Find this resource:

Belonoschkin, B. (1929a). Die Geschlechtswege von Octopus vulgaris und ihre Bedeutung fiir die Bewegung der Spermatozoen. Zeitschrift für Zellforschung und mikroskopische Anatomie, 9, 643–662.Find this resource:

Belonoschkin, B. (1929b). Das Verhalten der Spermatozoen zwischen Begattung und Befruchtung bei Octopus vulgaris. Zeitschrift für Zellforschung und mikroskopische Anatomie, 9, 750–753.Find this resource:

Benjamin, P. R., & Peat, A. (1971). On the structure of the pulmonate osphradium. Zeitschrift für Zellforschung und Mikroskopische Anatomie, 118, 168–189.Find this resource:

Bentley, P. J. (2001). Sex hormones in vertebrates. In Encyclopedia of Life Science (pp. 1–5). Hoboken, NJ: Wiley.Find this resource:

Bernay, B., Baudy-Floc’h, M., Zanuttini, B., Zatylny, C., Pouvreau, S., & Henry, J. (2006a). Ovarian and sperm regulatory peptides regulate ovulation in the oyster Crassostrea gigas. Molecular Reproduction and Development, 73, 607–616.Find this resource:

Bernay, B., Baudy-Floc’h, M., Gagnon, J., & Henry, J. (2006b). Ovarian jelly-peptides (OJPs), a new family of regulatory peptides identified in the cephalopod Sepia officinalis. Peptides, 27(6), 1259–1268.Find this resource:

Bernay, B., Gagnon, J., & Henry, J. (2004). Egg capsule secretion in invertebrates: A new ovarian regulatory peptide identified by mass spectrometry comparative screening in Sepia officinalis. Biochemical and Biophysical Research Communications, 314(1), 215–222.Find this resource:

Berry, C. F., & Cottrell, G. A. (1970). Neurosecretion in the vena cava of the cephalopod Eledone cirrosa. Zeitschrift fur Zellforschung und mikroskopische Anatomie, 104, 107–115.Find this resource:

Bianchi, D. (1969). Esperimenti sulla funzione del sistema neurosecretorio della vena cava nei cefalopodi. Bollettino della Societa italiana di biologia sperimentale, 45, 1615–1619.Find this resource:

Bianchi, D., & De Prisco, R. (1971). Esperimenti sulla funzione del sistema neurosecretorio della vena cava nei cefalopodi. III. Purificazione e caratterizazione parziale del principio attivo. Bollettino della Societa italina di biologia sperimentale, 47, 477–480.Find this resource:

Bigot, L., Zatylny-Gaudin, C., Rodetc, F., Bernay, B., Boudryd, P., & Favrel, P. (2012). Characterization of GnRH-related peptides from the Pacific oyster Crassostrea gigas. Peptides, 34(2), 303–310.Find this resource:

Bjorkman, N. (1963). On the ultrastructure of the optic gland in Octopus. Journal of Ultrastructure Research, 8, 195.Find this resource:

Blankenship, J. E., Rock, M. K., Robbins, L. C., Livingston, C. A., & Lehman, H. K. (1983). Aspects of copulatory behavior and peptide control of egg laying in Aplysia. Federation Proceedings, 42, 96–100.Find this resource:

Boer, H. H., Roubos, E. W., Van Dalen, H., & Groesbeek, J. R. F. Th. (1977). Neurosecretion in the basommatophoran snail Bulinus truncatus (Gastropoda, Pulmonata). Cell and Tissue Research, 176, 57–67.Find this resource:

Boer, H. H., Slot, J. W., & Van Andel, J. (1968). Electron microscopical and histochemical observations on the relation between medio-dorsal bodies and neurosecretory cells in the basommatophoran snails Lymnaea stagnalis, Ancylus fluviatilis, Australorbis glabratus and Planorbarius corneus. Zeitschrift fur Zellforschung und mikroskopische Anatomie, 87, 435–450.Find this resource:

Boletzky, S. v. (1983). Sepiola robusta. In P. R. Boyle (Ed.), Cephalopod life cycles (Vol. 1, pp. 53–67). London, England: Academic Press.Find this resource:

Boletzky, S. v. (1987). Fecundity variation in relation to intermittent or chronic spawning in the cuttlefish, Sepia officinalis. (Mollusca, Cephalopoda). Bulletin of Marine Science, 40, 382–338.Find this resource:

Boletzky, S. v. (1988). A new record of long continued spawning of Sepia officinalis (Mollusca: Cephalopoda). Rapport Commission International Mer Méditerranée, 31(2), 257.Find this resource:

Bolognari, A., Carmignai, M. P. A., & Zaccone, G. (1976). A cytochemical analysis of the follicular cells and the yolk in the crowing oocytes of Octopus vulgaris (Cephalopoda, Mollusca). Acta Histochemica et Cytochemica, 55, 167–175.Find this resource:

Bondesen, P. (1950). A comparative morphological-biological analyses of the eggcapsules of freshwater pulmonate gastropods. Acta Jutlandica, 3, 1–208.Find this resource:

Bonichon, A. (1967). Contribution a l’etude de la neurosecretion et de l’endocrinologie chez les Cephalopodes. I. Octopus vulgaris. Vie Milieu, 18, 227–263.Find this resource:

Bottke, W. (1974). The fine structure of the ovarian follicle of Alloteuthis subulata Lam (Mollusca, Cephalopoda). Cell and Tissue Research, 150, 463—479.Find this resource:

Bousfield, J. D., Gomm, J., McCapra, F., & Thomas, J. D. (1980). The molecular characteristics of chemoreception in the snail Biomphalaria glabrata (Say). Journal of Applied Ecology, 17, 631–639.Find this resource:

Bovbjerg, R. V. (1968). Responses to food in lymnaeid snails. Physiological Zoology, 41, 412–423.Find this resource:

Boycott, B. B., & Young, J. Z. (1956). The subpedunculate body and nerve and other organs associated with the optic tract of cephalopods. In K. G. Wingstrand (Ed.), Bertil Hanstrom: Zoological papers in honour of his sixty-fifth birthday (pp. 76–165). Lund, Sweden: Zoological Institute.Find this resource:

Boyle, P. R. (1983). Eledone cirrhosa. In P. R. Boyle (Ed.), Cephalopod life cycles (Vol. 1, pp. 365–386). London, England: Academic Press.Find this resource:

Boyle, P. R. (Ed.). (1987). Cephalopod life cycles. Comparative reviews (Vol. 2). London, England: Academic Press.Find this resource:

Boyle, P. R., & Chevis, D. (1991). Changes in follicle cell epithelium nuclei at the onset of vitellogenesis in the octopus Eledone cirrhosa. Bulletin of Marine Science, 49, 372–378.Find this resource:

Boyle, P. R., & Chevis, D. (1992). Egg development in the octopus Eledone cirrhosa. Journal of Zoology, 227, 623–628.Find this resource:

Boyle, P. R., & Rodhouse, P. G. (2005). Cephalopods: Ecology and fisheries. Oxford, England: Blackwell.Find this resource:

Bright, K., Kellett, E., Saunders, S. E., Brierley, M., Burke, J. F., & Benjamin, P. R. (1993). Mutually exclusive expression of alternatively spliced FMRFamide transcripts in identified neuronal systems of the snail Lymnaea. Journal of Neuroscience, 13, 2719–2729.Find this resource:

Brussaard, A. B., Schluter, N. C. M., Ebberink, R. H. M., Kits, K. S., & Ter Maat, A. (1990). Discharge induction in molluscan peptidergic cells requires a specific set of four autoexcitatory neuropeptides. Neuroscience, 39, 479–491.Find this resource:

Buckley, S. K. L. (1977). Oogenesis and its hormonal control in Octopus vulgaris. Unpublished Ph.D. dissertation, University of Cambridge, Cambridge, England.Find this resource:

Buresch, K. C., Boal, J. G., Nagle, G. T., Knowles, J., Nobuhara, R., Sweeney, K., & Hanlon, R. T. (2004). Experimental evidence that ovary and oviducal gland extracts influence male agonistic behavior in squids. Biological Bulletin, 206(1), 1–3.Find this resource:

Cain, G. L. (1956). Studies on cross-fertilization and self-fertilization in Lymnaea stagnalis appressa Say. Biological Bulletin, 111, 45–52.Find this resource:

Callan, H. G. (1940). The absence of a sex hormone controlling regeneration of the hectocotylus in Octopus vulgaris L. Pubblicazioni della Stazione Zoologica di Napoli, 18, 15–19.Find this resource:

Charnov, E. L. (1979). Simultaneous hermaphroditism and sexual selection. Proceedings of the National Academy of Sciences USA, 76, 2480–2484.Find this resource:

Charnov, E. L. (1982). Sex allocation. Princeton, NJ: Princeton University Press.Find this resource:

Chase, R. (2002). Behavior and its neural control in gastropod molluscs. Oxford, England: Oxford University Press.Find this resource:

Chase, R., & Blanchard, K. C. (2006). The snail’s love-dart delivers mucus to increase paternity. Proceedings of the Royal Society of London, B: Biological Sciences, 273, 1471–1475.Find this resource:

Cigliano, J. A. (1995). Assessment of the mating history of female pygmy octopuses and a possible sperm competition mechanism. Animal Behaviour, 49, 849–851.Find this resource:

Cobbs, J. S., & Pinsker, H. M. (1982b). Role of bag cells in egg deposition of Aplysia brasiliana—II. Contribution of egg movement to elicited behaviors. Journal of Comparative Physiology A, 147, 537–546.Find this resource:

Cobbs, J. S., & Pinsker, H. M. (1982a). Role of bag cells in egg deposition of Aplysia brasiliana—I. Comparison of normal and elicited behaviors Journal of Comparative Physiology A, 147, 523–535.Find this resource:

Cole, W. H. (1925). Egg laying in two species of Planorbis. American Naturalist, 59, 284–286.Find this resource:

Conn, P. J., & Kaczmarek, L. K. (1989). The bag cell neurons of Aplysia: A model for the study of the molecular mechanisms involved in the control of prolonged animal behaviors. Molecular Neurobiology, 3, 237–273.Find this resource:

Corner, B. D., & Moore, H. T. (1980). Field observations on the reproductive behavior of Sepia latimanus. Micronesica, 16, 235–260.Find this resource:

Crisp, M. (1973). Fine structure of some Prosobranch osphradia. Marine Biology, 22, 231–240.Find this resource:

Croll, R. P., & Chiasson, B. J. (1989). Postembryonic development of serotoninlike immunoreactivity in the central nervous system of the snail, Lymnaea stagnalis. Journal of Comparative Neurology, 280, 122–142.Find this resource:

Croll, R. P., & Van Minnen, J. (1992). Distribution of the peptide Ala-Pro-Gly-Trp-NH2 (APGWamide) in the nervous system and periphery of the snail Lymnaea stagnalis as revealed by immunocytochemistry and in situ hybridization. Journal of Comparative Neurology, 324, 567–574.Find this resource:

Croll, R. P., Van Minnen, J., Kits, K. S., & Smid, A. B. (1991). APGWamide: Molecular, histological and physiological examination of the novel neuropeptide involved with reproduction in the snail, Lymnaea stagnalis. In K. S. Kits, H. H. Boer, & J. Joosse (Eds.), Molluscan neurobiology (pp. 248–254). Amsterdam, The Netherlands: North-Holland.Find this resource:

Crown, A., Clifton, D. K., & Steiner, R. A. (2007). Neuropeptide signaling in the integration of metabolism and reproduction. Neuroendocrinology, 86(3), 175–182.Find this resource:

Cummins, S. F., Boal, J. G., Buresch, K. C., Kuanpradit, C., Sobhon, P., Holm, J. B., … Hanlon, R. T. (2011). Extreme aggression in male squid induced by a β-MSP-like pheromone. Current Biology, 21(4), 322–327.Find this resource:

Cummins, S. F., & Degnan, B. M. (2010). Sensory sea slugs—Towards decoding the molecular toolkit required for a mollusc to smell. Communicative and Integrative Biology, 3, 423–426.Find this resource:

Cummins, S. F., Nichols, A. E., Amare, A., Hummon, A. B., Sweedler, J. V., & Nagle, G. T. (2004). Characterization of Aplysia enticin and temptin, two novel water-borne protein pheromones that act in concert with attractin to stimulate mate attraction. Journal of Biological Chemistry, 279, 25614–25622.Find this resource:

Cummins, S. F., Nuurai, P., Nagle, G. T., & Degnan, B. M. (2010). Conservation of the egg-laying hormone neuropeptide and attractin pheromone in the spotted sea hare, Aplysia dactylomela. Peptides, 31, 394–401.Find this resource:

Cummins, S. F., & Wyeth, R. C. (2014). Olfaction in gastropods. In A. Di Cosmo (Ed.), Neuroecology and neuroethology in molluscs: The interface between behaviour and environment (pp. 45–71). Hauppauge, NY: Nova Science Publishers.Find this resource:

Cuomo, A., Di Cristo, C., Paolucci, M., Di Cosmo, A., & Tosti, E. (2005). Calcium currents correlate with oocyte maturation during the reproductive cycle in Octopus vulgaris. Journal of Experimental Zoology, A: Comparative Experimental Biology, 303(3), 193–202.Find this resource:

D’Aniello, A., Di Cosmo, A., Di Cristo, C., Assisi, L., Botte, V., & Di Fiore, M. M. (1996). Occurrence of sex steroid hormones and their binding proteins in Octopus vulgaris lam. Biochemical and Biophysical Research Communications, 227(3), 782–788.Find this resource:

Dalesman, S., Karnik, V., & Lukowiak, K. (2011). Sensory mediation of memory blocking stressors in the pond snail Lymnaea stagnalis. Journal of Experimental Biology, 214, 2528–2533.Find this resource:

Dalesman, S., & Lukowiak, K. (2011). Social snails: The effect of social isolation on cognition is dependent on environmental context. Journal of Experimental Biology, 214, 4179–4185.Find this resource:

Dalesman, S., Rundle, S. D., Coleman, R. A., & Cotton, P. A. (2006). Cue association and antipredator behaviour in a pulmonate snail, Lymnaea stagnalis. Animal Behaviour, 71, 789–797.Find this resource:

Dargaei, Z., Colmers, P. L. W., Hodgson, H. M., & Magoski, N. S. (2014). Electrical coupling between Aplysia bag cell neurons: Characterization and role in synchronous firing. Journal of Neurophysiology, 112, 2680–2696.Find this resource:

Davies, M. S., & Hawkins, J. (1998). Mucus from marine molluscs. Advances in Marine Biology, 34, 1–71.Find this resource:

Davison, A., Frend, H. T., Moray, C., Wheatley, H., Searle, L. J., & Eichhorn, M. P. (2009). Mating behaviour in Lymnaea stagnalis pond snails is a maternally inherited, lateralised trait. Biology Letters, 5, 20–22.Find this resource:

De Boer, P. A. C. M., Jansen, R. F., Koene, J. M., & Ter Maat, A. (1997). Nervous control of male sexual drive in the hermaphroditic snail Lymnaea stagnalis. Journal of Experimental Biology, 200, 941–951.Find this resource:

De Boer, P. A. C. M., Jansen, R. F., & Ter Maat, A. (1996). Copulation in the hermaphrodite snail Lymnaea stagnalis: A review. Invertebrate Reproduction and Development, 30, 167–176.Find this resource:

De Boer, P. A. C. M., Jansen, R. F., Ter Maat, A., Van Straalen, N. M., & Koene, J. M. (2010). The distinction between retractor and protractor muscles of the freshwater snail’s male organ has no physiological basis. Journal of Experimental Biology, 213, 40–44.Find this resource:

De Jong-Brink, M. (1969). Histochemical and electron microscope observations on the reproductive tract of Biomphalaria glabrata (Australorbis glabratus), intermediate host of Schistosoma mansoni. Zeitschrift für Zellforschung und mikroskopische Anatomie, 102, 507–542.Find this resource:

De Jong-Brink, M., Boer, H. H., Hommes, T. G., & Kodde, A. (1977). Spermatogenesis and the role of Sertoli cells in the freshwater snail Biomphalaria glabrata. Cell and Tissue Research, 181, 37–58.Find this resource:

De Jong-Brink, M., Reid, C. N., Tensen, C. P., & ter Maat, A. (1999). Parasites flicking the NPY gene on the host’s switchboard: Why NPY? FASEB Journal, 13(14), 1972–1984.Find this resource:

De Jong-Brink, M., ter Maat, A., & Tensen, C. P. (2001). NPY in invertebrates: Molecular answers to altered functions during evolution. Peptides, 22(3), 309–315.Find this resource:

De Lange, R. P. J., De Boer, P. A. C. M., Ter Maat, A., Tensen, C. P., & Van Minnen, J. (1998b). Transmitter identification in neurons involved in male copulation behavior in Lymnaea stagnalis. Journal of Comparative Neurology, 395, 440–449.Find this resource:

De Lange, R. P. J., Joosse, J., & Van Minnen, J. (1998a). Multi-messenger innervation of the male sexual system of Lymnaea stagnalis. Journal of Comparative Neurology, 390, 564–577.Find this resource:

De Lange, R. P. J., Van Golen, F. A., & Van Minnen, J. (1997). Diversity in cell specific co-expression of four neuropeptide genes involved in control of male copulation behaviour in Lymnaea stagnalis. Neuroscience, 78, 289–299.Find this resource:

De Lange, R. P. J., & Van Minnen, J. (1998). Localization of the neuropeptide APGWamide in gastropod molluscs by in situ hybridization and immunocytochemistry. General and Comparative Endocrinology, 109(2), 166–174.Find this resource:

De Lisa, E., Carella, F., De Vico, G., & Di Cosmo, A. (2013). The gonadotropin releasing hormone (GnRH)-like molecule in prosobranch Patella caerulea: Potential biomarker of endocrine-disrupting compounds in marine environments. Zoological Science, 30, 135–140.Find this resource:

De Lisa, E., Paolucci, M., & Di Cosmo, A. (2012). Conservative nature of oestradiol signalling pathways in the brain lobes of Octopus vulgaris involved in reproduction, learning and motor coordination. Journal of Neuroendocrinology, 24(2), 275–284.Find this resource:

De Visser, J. A. G. M., Ter Maat, A., & Zonneveld, C. (1994). Energy budgets and reproductive allocation in the simultaneous hermaphrodite pond snail, Lymnaea stagnalis (L.): A trade-off between male and female function. American Naturalist, 144, 861–867.Find this resource:

De Vlieger, T. A., Kits, K. S., Ter Maat, A., & Lodder, J. C. (1980). Morphology and electrophysiology of the ovulation hormone producing neuro-endocrine cells of the freshwater snail Lymnaea stagnalis (L.). Journal of Experimental Biology, 84, 239–271.Find this resource:

Defretin, R., & Richard, A. (1967). Ultrastructure de la glande optique de Sepia officinalis L. (Mollusques; Céphalopode). Mise en evidence de la sécrétion et de son controle photopériodique. Comptes Rendus de l’Académie des Sciences, 265, 1415–1418.Find this resource:

Delle Chiaie, S. (1828). Memorie sulla storia e notomia degli animali senza vertebre del Regno di Napoli (Vol. 3, pp. 1–232). Napoli, Italy: Società Tipografica.Find this resource:

DeWitt, T. J. (1996). Gender contests in a simultaneous hermaphrodite snail: A size-advantage model for behaviour. Animal Behavior, 51, 345–351.Find this resource:

Dewsbury, D. A. (1982). Ejaculate cost and male choice. American Naturalist, 119, 601–610.Find this resource:

Dhainaut, A., & Richard, A. (1976). Vitellogenese chez les céphalopodes decapodes, évolution de l’ovocyte et des cellules folliculaires au cours de la maturation génitale. Archives of Microbiology, 65, 183–208.Find this resource:

Di Cosmo, A., & Di Cristo, C. (1998). Neuropeptidergic control of the optic gland of Octopus vulgaris: FMRF-amide and GnRH immunoreactivity. Journal of Comparative Neurology, 398(1), 1–12.Find this resource:

Di Cosmo, A., Di Cristo, C., & Paolucci, M. (2001). Sex steroid hormone fluctuations and morphological changes of the reproductive system of the female of Octopus vulgaris throughout the Annual Cycle. Journal of Experimental Zoology, 289(1), 33–47.Find this resource:

Di Cosmo, A., Di Cristo, C., & Paolucci, M. (2002). A estradiol-17beta receptor in the reproductive system of the female of Octopus vulgaris: Characterization and immunolocalization. Molecular Reproduction and Development, 61(3), 367–375.Find this resource:

Di Cosmo, A., Paolucci, M. & Di Cristo, C. (2004). N-methyl-D-aspartate receptor-like immunoreactivity in the brain of Sepia and Octopus. Journal of Comparative Neurology, 477(2), 202–219.Find this resource:

Di Cosmo, A., Paolucci, M., Di Cristo, C., Botte, V., & Ciarcia, G. (1998). Progesterone receptor in the reproductive system of the female of Octopus vulgaris: Characterization and immunolocalization. Molecular Reproduction and Development, 50(4), 451–460.Find this resource:

Di Cristo, C. (2013). Nervous control of reproduction in Octopus vulgaris: A new model. Invertebrate Neuroscience, 13(1), 27–34.Find this resource:

Di Cristo, C., De Lisa, E., & Di Cosmo, A. (2009). GnRH in the brain and ovary of Sepia officinalis. Peptides, 30(3), 531–537.Find this resource:

Di Cristo, C., Delli Bovi, P., & Di Cosmo, A. (2003). Role of FMRFamide in the reproduction of Octopus vulgaris: Molecular analysis and effect on visual input. Peptides, 24(10), 1525–1532.Find this resource:

Di Cristo, C., Di Donato, P., Palumbo, A., d’Ischia, M., Paolucci, M., & Di Cosmo, A. (2010). Steroidogenesis in the brain of Sepia officinalis and Octopus vulgaris. Frontiers in Bioscience (Elite edition), 2, 673–683.Find this resource:

Di Cristo, C., Paolucci, M., & Di Cosmo, A. (2008). Progesterone affects vitellogenesis in Octopus vulgaris. Open Zoology Journal, 1, 29–36.Find this resource:

Di Cristo, C., Paolucci, M., Iglesias, J., Sanchez, J., & Di Cosmo A. (2002). Presence of two neuropeptides in the fusiform ganglion and reproductive ducts of Octopus vulgaris: FMRFamide and gonadotropin-releasing hormone (GnRH). Journal of Experimental Zoology, 292(3), 267–276.Find this resource:

Di Cristo, C., Van Minnen, J., Di Cosmo, A. (2005). The presence of APGWamide in Octopus vulgaris: A possible role in the reproductive behavior. Peptides, 26(1), 53–62.Find this resource:

Dogterom, G. E., & Van Loenhout, H. (1983). Specificity of ovulation hormones of some basommatophoran species studied by means of iso- and heterospecific injections. General and Comparative Endocrinology, 52, 121–125.Find this resource:

Dogterom, G. E., van Loenhout, H., Koomen, W., Roubos, E. W., & Geraerts, W. P. (1984). Ovulation hormone, nutritive state, and female reproductive activity in Lymnaea stagnalis. General and Comparative Endocrinology, 55, 29–35.Find this resource:

Drew, G. A. (1911). Sexual activities of the squid Loligo pealii(Les). I. Copulation. Egg-laying and fertilization. Journal of Morphology, 22, 327–359.Find this resource:

Durchon, M., & Richard, A. (1967). Étude en culture organotypique du role endocrine de la glandule optique dans la maturation ovarienne chez Sepia officinalís L. (Mollusque: Céphalopode). Comptes Rendus de l’Académie des Sciences, 264, 1497–1500.Find this resource:

Eberhardt, B., & Wabnitz, R. W. (1979). Morphological identification and functional analysis of central neurons innervating the penis retractor muscle of Helix pomatia. Comparative Biochemistry and Physiology, 63A, 599–613.Find this resource:

El Filali, Z., De Boer, P. A. C. M., Pieneman, A. W., De Lange, R. P. J., Jansen, R. F., Ter Maat, A., … Koene, J. M. (2015). Single-cell analysis of peptide expression and electrophysiology of right parietal neurons involved in male copulation behavior of a simultaneous hermaphrodite. Invertebrate Neuroscience 15, 7.Find this resource:

El Filali, Z., Hornshaw, M., Smit, A. B., & Li, K. W. (2003). Retrograde labeling of single neurons in conjunction with MALDI high-energy collision-induced dissociation MS/MS analysis for peptide profiling and structural characterization. Analytical Chemistry, 75, 2996–3000.Find this resource:

Elekes, K., Hernádi, L., & Kemenes, G. (1988). Serotonin immunoreactive neurons in the CNS of Helix and Lymnaea. In J. Salánki & K. S. -Rózsa (Eds.), Neurobiology of invertebrates, transmitters, modulators and receptors (pp. 703–711). Budapest, Hungary: Akademiai Kiad.Find this resource:

Emery, D. G. (1992). Fine-structure of olfactory epithelia of gastropod molluscs. Microscopy Research and Techniques, 22, 307–324.Find this resource:

Enault, J., Zatylny-Gaudin, C., Bernay, B., Lefranc, B., Leprince, J., Baudy-Floc’h, M., & Henry, J. (2012). A complex set of sex pheromones identified in the cuttlefish Sepia officinalis. PLoS ONE, 7, e46531.Find this resource:

Escobar, J. S., Auld, J. R., Correa, A. C., Alonso, J. M., Bony, Y. K., Coutellec, M-A., … David, P. (2011). Patterns of mating-system evolution in hermaphroditic animals: Correlations among selfing rate, inbreeding depression and the timing of reproduction. Evolution, 65, 1233–1253.Find this resource:

Facon, B., Ravigné, V., & Goudet, J. (2006). Experimental evidence of inbreeding avoidance in the hermaphroditic snail Physa acuta. Evolutionary Ecology, 20, 395–406.Find this resource:

Ferguson, G. P., Pieneman, A. W., Jansen, R. T., & Ter Maat, A. (1993). Neuronal feedback in egg-laying behaviour of the pond snail Lymnaea stagnalis. Journal of Experimental Biology, 178, 251–259.Find this resource:

Ferguson, G. P., Ter Maat, A., Parsons, D. W., & Pinsker, H. M. (1989). Egg laying in Aplysia: I. Behavioral patterns and muscle activity of freely behaving animals after selectively elicited bag cell discharges. Journal of Comparative Physiology A, 164, 835–847.Find this resource:

Fields, W. G., & Thompson, K. A. (1976). Ultrastructure and functional morphology of spermatozoa of Rossia pacifica (Cephalopoda: Decapoda). Canadian Journal of Zoology, 54, 908–932.Find this resource:

Fiorito, G., Affuso, A., Anderson, D. B., Basil, J., Bonnaud, L., Botta, G., … Andrews, P. (2014). Cephalopods in neuroscience: Regulations, research and the 3Rs. Invertebrate Neuroscience, 14, 13–36.Find this resource:

Fong, P. P., Olex, A. L., Farrell, J. E., Majchrzak, R. M., & Muschamp, J. W. (2005). Induction of preputium eversion by peptides, serotonin receptor antagonists, and selective serotonin reuptake inhibitors in Biomphalaria glabrata. Invertebrate Biology, 124, 296–302.Find this resource:

Franklin, A. M., Squires, Z. E., & Stuart-Fox, D. (2012). The energetic cost of mating in a promiscuous cephalopod. Biology Letters, 8, 754–756.Find this resource:

Franzen, A. (1955). Comparative morphological investigations into the spermiogenesis among Mollusca. Zoologiska bidrag från Uppsala, 30, 399–456.Find this resource:

Franzen, A. (1956). On spermiogenesis, morphology of the spermatozoa and biology of fertilization among invertebrates. Zoologiska bidrag från Uppsala, 31, 355–482.Find this resource:

Franzen, A. (1967). Spermiogenesis and spermatozoa of the Cephalopoda. Arkiv för Zoologi, 19, 323–334.Find this resource:

Froesch, D. (1974). The subpedunculate lobe of the octopus brain: Evidence for dual function. Brain Research, 75(2), 277–285.Find this resource:

Froesch, D., & Marthy, H. J. (1975). The structure and function of the oviducal gland in octopods (Cephalopoda). Proceedings of the Royal Society of London, B: Biological Sciences, 188, 95–101.Find this resource:

Geoffroy E., Hutcheson, R., & Chase, R. (2005). Nervous control of ovulation and ejaculation in Helix aspersa. Journal of Molluscan Studies 71, 393–399.Find this resource:

Geraerts, W. P. M., & Algera, L. H. (1976). The stimulating effect of the dorsal body hormone on cell differentiation in the female accessory sex organs of the hermaphrodite freshwater snail, Lymnaea stagnalis. General and Comparative Endocrinology, 29, 109–118.Find this resource:

Geraerts, W. P. M. & Bohlken, S. (1976). The control of ovulation in the hermaphrodite freshwater snail Lymnaea stagnalis by the neurohormone of the caudo-dorsal cells. General and Comparative Endocrinology, 28, 350–357.Find this resource:

Geraerts, W. P. M., & Hogenes, T. M. (1985). Heterogeneity of peptides released by electrically active neuroendocrine caudodorsal cells of Lymnaea stagnalis. Brain Research, 331, 51–61.Find this resource:

Geraerts, W. P. M., & Joosse, J. (1975). Control of vitellogenesis and of growth of female accessory sex organs by the dorsal body hormone (DBH) in the hermaphroditic freshwater snail Lymnaea stagnalis. General and Comparative Endocrinology, 27, 450–464.Find this resource:

Geraerts, W. P. M., Tensen, C., & Hogenes, T. M. (1983). Multiple release of peptides by electrically active neurosecretory caudo-dorsal cells of Lymnaea stagnalis. Neuroscience Letters, 41, 151–155.Find this resource:

Geraerts, W. P. M., Vreugdenhil, E., Ebberink, R. H. M., & Hogenes, T. M. (1985). Synthesis of multiple peptides from a larger precursor in the neuroendocrine caudo-dorsal cells of Lymnaea stagnalis. Neuroscience Letters, 56, 241–246.Find this resource:

Goldberg, J. I., Garofalo, R., Price, C. J., & Chang, J. P. (1993). Presence and biological activity of a GnRH-like factor in the nervous system of Helisoma trivolvis. Journal of Comparative Neurology, 336(4), 571–582.Find this resource:

Goldschmeding, J. T., Wilbrink, M., & Ter Maat, A. (1983). The role of the ovulation hormone in the control of egg-laying in Lymnaea stagnalis. In J. Lever & H. H. Boer (Eds.), Molluscan neuro-endocrinology (pp. 251–255). Amsterdam, The Netherlands: North-Holland.Find this resource:

Gorbman, A., Whiteley, A., & Kavanaugh, S. (2003). Pheromonal stimulation of spawning release of gametes by gonadotropin releasing hormone in the chiton, Mopalia sp. General and Comparative Endocrinology, 131(1), 62–65.Find this resource:

Graziadei, P. (1964). Electron microscopy of some primary receptors in the sucker of Octopus vulgaris. Zeitschrift für Zellforschung und mikroskopische Anatomie, 64, 510–522.Find this resource:

Häderer, I. K., Werminghausen, J., Michiels, N. K., Timmermeyer, N., & Anthes, N. (2009). No effect of mate novelty on sexual motivation in the freshwater snail Biomphalaria glabrata. Frontiers in Zoology, 6, 23.Find this resource:

Hanlon, R. T., Ament, S. A., & Gabr, H. (1999). Behavioural aspects of sperm competition in cuttlefish, Sepia officinalis (Sepioidea: Cephalopoda). Marine Biology, 134, 719–728.Find this resource:

Hanlon, R. T., Maxwell, M. R., & Shashar, N. (1997). Behavioral dynamics that would lead to multiple paternity within egg capsules of the squid Loligo pealeii. Biological Bulletin, 193, 212–214.Find this resource:

Hanlon, R. T., & Messenger, J. B. (1988). Adaptive coloration in young cuttlefish (Sepia officinalis L.): The morphology and development of body patterns and their relation to behaviour. Philosophical Transactions of the Royal Society of London, B: Biological Sciences, 320, 437–487.Find this resource:

Hanlon, R., & Messenger, J. (1996). Cephalopod behaviour. Cambridge, England: Cambridge University Press.Find this resource:

Harman, R. F., Young, R. E., Reid, S. B., Mangold, K. M., Suzuki, T., & Hixon, R. F. (1989). Evidence for multiple spawning in the tropical oceanic squid Stenoteuthis oualaniensis (Teuthoidea: Ommastrephidae). Marine Biology, 101, 513–519.Find this resource:

Haszprunar, G. (1987). The fine morphology of the osphradial sense organs of the Mollusca. III. Placophora and Bivalvia. Philosophical Transactions of the Royal Society of London, B: Biological Sciences, 315, 37–61.Find this resource:

Hathaway, J. J. M., Adema, C. M., Stout, B. A., Mobarak, C. D., & Loker, E. S. (2010). Identification of protein components of egg masses indicates parental investment in immunoprotection of offspring by Biomphalaria glabrata (Gastropoda, Mollusca). Developmental and Comparative Immunology, 34, 425–435.Find this resource:

Healy, J. M. (1989). Spermatozoa of the deep-sea cephalopod Vampyroteuthis infernalis Chun: Ultrastructure and possible evolutionary significance. Philosophical Transactions of the Royal Society of London, B: Biological Sciences, 323, 589–608.Find this resource:

Healy, J. M. (1990). Ultrastructure of spermatozoa in Spirula spirula (L.): Systematic importance and comparison with other cephalopods. Helgolander Meeresuntershungen, 44, 109–123.Find this resource:

Healy, J. M. (1993). Sperm and spermiogenesis in Opisthoteuthis persephone (Octopoda: Cirrata): Ultrastructure, comparison with other cephalopods and evolutionary significance. Journal of Molluscan Studies, 59, 105–115.Find this resource:

Heller, J. (1993). Hermaphroditism in molluscs. Biological Journal of the Linnean Society, 48, 19–42.Find this resource:

Henry, J., Cornet. V., Bernay, B., & Zatylny-Gaudin, C. (2013). Identification and expression of two oxytocin/vasopressin-related peptides in the cuttlefish Sepia officinalis. Peptides, 46, 159–166.Find this resource:

Henry, J., Favrel, P., & Boucaud-Camou, E. (1997). Isolation and identification of a novel Ala-Pro-Gly-Trp-amide-related peptide inhibiting the motility of the mature oviduct in the cuttlefish, Sepia officinalis. Peptides, 18, 1469–1474.Find this resource:

Hermann, P. M., De Lange, R. P. J., Pieneman, A. W., Ter Maat, A., & Jansen, R. F. (1997). Role of neuropeptides encoded on CDCH-1 gene in the organization of egg-laying behavior in the pond snail, Lymnaea stagnalis. Journal of Neurophysiology, 78, 2859–2869.Find this resource:

Hermann, P. M., Genereux, B., & Wildering, W. C. (2009). Evidence for age-dependent mating strategies in the simultaneous hermaphrodite snail, Lymnaea stagnalis (L.). Journal of Experimental Biology, 212, 3164–3173.Find this resource:

Hermann, P. M., Ter Maat, A., & Jansen, R. F. (1994). The neural control of egg-laying behavior in the pond snail Lymnaea stagnalis: motor control of shell turning, Journal of Experimental Biology, 197, 79–99.Find this resource:

Hetherington, M. S., McKenzie, J. D., Dean, H. G., & Winlow, W. (1994). A quantitative analysis of the biogenic amines in the central ganglia of the pond snail, Lymnaea stagnalis. Comparative Biochemistry and Physiology, 107C, 83–93.Find this resource:

Hodgkin, A. L., & Huxley, A. F. (1952a). Currents carried by sodium and potassium ions through the membrane of the giant axon of Loligo. Journal of Physiology, 116, 449–472.Find this resource:

Hodgkin, A. L., & Huxley, A. F. (1952b). The components of membrane conductance in the giant axon of Loligo. Journal of Physiology, 116, 473–496.Find this resource:

Hodgkin, A. L., & Huxley, A. F. (1952c). The dual effect of membrane potential on sodium conductance in the giant axon of Loligo. Journal of Physiology, 116, 497–506.Find this resource:

Hodgkin, A. L., & Huxley, A. F. (1952d). A quantitative description of membrane current and its application to conduction and excitation in nerve. Journal of Physiology, 117, 500–544.Find this resource:

Hodgkin, A. L., Huxley, A. F., & Katz, B. (1952). Measurement of current-voltage relations in the membrane of the giant axon of Loligo. Journal of Physiology, 116, 424–448.Find this resource:

Hoffer, J. N. A., Ellers, J., & Koene, J. M. (2010). Costs of receipt and donation of ejaculates in a simultaneous hermaphrodite. BMC Evolutionary Biology, 10, 393.Find this resource:

Hoffer, J. N. A., Schwegler, D., Ellers, J., & Koene, J. M. (2012). Mating rate influences female reproductive investment in a simultaneous hermaphrodite, Lymnaea stagnalis. Animal Behaviour, 84, 523–529.Find this resource:

Hoving, H. J. T., Roeleveld, M. A. C., Lipinski M. R., & Videler J. J. (2006). Nidamental glands in males of the oceanic squid Ancistrocheirus lesueurii (Cephalopoda: Ancistrocheiridae)—sex change or intersexuality? Journal of Zoology, 269, 341–348.Find this resource:

Ikeda, Y., Sakurai, Y., & Shimazaki, K. (1993). Fertilizing capacity of squid (Todarodes pacificus) spermatozoa collected from various sperm storage sites, with special reference to the role of gelatinous substance from oviducal gland in fertilization and embryonic development. Invertebrate Reproduction and Development, 23, 39–44.Find this resource:

Ito, E., Kojima, S., Lukowiak, K., & Sakakibara, M. (2013). From likes to dislikes: Conditioned taste aversion in the great pond snail (Lymnaea stagnalis). Canadian Journal of Zoology, 91, 405–412.Find this resource:

Iwakoshi, E., Takuwa-Kuroda, K., Fujisawa, Y., Hisada, M., Ukena, K., Tsutsui, K., & Minakata, H. (2002). Isolation and characterization of a GnRH-like peptide from Octopus vulgaris. Biochemical and Biophysical Research Communications, 291(5), 1187–1193.Find this resource:

Iwakoshi-Ukena, E., Ukena, K., Takuwa-Kuroda, K., Kanda, A., Tsutsui, K., & Minakata, H. (2004). Expression and distribution of octopus gonadotropin-releasing hormone in the central nervous system and peripheral organs of the octopus (Octopus vulgaris) by in situ hybridization and immunohistochemistry. Journal of Comparative Neurology, 477(3), 310–323.Find this resource:

Janse, C., Ter Maat, A., & Pieneman, A. W. (1990). Molluscan ovulation hormone containing neurons and age-related reproductive decline. Neurobiology of Aging, 11, 457–463.Find this resource:

Jansen R. F., & Ter Maat, A. (1985). Ring neuron control of columellar motor neurons during egg-laying behavior in the pond snail Lymnaea stagnalis. Journal of Neurobiology, 16, 1–14.Find this resource:

Jarne, P., David, P., Pointier, J-P., & Koene, J. M. (2010). Basommatophoran gastropods. In A. Córdoba-Aguilar & J. L. Leonard (Eds.), The evolution of primary sexual characters in animals (pp. 173–196). New York, NY: Oxford University Press.Find this resource:

Jiménez, C. R., Ter Maat, A., Pieneman, A., Burlingame, A. L., Smit, A. B., & Li, K. W. (2004). Spatio-temporal dynamics of the egg-laying-inducing peptides during an egg-laying cycle: A semi-quantitative matrix-assisted laser desorption/ionization mass spectrometry approach. Journal of Neurochemistry, 89, 865–875.Find this resource:

Joosse, J. (1964). Dorsal bodies and dorsal neurosecretory cells of the cerebral ganglia of Lymnaea stagnalis. Archives Néerlandaises de Zoologie, 16, 1–103.Find this resource:

Jordaens, K., Dillen, L., & Backeljau, T. (2007). Effects of mating, breeding system and parasites on reproduction in hermaphrodites: Pulmonate gastropods (Mollusca). Animal Biology, 57, 137–195.Find this resource:

Kamardin, N. N. (1983). Investigation of the homing behaviour of the lung snail Siponoria grisea L. Vestnik Leningradskovo Universiteta, 15, 101–104.Find this resource:

Kamardin, N. N. (1988). Le rôle probable de l’osphradium dans le homing des mollusques marins littoraux Acanthopleura gemmata Blainv. (Polyplacophora), Siphonaria grisea L. et Siphonaria sp. (Gastropoda, Pulmonata). Mesogèe, 48, 125–130.Find this resource:

Kamardin, N. N. (1995). The electrical responses of osphradial nerve and central neurons to chemical stimulation of Lymnaea osphradium. Acta Biologica Hungarica, 46, 315–320.Find this resource:

Kamardin, N. N., Shalanki, Y., Rozha, K. S., & Nozdrachev, A. D. (2001). Studies of chemoreceptor perception in mollusks. Neuroscience and Behavioral Physiology, 31, 227–235.Find this resource:

Kamardin, N. N., Szűcs, A., & Rosza, K. S. (1998). Distinct responses of osphradial neurons to chemical stimuli and neurotransmitters in Lymnaea stagnalis L. Cellular and Molecular Neurobiology, 19, 235–247.Find this resource:

Kanda, A., Satake, H., Kawada, T., & Minakata, H. (2005). Novel evolutionary lineages of the invertebrate oxytocin/sasopressin superfamily peptides and their receptors in the common octopus (Octopus vulgaris). Biochemical Journal, 387, 85–91.Find this resource:

Kanda, A., Takahashi, T., Satake, H., & Minakata, H. (2006). Molecular and functional characterization of a novel gonadotropin-releasing-hormone receptor isolated from the common octopus (Octopus vulgaris). Biochemical Journal, 395(1), 125–135.Find this resource:

Karnik, V., Braun, M., Dalesman, S., & Lukowiak, K. (2012). Sensory input from the osphradium modulates the response to memory-enhancing stressors in Lymnaea stagnalis. Journal of Experimental Biology, 215, 536–542.Find this resource:

Kawada, T., Kanda, A., Minakata, H., Matsushima, O., & Stake, H. (2004). Identification of a novel receptor for an invertebrate oxytocin/vasopressin superfamily peptide: Molecular and functional evolution of the oxytocin/vasopressin superfamily. Biochemical Journal, 382, 231–237.Find this resource:

Keay, J., Bridgham, J. T., & Thornton, J. W. (2006). The Octopus vulgaris estrogen receptor is a constitutive transcriptional activator: Evolutionary and functional implications. Endocrinology, 147(8), 3861–3869.Find this resource:

Kemenes, G., Elliott, C. J. H., & Benjamin, P. R. (1986). Chemical and tactile inputs to the Lymnaea feeding system: Effects on behaviour and neural circuitry. Journal of Experimental Biology, 122, 113–137.Find this resource:

Kemenes, G. Y., Elekes, K., Hiripi, L., & Benjamin, P. R. (1989). A comparison of four techniques for mapping the distribution of serotonin and serotonin-containing neurons in fixed and living ganglia of the snail, Lymnaea. Journal of Neurocytology, 18, 193–208.Find this resource:

Kimura, K., Chiba, S., & Koene, J. M. (2014). Common effect of the mucus transferred during mating in two dart-shooting snail species from different families. Journal of Experimental Biology, 217, 1150–1153.Find this resource:

Kits, K. S. (1980). States of excitability in ovulation hormone producing neuroendocrine cells of Lymnaea stagnalis (Gastropoda) and their relation to the-egg-laying cycle. Journal of Neurobiology, 11, 397–410.Find this resource:

Knipe, J. H., & Beeman, R. D. (1978). Histological observation on oogenesis in Loligo opalescens Berry. California Department of Fisheries and Game. Fisheries Bulletin, 169, 23–33.Find this resource:

Knott, K. E., Puurtinen, M., & Kaitala, V. (2003). Primers for nine microsatellite loci in the hermaphroditic snail Lymnaea stagnalis. Molecular Ecology Notes, 3, 333–335.Find this resource:

Koene, J. M. (2010). Neuro-endocrine control of reproduction in hermaphroditic freshwater snails: Mechanisms and evolution. Frontiers in Behavioral Neuroscience, 4, 167.Find this resource:

Koene, J. M. (2016). The physiology of pre- and post-copulatory sexual selection in simultaneously hermaphroditic freshwater snails. In A. S. M. Saleuddin & S. T. Mukai (Eds.), Physiology of molluscs (pp. 271–310). Apple Academic Press Inc., Waretown (NJ, USA)/ Oakville (ON, Canada).Find this resource:

Koene, J. M., Brouwer, A., & Hoffer, J. N. A. (2009a). Reduced egg laying caused by a male accessory gland product opens the possibility for sexual conflict in a simultaneous hermaphrodite. Animal Biology, 59, 435–448.Find this resource:

Koene, J. M. & Chase, R. (1998a). The love dart of Helix aspersa Müller is not a gift of calcium. Journal of Molluscan Studies, 64, 75–80.Find this resource:

Koene, J. M., Jansen, R. F., Ter Maat, A., & Chase, R. (2000). A conserved location for the central nervous system control of mating behaviour in gastropod molluscs: Evidence from a terrestrial snail. Journal of Experimental Biology, 203, 1071–1080.Find this resource:

Koene, J. M., Loose, M. J., & Wolters, L. (2008). Mate choice is not affected by mating history in the simultaneously hermaphroditic snail, Lymnaea stagnalis. Journal of Molluscan Studies, 74, 331–335.Find this resource:

Koene, J. M., Montagne-Wajer, K., Roelofs, D., & Ter Maat, A. (2009). The fate of received sperm in the reproductive tract of a hermaphroditic snail and its implications for fertilisation. Evolutionary Ecology, 23, 533–543.Find this resource:

Koene, J. M., Montagne-Wajer, K., & Ter Maat, A. (2006). Effects of frequent mating on sex allocation in the simultaneously hermaphroditic great pond snail. Behavioral Ecology and Sociobiology, 60, 332–338.Find this resource:

Koene, J. M., Montagne-Wajer, K., & Ter Maat, A. (2007). Aspects of body size and mate choice in the simultaneously hermaphroditic pond snail. Lymnaea stagnalis. Animal Biology, 57, 247–259.Find this resource:

Koene, J. M., Sloot, W., Montagne-Wajer, K., Cummins, S. F., Degnan, B. M., Smith, J. S., … Ter Maat, A. (2010). Male accessory gland protein reduces egg laying in a simultaneous hermaphrodite. PLoS ONE, 5, e10117.Find this resource:

Koene, J. M., & Ter Maat, A. (2001). “Allohormones”: A class of bioactive substances favoured by sexual selection. Journal of Comparative Physiology A, 187, 323–326.Find this resource:

Koene, J. M., & Ter Maat, A. (2005). Sex role alternation in the simultaneously hermaphroditic pond snail Lymnaea stagnalis is determined by the availability of seminal fluid. Animal Behaviour, 69, 845–850.Find this resource:

Koene, J. M., & Ter Maat, A. (2007). Coolidge effect in pond snails: Male motivation in a simultaneous hermaphrodite. BMC Evolutionary Biology, 7, 212.Find this resource:

Kohler, H. R., Kloas, W., Schirling, M., Lutz, I., Reye, A. L., Langen, J. S., … Schönfelder, G. (2007). Sex steroid receptor evolution and signalling in aquatic invertebrates. Ecotoxicology, 16(1), 131–143.Find this resource:

Kuanpradit, C., Stewart, M. J., York, P. S., Degnan, B. M., Sobhon, P., Hanna, P. J., … Cummins, S. F. (2012). Characterization of mucus-associated proteins from abalone (Haliotis)—candidates for chemical signaling. FEBS Journal, 279, 437–450.Find this resource:

Kupfermann, I. (1967). Stimulation of egg laying: Possible neuronedocrine function of bag cells of the abdominal ganglion of Aplysia californica. Nature, 216, 814–815.Find this resource:

Lafont, R., & Mathieu, M. (2007). Steroids in aquatic invertebrates. Ecotoxicology, 16(1), 109–130.Find this resource:

Landolfa, M. A., Green, D. M., & Chase, R. (2001). Dart shooting influences paternal reproductive success in the snail Helix aspersa (Pulmonata, Stylommatophora). Behavioral Ecology, 12, 773–777.Find this resource:

Lange, R., Werminghausen, J., & Anthes, N. (2014). Cephalo-traumatic secretion transfer in a hermaphrodite sea slug. Proceedings of the Royal Society of London, B: Biological Sciences, 281, 20132424.Find this resource:

Lankester, E. R. (1875). Observations on the development of the Cephalopoda. Quarterly Journal of Microscopical Science, 15, 37–47.Find this resource:

Laubier-Bonichon, A. (1973). Arguments experimentaux sur l’activite des cellules secretrices du lobe visceral d’un mollusque Cephalopode Octopus vulgaris. Comptes rendus de l’Académie des sciences, D, 276, 1593–1596.Find this resource:

Le Gall, S., FÈral, C., Van Minnen, J., & Marchand, C. R. (1988). Evidence for peptidergic innervation of the endocrine optic gland in Sepia by neurons showing FMRFamide-like immunoreactivity. Brain Research, 462(1), 83–88.Find this resource:

Lever, J., De Vries, C. M., & Jager, J. C. (1965). On the anatomy of the central nervous system and location of neurosecretory cells in Australorbis glabratus. Malacologia, 2, 219–230.Find this resource:

Levy, M., Blumberg, S., & Susswein, A. J. (1997). The rhinophores sense pheromones regulating multiple behaviors in Aplysia fasciata. Neuroscience Letters, 225, 113–116.Find this resource:

Li, G., & Chase, R. (1995). Correlation of axon projections and peptide immunoreactivity in mesocerebral neurons of the snail Helix aspersa, Journal of Comparative Neurology, 353, 9–17.Find this resource:

Li, K. W., Jimenez, C. R., Van Veelen, P., & Geraerts, W. P. M. (1999). Processing and targeting of a molluscan egg-laying peptide prohormone as revealed by mass spectrometric peptide fingerprinting and peptide sequencing. Endocrinology, 134, 1812–1819.Find this resource:

Li, K. W., Smit, A. B., & Geraerts, W. P. M. (1992). Structural and functional characterization of neuropeptides involved in the control of male mating behaviour of Lymnaea stagnalis. Peptides, 13, 633–638.Find this resource:

Li, K. W., Van Golen, F. A., Van Minnen, J., Van Veelen, P. A., Van der Greef, J., & Geraerts, W. P. M. (1994). Structural identification, neuronal synthesis, and role in male copulation of myomodulin-A of Lymnaea: A study involving direct peptide profiling of nervous tissue by mass spectrometry. Brain Research Molecular Brain Research, 25, 355–358.Find this resource:

Lindemans, M., Liu, F., Janssen, T., Husson, S. J., Mertens, I., Gäde, G., & Schoofs, L. (2009). Adipokinetic hormone signaling through the gonadotropin-releasing hormone receptor modulates egg-laying in Caenorhabditis elegans. Proceedings of the National Academy of Sciences USA, 106(5), 1642–1647.Find this resource:

Lipinski, M. R. (1979). Universal maturity scale for the commercially important squid (Cephalopoda: Teuthoidea). The results of maturity classifications of the Illex illecebrosus (LeSueur, 1821) populations for the years 1973–1977. International Commission for the Northwest Atlantic Fisheries Research Documents, 79/II/38, 40.Find this resource:

Lodi, M., & Koene, J. M. (2016a). The love-darts of land snails: Integrating physiology, morphology and behaviour. Journal of Molluscan Studies, 82, 1–10.Find this resource:

Lodi, M., & Koene, J. M. (2016b). On the effect specificity of accessory gland products transferred by the love-dart of land snails. BMC Evolutionary Biology, 16, 104.Find this resource:

Loose, M. J., & Koene, J. M. (2008). Sperm transfer is affected by mating history in the simultaneously hermaphroditic snail Lymnaea stagnalis. Invertebrate Biology, 127, 162–167.Find this resource:

Lucero, M. T., Gilly, W. F., Abbott, N. J., Williamson, R., & Maddock, L. (Eds.). (1995). Cephalopod neurobiology: Neuroscience studies in squid, octopus and cuttlefish. London, England: Oxford University Press.Find this resource:

Lucero, M. T., Horrigan, F. T., & Gilly, W. F. (1992). Electrical responses to chemical-stimulation of squid olfactory receptor cells. Journal of Experimental Biology, 162, 231–249.Find this resource:

Lucero, M. T., Huang, W., & Dang, T. (2000). Immunohistochemical evidence for the Na+/Ca2+ exchanger in squid olfactory neurons. Philosophical Transactions of the Royal Society of London, B: Biological Sciences, 355, 1215–1218.Find this resource:

MacGinitie, G. E. (1934). The egg-laying activities of the sea hare, Tethys californica (Cooper). Biological Bulletin, 67, 300–303.Find this resource:

Mann, T. (1970). Male reproductive tract, spermatophores and spermatophoric reaction in the giant octopus of the North Pacific, Octopus dofleini martini. Proceedings of the Royal Society of London, B: Biological Sciences, 175, 31–61.Find this resource:

Mann, T., Martin, A. W., & Thiersch, J. B. (1970). Male reproductive tract, spermatophores and spermatophoric reaction in the giant octopus of the North Pacific, Octopus dojleini martini. Proceedings of the Royal Sciety of London, B: Biological Sciences, 175, 31–61.Find this resource:

Marchand, W. (1913). Studien tiber Cephalopoden II. Ueber die Spermatophoren. Zoologica, 67, 171–200.Find this resource:

Markov, G. V., & Laudet, V. (2011). Origin and evolution of the ligand-binding ability of nuclear receptors. Molecular and Cellular Endocrinology, 334(1–2), 21–30.Find this resource:

Markov, G. V., Tavares, R., Dauphin-Villemant, C., Demeneix, B. A., Baker, M. E., & Laudet, V. (2009). Independent elaboration of steroid hormone signaling pathways in metazoans. Proceedings of the National Academy of Sciences USA, 106(29), 11913–11918.Find this resource:

Martin, R. (1968). Fine structure of the neurosecretory system of the vena cava in Octopus. Brain Research, 8, 201–205.Find this resource:

Matsumoto, T., Masaoka, T., Fujiwara, A., Nakamura, Y., Satoh, N., & Awaji, M. (2013). Reproduction-related genes in the pearl oyster genome. Zoological Science, 30, 826–850.Find this resource:

Maxwell, W. L. (1974). Spermiogenesis of Eledone cirrhosa Lamarck (Cephalopoda: Octopoda) Proceedings of the Royal Society of London, B: Biological Sciences, 186, 181–190.Find this resource:

Maxwell, W. L. (1975). Spermiogenesis of Eusepia officinalis (L.), Loligo forbesi (Steenstrup) and Alloteuthis subulata (L.) (Cephalopoda: Decapoda). Proceedings of the Royal Society of London, B: Biological Sciences, 191, 527–535.Find this resource:

McCarthy, T. A. (2004). Effects of pair-type and isolation time on mating interactions of a freshwater snail, Physa gyrina (Say, 1821). American Malacological Bulletin, 19, 47–55.Find this resource:

Messenger, J. B. (1967). The peduncle lobe: A visuo-motor centre in Octopus. Proceedings of the Royal Society of London, B: Biological Sciences, 167, 225–251.Find this resource:

Messenger, J. B. (1979). The nervous system of Loligo. IV. The peduncle and olfactory lobes. Philosophical Transactions of the Royal Society of London, B: Biological Sciences, 285, 275–308.Find this resource:

Michelson, E. H. (1960). Chemoreception in the snail Australorbis glabratus. American Journal of Tropical Medicine and Hygene, 9, 480–487.Find this resource:

Michiels, N. K. (1998). Mating conflicts and sperm competition in simultaneous hermaphrodites. In T. R. Birkhead & A. P. Møller (Eds.) Sperm competition and sexual selection (pp. 219–254). Academic Press Ltd.Find this resource:

Mobley, A. S., Mahendra, G., & Lucero, M. T. (2007). Evidence for multiple signaling pathways in single squid olfactory receptor neurons. Journal of Comparative Neurology, 501, 231–242.Find this resource:

Mobley, A. S., Lucero, M. T., & Michel, W. C. (2008a). Cross-species comparison of metabolite profiles in chemosensory epithelia: An indication of metabolite roles in chemosensory cells. Anatomical Record, 291, 410–432.Find this resource:

Mobley, A. S., Michel, W. C., & Lucero, M. T. (2008b). Odorant responsiveness of squid olfactory receptor neurons. Anatomical Record, 291, 763–774.Find this resource:

Morgan, K., & Millar, R. P. (2004). Evolution of GnRH ligand precursors and GnRH receptors in protochordate and vertebrate species. General and Comparative Endocrinology, 139(3), 191–197.Find this resource:

Moroz, L. L., Citarella, M., Yu, F., Di Cristo, C., Burbach, J. P. H., Di Cosmo, A., … Kohn, A. B. (2011). Genomics and neurogenomics of cephalopods: From genes to behavior. Journal of Shellfish Researcj, 30, 1014–1015.Find this resource:

Murata, M., Ishii, M., & Osako, M. (1982). Some information on copulation of the oceanic squid Onychoteuthis borealijaponica Okada. Bulletin of the Japanese Society of Scientific Fisheries, 48, 351–354.Find this resource:

Muschamp, J. W., & Fong, P. P. (2001). Effects of the serotonin receptor ligand methiothepin on reproductive behavior of the freshwater snail Biomphalaria glabrata: Reduction of egg laying and induction of penile erection. Journal of Experimental Zoology, 289, 202–207.Find this resource:

Nakadera, Y., & Koene, J. M. (2013). Reproductive strategies in hermaphroditic gastropods: Conceptual and empirical approaches. Canadian Journal of Zoology, 91, 367–381.Find this resource:

Nakadera, Y., Swart, E. M., Hoffer, J. N. A., Den Boon, O., Ellers, J., & Koene, J. M. (2014a). Receipt of seminal fluid proteins causes reduction of male investment in a simultaneous hermaphrodite. Current Biology, 24, 1–4.Find this resource:

Nakadera, Y., Blom, C. & Koene, J. M. (2014b). Duration of sperm storage in the simultaneous hermaphrodite Lymnaea stagnalis. Journal of Molluscan Studies, 80, 1–7.Find this resource:

Nakamura, H., Ito, I., Kojima, S., Fujito, Y., Suzuki, H., & Ito, E. (1999a). Histological characterization of lip and tentacle nerves in Lymnaea stagnalis. Neuroscience Research, 33, 127–136.Find this resource:

Nakamura, H., Kojima, S., Kobayashi, S., Ito, I., Fujito, Y., Suzuki, H., & Ito, E. (1999b). Physiological characterization of lip and tentacle nerves in Lymnaea stagnalis. Neuroscience Research, 33, 291–298.Find this resource:

Nakamura, S., Osada, M., & Kijima, A. (2007). Involvement of GnRH neuron in the spermatogonial proliferation of the scallop, Patinopecten yessoensiss. Molecular Reproduction and Development, 74, 108–115.Find this resource:

Naud, M. J., & Havenhand, J. N. (2006). Sperm motility and longevity in the giant cuttlefish, Sepia apama (Mollusca: Cephalopoda). Marine Biology, 148, 559–566.Find this resource:

Naud, M. J., Shaw, P. W., Hanlon, R. T., & Havenhand, J. N. (2005). Evidence for biased use of sperm sources in wild female giant cuttlefish (Sepia apama). Proceedings of the Royal Society of London, B: Biological Sciences, 272, 1047–1051.Find this resource:

Nezlin, L., Moroz, L., Elofsson, R., & Sakharov, D. (1994). Immunolabeled neuroactive substances in the osphradium of the pond snail Lymnaea stagnalis. Cell and Tissue Research, 275, 269–275.Find this resource:

Nezlin, L., & Voronezhskaya, E. (1997). GABA-immunoreactive neurones and interactions of GABA with serotonin and FMRFamide in a peripheral sensory ganglion of the pond snail Lymnaea stagnalis. Brain Research, 772, 217–225.Find this resource:

Nezlin, L. P. (1995). Primary sensory neurons and their central projections in the pond snail Lymnaea stagnalis. Acta Biologica Hungarica, 46, 305–313.Find this resource:

Nezlin, L. P. (1997). The osphradium is involved in the control of egg-laying in the pond snail Lymnaea stagnalis. Invertebrate Reproduction and Development, 32, 163–166.Find this resource:

Ng, T., Saltin, S. H., Davies, M. S., Johannesson, K., Stafford, R., & Williams, G. A. (2013). Snails and their trails: The multiple functions of trail‐following in gastropods. Biological Reviews, 88, 683–700.Find this resource:

Nhan, H. T., Jung, L. H., Ambak, M. A., Watson, G. J., & Siang, H. Y. (2010). Evidence for sexual attraction pheromones released by male tropical donkey’s ear abalone (Haliotis asinina), (L.). Invertebrate Reproduction and Development, 54, 169–176.Find this resource:

Nigmatullin, C. M., & Laptikhovsky, V. V. (1994). Reproductive strategies in the squid of the family Ommastrephidae (preliminary report). Ruthenica, 4, 79–82.Find this resource:

Nishioka, R. S., Bern, H. A., & Golding, D. W. (1970). Innervation of the cephalopod optic gland. In W. Bargman & B. Scharrer (Eds.), Aspects of neuroendocrinology (pp. 47–54). Berlin, Germany: Springer-Verlag.Find this resource:

O’Dor, R. K., & Wells, M. J. (1973). Yolk protein synthesis in the ovary of Octopus vulgaris and its control by the optic gland gonadotropin. Journal of Experimental Biology, 59(3), 665–674.Find this resource:

O’Dor, R. K., & Wells, M. J. (1975). Control of yolk protein synthesis by octopus conadotropin in vivo and in vitro (effects of octopus gonadotropin). General and Comparative Endocrinology, 27(2), 129–135.Find this resource:

O’Dor, R. K., & Wells, M. J. (1978). Reproduction versus somatic growth: Hormonal control in Octopus vulgaris. Journal of Experimental Biology, 77, 15–31.Find this resource:

Onitsuka, C., Yamaguchi, A., Kanamaru, H., Oikawa, S., Takeda, T., & Matsuyama, M. (2009). Molecular cloning and expression analysis of a GnRH-like dodecapeptide in the swordtip squid, Loligo edulis. Zoological Science, 26(3), 203–208.Find this resource:

Osada M., & Treen, N. (2013). Molluscan GnRH associated with reproduction. General and Comparative Endocrinology, 181, 254–258.Find this resource:

Owen, R. (1832). Memoir on the pearly nautilus (Nautilus pompilius Linn.) with illustrations of its external form and internal structure. London, England: Council of Royal College Surgeons.Find this resource:

Packard, A. (1961). Sucker display of octopus. Nature, 190, 736–737.Find this resource:

Painter, S. A., Clough, B., Black, S., & Nagle, G. T. (2003). Behavioral characterization of attractin, a watter-borne peptide pheromone in the genus Aplysia. Biological Bulletin, 205, 16–25.Find this resource:

Painter, S. D., Cummins, S. F., Nichols, A. E., Akalal, D-B. G., Schein, C. H., Braun, W., … Nagle, G. T. (2004). Structural and functional analysis of Aplysia attractins, a family of water-borne protein pheromones with interspecific attractiveness. Proceedings of the National Academy of Sciences USA, 101, 6929–6933.Find this resource:

Parker, G. A. (1970). Sperm competition and its evolutionary consequences in the insects. Biological Reviews of the Cambridge Philosophical Society, 45, 525–567.Find this resource:

Pazos, A. J., & Mathieu, M. (1999). Effects of five natural gonadotropin-releasing hormones on cell suspensions of marine bivalve gonad: Stimulation of gonial DNA synthesis. General and Comparative Endocrinology, 113(1), 112–120.Find this resource:

Perry, J. C., Sirot, L., & Stuart, W. (2013). The seminal symphony: How to compose an ejaculate. TREE, 28, 414–422.Find this resource:

Pinsker, H. M., & Parsons, D. W. (1985). Temperature dependence of egg laying in Aplysia brasiliana and A. californica. Journal of Comparative Physiology B, 156, 21–27.Find this resource:

Piper, D. R., & Lucero, M. T. (1999). Calcium signalling in squid olfactory receptor neurons. Biological Signals and Receptors, 8, 329–337.Find this resource:

Plesch, B., De Jong-Brink, M., & Boer H. H. (1971). Histological and histochemical observations on the reproductive tract of the hermaphrodite pond snail Lymnaea stagnalis (L.). Netherlands Journal of Zoology, 21, 180–201.Find this resource:

Polese, G., Bertapelle, C., & Di Cosmo, A. (2015). Role of olfaction in Octopus vulgaris reproduction. General and Comparative Endocrinology, 210, 55–62.Find this resource:

Polese, G., Bertapelle, C., & Di Cosmo, A. (2016). Olfactory organ of Octopus vulgaris: Morphology, plasticity, turnover and sensory characterization. Biology Open, 15, 611–619.Find this resource:

Reich, G. (1992). A new peptide of the oxytocin/vasopressin family isolated from nerves of the cephalopod Octopus vulgaris. Neuroscience Letters, 134, 191–194.Find this resource:

Richard, A. (1967). Action de la température sur ĺevolution génitale de Sepia officinalis. Comptes Rendus de l’Académie des Sciences, D, 263, 1998–2001.Find this resource:

Richard, A. (1970). Differentiation sexuelle des cephalopodes en culture in vitro. Annee Biologique, 9, 409–415.Find this resource:

Richard, A. (1971). Contribution a l’étude experimentale de la croissance et de la maturation sexuelle de Sepia officinalis L. (Mollusque, Cephalopode). Lille, France: Université de Lille.Find this resource:

Rigby, J. E. (1982). The fine structure of differentiating spermatozoa and Sertoli cells in the gonad of the pond snail, Lymnaea stagnalis. Journal of Molluscan Studies, 48, 111–123.Find this resource:

Roch, G. J., Busby, E. R., & Sherwood, N. M. (2011). Evolution of GnRH: Diving deeper. General and Comparative Endocrinology, 171(1), 1–16.Find this resource:

Rocha, F., Guerra, A. & Gonzale, A. F. (2001). A review of reproductive strategies in cephalopods. Biological Reviews, 76, 291–304.Find this resource:

Rock, M. K., Blankenship, J. E., & Lebeda, F. J. (1977). Penis retractor muscle of Aplysia: Excitatory motor neurones. Journal of Neurobiology, 8, 569–579.Find this resource:

Rodaniche, A. F. (1984). Iteroparity in the lesser Pacific striped octopus, Octopus chierchiae (Jatta, 1889). Bulletin of Marine Science, 35, 99–104.Find this resource:

Rogers, D., & Chase, R. (2001). Dart receipt promotes sperm storage in the garden snail Helix aspersa. Behavioral Ecology and Sociology, 50, 122–127.Find this resource:

Rogers, D., & Chase, R. (2002). Determinants of paternity in the garden snail Helix aspersa. Behavioral Ecology and Sociology, 52, 289–295.Find this resource:

Roubos, E. W. (1976). Neuronal and non-neuronal control of the neurosecretory caudo-dorsal cells of the freshwater snail Lymnaea stagnalis (L.). Cell and Tissue Research, 168, 11–31.Find this resource:

Roubos, E. W., Geraerts, W. P., Boerrigter, G. H., & van Kampen, G. P. (1980). Control of the activities of the neurosecretory light green and caudo-dorsal cells and of the endocrine dorsal bodies by the lateral lobes in the freshwater snail Lymnaea stagnalis (L.). General and Comparative Endocrinology, 40(4), 446–454.Find this resource:

Roubos, E. W., & Van der Ven, A. M. H. (1987). Morphology of neurosecretory cells in basommatophoran snails homologous with egg-laying and growth-hormone producing cells of Lymnaea stagnalis. General and Comparative Endocrinology, 67, 7–23.Find this resource:

Rudolph, P. H. (1979a). The strategy of copulation of Stagnicola elodes (Say) (Basommatophora: Lymnaeidea). Malacologia, 18, 381–389.Find this resource:

Rudolph, P. H. (1979b). An analysis of copulation in Bulinus (Physopsis) globosus (Gastropoda: Planorbidea). Malacologia, 19, 147–155.Find this resource:

Rudolph, P. H., & Bailey, J. B. (1985). Copulation as females and use of allosperm in the freshwater snail genus Bulinus (Gastropoda: Planorbidae). Journal of Molluscan Studies, 51, 267–275.Find this resource:

Rudolph, P. H., & White, J. K. (1979). Egg laying behaviour of Bulinus octoploidus Burch (Basommatophora: Planorbidae). Journal of Molluscan Studies, 45, 355–363.Find this resource:

Ruth, P., Schmidtberg, H., Westermann, B., & Schipp, R. (2002). The sensory epithelium of the tentacles and the rhinophore of Nautilus pompilius L. (Cephalopoda, Nautiloidea). Journal of Morphology, 251, 239–255.Find this resource:

Salman, A., & Akalin, M. (2012). A rare pelagic cephalopod Ocythoe tuberculata (Octopoda: Argonautoidea): The record fecundity for Octopoda and new data on morphometry. Turkish Journal of Fisheries and Aquatic Sciences, 12, 339–344.Find this resource:

Santama, N., Li, K. W., Bright, K. E., Yeoman, M. S., Geraerts, W. P. M., Benjamin, P. R., & Burke, J. F. (1993). Processing of the FMRFamide precursor protein in the snail Lymnaea stagnalis: Characterization and neuronal localization of a novel peptide, “SEEPLY.” European Journal of Neuroscience, 5, 1003–1016.Find this resource:

Santama, N., Wheeler, S. H., Skingsley, D. R., Yeoman, M. S., Bright, K., Kaye, I., Burke, J. F., & Benjamin, P. R. (1995). Identification, distribution and physiological activity of three neuropeptides of Lymnaea; EFLRIamide and pQFYRIamide encoded by the FMRFamide gene and a related peptide. European Journal of Neuroscience, 7, 234–246.Find this resource:

Schärer, L. (2009). Tests of sex allocation theory in simultaneously hermaphroditic animals. Evolution, 63, 1377–1405.Find this resource:

Schärer, L. (2014). Evolution: Don’t Be So Butch, Dear! Current Biology, 24, R311–R313.Find this resource:

Schnell, A. K., Smith, C. L., Hanlon, R. T., & Harcourt, R. T. (2015). Female receptivity, mating history, and familiarity influence the mating behavior of cuttlefish. Behavioral Ecology and Sociobiology, 69(2), 283–292.Find this resource:

Schot, L. P. C., & Boer, H. H. (1982). Immunocytochemical demonstration of peptidergic cells in the pond snail Lymnaea stagnalis with an antiserum to the molluscan cardioactive tetrapeptide FMRF-amide. Cell and Tissue Research, 225, 347–354.Find this resource:

Schuldt, M. (1979). Contribucion al conocimiento del ciclo reproductor de Illex argentinus (Cephalopoda: Ommastrephidae). Monographias, 10, 110.Find this resource:

Selman, K., & Arnold, J. M. (1977). An ultrastructural and cytochemical analysis of oogenesis in the squid, Loligo pealei. Journal of Morphology, 152, 381–400.Find this resource:

Selman, K., & Wallace, R. A. (1978). An autoradiographic study of vitellogenesis in the squid, Loligo pealei. Tissue Cell, 10, 599–608.Find this resource:

Sirinupong, P., Suwanjarat, J., & Van Minnen, J. (2011). Distribution of APGWamide-immunoreactivity in the brain and reproductive organs of adult Pygmy squid, Idiosepius pygmaeus. Invertebrate Neuroscience, 11, 97–102.Find this resource:

Smart, D., Shaw, C., Johnston, C., Thim, L., Halton, D., & Buchanan, K. (1992). Peptide tyrosine phenylalanine: A novel neuropeptide F-related nonapeptide from the brain of the squid, Loligo vulgaris. Biochemical and Biophysical Research Communications, 186(3), 1616–1623.Find this resource:

Smit, A. B., Jimenez, C. R., Dirks, R. W., Croll, R. P., & Geraerts, W. P. M. (1992). Characterization of a cDNA clone encoding multiple copies of the neuropeptide APGWamide in the mollusk Lymnaea stagnalis. Journal of Neuroscience, 12, 1709–1715.Find this resource:

Smith, R. L. (Ed.). (1984). Sperm competition and the evolution of animal mating system. Orlando, FL: Academic Press.Find this resource:

Smith, S. A., Nason, J., & Croll, R. P. (1997). Detection of APGWamide-like immunoreactivity in the sea scallop, Placopecten magellanicus. Neuropeptides, 31, 155–165.Find this resource:

Squires, Z. E., Norman, M. D., & Stuart-Fox, D. (2013). Mating behaviour and general spawning patterns of the southern dumpling squid Euprymna tasmanica (Sepiolidae): A laboratory study. Journal of Molluscan Studies, 79, 263–269.Find this resource:

Squires, Z. E., Wong, B. B. M., Norman, M. D., & Stuart-Fox, D. (2014). Multiple paternity but no evidence of biased sperm use in female dumpling squid Euprymna tasmanica. Marine Ecology Progress Series, 511, 93–103.Find this resource:

Squires, Z. E., Wong, B. B. M., Norman, M. D., & Stuart-Fox, D. (2015). Last male sperm precedence in a polygamous squid. Biological Journal of the Linnean Society, 116, 277–287.Find this resource:

Stewart, M. J., Favrel, P., Rotgans, B. A., Wang, T., Zhao, M., Sohail, M., … Cummins, S. F. (2014). Neuropeptides encoded by the genomes of the Akoya pearl oyster Pinctata fucata and Pacific oyster Crassostrea gigas: A bioinformatic and peptidomic survey. BMC Genomics, 15, 840.Find this resource:

Stewart, M. J., Wang, T., Koene, J. M., Storey, K. B., & Cummins, S. E. (2016a). A love dart allohormone identified in the mucous glands of hermaphroditic land snails. Journal of Biological Chemistry, 291, 7938–7950.Find this resource:

Stewart, M. J., Wang, T., Harding, B. I., Bose, U., Wyeth, R. C., Storey, K. B., & Cummins S. F. (2016b). Characterisation of reproduction-associated genes and peptides in the pest land snail, Theba pisana. PLoS ONE, 11, e0162355.Find this resource:

Strumwasser, F., Kaczmarek, L. K., Chiu, A. Y., Hiller, E., Jennings, K. R., & Viele, D. P. (1980). Peptides controlling behavior in Aplysia. In F. E. Bloom (Ed.), Peptides: Integrators of cell and tissue function (pp. 197–218). New York, NY: Raven Press.Find this resource:

Sun, B., & Tsai, P-S. (2011). A gonadotropin-releasing hormone-like molecule modulates the activity of diverse central neurons in a gastropod mollusk, Aplysia californica. Frontiers in Endocrinology, 2(36), 1–8.Find this resource:

Susswein, A. J., Gev, S., Achituv, Y., & Markovich, S. (1984). Behavioral patterns of Aplysia fasciata along the Mediterranean coast of Israel. Behavioral and Neural Biology, 41, 7–22.Find this resource:

Suzuki, H., Muraoka, T., & Yamamoto, T. (2003). Localization of corticotropin-releasing factor-immunoreactive nervous tissue and colocalization with neuropeptide Y-like substance in the optic lobe and peduncle complex of the octopus (Octopus vulgaris). Cell and Tissue Research, 313(1), 129–138.Find this resource:

Suzuki, H., Yamamoto, T., Inenaga, M., & Uemura, H. (2000). Galanin-immunoreactive neuronal system and colocalization with serotonin in the optic lobe and peduncle complex of the octopus (Octopus vulgaris). Brain Research, 865(2), 168–176.Find this resource:

Suzuki, H., Yamamoto, T., Nakagawa, M., & Uemura, H. (2002). Neuropeptide Y-immunoreactive neuronal system and colocalization with FMRFamide in the optic lobe and peduncle complex of the octopus (Octopus vulgaris). Cell and Tissue Research, 307(2), 255–264.Find this resource:

Tait, R. W. (1986). Aspects physiologigues de la senescence postreproductive chez Octopus vulgaris. Paris, France: Université de Paris II.Find this resource:

Takahashi, N. (1978). Ultrastructural characteristics of the proteid yolk formation in the ovary of the squid, Todarodes pacificus. Bulletin of the Faculty of Fisheries Hokkaido University, 29, 89–99.Find this resource:

Takuwa-Kuroda, K., Iwakoshi-Ukena, E., Kanda, A., & Minakata, H. (2003). Octopus, which owns the most advanced brain in invertebrates, has two members of vasopressin/oxytocin superfamily as in vertebrates. Regulatory Peptides, 115, 139–419.Find this resource:

Ter Maat, A. (1979). Neuronal input on the ovulation hormone producing neuro-endocrine caudo-dorsal cells of the freshwater snail Lymnaea stagnalis. Proceedings of the Koninklijke Nederlandse Akademie van Wetenschappen, 82C, 333–342.Find this resource:

Ter Maat, A., Dijcks, F. A., & Bos, N. P. A. (1986). In vivo recording of neuroendocrine cells (caudo-dorsal cells) in the pond snail. Journal of Comparative Physiology A, 158, 853–859.Find this resource:

Ter Maat, A., Geraerts, W. P. M., Jansen, R. F., & Bos, N. P. A. (1988). Chemically mediated positive feedback generates long-lasting discharge in the molluscan neuroendocrine system. Brain Research, 438, 77–82.Find this resource:

Ter Maat, A., & Lodder, J. C. (1980). A biphasic cholinergic input on the ovulation hormone producing caudo-dorsal cells of the freshwater snail Lymnaea stagnalis, Comparative Biochemistry and Physiology, 66, 115–119.Find this resource:

Ter Maat, A., Lodder, J. C., Veenstra, J., & Goldschmeding, J. T. (1982). Suppression of egg-laying during starvation in the snail Lymnaea stagnalis by inhibition of the ovulation hormone producing caudo-dorsal cells. Brain Research, 239, 535–542.Find this resource:

Ter Maat, A., Lodder, J. C., & Wilbrink, M. (1983). Induction of egg-laying in the pond snail Lymnaea stagnalis by environmental stimulation of the release of ovulation hormone from the caudo-dorsal cells. International Journal of Invertebrate Reproduction, 6, 239–247.Find this resource:

Ter Maat, A., Pieneman, A. W., Goldschmeding, J. T., Smelik, W. F. E., & Ferguson, G. P. (1989). Spontaneous and induced egg laying behavior of the pond snail, Lymnaea stagnalis. Journal of Comparative Physiology A, 164, 673–683.Find this resource:

Ter Maat, A., Pieneman, A. W., & Koene, J. M. (2012). The effect of light on induced egg laying in the simultaneous hermaphrodite Lymnaea stagnalis. Journal of Molluscan Studies, 78, 262–267.Find this resource:

Ter Maat, A., Van Duivenboden, Y. A., & Jansen, R. F. (1987). Copulation and egg-laying behavior in the pond snail. In H. H. Boer, W. P. M. Geraerts, & J. Joosse (Eds.), Neurobiology: Molluscan models (pp. 255–261). Amsterdam, The Netherlands: North-Holland.Find this resource:

Ter Maat, A., Zonneveld, C., De Visser, J. A. G. M., Jansen, R. F., Montagne-Wajer, K., & Koene, J. M. (2007). Food intake, growth, and reproduction as affected by day length and food availability in the pond snail Lymnaea stagnalis. American Malacological Bulletin, 23, 113–120.Find this resource:

Tosti, E., Di Cosmo, A., Cuomo, A., Di Cristo, C., Gragnaniello, G. (2001). Progesterone induces activation in Octopus vulgaris spermatozoa. Molecular Reproduction and Development, 59(1), 97–105.Find this resource:

Townsend, C. R. (1973). The role of the osphradium in chemoreception by the snail Biomphalaria glabrata (Say). Animal Behavior, 21, 549–556.Find this resource:

Townsend, C. R. (1974). Mucus trail following by the snail Biomphalaria glabrata (Say), Animal Behavior, 22, 170–177.Find this resource:

Treen, N., Itoh, N., Miura, H., Kikuchi, I., Ueda, T., Takahashi, K. G., … Osada, M. (2012). Mollusc gonadotropin-releasing hormone directly regulates gonadal functions: A primitive endocrine system controlling reproduction. General and Comparative Endocrinology, 176(2), 167–172.Find this resource:

Tsai, P. S., Sun, B., Rochester, J. R., & Wayne, N. L. (2010). Gonadotropin-releasing hormone-like molecule is not an acute reproductive activator in the gastropod, Aplysia californica. General and Comparative Endocrinology, 166(2), 280–288.Find this resource:

Tsai, P. S., & Zhang, L. (2008). The emergence and loss of gonadotropin-releasing hormone in protostomes: Orthology, phylogeny, structure, and function. Biology of Reproduction, 79(5), 798–805.Find this resource:

Van Duivenboden, Y. A., Pieneman, A. W., & Ter Maat, A. (1985). Multiple mating suppresses fecundity in the hermaphrodite freshwater snail Lymnaea stagnalis: A laboratory study. Animal Behavior, 33, 1184–1191.Find this resource:

Van Duivenboden, Y. A., & Ter Maat, A. (1985). Masculinity and receptivity in the hermaphrodite pond snail, Lymnaea stagnalis. Animal Behavior, 33, 885–891.Find this resource:

Van Duivenboden, Y. A., & Ter Maat, A. (1988). Mating behaviour of Lymnaea stagnalis. Malacologia, 28, 23–64.Find this resource:

Van Golen, F. A., Li, K. W., De Lange, R. P., Van Kesteren, R. E., Van Der Schors, R. C., & Geraerts, W. P. M. (1995). Co-localized neuropeptides conopressin and ALA-PRO-GLY-TRP-NH2 have antagonistic effects on the vas deferens of Lymnaea. Neuroscience, 69, 1275–1287.Find this resource:

Van Heukelem, W. F. (1973). Growth and lifespan of Octopus cyanea (Mollusca: Cephalopoda). Journal of Zoology, 169, 299–315.Find this resource:

van Kesteren, R. E., Smit, A. B., de With, N. D., Van Minnen, J., Dirks, R. W., van der Schors, R. C., & Joosse, J. (1992). A vasopressin-related peptide in the mollusc Lymnaea stagnalis: peptide structure, prohormone organization, evolutionary and functional aspects of Lymnaea conopressin. Progress in Brain Research, 92, 47–57.Find this resource:

Van Kesteren, R. E., Tensen, C. P., Smit, A. B., Van Minnen, J., Kolakowski, L. F., Meyerhof, W., … Geraerts, W. P. (1996). Co-evolution of ligand-receptor pairs in the vasopressin/oxytocin superfamily of bioactive peptides. Journal of Biological Chemistry, 271(7), 3619–3626.Find this resource:

Van Minnen, J., Dirks, R. W., Vreugdenhil, E., & Van Diepen, J. (1989). Expression of the egg-laying hormone genes in peripheral neurons and exocrine cells in the reproductive tract of the mollusc Lymnaea stagnalis. Neuroscience, 33, 35–46.Find this resource:

Van Minnen, J., Schallig, H. D. F. H., & Ramkema, M. D. (1992). Identification of putative egg-laying hormone containing neuronal systems in gastropod molluscs. General and Comparative Endocrinology, 86, 96–102.Find this resource:

Veenstra, J. A. (2010). Neurohormones and neuropeptides encoded by the genome of Lottia gigantea with reference to other mollusks and insects. General and Comparative Endocrinology, 167, 86–103.Find this resource:

Vernon, J. G., & Taylor, J. K. (1996). Patterns of sexual roles adopted by the schistosome-vector snail Biomphalaria glabrata (Planorbidae). Journal of Molluscan Studies, 62, 235–241.Find this resource:

Villanueva, R. (1992). Continuous spawning in the cirrate octopods Opisthoteuthis agassizii and O.vossi: features of sexual maturation defining a reproductive strategy in cephalopods. Marine Biology, 114, 265–275.Find this resource:

Villanueva, R., & Norman, M. D. (2008). Biology of the planktonic stages of benthic octopuses. Oceanographic and Marine Biology Annual Review, 46, 105–202.Find this resource:

Voight, J. R. (1991). Ligula length and courtship in Octopus digeti: A potential mechanism of mate choice. Evolution, 45, 1726–1730.Find this resource:

Vreugdenhil, E., Geraerts, W. M. P., Jackson, J. F., & Joosse J. (1985). The molecular basis of the neuro-endocrine control of egg-laying behaviour in Lymnaea. Peptides, 6, 465–470.Find this resource:

Walderon, M. D., Nolt, K. J., Haas, R. E., Prosser, K. N., Holm, J. B., Nagle, G. T., & Boal, J. G. (2011). Distance chemoreception and the detection of conspecifics in Octopus bimaculoides. Journal of Molluscan Studies, 77, 309–311.Find this resource:

Wedemeyer, H., & Schild, D. (1995). Chemosensitivity of the osphradium of the pond snail Lymnaea stagnalis. Journal of Experimental Biology, 198(8), 1743–1754.Find this resource:

Wells, M. J. (1960). Optic glands and the ovary of Octopus. Symposia of the Zoological Society of Lond, 2, 87–101.Find this resource:

Wells, M. J. (1962). Taste by touch: Some experiments with Octopus. Journal of Experimental Biology, 40, 187–193.Find this resource:

Wells, M. J. (1978). Octopus—Physiology and behaviour of an advanced invertebrate. London, England: Chapman and Hall.Find this resource:

Wells, M. J., & Buckley, S. K. L. (1972). Snails and trails. Animal Behavior, 20, 345–355.Find this resource:

Wells, M. J., O’Dor, R. K., & Buckley, S. K. (1975). An in vitro bioassay for a mulluscan gonadotropin. Journal of Experimental Biology, 62(2), 433–446.Find this resource:

Wells, M. J., & Wells, J. (1959). Hormonal control of sexual maturity in Octopus. Journal of Experimental Biology, 36, 1–33.Find this resource:

Wells, M. J., & Wells, J. (1972). Sexual displays and mating of Octopus vulgaris Cuvier and Octopus cyanea Gray and attempts to alter performance by manipulating the glandular condition of the animals. Animal Behavior, 20, 293–308.Find this resource:

Wells, M. J., & Wells, J. (1975). Optic gland implants and their effects on the gonads of Octopus. Journal of Experimental Biology, 62(3), 579–588.Find this resource:

Wendelaar Bonga, S. E. (1971). Formation, storage, and release of neurosecretory material studied by quantitative electron microscopy in the fresh water snail Lymnaea stagnalis (L.). Zeitschrift für Zellforschung und mikroskopische Anatomie, 113, 490–517.Find this resource:

Werminghausen, J., Lange, R., & Anthes, N. (2013). Seeking a sex-specific Coolidge effect in a simultaneous hermaphrodite. Ethology, 119, 541–551.Find this resource:

Wethington, A. R., & Dillon, R. T., Jr. (1996). Gender choice and gender conflict in a non-reciprocally mating simultaneous hermaphrodite, the freshwater snail, Physa. Animal Behavior, 51, 1107–1118.Find this resource:

Wijdenes, J., Van Elk, R., & Joosse, J. (1983). Effects of two gonad-otropic hormones on polysaccharide synthesis in the albumen gland of Lymnaea stagnalis studied with the organ culture technique. General and Comparative Endocrinology, 51, 263–271.Find this resource:

Wijsman, T. C. M. (1989). Glycogen and galactogen in the albumin gland of the fresh-water snail Lymnaea stagnalis—effects of egg laying, photo period and starvation. Comparative Biochemistry and Physiology A—Physiology, 92, 53–59.Find this resource:

Wilson, J. R., Kuehn, R. E., & Beach, F. A. (1963). Modification in the sexual behavior of male rats produced by changing the stimulus female. Journal of Comparative and Physiological Psychology, 56, 636–644.Find this resource:

Wodinsky, J. (1977). Hormonal inhibition of feeding and death in Octopus: Control by optic gland secretion. Science, 198, 948–951.Find this resource:

Wood, J. F. G. (1963). Observations on the behavior of octopus. International Congress of Zoology, 16, 73.Find this resource:

Woodhams, P. L., & Messenger, J. B. (1974). A note on the ultrastructure of the Octopus olfactory organ. Cell and Tissue Research, 152(2), 253–258.Find this resource:

Wyeth, R. C., & Croll, R. P. (2011). Peripheral sensory cells in the cephalic sensory organs of Lymnaea stagnalis. Journal of Comparative Neurology—Research in Systems Neuroscience, 519, 1894–1913.Find this resource:

Young, J. Z. (1971). The anatomy of the nervous system of Octopus vulgaris. Oxford, England: Clarendon.Find this resource:

Young, K. G., Chang, J. P., & Goldberg, J. I. (1999). Gonadotropin-releasing hormone neuronal system of the freshwater snails Helisoma trivolvis and Lymnaea stagnalis: Possible involvement in reproduction. Journal of Comparative Neurology, 404, 427–437.Find this resource:

Yung Ko Ching, M. (1930). Contribution á l’étude cytologique de l’ovogénèse, du développement et de quelques organes chez les céphalopodes. Annales de l’ Institute Océanographique, 7, 299–364.Find this resource:

Zaitseva, O. V. (1994). Structural organization of the sensory systems of the snail. Neuroscience and Behavioral Physiology, 24, 47–57.Find this resource:

Zaitseva, O. V. (1999). Principles of the structural organization of the chemosensory systems of freshwater gastropod mollusks. Neuroscience and Behavioral Physiology, 29, 581–593.Find this resource:

Zaitseva, O. V., & Bocharova, L. S. (1981). Sensory cells in the head skin of pond snails. Cell and Tissue Research, 220, 797–807.Find this resource:

Zatylny, C., Gagnon, J., Boucaud-Camou, E., & Henry, J. (2000a). The SepOvotropin: A new ovarian peptide regulating oocyte transport in Sepia officinalis. Biochemical and Biophysical Research Communications, 276(3), 1013–1018.Find this resource:

Zatylny, C., Gagnon, J., Boucaud-Camou, E., & Henry, J. (2000b). ILME: A waterborne pheromonal peptide released by the eggs of Sepia officinalis. Biochemical and Biophysical Research Communications, 275(1), 217–222.Find this resource:

Zatylny, C., Marvin, L., Gagnon, J., & Henry, J. L. (2002). Fertilization in Sepia officinalis: The first mollusk sperm-attracting peptide. Biochemical and Biophysical Research Communications, 296, 1186–1193.Find this resource:

Zatylny-Gaudin, C., Cornet, V., Leduc, A., Zanuttini, B., Corre, E., Le Corguillé, G., … Henry, J. (2016). Neuropeptidome of the cephalopod Sepia officinalis: Identification, tissue mapping, and expression pattern of neuropeptides and neurohormones during egg laying. Journal of Proteome Research, 15, 48–67.Find this resource:

Zhang, X., Mao, Y., Huang, Z., Qu, M., Chen, J., Ding, S., … Sun, T. (2012). Transcriptome analysis of the Octopus vulgaris central nervous system. PLoS ONE, 7(6), e40320.Find this resource:

Zhang, L., Tello, J. A., Zhang, W., & Tsai, P. S. (2008). Molecular cloning, expression pattern, and immunocytochemical localization of a gonadotropin-releasing hormone-like molecule in the gastropod mollusk, Aplysia californica. General and Comparative Endocrinology, 156(2), 201–209.Find this resource:

Zhang, L., Wayne, N. L., Sherwood, N. M., Postigo, H. R., & Tsai, P. S. (2000). Biological and immunological characterization of multiple GnRH in an opisthobranch mollusk, Aplysia californica. General and Comparative Endocrinology, 118, 77–89.Find this resource:

Zijlstra, U., (1972). Distribution and ultrastructure of epidermal sensory cells in the freshwater snails Lymnaea stagnalis and Biomphalaria pfeifferi. Netherlands Journal of Zoology, 22, 283–298.Find this resource:

Ziv, I., Benni, M., Markovich, S., & Susswein, A. J. (1989). Motivational control of sexual behavior in Aplysia fasciata: Sequencing and modulation by sexual deprivation and by addition of partners. Behavioral and Neural Biology, 52, 180–193.Find this resource:

Zizzari, Z. V., Smolders, I., & Koene, J. M. (2014). Alternative delivery of male accessory gland products. Frontiers in Zoology, 11, 32.Find this resource: