Morphological grounds for the obligate aerial respiration of an aquatic snail: functional and evolutionary perspectives

Animals and dissections

Animals (young-adult males, 20 mm in shell length) from our cultured Rosedal strain (see Rodriguez et al., 2019 for details) were kept in a water bath at 4°C for 20–30 min to provoke relaxation and minimize nociception. After carefully cracking the shell, we dissected out the lung and surrounding tissues both for histological examinations and 3D reconstruction of the blood and nerve systems of the lung.

We dissected out lung sacs for routine light microscopy (from N = 6 animals); samples that contained most of the mantle edge and the copulatory apparatus, the lung, as well as parts of the gill, ureter, and pericardium, for 3D reconstructions (from N = 2 animals); and pieces of the lung floor and roof for scanning and transmission electron microscopy (SEM and TEM, from N = 4 and N = 10 animals, respectively).

The Institutional Committee for the Care and Use of Laboratory Animals (Comité Institucional para el Cuidado y Uso de Animales de Laboratorio, Facultad de Ciencias Médicas, Universidad Nacional de Cuyo) approved the procedures for animal culture, sacrifice and tissue sampling (approval protocol N° 55/2015).

Light microscopy and 3D reconstruction of the lung’s blood system and innervation

The histological processing involved sample fixation in dilute (1:2) Bouin’s fluid, dehydration in a graded ethanol series, clearing in xylene, paraffin-embedding (Histoplast, Argentina), and sectioning (5–10 μm-thick) on a rotary microtome (Microm). After staining with Gill’s hematoxylin-eosin, we examined and photographed the samples under a Nikon Eclipse 80i microscope using Nikon DS-Fi1-U3 camera and Nikon NIS-ELEMENT Image Software.

Also, we examined serially sectioned (10 μm-thick) Harris hematoxylin-eosin-stained lung samples under a Nikon Alphaphot-2 YS2 microscope equipped with a Nikon Digital Sight DS-5M camera. We selected the images of every fifth section for the 3D-reconstruction of the blood system and innervation of the lung. For this purpose, we used the software Reconstruct 1.1.0.0 (Fiala, 2005) and followed the procedure described by Ruthensteiner & Heß (2008) to assemble the final PDF-model, as described in Rodriguez et al. (2019).

Scanning and transmission electron microscopy

For SEM examination, we cut-opened the lung sacs to wash the lung cavity out of mucus, and then we fixed them with dilute Bouin’s fluid. After that, sample processing involved dehydration in a graded acetone series, critical point-drying, mounting on aluminum stubs, coating with gold, and examination with a Jeol/EO JSM-6490LV electron microscope.

The preparation of both lung roof and floor samples for TEM involved fixation in Karnovsky’s fluid, post-fixation in 1% osmium tetroxide overnight, dehydration in a graded acetone series, and embedding in Spurr’s resin. We obtained silver-grey, ultrathin sections, which we mounted on copper grids, stained them with uranyl acetate and lead citrate, and examined them with a Zeiss EM 900 microscope. Besides, we used ~200 nm-thick ultramicrotome sections stained with toluidine blue and mounted with DPX medium (Cat. #44581; Sigma–Aldrich, Sigma–Aldrich, St. Louis, MO, USA) for topographical orientation and examination under light microscopy.

General organization of the lung

The lung of P. canaliculata (Fig. 1A) is a single sac that occupies most of the roof of the mantle cavity. It abuts the gill on its posterior and right margins. An opening in the lung floor, namely the pneumostome, lies anteriorly, near the osphradium and the left mantle edge, and communicates the lung cavity with the mantle cavity. The pneumostome shows two lips (anterior and posterior) and can close by their apposition. Prominent blood sinuses traverse both the lung roof and floor. The muscular layer of the lung floor is formed of muscle strands arranged in two directions, thus is thicker than that of the roof, particularly in the region rostral to the pneumostome.

