TY - JOUR
AU1 - Yu,, Meng-Chen
AU2 - Kolbasov, Gregory, A
AU3 - Høeg, Jens, T
AU4 - Chan, Benny K, K
AB - Abstract Sponges are common in coral reefs and provide secondary habitats and shelter to a very diverse associated biota. To examine the symbiotic relationships between crustacean associates and their sponge hosts, the most important step is to collect live crustaceans and sponges for subsequent taxonomic identification as well as for larval rearing and experiments on larval biology. Using sponge-inhabiting barnacles as a model, we describe a set of collection procedures, identification methods, and laboratory-rearing systems for maintaining living barnacles and their host sponges. These methods also permit observing the behavior of the barnacle symbionts, including feeding, mating, as well as larval development and settlement, information that can be applied to the study of host-specificity, larval biology, and host selection. INTRODUCTION Sponges (Porifera) are active filter feeders that usually have complicated networks of internal chambers and channels and house a huge diversity of associated organisms, from microbes to mollusks and crustaceans (Pawlik, 1983; Klitgaard, 1991; Iseto et al., 2008; Ďuriš et al., 2011; Dahihande & Thakur, 2017; Steinert et al., 2017; Hiller & Werding, 2018). Most studies on sponge-associated crustaceans have focused on the taxonomy of adult stages based on photographs of living specimens and morphological examination of preserved specimens (MacDonald et al., 2006, 2009; Winfield et al., 2009; White & Reimer, 2012; Horká et al., 2016; Lee & Kim, 2017; Kou et al., 2018). Successful rearing of sponge-inhabiting invertebrates is important for studying the symbiosis between the hosts and their associates (Wägele, 1988). Studies on such topics as host-specificity, behavior, and larval biology involving associated crustaceans have been few (Crisp & Southward, 1961; Kolbasov, 1993; van Syoc et al., 2015) because it is difficult to 1) identify the host sponges (requiring both in-situ observation and laboratory-based examination) and their crustacean associates and 2) maintain living sponges in the laboratory and rear larvae from their associated crustaceans. Sponge barnacles, one of the most common sponge-associated crustaceans (van Syoc et al., 2015), are specialised barnacles that live within the body or embedded in the surface layer of their sponge hosts, usually located on/in the inhalant side to catch zooplankton from the sponge’s inhalant currents. The sponge’s body also protects the barnacles from predators (Ilan et al., 1999; Magnino et al., 1999; Wulff, 2006; Bell, 2008). Sponges and their associated barnacles are widely distributed in marine ecosystems, from the deep sea to shallow-water coral reefs and tide pools on rocky intertidal shores. The shells of sponge barnacles are highly variable; the shell bases can be calcareous or membranous and the outer shell surface can be smooth, pectinate, or spiny. Sponge barnacles are classified into four taxa based on the morphological features of their shells: the subfamilies Acastinae and Bryozobiinae (both in Archaeobalanidae) and the genera MembranobalanusPilsbry, 1916 (also in Archaeobalanidae, subfamily Archaeobalaninae) and PyrgospongiaAchituv & Simon-Blecher, 2006 (in Pyrgomatidae, subfamily Pyrgomatinae) (Kolbasov, 1993; van Syoc & Newman, 2010; van Syoc et al., 2015; Yu et al., 2017a, b). We use the presently established nomenclature, although several taxa such as Archaeobalanidae may not be monophyletic (Pérez-Losada et al., 2014). As examples of sponge-associated crustaceans with a potential for multi-faceted use in research, we show how sponge barnacles can be located and collected for use in studies of behavior of adult barnacles and larval biology. We also introduce the standard morphological and molecular methods used to identify sponge barnacles and their sponge hosts for host-specificity studies and we explain how recent methodological advances of various kinds may lead to breakthroughs in studies of the larval biology, including larval settlement, of these sponge-associated crustaceans. COLLECTION & PRESERVATION Collection of adult sponge barnacles and their sponge hosts Sponge barnacles can be collected by snorkeling and scuba diving in accessible depths (0−50 m). The GPS data for all collection sites should be recorded. Prior to diving, a sharp diving knife, forceps (AA grade), a large number of 50 ml collection tubes (e.g., Falcon® tubes), and waterproof paper labels (about 4 × 5 cm) with pre-printed ID-codes (i.e. numbers, hereafter referred as labels) should be prepared. The first step during collection is to