Functional neuroanatomy of the rhinophore of Archidoris pseudoargus
© Springer-Verlag and AWI 2007
Received: 4 December 2006
Accepted: 18 January 2007
Published: 15 March 2007
For sea slugs, chemosensory information represents an important sensory modality, because optical and acoustical information are limited. In the present study, we focussed on the neuroanatomy of the rhinophores and processing of olfactory stimuli in the rhinophore ganglion of Archidoris pseudoargus, belonging to the order of Nudibranchia in the subclass of Opisthobranchia. Histological techniques, fluorescent markers, and immunohistochemistry were used to analyse neuroanatomical features of the rhinophore. A large ganglion and a prominent central lymphatic channel are surrounded by longitudinal muscles. Many serotonin-immunoreactive (IR) processes were found around the centre and between the ganglion and the highly folded lobes of the rhinophore, but serotonin-IR cell bodies were absent inside the rhinophore. In contrast to the conditions recently found in Aplysia punctata, we found no evidence for the presence of olfactory glomeruli within the rhinophore. Using calcium-imaging techniques with Fura II as a calcium indicator, we found differential calcium responses in various regions within the ganglion to stimulation of the rhinophore with different amino acids. The lack of glomeruli in the rhinophores induces functional questions about processing of chemical information in the rhinophore.
Chemical signals play a prominent role in most organisms. Sea slugs, living in shallow waters at the North sea should primarily rely on chemical and mechanical senses, as optical information is limited and the sensitivity of long-range acoustic stimuli without a swim bladder or a sophisticated ear is probably low. Archidoris belongs to the order of nudibranchia, and its rhinophores were shown to be sensitive to mechanical stimulation (Agersborg 1922). Anatomical studies by Storch and Welsch (1969), for the first time, suggested similarities to the osphradium (Welsch and Storch 1969; Wedemeyer and Schild 1995) and that the posterior tentacles, the rhinophores, may serve as chemoreceptive organs. Using electrophysiological recordings and neuroanatomical backfill techniques, Bicker et al. (1982) investigated mechano- and chemoreception in Pleurobranchaea californica. In Aplysia, the rhinophore epithelium was suggested to be chemoreceptive (Audesirk 1975; Emery and Audesirk 1978), and the function of the rhinophore as an olfactory organ was described by Audesirk and Audesirk (1977). Ablation of the rhinophores of Aplysia was shown to cause a decrease in the time spent for mating and the egg laying behaviour, suggesting the sensitivity of the rhinophores for pheromones (Levy et al. 1997; Susswein and Nagle 2004; Cummins et al. 2004). In a recent study using calcium-imaging techniques, Wertz et al. (2006) showed that amino acids are detected and processed by the rhinophores of Aplysia punctata supporting their olfactory function.
Compared to vertebrates and insects, the olfactory pathway of molluscs has not been investigated deeply. In terrestrial gastropods, the importance of the procerebrum for olfactory information processing was shown (Delaney et al. 1994; Gelperin 1999; Gelperin and Tank, 1990), and the neuroanatomy and function of the cerebrum with respect to olfactory information processing was reviewed by Chase (2000). However, only little is known about the olfactory pathway from the sensory cells to higher centres. In the stylommatophoran terrestric pulmonate Achatina fulica, Chase and Tolloczko (1993) described the anatomy of the posterior tentacle and found olfactory glomeruli with similarities to glomeruli found in vertebrates and arthropods (Hildebrand and Shepherd 1997) and Croll et al. (2003) described glomeruli-like structures in the nudibranch Phestilla. Likewise, the rhinophore of A. punctata possesses a rhinophore ganglion and glomeruli, which are arranged around the rhinophore groove (Wertz et al. 2006). For the genus Archidoris, there is no information available about processing of chemosensory information or the underlying neuroanatomy within the rhinophore. In A. punctata, the rhinophore ganglion and the glomeruli are innervated by centrifugal serotonergic processes (Wertz et al. 2006). Similar to results in the opisthobranchia Pleurobranchaea and Tritonia (Moroz et al. 1997) and sensory organs in Phestilla, serotonergic cell bodies were shown to be absent in the periphery (Croll et al. 2003).