From dorsal to ventral, the lung roof (Fig. 1B) is composed of (1) the outer mantle epithelium, formed by columnar pigment and mucous cells; (2) a single muscle layer; (3) a vascular layer containing the main blood sinuses supplying the lung, which are surrounded by the perivascular storage tissue (Giraud-Billoud et al., 2011); and (4) the respiratory lamina. Almost mirroring that arrangement, the lung floor (Fig. 1C) is composed, from dorsal to ventral, of (1) the respiratory lamina; (2) the vascular and storage tissue layer, which is thicker than that of the roof; (3) a double muscle layer with fibers arranged in perpendicular directions; and (4) the inner mantle epithelium that lines the mantle cavity, and which is formed by columnar ciliated and mucous cells.

Blood supply and innervation of the lung

The 3D reconstruction of the lung and related pallial organs showed the blood supply to the roof and the floor comes from different origins (Fig. 2; see the interactive 3D model in Fig. S1). The afferent pulmonary vein, which is a prolongation of the afferent branchial vein, runs along the anterior and left margins of the lung, branching repeatedly and supplying the lung roof all along its course. Upon irrigating the lung roof, the blood reaches the efferent pulmobranchial vein, which conveys blood also from the gill leaflets and carries it to the heart auricle (Figs. 2B and 2E).

In turn, the lung floor receives blood from several origins and, for clarity, will be treated separately as the right (Fig. 2C) and the left afferent systems (Fig. 2D). Connection of the gill and lung circulation occurs through the right afferent sinuses. The ventral afferent pulmonary vein is the main right afferent and runs parallel to the already mentioned efferent pulmobranchial vein. It receives blood from the gill leaflets and divides within the right half of the lung floor into numerous right afferent sinuses (Figs. 2C and 2F). All the latter will end at the ventral efferent pulmonary vein, which follows a diagonal path within the lung floor. This large vein arises as a branch of the efferent pulmobranchial vein, and finally joins it again close to its entrance into the auricle. Also, a branch arising from the afferent pulmonary vein, near the osphradium, turns to the right of the snail and ramifies into the anterior part of the lung floor. The latter branches will join the efferent pulmobranchial vein (Figs. 2C and 2F).

The left part of the lung floor is irrigated by numerous left afferent sinuses that arise from left pallial sinuses (Figs. 2D and 2F). All these left afferent sinuses end in the ventral efferent pulmonary vein. Finally, the two largest efferent veins (the ventral efferent pulmonary vein and the efferent pulmobranchial vein) merge upon entering the auricle (Figs. 2D and 2F).

The lung roof is innervated by branches arising from the supraesophageal ganglion while the floor is innervated from the accessory visceral ganglion (Fig. 3; Fig. S1). In addition, the supraesophageal ganglion sends two branches to the pneumostome. The smaller nerve branches could not be followed in hematoxylin–eosin preparations, but neurites were found spread in TEM preparations of the respiratory lamina, which will be described below.

The inner surface of the lung and the pneumostome

Under SEM, the lung cavity appears covered by pavement cells, with irregularly interspersed tufts of ciliated columnar cells (Fig. 4A). Numerous microvilli occur in the shallow grooves between pavement cells and some of them show enlarged tips (Fig. 4B); occasionally, pavement cells show microvilli on its central surface too, but these are shorter than those found in the grooves. The ciliary tufts (Fig. 4C) show cells with different apical specializations besides their long cilia, such as short microvilli and partially fused secretory globules (Fig. 4D). These apical specializations are further described in the corresponding section, as seen under TEM.

The pneumostome communicates the lung and the mantle cavities and an epithelium of ciliated and mucous cells lines it, which is continuous with the epithelium lining the rest of the mantle cavity (see Fig. 1C). In contrast, the epithelium covering the lung cavity side of the pneumostome lips (Fig. 5) shows ciliated cells arranged in single or double parallel rows, which gives the lips a striated appearance (Figs. 5A and 5B). Around this region of parallel rows, there are fields of densely packed ciliated cells (Fig. 5C) which are in turn surrounded by pavement cells. These cells are not separated by grooves (as in the respiratory lamina), but by ciliary rows (Fig. 5D). This zone marks the transition to the typical pavement of the respiratory lamina (Figs. 4A and 4B).