determine whether a sponge actually contains barnacles. While closely observing the surface of a sponge, a diver should search for small, rounded or oval openings (Fig. 1). Sponge barnacles are filter feeders and will usually extend their cirri through these openings to catch plankton (Fig. 1I, L, O). In addition, their opercular valves can sometimes be seen through these openings (Fig. 1C, F). Figure 1. View largeDownload slide Diversity of openings of sponge barnacles on sponge surfaces and the locations of sponge barnacles. Sponge Mycale sp. (A−C) with enlargement of region with openings marking locations of barnacle Acasta sp. (B) and opercular valves of barnacle Acasta sp. (C). Sponge Xestospongia testudinaria (Lamarck, 1815) (D−F) with enlargement of region with openings of barnacle Acasta sandwichi Yu, Chan, Achituv & Kolbasov, 2017 (Yu et al., 2017a) (E), and opercular valves and shell plates of A. sandwichi as viewed within opening in sponge (F). Sponge Spheciospongia confoederata Laubenfels, 1930 (G−I) with enlargement of region with openings of barnacle Membranobalanus brachialis (Rosell, 1972) (H), also showing cirri of M. brachialis extended via one such opening (I). Sponge Callyspongia sp. (J−L) with enlargement of area with openings of Acasta sulcataLamarck, 1818 (K), and detail of one partially exposed specimen of A. sulcata embedded in sponge with cirri extended via such an opening (L). Sponge Spheciospongia vagabunda (Ridley, 1884) (M−O) with enlargement of area showing openings of barnacle Pyrgospongia stellula (Ross & Newman, 1973) (N) and detail of one specimen of P. stellula embedded in sponge with cirri extended via the opening (O). Openings of sponge barnacles and oscula of sponges indicated by arrows and arrowheads, respectively. This figure is available in colour at Journal of Crustacean Biology online. Figure 1. View largeDownload slide Diversity of openings of sponge barnacles on sponge surfaces and the locations of sponge barnacles. Sponge Mycale sp. (A−C) with enlargement of region with openings marking locations of barnacle Acasta sp. (B) and opercular valves of barnacle Acasta sp. (C). Sponge Xestospongia testudinaria (Lamarck, 1815) (D−F) with enlargement of region with openings of barnacle Acasta sandwichi Yu, Chan, Achituv & Kolbasov, 2017 (Yu et al., 2017a) (E), and opercular valves and shell plates of A. sandwichi as viewed within opening in sponge (F). Sponge Spheciospongia confoederata Laubenfels, 1930 (G−I) with enlargement of region with openings of barnacle Membranobalanus brachialis (Rosell, 1972) (H), also showing cirri of M. brachialis extended via one such opening (I). Sponge Callyspongia sp. (J−L) with enlargement of area with openings of Acasta sulcataLamarck, 1818 (K), and detail of one partially exposed specimen of A. sulcata embedded in sponge with cirri extended via such an opening (L). Sponge Spheciospongia vagabunda (Ridley, 1884) (M−O) with enlargement of area showing openings of barnacle Pyrgospongia stellula (Ross & Newman, 1973) (N) and detail of one specimen of P. stellula embedded in sponge with cirri extended via the opening (O). Openings of sponge barnacles and oscula of sponges indicated by arrows and arrowheads, respectively. This figure is available in colour at Journal of Crustacean Biology online. Accurate identification of both the sponges and their associated barnacles is obviously important for studying symbiotic relationships. Both in-situ information (color and form) and preserved specimens of both partners are essential. Before collecting any barnacles, in-situ photographs of the sponge (the whole sponge and detailed close-ups of its surface structure, oscula, and site of excision) should be taken with a flash or under adequate lighting (Figs. 1, 2). In-situ photographs of sponges should also show the sample label(s) that will be packaged, together with the collected specimens (Fig. 2C). These photographs will also be useful for recording the coloration in life and other morphological characteristics of the sponges, which may change after collection, especially after preservation. Figure 2. View largeDownload slide The collection of sponge barnacles includes selecting a sponge (A), preparing a collection tube and label (B) taking in- situ photographs with a label (C), excising the sponge barnacle using forceps or a diving knife (D, E), and placing the barnacle and voucher sponge material in a collection tube (F). Scale bars: A−F = 5 cm. This figure is available in colour at Journal of Crustacean Biology online. Figure 2. View largeDownload slide The collection of sponge barnacles includes selecting a sponge (A), preparing a collection tube and