In the present study, we investigated the rhinophore of Archidoris pseudoargus using histological techniques, serotonin immunohistochemistry, and fluorescent tracers to reveal general neuroanatomical features. Responses to amino acids as potential odorants were measured in the rhinophore ganglion using fluorimetric calcium imaging. The results of this study give a first account of the chemosensory structure and function of the rhinophores of A. pseudoargus and open up new avenues for further studies of chemoreception in sea slugs. Differences with recent findings in A. punctata are discussed.
Materials and methods
Tissue preparation, immunohistochemistry, and fluorescent tracers
Specimens of A. pseudoargus were collected from shallow waters around Helgoland. Animals were of different ages and body sizes. For preparation, the animals were cooled on ice and fixed in 4% formaldehyde in artificial sea water (ASW; pH 7.5; in mM: 460 NaCl, 104 KCl, 55 MgCl, 11 CaCl2, and 15 Na–HEPES (N-2-hydroxyethylpiperazine-N’-2-ethanesulfonic acid Na-salt). Before further treatments, the rhinophores were washed three times in 0.1 M phosphate buffered saline (PBS, pH 7.2). For labelling with fluophore-conjugated phalloidin, immunohistochemistry, tracing with biocytin markers and nuclear staining, the rhinophores were embedded in 5% low-melting point agarose (Agarose II, Amresco, Solon, OH, USA, No. 210-815) and sectioned in a frontal or sagittal plane at 150 μm with a vibrating microtome (Leica VT 1000S, Wetzlar, Germany). Free-floating agarose sections were preincubated in PBS with 0.2% Triton X-100 and 2% normal goat serum (NGS: ICN, Biomedicals, Orsay, France, Cat. No. 191356) for 1 h at room temperature. Different combinations of double labellings were performed. To label serotonergic neurons, sections were incubated with a primary antibody against serotonin derived from rabbit (1:4000, DiaSorin, Stillwater, MN, USA, Cat. No. 20080, Lot No. 051007) in PBS with 0.2% Triton X-100 and 2% NGS overnight at room temperature. This antibody was used successfully in previous studies in gastropod molluscs (Croll et al. 2003; Moroz et al. 1997; Wertz et al. 2006). After five rinses in PBS, sections were incubated in Alexa Fluor 488-conjugated goat anti-rabbit secondary antibody (1:250, Molecular Probes, Eugene, OR, USA, Cat. No. A -11008). To label filamentous (f)-actin in muscles and nervous tissue, sections were incubated in 0.2 units of Alexa Fluor 568 phalloidin (Molecular probes, A-12380) in PBS overnight at 4°C. To stain cell nuclei, sections were incubated for 15 min in 25 μg/ml propidium iodide (Molecular probes, P-1304) in PBS with 0.2% Triton X-100 at room temperature. Sections were finally washed at least five times with PBS, transferred into 60% glycerol/PBS for 30 min, and mounted on microscopic slides in 80% glycerol in PBS.
Staining with biocytin
For live staining with biocytin (Molecular Probes, B1592), rhinophores were transferred to a dish containing ASW, which was then removed by a dry paper. Small amounts of biocytin crystals were applied into the tentacle nerve using a minuten pin. After 2–4 h incubation at room temperature, the rhinophores were fixed in 4% formaldehyde in ASW for 1 day at 4°C. After rinsing three times in PBS, rhinophores were embedded in agarose and sliced at 400 μm thickness with a vibratome. To visualize biocytin, slices were incubated with streptavidin conjugated with Alexa 488 Fluorophore (1:125, Molecular Probes, S11223) overnight. Following dehydration in an ascending series of ethanol, slices were cleared and mounted in methyl salicylate.
Histology and confocal microscopy
Rhinophores were fixed in Bouin’s fixative solution for 2 days, washed with ethanol, embedded in Spurr’s resin, and sectioned in the sagittal and frontal planes (6 μm). After standard histological procedures, plastic sections were stained on a hotplate after Mallory (1900). Stained sections were washed with distilled water, dried on the hotplate, and mounted in Entellan (Merck, Darmstadt, Germany). Images were taken with a digital Camera (Spotinsight Color, Vistron Systems, Puchheim, Germany) mounted on a microscope (Zeiss Axiophot, Carl Zeiss GmbH, Jena, Germany). Image processing was performed with CorelDRAW (Corel Corporation, Ottawa, Ontario, Canada).
Fluorescent tracers and antibodies were visualized with a laser-scanning confocal microscope (Leica TCS SP) using appropriate lasers, filter settings, and objectives. Image processing was performed with the following software: Zeiss Image Browser (Zeiss GmbH, Jena, Germany), Corel Photopaint and CorelDRAW Graphics Suite (Corel Corporation, Ottawa, Ontario, Canada), and Adobe Photoshop (Adobe, San Jose, USA). Three rhinophores were used for frontal sections and two for sagittal sections applying Mallory stains. Biocytin stainings were perfomed in five rhinophores and all double stainings were replicated at least three times.
Fluorimetric measurements of intracellular Ca2+ levels
The rhinophores were dissected as described above and cut longitudinally using a razor blaze. Sliced rhinophores were incubated for 60 min at 4°C in ASW containing 5 μM Fura II acetoxymethylester (AM) (Sigma-Aldrich). After removal of the incubation buffer, the rhinophores were washed for 10 min. Changes in fluorescence were monitored with an imaging system (Visitron, Puchheim) and a CCD camera (Coolsnap cF, Photometrics) mounted on an inverted microscope (Zeiss Axiovert 100) equipped with a UV objective (Zeiss NeoFluar 20×). Different regions within the rhinophore ganglion were measured using the “region” function of the software (Metafluor, Meta Imaging Series, Universal Imaging Corporation). Changes in fluorescence were obtained by ratiometric measurements with excitations at 340 and 380 nm. Values were presented as relative changes in ratios representing alterations in intracellular Ca2+ levels. Fluorescence images were acquired with an interval of 5 s and an exposure time of 50 ms per image.
For odour stimulation the recording chamber (volume 3 ml) was mounted on the microscope stage, and the bath flow was adjusted to 4 ml/min with a peristaltic pump. The chamber volume was exchanged in less than 1 min. After addition of amino acids to the source beaker the solutions reached the bath chamber after 120 ± 10 s. Amino acids, which induced the highest responses in P. californica (Bicker et al. 1982) and in previous experiments with A. punctata (Wertz et al. 2006) were chosen as olfactory stimuli (Alanine, Arginine, Glutamine, Methionine, and Isoleucine; all purchased at Sigma-Aldrich, Munich, Germany). Amino acids were applied for 1 min at various concentrations with the peristaltic pump system. Each amino acid (1 M stock each) was dissolved in ASW and final concentrations ranged between 2 and 20 mM. Stimulus solutions were prepared immediately before use by dissolving the respective stock solution in ASW. After stimulation, ASW was pumped through the recording chamber for at least 10 min to wash out all amino acids. In most cases, 50 regions of interest were measured simultaneously. To test for the viability of the preparations, the last stimulus at the end of an experiment always was a high K+ buffer stimulation (400 mM NaCl was replaced by 400 mM KCl), which elicited a strong response. Calcium-imaging experiments were performed with 15 (K+)-responsive rhinophores from eight animals using the following application of amino acids. Rhinophore 1: Alanine (Ala), Valine (Val), Histidine (His); 2: Ala, Arginine (Arg), Isoleucine (Ile), Methionine (Met), Glutamine (Gln); 3: Met, Met, Met; 4: Met, Met, Met, Met; 5. Ala, Ala, Arg; 6: Met, His, Val; 7: Ala, Ala, Ala; 7: His, Met, Ala; 8: Met, His, His; 9: Met, Ala, His; 10: Gln, Phenylalanine (Phe), Val, His; 11: Met, Arg, Gln, Gln; 12: Met, Phe, Val; 13: Met, Met, Gln, Ala; 14: repetitive (K+); 15: Ala, Met, Ala, Met. Representative experiments are shown in the figures, but not pooled for responses to specific amino acids because of differential responses.
Neuroanatomy of the rhinophore
Figure 3c–f shows double stainings with the nucleic acid marker propidium idodide and serotonin-immunoreactivity. Sagittal and frontal slices revealed many cell nuclei in the epithelium as well in and around the rhinophore ganglion. Serotonergic fibres extended from the tentacle ganglion to the lobes in the periphery and were found across the rhinophore. No serotonergic cell bodies were found within the rhinophore. The lymphatic channels (LC) were associated with serotonin-immunoreactivity (Fig. 3f) indicating that these structures are potentially innervated by 5HT IR processes. Most importantly, no glomeruli or glomerulus-like structures were found within the rhinophore or asscociated with the rhinophore ganglion of A. pseudoargus.
Calcium imaging of amino acid evoked responses within the tentacle ganglion
In Fig. 4, five regions are demonstrated which showed responses to the application of amino acids. Strongest responses were recorded to alanine and arginine. All measured regions responded with a change in intracellular calcium levels to arginine, whereas to alanine, region V did not respond clearly. In this experiment isoleucine, methionine, and glutamine induced no observable changes in intracellular calcium levels. Application of ASW containing high K+ induced a calcium level elevation in regions III, IV, and V, whereas region I responded with a calcium level decrease. In region II the calcium level decrease was followed by an increase. With application of arginine such differential responses were observed as well. In the experiment shown in Fig. 4c the same amino acids were applied. The positions of three responding regions are shown. Region I responded with a decrease of intracellular calcium levels, whereas regions II and III showed an elevation. High K+ induced a decrease in region I and an increase in II and III. In Fig. 4e, three calcium responses measured in three rhinophores are shown. An increase and a decrease of calcium levels in response to the application of alanine, isoleucine, and glutamine were measured, suggesting complex neuronal processing within the ganglion following stimulation with amino acids.
The rhinophore of A. pseudoargus comprises a prominent rhinophore ganglion. The ganglion is surrounded by a perineurial glial-like sheath. In contrast to the conditions in Aplysia (Wertz et al. 2006) and results obtained in Achatina (Chase and Tolloczko 1993) in histological investigations, serotonin-immunostaining and fluorescent tracing with biocytin did not reveal any olfactory glomeruli inside the rhinophore. In the same line, the homogeneous texture of nervous tissue inside the rhinophore ganglion gave no indications of glomerular structures inside the ganglion. Our results are corroborated by new work of the group of Klussmann-Kolb (Faller and Klussmann-Kolb 2006). The findings are in contrast to the presence of glomeruli in most vertebrate and insect primary olfactory centres (e.g. Hildebrand and Shepherd 1997; Rössler et al. 2002). Many serotonergic processes were found to innervate the rhinophore. The origin of these centrifugal serotonergic neurons is not clear, as no serotonergic cell bodies could be determined in the rhinophore, similar to results in other gastropods (Croll et al. 2003; Boudko et al. 1999; Wertz et al. 2006).
The rhinophore contains a lymphatic channel system surrounded by muscular layers, as demonstrated by phalloidin stainings of f-actin. Interestingly the lymphatic channel system appears to be associated with serotonin-IR, which may indicate that the hydraulic system for fast longitudinal movement of the tentacles is under control of serotonergic neurons. Further investigations at the ultrastructural level are needed to resolve the fine structure of potential release sites close to the lymphatic channels. We found no transversal muscles, which otherwise are necessary for expansion of the rhinophores.
Differential alterations (elevation–decrease) of intracellular calcium levels in the presence of single amino acids indicate that chemosensory input is processed in the rhinophore ganglion. The results also demonstrate that various amino acids are detected and differentially processed within the rhinophore ganglion. The lack of any obvious olfactory glomeruli in Archidoris may indicate that the rhinophore may not serve in the first line as an olfactory organ for long distance reception of odorants and may be a primarily tactile or rheotaxic organ. In favour of the rhinophores as an olfactory organ Wyeth and Willows (2006) showed the importance of predator or prey odour plumes for navigational response in Tritonia. Receptor neurons within the rhinophore epithelium of Archidoris may not be very sensitive to stimulation with amino acids. Responses to tactile stimuli and other chemical substances such as alkaloids used in Aplysia (Bickmeyer et al. 2004) need to be investigated in the future to clarify this issue.
Stimulation with alanine, isoleucine, and glutamine induced calcium elevations as well as decreases in the ganglion possibly reflecting inhibitory and excitatory influences and neuronal processing of chemosensory information in the ganglion. Another possibility could be direct inhibition and excitation of sensory cells by chemical stimuli, similar to results found in crustaceans (Michel et al. 1991) and squids (Danaceau and Lucero 1998; Lucero et al. 1992). Further studies of the sensory neurons within the rhinophore epithelium are needed to clarify these aspects of sensory reception and processing of chemical stimuli in Archidoris.
In conclusion, a large ganglion and a prominent central lymphatic channel surrounded by longitudinal muscles are present in the rhinophore. Many serotonin-immunoreactive (IR) processes but no serotonin-IR cell bodies were found inside the rhinophore. No evidence for the presence of olfactory glomeruli within the rhinophore can be presented. Different amino acids are detected by the rhinophore.
We thank the Helgoland diving group, in particular Carsten Wanke, Saskia Brandt and the late Udo Schilling for collecting Archidoris.
- Agersborg HPK (1922) Some observations on qualitative chemical and physical stimulations in nudibranchiate mollusks with special reference to the role of the ‘rhinophores’. J Exp Zool 36:423–445View ArticleGoogle Scholar
- Audesirk TE (1975) Chemoreception in Aplysia californica. I. Behavioral localization of distance chemoreceptors used in food-finding. Behav Biol 15:45–55PubMedView ArticleGoogle Scholar
- Audesirk TE, Audesirk GJ (1977) Chemoreception in Aplysia californica. 2. Electrophysiological evidence for detection of odor of conspecifics. Comp Biochem Physiol A 56:267–270View ArticleGoogle Scholar
- Bicker G, Davis WJ, Matera EM (1982). Chemoreception and mechanoreception in the gastropod mollusk Pleurobranchaea californica. 2. Neuroanatomical and intracellular analysis of afferent pathways. J Comp Physiol 149:235–250View ArticleGoogle Scholar
- Bicker G, Davis WJ, Matera EM, Kovac MP, Stormogipson DJ (1982) Chemoreception and mechanoreception in the gastropod mollusk Pleurobranchaea californica. 1. Extracellular analysis of afferent pathways. J Comp Physiol 149:221–234View ArticleGoogle Scholar
- Bickmeyer U, Drechsler C, Köck M, Assmann M (2004) Brominated pyrrole alkaloids from marine Agelas sponges reduce depolarization-induced cellular calcium elevation. Toxicon 44:45–51PubMedView ArticleGoogle Scholar
- Boudko DY, Switzer-Dunlap M, Hadfield MG (1999) Cellular and subcellular structure of anterior sensory pathways in Phestilla sibogae (Gastropoda, Nudibranchia). J Comp Neurol 403:39–52PubMedView ArticleGoogle Scholar
- Chase R (2000) Structure and function in the cerebral ganglion. Microsc Res Techn 49:511–520View ArticleGoogle Scholar
- Chase R, Tolloczko B (1993) Tracing neural pathways in snail olfaction—from the tip of the tentacles to the brain and beyond. Microsc Res Techn 24:214–230View ArticleGoogle Scholar
- Croll RP, Boudko DY, Pires A, Hadfield MG (2003) Transmitter contents of cells and fibers in the cephalic sensory organs of the gastropod mollusc Phestilla sibogae. Cell Tiss Res 314:437–448View ArticleGoogle Scholar
- Cummins SF, Nichols AE, Rajarathnam K, Nagle GT (2004) A conserved heptapeptide sequence in the waterborne attractin pheromone stimulates mate attraction in Aplysia. Peptides 25:185–189PubMedView ArticleGoogle Scholar
- Danaceau JP, Lucero MT (1998) Betaine activates a hyperpolarizing chloride conductance in squid olfactory receptor neurons. Journal of Comparative Physiology A-Sensory Neural and Behavioral Physiology 183:225–235View ArticleGoogle Scholar
- Delaney KR, Gelperin A, Fee MS, Flores JA, Gervais R, Tank DW, Kleinfeld D (1994) Waves and stimulus-modulated dynamics in an oscillating olfactory network. Proc Natl Acad Sci U S A 91:669–673PubMedView ArticleGoogle Scholar
- Emery DG, Audesirk TE (1978) Sensory cells in Aplysia. J Neurobiol 9:173–179PubMedView ArticleGoogle Scholar
- Faller S, Klussmann-Kolb A (2006) Transmitter distribution in the cephalic sensory organs of Ophistobranchia. In: 2nd international ophistobranch workshop (Bonn) Abstract p 21Google Scholar
- Gelperin A (1999) Oscillatory dynamics and information processing in olfactory systems. J Exp Biol 202:1855–1864PubMedGoogle Scholar
- Gelperin A, Tank DW (1990) Odor-modulated collective network oscillations of olfactory interneurons in a terrestrial mollusk. Nature 345:437–440PubMedView ArticleGoogle Scholar
- Hildebrand JG, Shepherd GM (1997) Mechanisms of olfactory discrimination: converging evidence for common principles across phyla. Annu Rev Neurosci 20:595–631PubMedView ArticleGoogle Scholar
- Levy M, Blumberg S, Susswein AJ (1997) The rhinophores sense pheromones regulating multiple behaviors in Aplysia fasciata. Neurosci Lett 225:113–116PubMedView ArticleGoogle Scholar
- Lucero MT, Horrigan FT, Gilly WF (1992) Electrical responses to chemical-stimulation of squid olfactory receptor-cells. J Exp Biol 162:231–249Google Scholar
- Mallory FB (1900) A contribution to staining methods: I. A differential stain for connective-tissue fibrillae and reticulum. II. Chloride of iron haematoxylin for nuclei and fibrin. III. Phosphotungstic acid haematoxylin for neuroglia fibres. J Exp Med 15–20Google Scholar
- Michel WC, Mcclintock TS, Ache BW (1991) Inhibition of lobster olfactory receptor-cells by an odor-activated potassium conductance. J Neurophysiol 65:446–453PubMedGoogle Scholar
- Moroz LL, Sudlow LC, Jing J, Gillette R (1997) Serotonin-immunoreactivity in peripheral tissues of the opisthobranch molluscs Pleurobranchaea californica and Tritonia diomedea. J Comp Neurol 382:176–188PubMedView ArticleGoogle Scholar
- Murphy BF, Hadfield MG (1997) Chemoreception in the nudibranch gastropod Phestilla sibogae. Comp Biochem Physiol A 118:727–735View ArticleGoogle Scholar
- Rössler W, Kuduz J, Schurmann FW, Schild D (2002) Aggregation of F-actin in olfactory glomeruli: a common feature of glomeruli across phyla. Chem Senses 27:803–810PubMedView ArticleGoogle Scholar
- Storch V, Welsch U (1969) Cytology and function of the nudibranch rhinophores. Z Zellforsch 97:528–536PubMedView ArticleGoogle Scholar
- Susswein AJ, Nagle GT (2004) Peptide and protein pheromones in molluscs. Peptides 25:1523–1530PubMedView ArticleGoogle Scholar
- Wedemeyer H, Schild D (1995) Chemosensitivity of the osphradium of the pond snail Lymnaea stagnalis. J Exp Biol 198:1743–1754PubMedGoogle Scholar
- Welsch U, Storch V (1969) Osphradium of prosobranch gastropods Buccinum undatum and Neptunea antiqua. Z Zellforsch 95:317–330PubMedView ArticleGoogle Scholar
- Wertz A, Rössler W, Obermayer M, Bickmeyer U (2006) Functional neuroanatomy of the rhinophore of Aplysia punctata. Front Zool 3:6PubMedView ArticleGoogle Scholar
- Wyeth RC, Willows AO (2006) Odours detected by rhinophores mediate orientation to flow in the nudibranch mollusc, Tritonia diomedea. J Exp Biol 209:1441–1453PubMedView ArticleGoogle Scholar