The respiratory lamina: the gas-exchange barrier

A diagram overviewing our findings on the cellular components of the respiratory lamina and underlying structures is presented in Fig. 6. Pavement cells (Figs. 7A–7D), which correspond to those seen under SEM in Figs. 4A and 4B, cover the inner surface of the lung cavity. They show rather electron-dense and middle-sized nuclei, usually located near the intercellular grooves or in the grooves themselves. Both desmosome-like and septate junctions join the pavement cells, which emit numerous and short microvilli, particularly at or near the grooves (Figs. 7A and 7B). The cytoplasm of pavement cells has a low organelle content and extends as a thin layer overlying an anastomosing network of small blood spaces that rests over a fibromuscular layer made of muscle fibers in a collagen matrix. The resulting blood–gas barrier (80–150 nm thick) is made by the thin cytoplasmic layer and its corresponding lamina densa (Figs. 7A–7D). Also, irregular cytoplasmic extensions deepen at the grooves, sometimes reaching the fibromuscular layer, so that dome-shaped interconnected microsinuses are formed (Fig. S2). Hemocytes in the sinus blood show prominent pseudopodia and phagosomes (Fig. 7D), which seem involved in the phagocytosis of a particulate material of variable size and appearance that occurs in the sinus blood.

Muscle and collagen fibers appear sectioned both transversally and longitudinally in the fibromuscular layer (Figs. 7E and 7F). Neurite bundles with vesicles of moderate to high electron density and glial cells with granules of high electron density (Fig. 7F) occur in this fibromuscular layer, frequently in the proximity of the ciliary tufts, but no evidence of intraepithelial innervation (as that seen in the gill, Rodriguez et al., 2019) could be discerned in the lung.

It should be noted that this layer of fibromuscular tissue separates the respiratory lamina from the large blood sinuses that supply the lung. Radial sinuses originating from them traverse the fibromuscular layer conveying blood to the sinus of the respiratory lamina (Figs. 1B and 1C). These radial sinuses are of varying length, but they are much narrower than the main sinuses, and in certain occasions, there are direct connections between the main blood sinuses and the blood spaces of the respiratory lamina (Fig. 1A). We found sparse endothelial-like cells lining the radial sinuses (Fig. 6).

The respiratory lamina: the ciliary tufts

Ciliated cells are the main cell type composing the tufts; they bear both long cilia and short microvilli (Fig. 8A). Secretory cells show granules with varying electron density (Fig. 8B). Some of these secretory cells show metachromatic clumps when stained with toluidine blue (presumably mucin, Fig. S2). Another type of secretory cells shows thick, globule-forming protrusions of the apical cytoplasm that detach to the pulmonary cavity (Figs. 8C and 8D; Fig. S2B), that is, as a kind of apocrine secretion. Under SEM (Fig. 4D), these globules are seen merging into larger masses. There are also microvillar cells in the tufts, which show rather long and ramified microvilli (Figs. 8A and 8B), and whose cytoplasm contains only some mitochondria, which are more abundant in the subnuclear region. These cells resemble the β-cells found in the gill epithelium of P. canaliculata (Rodriguez et al., 2019), although the cytoplasm of those is packed with mitochondria. Cell-to-cell contacts comprise both desmosome-like and septate junctions, which keep the tuft cells together, between them and with the pavement cells of the respiratory lamina.