label (B) taking in- situ photographs with a label (C), excising the sponge barnacle using forceps or a diving knife (D, E), and placing the barnacle and voucher sponge material in a collection tube (F). Scale bars: A−F = 5 cm. This figure is available in colour at Journal of Crustacean Biology online. After locating a rounded or oval opening corresponding to the aperture of a sponge barnacle, the diver should feel the sponge lightly with gloved hands to locate the exact position of the barnacle. For sponges with a soft texture (e.g., species of Haliclona Grant, 1841), forceps can be used to rip the sponge open to locate the barnacle (Fig. 2D). For sponges with a hard texture (e.g., species of Xestospongia Laubenfels, 1932), a sharp diving knife can be used to excise a small cube (about 2 cm3) of sponge specimen that includes the sponge barnacle (Fig. 2E). Another piece of the host sponge with an intact outer surface layer (pinacoderm) of at least 2 cm × 2 cm should also be taken as voucher material for later identification in the laboratory by examination of spicules and microscopic examination of structure. Sponge cubes containing barnacles from the same individual sponge, along with any samples from that sponge, should all be stored in the same collection tube, together with any labels that were photographed in situ. It is advisable to place the barnacles and material from each separate sponge in a different collection tube to prevent contamination of the samples with material and spicules of other sponges and to prevent mixing labels. This arrangement will also make it easier to match specimens to photographs in the laboratory. Field-collected samples should be transported to the laboratory as quickly as possible, taking no longer than 6 h. During transport, all live samples should be kept at the same temperature as that recorded at the collecting site. Large quantities of chemicals and mucus often leak from living sponges in transit, and such materials can contribute to barnacle mortality. If long-distance transportation is necessary, each barnacle should be dissected on site free of its sponge cube using forceps, and thereafter be kept in a vial of seawater that is changed every 6 h. Sponge voucher material, however, should be preserved in a fixative or preservative (e.g., 10% formalin or 95% ethanol) before any long-distance transport. Once at the laboratory, the live material should be emptied from the collection tubes into a container with flowing seawater while taking care not to mix the samples, and the sponge specimen and some sponge barnacles from each sample should be fixed for species identification. The remaining barnacles should be extracted from their sponge cubes under seawater in a Petri dish and thereafter be kept in seawater for culturing. Such cleaning should be done with forceps under a dissecting microscope to reduce the chance of damage to the fragile barnacle shells. Preservation of adult barnacles and sponges The selection of a suitable fixative medium for sponges depends on its indented use. For taxonomic studies involving morphological and molecular analysis, it is crucial to preserve the shape and integrity of the samples and to extract good-quality DNA. There are various methods for fixing and preserving sponges. Salgado et al. (2007) evaluated the efficacy of six fixatives and four DNA extraction protocols for marine sponges. Although there are variations in the quality and quantity of DNA extracted from different protocols/fixatives, single PCR products can be successfully derived from all protocols/fixatives used. In consideration of specimen integrity, shipping procedures, convenience, and the large scale of many faunistic surveys, ethanol of 95% or greater concentration is recommended for preserving sponge samples (Addis & Peterson, 2005; Erpenbeck et al., 2005; Escobar et al., 2012; Morrow et al., 2012; Vargas et al., 2015; Plotkin et al., 2017) as also being suitable as a preservative for barnacles (Yu et al., 2017a, b; Chan et al., 2018; Jaberimanesh et al., 2019; Kim et al., 2019). CULTURE Laboratory culture and maintenance of sponge barnacles Extracted barnacles should be rinsed twice in seawater that has been filtered by syringe or membrane filter (0.22 μm) to remove all plankton as well as bacterial and protozoan contaminants. Barnacles can be cultured in 600 ml glass beakers (e.g., 600 ml Pyrex low-form Griffin beaker). Different species of barnacles should be kept in different beakers and cultured in filtered seawater (400 ml). Weak aeration should be applied at the bottom of the beaker by attaching the plastic tube of the air pump to a sterilised glass pipette via a syringe filter (0.22 μm) (Fig. 3). Figure 3. View largeDownload slide Culturing of adult sponge barnacles in the laboratory. Sponge barnacles cultured in beakers (A). A narrow slit incised with forceps in a strip of artificial sponge to house a barnacle (B). U-folded overhead-projector transparency film with small, circular holes cut into its basal part to house sponge barnacles (C). Diagram of barnacles embedded in an artificial sponge (D). Top view of barnacles mounted in a strip of artificial sponge in a beaker (E). Side views (diagram and photograph) of barnacles in culture stabilised by insertion in an artificial sponge (F, G). Diagram of sponge barnacles mounted on transparency film (H). Top view of cultured barnacles mounted in a beaker on a strip of transparency film (I). Side views (diagram and photograph) of cultured barnacles fixed in a strip of transparency film (J, K). Sponge barnacles mounted in strips of artificial sponge with cirri (in one case, also penis) extended (L−N). Sponge barnacles mounted in holes in transparency films, with cirri (in one case, also the penis) extended (O−Q). All beakers of 600 ml volume. This figure is available in colour at Journal of Crustacean Biology online. Figure 3. View largeDownload slide Culturing of adult sponge barnacles in the laboratory. Sponge barnacles cultured in beakers (A). A narrow slit incised with forceps in a strip of artificial sponge to house a barnacle (B). U-folded overhead-projector transparency film with small, circular holes cut into its basal part to house sponge barnacles (C). Diagram of barnacles embedded in an artificial sponge (D). Top view of barnacles mounted in a strip of artificial sponge in a beaker (E). Side views (diagram and photograph) of barnacles in culture stabilised by insertion in an artificial sponge (F, G). Diagram of sponge barnacles mounted on transparency film (H). Top view of cultured barnacles mounted in a beaker on a strip of transparency film (I). Side views (diagram and photograph) of cultured barnacles fixed in a strip of transparency film (J, K). Sponge barnacles mounted in strips of artificial sponge with cirri (in one case, also penis) extended (L−N). Sponge barnacles mounted in holes in transparency films, with cirri (in one case, also the penis) extended (O−Q). All beakers of 600 ml volume. This figure is available in colour at Journal of Crustacean Biology online. Artificial spongy material for aquarium filters can be used to support the base of the barnacle and stabilise its position and orientation in the beaker (Fig. 3D−G, L−N). A narrow slit to house the barnacle (the width depending on the size of the barnacle) is incised in a rectangular strip of this material (Fig. 3B, D) and the ends of the strip are curled so as to anchor the strip against the inner wall of the beaker (Fig. 3E−G). An alternative way to stabilise the barnacles is to cut a rectangular strip of overhead-projector transparency film to make a square-bottomed U-shaped fold, with small, circular holes (smaller than the diameter of the barnacles) cut into the basal side of the U to house the sponge barnacles (Fig. 3H−K, O−Q), and with the tips of the strip bent to hang from the lip of the beaker (Fig. 3C, H, J−K). The width of the strip should be similar to the radius of the beaker (Fig. 3I−K). The barnacles can be mounted close to each other to facilitate mating if needed. The culture conditions (temperature, salinity, and light-dark cycles) can be adjusted according to the collection sites. In Taiwan, the seawater temperature in the culture should be maintained at 25 °C (mean seawater temperature at the collection site), with a 12:12 h light-dark cycle (mean light-dark cycle at the collection site). Daily feeding with rotifers or brine shrimp larvae (Artemia) is recommended. The seawater should be changed every two days to prevent waste products from accumulating. The spongy material and transparency film used to support the barnacles should be changed when biofilm or algae starts to grow on them. This laboratory set-up allows for the observation of feeding behavior, cirral activity, and mating behavior of adult barnacles. Larval cultures As is true for most thoracican barnacles, sponge barnacles develop through a series of planktotrophic naupliar stages (I−VI) (Fig. 4A−G) and a final lecithotrophic cypris stage (Fig. 4H−J), involving a total of six molts (Chan et al., 2014). Figure 4. View largeDownload slide Larval stages of sponge barnacles with photopositive nauplii. Nauplii swimming into the light beam (A). The six naupliar stages of A. sulcata (B−G). Cypris larvae of Pectinoacasta sulpturata (Broch, 1931), M. brachialis, and A. sandwichi (respectively H−J). Scale bars: B−J in μm. This figure is available in colour at Journal of Crustacean Biology online. Figure 4. View largeDownload slide Larval stages of sponge barnacles with photopositive nauplii. Nauplii swimming into the light beam (A). The six naupliar stages of A. sulcata (B−G). Cypris larvae of Pectinoacasta sulpturata (Broch, 1931), M. brachialis, and A. sandwichi (respectively H−J). Scale bars: B−J in μm. This figure is available in colour at Journal of Crustacean Biology online. All equipment and materials for larval rearing should be sterilised, including glass pipettes and beakers, and the mouths of beakers should be covered with aluminum foil. Seawater of 34−36‰ salinity that has been filtered through a 0.22 μm filter to remove all plankton, bacterial, and protozoan contamination should be used. A suspension of microalgal cells 2–25 μm (e.g., Chaetoceros muelleri Lemmermann, Tisochrysis lutea Bendif & Probert (in Bendif et al., 2013), Nannochloropsis oculata (Droop) Hibberd, Skeletonema costatum (Greville) Cleve, and/or Tetraselmis chui Butcher) should be prepared as food for the nauplii (Fig. 5). The plankton provided as food should match the size of the mouth opening of larvae (Anderson, 1994; Vargas et al., 2006). Microalgal strains can be purchased from microalgae culture collection centers, such as the National Center for Marine Algae and Microbiota (Bigelow Laboratory for Ocean Sciences, East Boothbay, ME, USA). The concentration of microalgae used for culturing nauplii should be enough to make the seawater slightly cloudy (2−5 × 105 cells ml–1) to ensure adequate food supply (Moyse, 1960; Freiberger & Cologer, 1966; Tighe-Ford et al., 1970; Barker, 1976; Nunes et al., 2017). Figure 5. View largeDownload slide Cell size of microalgae Chaetoceros muelleri Lemmermann, Tisochrysis lutea Bendif & Probert, Nannochloropsis oculata (Droop) Hibberd, Skeletonema costatum (Greville) Cleve, and Tetraselmis chui Butcher, used for feeding nauplius larvae of sponge barnacles. Figure 5. View largeDownload slide Cell size of microalgae Chaetoceros muelleri Lemmermann, Tisochrysis lutea Bendif & Probert, Nannochloropsis oculata (Droop) Hibberd, Skeletonema costatum (Greville) Cleve, and Tetraselmis chui Butcher, used for feeding nauplius larvae of sponge barnacles. Adult barnacles cultured as described above often release nauplii either soon after collection or at intervals for some days afterward. Constant checks should be made for the presence of larvae in the aquarium system. These larvae are strongly photopositive (Barnes & Klepal, 1972; Yule, 1986); if a directed light source is placed close to the beaker, nauplii will swim into the light beam and can gradually be transferred into another beaker (one filled with filtered seawater) by means of a glass pipette (Tu et al., 2009; Brickner & Høeg, 2010; Liu et al., 2016). The density of nauplii in culture should be kept at approximately 250 nauplii l–1 or lower; a low density serves to maintain water quality during the period of larval rearing. Feeding with a mixture of microalgae added into the beaker should be done daily after the seawater has been changed. Twenty-four hours after the addition of food, some nauplii should be transferred by pipette into a Petri dish to determine under a microscope the instar they have reached and to observe their gut contents to confirm whether they have ingested the microalgae. The seawater in the rearing beaker should be changed daily as described above to avoid the accumulation of waste products, including the naupliar cuticles shed at each molt because these might encourage the growth of deleterious bacteria. As is also done for adult barnacles, the culture temperature for nauplii should generally be maintained at 25 °C in a 12:12 h light-dark cycle. Weak aeration should be applied by inserting the air pump’s plastic tubing into a sterilised glass pipette through a syringe filter (0.22 μm) to provide aeration at the bottom of the beaker and to keep the food suspended. After the nauplii have molted into cypris larvae, a glass pipette can be used to transfer the cyprids from the beaker into a petri dish (60 × 20 mm), where they can be maintained in filtered seawater for morphological studies and behavioral and settlement experiments (Fig. 4H−J). For scanning electron microscopy (SEM), cyprids should be placed into 30% ethanol for 2 h to relax them, followed by transfer into 70% ethanol for permanent preservation. The SEM pre-treatment process includes dehydration in a series of graded ethanol solutions (e.g., 30%, 50%, 70%, 90% and 95%, each concentration for 2 h) and infiltration in acetone for 2 h before critical-point drying with CO2. The dried cyprids are transferred by forceps and mounted onto brass stubs by means of double-sided sticky tape, sputter-coated with gold, and observed under SEM (Chan et al., 2013; Murtey & Ramasamy, 2016). Collection and culture of live sponges for larval-settlement experiments Living, healthy sponges are essential for successful larval-settlement experiments. For laboratory culture, it is advisable to collect sponges small enough for observation under the dissecting microscope. Prior to the collection by scuba diving, one should prepare a hammer, chisel, and a large plastic box with a cover (e.g., 310 × 215 × 221 mm; Lock & Lock, Seoul, South Korea). Sponges are generally attached on substrates such as coral fragments, rocks, and shells. After photographing the sponge as described above, it can be collected whole by chiseling loose the hard substrate to which it is attached (Fig. 6A). The sponge specimen should not be squeezed to minimise trauma. Using the detached bit of substrate as a handle (Fig. 6B), the diver can place the sponge into the plastic box in situ, and then close and lock the lid (Fig. 6C). One box should be prepared for each sponge specimen. All boxes must be filled with seawater with no remaining air because air bubbles that enter the chambers or water channels of the sponge will cause necrosis and adversely affect the symbiotic organisms in the sponge (Osinga et al., 1999; Hoffmann et al., 2003,Luter et al., 2011). The boxes should be moved gently to avoid shaking and damaging the sponges. Figure 6. View largeDownload slide Collection of live sponges for laboratory experiments: chiseling a piece of the hard substrate to which the sponge is attached (A); handling the substrate, not the sponge, to avoid any damage to the sponge (B); placing the sponge with its substrate and a label in a plastic box in-situ and lock the cover while the box is still underwater (note the box is fully filled with seawater) (C). Scale bars = 10 cm. This figure is available in colour at Journal of Crustacean Biology online. Figure 6. View largeDownload slide Collection of live sponges for laboratory experiments: chiseling a piece of the hard substrate to which the sponge is attached (A); handling the substrate, not the sponge, to avoid any damage to the sponge (B); placing the sponge with its substrate and a label in a plastic box in-situ and lock the cover while the box is still underwater (note the box is fully filled with seawater) (C). Scale bars = 10 cm. This figure is available in colour at Journal of Crustacean Biology online. Once the boxes with their sponges have arrived at the laboratory, the collected sponges should be transferred to a larger aquarium provided with flowing seawater. The boxes must be totally immersed in seawater before the lids are opened in order not to expose the sponges to air for even a short time. After two days, if the sponge has not rotted (detectable as a bad smell) or if it has not produced quantities of mucus, it can be transferred to a new aquarium provided with a filter system. The sponges should be fed daily with microalgae, rotifers, or brine shrimp (Artemia) while stopping the filter system for 2 h at feeding time. IDENTIFICATION Morphological analysis of sponge barnacles Shell parts (basis, wall plates, scuta, and terga) and parts of the somatic body (six pairs of cirri, penis, and oral cone with mouthparts or trophi) provide important morphological characters for identifying sponge barnacles to species (Fig. 7). Figure 7. View largeDownload slide Taxonomic features of sponge barnacles. Complete shell, including basis, wall, and operculum, of Acasta crucibasis Yu, Chan, Achituv & Kolbasov, 2017 (Yu et al, 2017a) in rostral view (A), in top view showing entire shell (B), and in lateral view (C). Disassembled shell plates of A. crucibasis after bleach treatment, including external views of scuta and terga (D), internal views of scuta and terga (E), external and internal views of rostrum (F, G), external views of rostrolaterals (H, I), internal views of rostrolaterals (J, K), external views of carinolaterals (L, M), internal views of carinolaterals (N, O), external and internal views of carina (P, Q), and external and internal views of basis (R, S). Somatic body of the same species removed from shell, in lateral view showing the cirri I to cirri VI (T). Mouthparts of same species, with inner and outer views of oral cone (U, V, respectively). Scale bars: A−S = 1 mm; T−V = 100 μm. This figure is available in colour at Journal of Crustacean Biology online. Figure 7. View largeDownload slide Taxonomic features of sponge barnacles. Complete shell, including basis, wall, and operculum, of Acasta crucibasis Yu, Chan, Achituv & Kolbasov, 2017 (Yu et al, 2017a) in rostral view (A), in top view showing entire shell (B), and in lateral view (C). Disassembled shell plates of A. crucibasis after bleach treatment, including external views of scuta and terga (D), internal views of scuta and terga (E), external and internal views of rostrum (F, G), external views of rostrolaterals (H, I), internal views of rostrolaterals (J, K), external views of carinolaterals (L, M), internal views of carinolaterals (N, O), external and internal views of carina (P, Q), and external and internal views of basis (R, S). Somatic body of the same species removed from shell, in lateral view showing the cirri I to cirri VI (T). Mouthparts of same species, with inner and outer views of oral cone (U, V, respectively). Scale bars: A−S = 1 mm; T−V = 100 μm. This figure is available in colour at Journal of Crustacean Biology online. Shell plates should be cleaned and separated to observe shell structure. Forceps can be used to carefully remove sponge material from the outer surface of the plates, which should be immersed in 2% sodium hypochlorite (household bleach) for approximately 2 h to completely digest the organic material; the shell parts should then be rinsed in purified water five times and air-dried for species identification. Shell structure can be examined directly under a dissecting microscope or shell plates can be photographed with a digital single-lens-reflex camera equipped with a macro lens. All six pairs of cirri, the penis, and the oral cone should be dissected free from the somatic body; after the removal of organic debris from them with forceps and ultrasonic cleaning for 1−3 sec, these can be mounted temporarily or permanently on glass slides for examination under a light microscope with high-definition lenses, or they can be critical-point dried and mounted on metal stubs for examination under a scanning electron microscope. Kolbasov (1993) and van Syoc & Newman (2010) provide comprehensive guides to the systematics and classification of sponge-inhabiting barnacles of the subfamilies Acastinae and Bryozobiinae, respectively. Morphological analysis of sponges The morphological identification of sponge species can be accomplished by reference to in-situ photographs, adequate information, and skeletal structure (Fig. 8). Once this data is at hand, studies of systematics and classification can begin, based at first on Hooper et al. (2002) and van Soest et al. (2019). Figure 8. View largeDownload slide In-situ photograph and cut sponge showing skeletal layers of sponge Aaptos suberitoides (Brøndsted, 1934) and sponge barnacle Euacasta sporillusDarwin, 1854 embedded within it (A), as well as location of hand-cut, cross-section perpendicular to surface of same sponge, showing skeletal components (spicules and spongin fibers) and integument (pinacoderm) (B). This figure is available in colour at Journal of Crustacean Biology online. Figure 8. View largeDownload slide In-situ photograph and cut sponge showing skeletal layers of sponge Aaptos suberitoides (Brøndsted, 1934) and sponge barnacle Euacasta sporillusDarwin, 1854 embedded within it (A), as well as location of hand-cut, cross-section perpendicular to surface of same sponge, showing skeletal components (spicules and spongin fibers) and integument (pinacoderm) (B). This figure is available in colour at Journal of Crustacean Biology online. Morphological features of a sponge, especially its color, shape, and surface microstructure, may change or be damaged during collection and fixation. All information gathered in-situ, including the physical attributes of the site (such as locality, habitat, depth), the morphological features of the sponge (shape, size, color, surface features and texture), mucus production, smell, and body structure (presence or absence of spongin fibers, spicules, a skeletal framework) are therefore essential. In-situ color photographs are especially helpful for the species identification of sponges. Details of the skeletal structures, including the arrangement of spongin fibers and the type, size, and distribution of the mineralised components, along with various kinds of microscopic observation of skeletal structures (especially spicules) and SEM of the surface structures are essential for identifying sponges (Boury-Esnault & Rützler, 1997; Jain, 2017). The sponge material should be preserved in 95% ethanol, which should be replaced more than five times to prevent dilution of the ethanol. The time between changes of ethanol is contingent upon the size and structure of the specimen. The following methods have been modified from Lim et al. (2017). To make preparations of skeletal elements and spongin fibers, thick sections of sponge material should be cut by hand perpendicular to (cross-section) and parallel to (pinacoderm layer) the sponge surface using a surgical scalpel blade. The perpendicular sections must include both the choanosome and the ectosome (see Fig. 8). A new scalpel blade should be used for every specimen to prevent cross-contamination of spicules. A xylene:phenol (1:1) solution should be used to clean and dehydrate the sections; xylene and phenol are volatile, toxic compounds that must be handled under a fume hood. Cleared sections can be mounted on glass slides in a mixture of distyrene (a polystyrene), a plasticiser (tricresyl phosphate), and xylene (DPX mounting medium), then observed under a microscope. Digestion by either bleach or acid (in a fume hood) is useful for preparing siliceous spicules. Acid digestion can provide a cleaner result, but it cannot be used with calcareous spicules. Concentrated nitric acid (2 ml) can be used to dissolve sponge material (1 cm3; including both the choanosomal and ectosomal parts of the skeleton) in a glass tube (16 × 100 mm). A test tube holder should be used in a fume hood to hold the tube with its opening directed away from the face while adding nitric acid into it with a glass pipette. The tube should be heated over an alcohol flame until the acid solution has completely evaporated, and then be allowed to cool to room temperature. The spicules should be washed three times, each time first adding ddH2O (10 ml) into the tube to wash the spicules and waiting for 5 min for the spicules to settle, then discarding the supernatant; 95% ethanol is added at the end for permanent preservation. Suspensions of cleaned spicules can be pipetted out onto brass stubs and sputter-coated with gold in order to measure spicule dimensions under SEM. Preparation of sponge (including the surface layer) for SEM involves drying it at 40 oC in an oven, sputter-coating with gold, and observing it under SEM. Recorded spicule data should include their types and size and recorded as minimum-average-maximum size based on a minimum of 25 measurements for each type unless otherwise indicated. DNA sequencing of sponge barnacles and sponges DNA can be extracted from tissues of individual specimens using a commercial DNA extraction kit (e.g., QIAamp Tissue Kit; Qiagen, Hilden, Germany) following the manufacturer’s instructions. A small piece (2.5 mm3) of barnacle muscle tissue can be used for DNA extraction. Sponges house many associated organisms, especially in their surface layer, where many microbes and algae can be found, so the material for DNA extraction should be taken from the central part of the sponge to avoid contamination. As is true for many organisms, mitochondrial 12S rRNA and cytochrome oxidase subunit (COI) genes of sponge barnacles are useful for species identification. These markers have been widely applied to discriminate other types of barnacles (Tsang et al., 2014; Lin et al., 2016; Chan et al., 2017; Yu et al., 2017b), and 28S rRNA and COI are chosen for sponges (Morrow & Cárdenas, 2015; Erpenbeck et al., 2016; Lim et al., 2017). The procedure for undertaking of polymerase chain reaction (PCR) and nucleotide-chain primers that can be used has been described elsewhere (Folmer et al., 1994; Chombard et al., 1998; Tsang et al., 2009; Morrow et al., 2012) and are listed in supplementary material Table S1 The COI primers for sponge in supplementary material Table S1 also are suitable for barnacles. PCR products should be purified using a purification kit (e.g., QIAquick PCR Purification Kit, Qiagen) and sequenced by a DNA analyzer (e.g., Applied Biosystems 3730 DNA Analyzer; Thermo Fisher Scientific, Waltham, MA, USA). The DNA sequences should be tested in a sequence database, e.g., GenBank, National Center for Biotechnology Information (Benson et al., 2005) or the Sponge Barcoding Database (Wörheide et al., 2008). SUPPLEMENTARY MATERIAL Supplementary material is available at Journal of Crustacean Biology online. S1 Table. Primer sequences and annealing temperature used for PCR amplification and sources. 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TI - Crustacean-sponge symbiosis: collecting and maintaining sponge-inhabiting barnacles (Cirripedia: Thoracica: Acastinae) for studies on host specificity and larval biology
JF - The Journal of Crustacean Biology
DO - 10.1093/jcbiol/ruz025
DA - 2019-07-24
UR - https://www.deepdyve.com/lp/oxford-university-press/crustacean-sponge-symbiosis-collecting-and-maintaining-sponge-yTvcm9dnKF
SP - 522
VL - 39
IS - 4
DP - DeepDyve
ER -