Below the respiratory lamina: rhogocytes and storage tissue

We found rhogocytes either isolated or grouped, near the roots of the ciliary tufts and associated with radial sinuses and cells of the storage tissue (Fig. 9A) (Giraud-Billoud et al., 2008). Rhogocytes varied in size and shape and exhibited the characteristic extracellular lacunae enclosed by cytoplasmic bars separated by slits (Haszprunar, 1996; Kokkinopoulou et al., 2014) (Figs. 9B, 9C and 10). Rhogocytes were enclosed within a lamina of the extracellular matrix (Fig. 9C, arrowheads) and often surrounded by dense bundles of collagen fibers (Figs. 10A–10C). They also showed long cytoplasmic extensions that frequently run parallel to each other (Fig. 10B). Rhogocytes’ nuclei were large, with heterochromatic clumps and nucleoli (Figs. 9B, 10B, 10C and 10E); the cytoplasm showed a remarkable arrangement of both smooth and rough endoplasmic reticula (SER and RER; Fig. 10). The SER was made of rather wide tubuli containing a flocculent material and that communicated with the extracellular lacunae. Notably, the slits of these lacunae do not open directly to the surrounding extracellular fluid or the hemocoel, because rhogocytes and their cytoplasmic projections are encased by an external lamina or are in contact with dense collagen tissue (Figs. 9C and 10). The content of the RER tubuli was of an electron density similar to that of the surrounding cytoplasm (Fig. 10A). Wide cisternae formed in the course of both SER (Fig. 10E) and RER tubuli (Fig. 10B). Groups of glycogen clumps (Fig. 10A), as well as some well-delimited cytoplasmic globules, with varying sizes and electron densities (Figs. 10A, 10C and 10D), also occurred in the rhogocytes of the examined P. canaliculata individuals.

A summary of the lung structures in P. canaliculata and their proposed functions is presented in Table 1.

The lungs of the Ampullariidae in the context of the Gastropoda

Cavities regarded as lungs occur in the major gastropod clades Neritimorpha, Caenogastropoda and Heterobranchia (Ponder, Lindberg & Ponder, 2019). The majority of lung-bearing gastropods, however, are comprised within the Stylommatophora (superorder Eupulmonata) and the “basommatophora” (superorder Hygrophila) (Bouchet et al., 2017), in which the pallial cavity has acquired the role of an air-breathing organ and is partly closed by extensions of the mantle edge. In these cases, the lung and the pallial cavity are homologous structures (Ruthensteiner, 1997) and the pneumostome opens to the exterior (Fig. 11A).

In the particular case of the Ampullariidae, however, the lung is a flattened sac exhibiting distinct roof and floor regions, and a pneumostome that opens through the floor, connecting the lung cavity with the mantle cavity (Fig. 11B) (Hylton Scott, 1957). Thus, this pneumostome is not homologous to the pneumostome of other lung-bearing gastropods. Indeed, the ampullariid lung arises during development as an invagination of the mantle cavity roof, between the primordia of the gill and the osphradium, at least in the genera Pomacea (Brooks & McGlone, 1908; Koch, Winik & Castro-Vazquez, 2009), Marisa (Demian & Yousif, 1973) and Pila (Ranjah, 1942). Hence, the embryonic inner mantle epithelium lines the lung floor on both sides, but only the inner side differentiates as a “respiratory lamina” (sensu Rodriguez et al., 2018) (see Figs. 11A and 11B for comparison).

To our knowledge, and before the current study, the detailed histology of these presumptive gas exchanging areas had being adequately studied in just one tropical slug (Maina, 1989; Polytoxon robustum, Urocyclidae, as Trichotoxon copleyi, a junior synonym) and one ampullariid snail (Lutfy & Demian, 1965; Marisa cornuarietis). The study in P. robustum was made using SEM and TEM, and the inner surface of the lung (the presumptive gas exchanger) appears covered by approximately parallel ridges, which appear hollow in TEM sections and that Maina (1989) called “capillaries”. They are lined externally by pavement cells, much similar to those lining the blood respiratory sinuses in P. canaliculata. Also, the blood sinuses contain a particulate material among the hemocytes, similar to that seen in P. canaliculata. We interpret they are cell debris that the slow blood flow accumulates in the sinuses.

Instead, the study in M. cornuarietis (Lutfy & Demian, 1965) was made exclusively using light microscopy of paraffin sections, which we think adds merit to that outstanding paper, in which they accurately described the blood sinuses of the respiratory lamina but, unfortunately, they mistakenly equated the storage cells of the lung wall with blood lacunae. In fact, the cytoplasmic contents of these large cells may be extracted during histological processing, and the remaining cytoplasmic rim may give the false appearance of the wall of blood sinuses. Another important limitation of that paper is its strictly descriptive character and, consequently, the fact that the authors did not advance any functional interpretation. In both studied Ampullariidae (Lutfy & Demian, 1965; and this article), the vascularized area involves all the inner surface of the lung cavity.

The lung floor, the gill, and the adaptation of Pomacea canaliculata to air breathing

The lung roof and the lung floor receive blood from partly different origins. The lung roof receives blood from right afferents (Fig. 2E), while the lung floor receives blood from right afferents that have already traversed the gill and from left afferents that come directly (Fig. 2F). Deoxygenated blood from the visceral hump reaches the afferent branchial vein through the right pallial and rectal sinuses. That vein not only supplies the gill but, after leaving it, this vein is termed the afferent pulmonary vein (Andrews, 1965) and distributes blood to all of the lung roof and to the anterior and right side of the lung floor. Deoxygenated blood conveyed by blood sinuses from the mantle edge also reaches the lung roof through the afferent pulmonary vein. After oxygenation in the respiratory lamina of the lung roof, the blood is drained with that coming from the gill through a common efferent route, that is, the efferent pulmobranchial vein (Fig. 2A). Instead, the lung floor receives oxygenated blood from the gill leaflets through the ventral afferent pulmonary vein (Fig. 2B, right sinuses). It also receives deoxygenated blood from the left side of the body, through several slender left sinuses and from a large sinus that collects blood from the siphon and the left mantle border (Fig. 2B, left sinuses).

When the animal is submerged and the gill leaflets are deployed, oxygenated blood may reach the auricle directly through the efferent pulmobranchial vein. However, when the animal is outside of the water, the gill leaflets collapse and stick together, thus drastically reducing both the circulation through the leaflet sinuses and the available surface for gas exchange. Then, the blood would shunt through the basal leaflet sinuses (Rodriguez et al., 2019) converging into the ventral afferent pulmonary vein, which would distribute blood in the right side of the lung floor. Also, even when breathing in water, blood conveyed by the rectal and right pallial sinuses may traverse the gill leaflets, also reaching the lung floor through the ventral afferent pulmonary vein.

A possible mechanism in the evolution towards water independence, might have been the development of the ventral afferent pulmonary vein (see Figs. 1A and 2C) which, as we have commented above, would allow the shunt of blood from the gill towards the lung floor, and that it may have derived from the longitudinal division of the efferent pulmobranchial vein (Andrews, 1965). Also, Simone (2004) found the longitudinal splitting of this vein in Pomacea lineata and Pomacea sordida. Other ampullariid species this author investigated with negative results were Pomacea crosseana, Pomacea curumim, Pomacea megastoma and Pomacea scalaris, as well as Marisa planogyra and Felipponea neritiniformis (Simone, 2004). Also, Demian (1965) obtained negative results for M. cornuarietis. Admittedly, this number of species is far from representative of this highly diverse family. And also, it should be considered that only P. canaliculata (Seuffert & Martín, 2009; Seuffert & Martín, 2010) and, to some extent, P. lineata (Little, 1968) have been studied ecophysiologically. Thus, further investigations in a much larger number of species, from both morphological and functional viewpoints, would be needed to test this hypothesis.

The respiratory lamina: how it can be both thin and strong

In general, a blood–gas barrier must have both a thin and an extensive surface to facilitate gas diffusion, but it should be sturdy enough to withstand the rough work of respiratory movements and currents. Comparative studies have stressed that this is achieved by structures that combine a “soft” and a “hard part”, the former to allow gas-exchange, and the latter to keep the soft part in place, and in functional conditions (Maina & West, 2005).

In P. canaliculata, the thin gas exchange membrane (80–150 nm width, Fig. 7D) needs to be anchored to a “hard part” (Maina & West, 2005), which in this case is the fibromuscular layer (Figs. 7A–7D). This is achieved by the intercellular junctions that bind the pavement cells to the epithelial cells of the ciliary tufts, which are themselves deeply rooted in the fibromuscular layer at frequent intervals (Fig. 4A), and thereby may be acting as rivets that keep the epithelial and connective parts together (Fig. 4A). We have also occasionally observed some fibroblast cells in direct contact with the basal part of pavement cells, and that may be an additional structural component that needs to be explored further.

However, other important functions of the ciliary tufts should be those related to their secretions and to the ability of their cilia to spread them over the surface of pavement cells. Possible roles would be the agglutination of dust particles that have entered with air, and that would be expelled by the ciliary rows of the pneumostome (Fig. 5), as well as the lubrication of the delicate respiratory lamina during breathing movements. Besides that, a phospholipidic surfactant, found in the lungs or tracheoles of a wide variety of animals (Orgeig et al., 2007), has been reported in the lung of the garden snail Cornu aspersum (Stylommatophora, Helicidae) (Daniels et al., 1999). This should encourage the search for compounds with a similar function in the ciliary tufts’ secretions in P. canaliculata.

Taken together, our histological and ultrastructural findings on the respiratory organs may constitute the morphological basis to explain why P. canaliculata is an obligate air-breather.

The respiratory lamina in relation to hemocyte nodulation, rhogocytes and storage tissue

The particulate material occurring in the sinuses of the respiratory lamina is likely composed of cell debris collected when blood traverses the body vascular network. Crossing these wide spaces before reaching the lung should cause the blood to flow at low pressure and speed in the respiratory lamina. That, along with the intricacy of this structure, may promote the contact of circulating phagocytes (Cueto et al., 2015) with blood-borne microorganisms and foreign particles. In turn, this may initiate the defensive reaction of hemocyte nodulation we have shown elsewhere (Rodriguez et al., 2018). Also, both hematopoietic stem cells and proliferating cells are known to occur in the circulation of P. canaliculata (Rodriguez et al., 2020) and these cells may be attracted, or even trapped in these narrow spaces, where nodulation may start. In this way, both the kidney islets and the lung’s respiratory lamina may act as immune barriers and hematopoietic sites (Rodriguez et al., 2018).

Rhogocytes occurred near the roots of the ciliary tufts and in the storage tissue underneath the respiratory lamina. Storage cells have been proposed to accumulate glycogen (Andrews, 1965) and urates (Giraud-Billoud et al., 2008; Vega et al., 2007) in P. canaliculata or, more recently, calcium carbonate in P. maculata (Mueck, Deaton & Lee, 2020a). Instead, rhogocytes have been implicated in metal detoxification and hemocyanin synthesis (Haszprunar, 1996; Kokkinopoulou et al., 2015). Like rhogocytes, storage cells have received several names in the past, and at least one of them was shared (vesicular cells, =Blasenzellen), suggesting they have been sometimes considered the same under the light microscope. However, they are clearly different under the electron microscope, and the existence of the “slit-apparatus”, which delimits the extracellular lacunae, is distinctive for rhogocytes (Kokkinopoulou et al., 2015). In P. canaliculata, Giraud-Billoud et al. (2008) have described the location of storage cells in several organs and tissues, as well as their association with high tissue levels of uric acid. Vega et al. (2007) were able to isolate crystalloids from perivascular storage tissue of the digestive gland and found they were mainly composed of urates and protein. The close association of rhogocytes with storage cells has been intriguing to us, because we have occasionally observed rhogocytes in other tissues too, such as the aortic ampulla and the perivascular tissue of the digestive gland, and all of these organs also contain storage cells (Giraud-Billoud et al., 2008). In P. canaliculata, there is substantial evidence for a role of the accumulated uric acid in counteracting the oxidative burst that accompanies the arousal from estivation and hibernation (Giraud-Billoud et al., 2011; Giraud-Billoud et al., 2018; Giraud-Billoud et al., 2013), while Mueck, Deaton & Lee (2020a) have proposed a role of the calcareous crystalloids in buffering the lowering of pH that occurs during estivation in P. maculata.

Regarding the possible role of rhogocytes in hemocyanin synthesis, we must first say we could not discern any images of hemocyanin polymers in lung rhogocytes or in their vicinity, like those described in other gastropods (Albrecht et al., 2001; Kokkinopoulou et al., 2015). Instead, in a preceding publication (Rodriguez et al., 2018), we did observe hemocyanin polymers replenishing the lacunae of the renal hemocyte islets, even though we observed no rhogocytes in renal tissue. In the current study, a flocculent material was contained in SER vesicles and tubuli and was apparently secreted into the extracellular lacunae (Figs. 10E and 10F), and one may wonder whether this material could correspond to hemocyanin monomers or oligomers to be transported in blood to the renal hemocyte islets, where they would be polymerized. However, further research is needed to elucidate the exact pathways for the synthesis, polymerization, storage and bioavailability of hemocyanin.

The lung of P. canaliculata in the evolutionary framework of the Ampullariidae

Our knowledge of the phylogenesis of the Ampullariidae is based on morphological (Berthold, 1991), behavioral (Hayes et al., 2009) and molecular studies (Hayes, Cowie & Thiengo, 2009) that have disclosed much of the intergeneric relationships within the family. However, an all-encompassing, comparative study of the respiratory organs in Ampullariidae is still lacking. That said, some general trends might be identified with the available data within the framework of the intergeneric relationships.

The lungs of the basal, Old World ampullariid genera Afropomus and Saulea are less elaborate than those of the derived, New World genera Pomacea and Marisa (Berthold, 1988, 1991). This evolutionary trend in lung morphology may parallel the evolution of the left nuchal lobe into an extensible siphon, and of the corresponding siphonal mode of aerial respiration, which has not been reported for the basal genera. It also correlates with the oviposition behavior and the characteristics of the egg clutches in different genera (Hayes et al., 2009), as well as with the occurrence of other adaptations in the eggs related to oviposition outside water, such as a calcareous capsule and defensive perivitelline proteins (Brola et al., 2020; Dreon et al., 2013; Hayes et al., 2015; Sun et al., 2019).

Another aspect that is worth noting is the complexity of the vascular network. “Highly vascularized lungs” have been attributed by authors to species of the derived genera, though this expression lacks the needed precision. For instance, species showing large sinuses traversing the lung floor may impress the observer as being “highly vascularized”, but what really matters for respiration is the organization of the respiratory lamina. Notably, however, such structure has only been studied to some extent in M. cornuarietis (Lutfy & Demian, 1965) and, more in depth, in P. canaliculata (Rodriguez et al., 2018; and this article).

The varying thickness of the lung floor of the Ampullaridae, with a tendency to get thicker in derived species, was noted by some authors (Berthold, 1991; Simone, 2004); however, more data are needed to make meaningful comparisons. In fact, the main tissues affecting the thickness of the lung floor are (1) the thick muscle layer that underlies the mantle cavity epithelium in front of the pneumostome (see Fig. 1A), and (2) the layer of storage cells (see Figs. 1B and 1C). Only the first one may have a functional role in respiration, because its contraction or relaxation may affect the volume of the lung cavity. The second one has, at least in P. canaliculata, a role in the antioxidant protection during arousal from estivation (Giraud-Billoud et al., 2011; Giraud-Billoud et al., 2013) and hibernation (Giraud-Billoud et al., 2018).

Regarding the pneumostome, the ancestral condition among the Ampullariidae may be represented by a round, broad and thus presumably non-closable structure, placed centrally and slightly to the right of the lung floor, as it is represented in the basal genus Afropomus (Berthold, 1988). Instead, the derived condition may be characterized by a slit-like, small, and closable pneumostome, placed anteriorly and to the left, near the siphon base, as it occurs in the genera Marisa and Pomacea (Berthold, 1991). It is worth noting that in Saulea, the pneumostome is rounded but small and controlled by a muscular ring, and therefore it may represent an intermediate condition (Berthold, 1991).

The lungs of Pomacea species are likely among the largest and most elaborate within the family Ampullariidae. In P. canaliculata, the lung occupies most of the roof of the pallial cavity (Fig. 1A) and has a respiratory lamina with a prominent network of supplying and draining vessels (Figs. 1A and 2). The pneumostome is small, slit-like, and can be closed by overlapping its two lips. It is muscular and is innervated by two nerve branches from the supraesophageal ganglion (Fig. 3). As already mentioned, the musculature at the left anterior part of the lung floor is thicker than in the rest of the lung (Fig. 1A), and may be engaged in bringing the pneumostome forward (Andrews, 1965), close to the siphon base during the siphonal mode of respiration. The abundance of cilia near the pneumostome and its orderly arrangement suggest they serve the function of expelling the small particles that enter with air and are agglutinated by mucus from the ciliary tufts.

The embryonic development of the lung has been studied in representatives of three of the major clades of Ampullariidae, namely the Pila clade (Asian), and the Marisa and Pomacea clades (South American) (Hayes, Cowie & Thiengo, 2009) and this may also help to put our findings in an evolutionary framework. The lung develops in Pila globosa, P. canaliculata and M. cornuarietis as an invagination of the dorsal wall of the mantle cavity (Brooks & McGlone, 1908; Demian & Yousif, 1973; Koch, Winik & Castro-Vazquez, 2009; Ranjah, 1942). However, while the mantle cavity in Pila globosa and P. canaliculata originates at the posterior end of the embryo, long before torsion begins, it originates in M. cornuarietis after the onset of torsion as a depression on the right dorsolateral side of the embryo (Demian & Yousif, 1973). Other marked differences in the developmental timing of the respiratory organs are brought out by the findings of Demian & Yousif (1973) and Koch, Winik & Castro-Vazquez (2009) who found the lung develops after the gill and the osphradium in P. canaliculata and M. cornuarietis, in the initially narrow space between those organs, while Brooks & McGlone (1908) and Ranjah (1942) reported the lung, gill, and osphradium develop simultaneously in P. paludosa and Pila globosa. A greater asynchronicity occurs in the development of the lung in Asolene platae, which according to Tiecher, Burela & Martín (2013), was not detectable in newly hatched juveniles and only became functional two months after hatching.

Nerves controlling branchial and pulmonary respiration in P. canaliculata

The lung and the gill share innervation from both the supraesophageal and the accessory visceral ganglia. The former ganglion gives off branches that innervate the lung roof, the pneumostome, the osphradium, and each of the gill leaflets through the leaflet nerves (Fig. 3). The accessory visceral ganglion gives off several branches that innervate the lung floor and a single branch that runs all the way through the branchial base, namely the branchial base nerve (Fig. 3C).

The shared innervation as well as the interconnections between the supraesophageal and the accessory visceral ganglia might also provide the neural mechanisms for the behavioral shifts between branchial and pulmonary breathing, which is coupled with the deployment of the siphon for air breathing (Bavay, 1873, 1875; Bouvier, 1888, 1891; Fischer & Bouvier, 1890; Jourdain, 1879; Sabatier, 1879). Also, there may be local responses in the gill and lung. Responses in the gill may involve regulation of blood flow by the contraction of muscular trabeculae controlled by subepithelial innervation, and the modulation of the ionic/osmotic regulation function of the gill epithelium by intraepithelial innervation (Rodriguez et al., 2019). Such intraepithelial innervation does not occur in the lung’s respiratory lamina. Indeed, neurite bundles and glial cell processes are only found near the roots of the ciliary tufts and in the fibromuscular layer that underlies the blood sinuses of the respiratory lamina (Figs. 7E and 7F). Those neurite bundles show vesicles of moderate to high electron density and are associated with glial cells bearing larger granules like those observed in the gill leaflets of P. canaliculata (Rodriguez et al., 2019). The responses in the lung would involve the control of secretory cells in the ciliary tufts, the contraction of the lung floor, or the closure of the pneumostome. These morphological studies may pave the way to the needed experimental physiological studies.