Collecting and Culturing Bryozoans for Regenerative Studies

Abstract

Among marine invertebrates, bryozoans are small, not well known, and complex to identify. Nevertheless, they offer unique opportunities for whole-body generation research, because of their colonial, modular mode of growth. Here, we describe detailed methods for collection of bryozoans from a range of environments, sample preparation and identification, culture and feeding, spawning and breeding, marking colonies for growth studies, and histological preparation.

1 Introduction

The Bryozoa (moss animals) is a diverse phylum of colonial aquatic invertebrates found in almost all freshwater and marine environments. The phylum comprises ~6000 living species [1] which grow into a bewildering array of colony types, including soft (weedy or gelatinous) and hard (calcified) forms, which may be moss-, sponge-, or coral-like in overall appearance. Numerous taxa grow as thin crusts or delicate lace-like encrustations over suitable substrates (Fig. 1) [2]. Although often overlooked, bryozoans are often among the most diverse and abundant members of marine communities, especially in the Southern Hemisphere. All bryozoans are suspension feeders, extracting small food particles from the water column, and colonies typically live attached to seafloor substrates (e.g., shells, rocks, algae) or on surfaces in freshwater ponds, rivers, and lakes [3]. Three of the main extant clades are the freshwater Phylactolaemata, the marine Stenolaemata, and the predominantly marine Gymnolaemata (Fig. 2). All three offer possibilities for the study of WBR and related phenomena.

Fig. 1

Morphology of bryozoans. (a) An encrusting colony of the marine cheilostome bryozoan Watersipora subatra. (b) SEM image of the calcified autozooids of a Microporella discors colony (marine Cheilostomatida) showing ~6 polygonal autozooids; black arrowhead—autozooidal aperture; white arrowhead—avicularium (defensive polymorphic zooid). (c) Part of a living colony of the marine cheilostome Hippomenella vellicata showing feeding autozooids with extended lophophores (top); retracted autozooids with closed lid-like opercula (middle); and developing asexually budded autozooids at the colony margin (bottom). (d) Living colony of Hastingsia sp. This well-calcified continental shelf cyclostome was successfully grown in a laboratory culture system using natural seawater supplemented by cultured microalgae. (e) Two polypide regression products (brown bodies), indicated with white arrowheads; the adjacent zooidal chamber contains a developing polypide that will replace the previous polypide, which has degenerated (Hornera sp., marine cyclostomate, H&E stained). (f) Large colony of the gelatinous freshwater phylactolaemate Pectinatella magnifica. (g) Living colony of Cristatella mucedo, a freshwater phylactolaemate bryozoan; several rows of horseshoe-shaped lophophores line the periphery of the colony, which is capable of creeping along the substratum. (h) Statoblast (asexually produced resting propagule) of the freshwater bryozoan Plumatella cf. geimermassardi (fixed but unstained whole mount, imaged under compound microscope). Scale bars: a, 10 mm; b, 200 μm; c, 1 mm; d, 1 mm; e, 50 μm; f, 5 cm; g, 1 mm; h, 100 μm

Fig. 2

Generalized phylogeny and relationships of the phylum Bryozoa (includes only extant taxa)

Bryozoan colonies are composed of iterated (mostly) submillimeter animals called zooids, which are budded as asexual clones from a single founder zooid, the ancestrula, itself derived from a free-swimming larva [4]. Depending on the species, a single colony may contain several to many hundreds of thousands of zooids. Autozooids are the zooid polymorphs responsible for feeding within a bryozoan colony; each has a lophophore bearing a crown of ciliated tentacles that captures microscopic food particles [5], typically microalgae. This feeding apparatus is normally extended into the water column on a flexible sheath, but can be retracted into a protective box-like or tubular zooid chamber, which may be gelatinous, leathery, or rigid in marine species that secrete a calcified skeleton [3]. The remaining parts of an autozooid include the polypide (comprising the lophophore, u-shaped unidirectional gut, a ganglion, and retractor muscles), and the cystid, the living and nonliving structural parts of the body wall (Fig. 3) [5]. Species identification of bryozoans often relies upon examination of the individual zooid architecture, and commonly requires the use of a dissecting microscope.

Fig. 3

Generalized anatomy of a cheilostomate bryozoan individual zooid. Scale: zooids generally range from 0.1 to 1.0 mm in length

Zooids are physiologically interconnected via tissue strands (funiculus) which pass through pores in shared body walls, or via shared body cavities in budding zones [6, 7]. Autozooids possess variable degrees of physiological integration within the colony, while retaining a basic functional autonomy. Nonfeeding polymorphic zooids are common in marine bryozoans, and include reproductive, defensive, and structural modules [8] that rely on autozooids for nutrition. Bryozoans may undergo seasonal sexual reproduction, while asexual budding occurs all year. Freshwater and a few marine bryozoans produce clonal resting propagules (statoblasts and hibernacula) containing stem cells [9].

This group offers unique opportunities for whole-body regeneration (WBR) research, but has been underutilized compared to other invertebrate models. Studies of WBR in this phylum could focus on zooid-scale processes in the context of the whole colony. Autozooids undergo agametic cloning to produce new zooids by budding, resulting in new colony growth, and subsequently undergo one or more polypide degeneration–regeneration cycles, which replace senescent polypides within existing zooids of the colony. The latter process occurs throughout the functional life of a zooid and involves full breakdown of the incumbent polypide into a residual “brown body,” and development of a polypide replacement, which arises from a blastema on the cystid [10]. Individual polypides typically last 1–10 weeks before regression commences, and the regression phase lasts 3–20 days, depending on the species [10].

Little is known of bryozoan regenerative processes at the subzooidal scale, for example, following partial injury to a polypide, but both WBR and body organ/tissue (= “structure”) regeneration has been reported for this phylum by Bely & Nyberg [11]. In Cristatella, at least, surgical damage to zooids is repaired rapidly without apparent damage. It is relatively easy to surgically separate living colonies into multiple ramets (clonal subcolonies), which will heal and continue to grow by budding [12]. Some bryozoans (e.g., Cupuladria exfragminis) are known to naturally autofragment as a clonal propagation strategy [13]. Bryozoans usually maintain budding along a given body axis during normal growth [14], but some taxa can undergo reversed-polarity budding and lateral budding during repair of individual zooids or during regeneration of mechanically broken branches [15]. At the colony scale, reversed-polarity budding can happen following breakage in branching forms. The precise extent of WBR in bryozoans remains to be determined.

In this chapter we present methods to study bryozoans, starting with how to find and collect bryozoans. Intertidal and shallow subtidal bryozoans can be scraped off rocks, picked from macroalgae or collected by divers, whereas shelf and deep-sea bryozoans are commonly collected via dredge or grab sampling. Preliminary on-board classification and sorting of bryozoans is achieved using overall colony form and colour, but proper species determination requires either light microscopy (difficult but can be nondestructive if done carefully) or scanning electron microscopy (lethal). Keeping living bryozoans in tanks requires careful preparation and maintenance of water quality, regular feeding of mixed phytoplankton cultures and, in some cases, regular gentle cleaning of colonies. Some encrusting bryozoans grow well in culture, but many are capable of surviving a long time without growing at all. It is possible to spawn at least some species of bryozoans, settle larvae, and raise colonies in a laboratory setting. Growth in bryozoans can be difficult to ascertain [16], but nontoxic marking of colonies using Calcein can be effective [17]. Deeper study into the structure of soft-parts (histology) and hard parts (micro-CT, SEM) allows for evaluation of life-cycle and effect of experimentation. In this contribution we provide the basic techniques for bryozoan collection , culture, and maintenance.

2 Materials

Not all materials presented in this section are required for every study. Researchers will need to choose the required materials based on their project specifications.

2.1 Colony Collection

• 1. Cool bag or insulated box for transport.

• 2. Airtight container(s) with lids, 1–3 L capacity.

• 3. Frozen cold-packs.

• 4. Thin wet protective layer (e.g., fresh seaweed, damp blotting paper).

• 5. Sampling kit: pocket knife, chisel, forceps, dropper.

• 6. Recording equipment: log book, waterproof paper.

• 7. Imaging equipment: camera with macrolens, scale bar, 10 × 10 cm black velvet.

• 8. Hand lens.

• 9. Collection permits (if necessary).

• 10. Underwater sampling equipment: dredge, grab sampler, grapnel.

• 11. Sorting and storage equipment: trays, 2 cm deep, about 30 × 20 cm², buckets, padding material (polystyrene or Styrofoam).

• 12. 95% analytical grade ethanol in dispensing bottle, triple-contained, except when in use.

• 13. 6–10 shallow trays.

• 14. Cool freshly collected seawater.

• 15. Small lab equipment: tweezers, scissors, pencil.

• 16. Blotting paper.

2.2 Identification of Bryozoans

• 1. Small fragments of dried bryozoans.

• 2. 12% NaClO solution: commercially available bleach.

• 3. 0.6% (w/v) bleaching solution: 5 mL 12% NaClO solution in 95 mL water.

• 4. 2.5 cm diameter mounting stubs for scanning electron microscope.

• 5. Mount-holding tweezers.

• 6. Lint-free cotton gloves.

• 7. 1 cm-wide double-sided carbon tape.

• 8. Silver paint.

• 9. Thin paintbrush.

• 10. Spray air duster.

• 11. Pick or fine tweezers.

• 12. Microscope.

• 13. Fine black indelible pen.

• 14. Sputter coater with gold-palladium source.

• 15. Scanning electron microscope (SEM).

2.3 Culture and Feeding

• 1. Tanks (e.g., glass, acrylic, PVC or opaque plastic).

• 2. Water circulation systems: pumps, hoses, spigots.

• 3. Aeration or bubble-free stirrers (not in all cases).

• 4. Residual-current device (RCD).

• 5. Temperature logger.

• 6. Temperature control mechanism (e.g., controller + heaters, chillers, or both).

• 7. Water supply (e.g., flow-through, recirculating or semiclosed recirculating) for filtered or artificial seawater.

• 8. Food supply (e.g., natural or cultured sources of microalgae).

• 9. Cell counter.

• 10. Tank cleaning tools (e.g., large-bore pipette).

• 11. Growth media for culture food and/or bryozoans.

• 12. Colony-cleaning tools: ultrafine brushes, disposable pipettes.

• 13. Black plastic containers (2–5 L).

• 14. Black plastic sheets.

• 15. Material for settlement substrates (e.g., acetate or glass slides).

• 16. Attachment system for substrates (rails, clips, etc.; avoid metals, especially copper).

• 17. Glass tank.

• 18. Intense light source (e.g., fluorescent or incandescent).

• 19. Disposable 3 mL pipette.

2.4 Marking

• 1. Living calcified bryozoans.

• 2. 10 L tank with bubbler.

• 3. Seawater at same temperature as bryozoans in squeeze bottle.

• 4. Sheets of black plastic.

• 5. Calcein solution: C₃₀H₂₆N₂O₁₃ (Fluorexon) at 50 to 150 mg/L in seawater (see Note 1).

• 6. 5% (v/v) diluted bleach: 417 mL 12% NaClO solution in 583 mL water.

• 7. Fluorescent light-source at wavelength 495 nm (see Note 2).

2.5 Anesthetizing and Fixing for Histology

Solutions should be prepared using ultrapure water and analytical grade reagents. Solutions should be stored at room temperature unless otherwise stated. Chemicals used in this section, with a single exception of MgCl₂, are toxic. Use fume hood for preparation and store tightly stoppered.

• 1. MgCl₂ solution: 73.2 g MgCl₂•6H₂O in 1 L H₂O.

• 2. Filtered sea water (FSW): 20 μm filtered fresh sea water.

• 3. Marine buffered formalin: 40 mL formalin, 100 g chalk, 960 mL FSW. Prepare 2 days before use.

• 4. 1–3 mL pipette.

• 5. 4% buffered formalin: 40 mL formalin, 100 g chalk, 960 mL H₂O. Prepare 2 days before use.

• 6. 0.1 M NaOH.

• 7. 16% PFA stock solution: Stir 70 mL distilled water at 60 °C with 16 g paraformaldehyde (PFA) for 1–2 h. Add one drop of 0.1 M NaOH at a time until the solution stops being opalescent. Bring the volume to 100 mL with distilled water.

• 8. 0.2 M phosphate buffered saline (PBS): one commercially available 2 g tablet in 200 mL purified water. Dissolve for 10 min to achieve a pH of 7.4.

• 9. 4% PFA fixative: 50% (v/v) 0.2 M PBS, 25% (v/v) 16% PFA stock solution (see Note 3).

• 10. PFA–glutaraldehyde (GA) mixture in buffer: 5 mL commercially available 25% GA, 12.5 mL 16% PFA stock solution, 25 mL 0.2 M PBS, 7.5 mL distilled water (see Note 4). Store refrigerated and use within days of preparation.

3 Methods

3.1 Onshore Collection

Bryozoan colonies are very diverse in terms of external appearance (Fig. 1). Encrusting bryozoans are generally small roundish patches, 1–3 cm in diameter, and variable in colour. When poked, they are predominantly hard to the touch, but some species are filamentous, weedy or gelatinous. It is easy to confuse these bryozoans with patches of algae—examination with a hand lens will show the regular openings (like pinpricks) on the surface of bryozoans, whereas most algae are smooth. Bryozoans prefer undersides of hard substrate, while algae need sunlight. Erect bryozoans tend to occur in deeper waters, and come in a multitude of shapes: fans, nets, fingers, trees, feathers. They can be confused with corals, macroalgae. Again, if they are hard/rigid and covered with small openings, found on undersides and overhangs, they are likely to be bryozoans.

• 1. Select an appropriate sampling site (see Note 5).

• 2. Record location details in log book, including time, date, latitude, longitude, water depth and relevant features or observations.

• 3. Fill an airtight container with water from the sampling site.

• 4. Scout around looking for orangish erect or encrusting roundish patches.

• 5. Look at piers, piles, bottles, cans, pieces of plastic and other human-made items.

• 6. Photograph specimens in situ, using macro lens and a ruler or scale bar.

• 7. Collect specimens with their substrate if possible (see Note 6).

• 8. Sample animals attached to large objects by gently scraping them off the surface using tools from the sampling kit (see Note 7).

• 9. Place the specimens into airtight container, covered with seaweed or blotting paper.

• 10. Repeat steps 3 to 9 until sufficient colonies are collected.

• 11. Add a paper label indicating the sampling location into airtight containers (see Note 8).

• 12. Transport the colonies back to the lab within 3 h if possible (see Note 9).

3.2 Offshore Collection

• 1. Sail to a sampling location of interest (see Note 10).

• 2. Once on station, record location details in log book, including time, date, latitude, longitude, water depth, relevant features or observations.

• 3. Deploy a dredge, grab or grapnel (see Note 11).

• 4. Turn on deck hose (see Note 12).

• 5. Retrieve collection device without dumping its content on the deck.

• 6. Gently run deck hose at low pressure over top surface of the contents.

• 7. Follow steps 7 to 9 of Subheading 3.1, choosing complete unbroken colonies (see Note 13).

• 8. Dump contents onto the deck.

• 9. Photograph the material on deck with a label indicating location number (see Note 14).

• 10. Follow steps 7 to 11 of Subheading 3.1.

• 11. Keep living colonies, cool, wet and aerated until able to sort them out (Subheading 3.3).

• 12. Store colonies for molecular analysis in ethanol, triple-contained while on boat and in transit.

3.3 Postcollection Sorting

• 1. Fill sorting trays with cool water from the sampling site.

• 2. Transfer morphologically similar samples from the transport container into the sorting trays using tweezers, one group per tray (see Note 15).

• 3. Repeat steps 1 and 2 until the contents of the transport container has been processed.

• 4. Examine specimens with hand lens to ensure each tray contains similar-looking specimens.

• 5. Prepare two location labels for each group of bryozoans with pencil and small pieces of waterproof paper (see Note 16).

• 6. Place one location label inside each tray.

• 7. Transfer one complete representative colony onto a piece of black velvet and blot dry.

• 8. Place the second location label next to the sample.

• 9. Place a ruler or scale bar next to the sample.

• 10. Take one or more pictures with the camera using macro lens, including the label and scale bar in each photo.

• 11. Place this sample and its paper label in a labeled dry 15 mL tube for identification using SEM.

• 12. Place the SEM tube into a plastic bag labeled with sample location number and the note “For SEM.”

• 13. Transfer a second representative colony into a labeled 10 mL tube filled with 95% AR grade ethanol for Genetic Archive.

• 14. Place the Genetic Archive sample into a plastic bag labeled with sample location number and the note “Genetic Archive.”

• 15. Transfer remaining colonies and the paper location label into a labeled 50 mL tube filled with clean water from sample locations.

• 16. Place the container into a cool and dark container.

• 17. Repeat steps 4 to 16 until all sorting trays have been processed.

• 18. Place the bag for Genetic Archive into a second plastic bag.

• 19. Store this Genetic Archive at 4 °C.

• 20. Transfer the live colonies from their seawater container into the aquarium system within the next hour.

3.4 Identification

Bryozoans are difficult to identify in the field and in hand specimen. There are many species which are superficially similar. Acquiring a bryozoan expert for confirming IDs is an excellent plan. If you need to send images for ID, scanning electron microscope images are best. Methodical comparison of SEM photos with species descriptions in specialist literature is, unfortunately, the only reliable way to identify bryozoans.

• 1. Pick one SEM tube from the plastic bag.

• 2. Transfer the sample into a plastic dish using a pair of tweezers.

• 3. Divide the specimen in half.

• 4. If the specimen is rigid and heavily calcified, soak one half in bleaching solution for 1 h. If it is soft, fluffy, goopy, or pliable, omit this step.

• 5. Rinse the bleached sample for a few seconds with tap water.

• 6. Transfer both halves of the colony onto a labeled glass dish.

• 7. Warm the glass dish to no more than 60 °C.

• 8. Wait 1 h for the samples to dry slowly.

• 9. Cut two pieces of double-sided carbon tape, each 2.5 cm long.

• 10. Wear fabric gloves.

• 11. Label a mounting stub on the bottom with the location (on the label in the tube) using indelible pen.

• 12. Hold the mounting stub using mount-holding tweezers (see Note 17).

• 13. Stick one piece of tape down to cover half of the mounting stub, without touching the mounting stub.

• 14. Stick the second piece of tape down so that there is a small gap between the two tapes, running across the middle of the stub surface.

• 15. Under binocular microscope, position the bryozoan specimens on either side of the seam (see Note 18).

• 16. Use spray air duster to remove any dust or crumbs..

• 17. Use silver paint to fill any gaps under large specimens, or to connect any loose branches to the carbon tape.

• 18. Allow 10 min to dry.

• 19. Coat the specimen with a thin layer of metal conductor (e.g., C, Au:Pd) using sputter coater according to instructions (see Note 19).

• 20. Draw the stub in the Notebook, showing the orientation and shape of each fragment relative to the seam, and label them.

• 21. Load the stub(s) into the SEM and pump down.

• 22. Modify your working distance, focus, and contrast to obtain high-contrast well-focused images (see Note 20).

• 23. For each species, take a photomicrograph of three different areas at magnification of 30×, making sure to show numerous zooids and heterozooids and their orientation to each other.

• 24. Take photomicrographs of ten single zooids at a magnification of 100–150× (so that the zooid fills the photo). Include both autozooids and heterozooids (see Note 21).

• 25. Every SEM micrograph should be recorded in the notebook, with photo date, photo number, specimen details, voltage, working distance and magnification.

• 26. Download photos onto flash drive and name each file with the photo number and date.

3.5 Culture and Feeding

Bryozoans are not easy to culture, so careful planning needs to be undertaken to design and run a laboratory-based culturing set-up (see Note 22). Useful summaries for culturing of bryozoans include [15] for marine and freshwater species, and [18] for freshwater species.

• 1. Select species to be cultured (see Note 23).

• 2. Develop a multitank system, based on environmental parameters (e.g., light levels, circulations, aeration, temperature) where species is found naturally (see Note 24). Examples of multitank systems are given in Fig. 4.

• 3. Set up the tank and allow it to run without bryozoans for at least 48 h.

• 4. Decide on a feeding strategy (see Note 25), and ensure a good supply of the food source.

• 5. Collect living bryozoans (see Subheadings 3.1 and 3.2).

• 6. Position colonies carefully in the tank, away from aeration systems (see Note 26).

• 7. Establish feeding and tank maintenance schedule for 2 weeks before beginning experiments or measurements (see Note 27).

Fig. 4

Tank-based culture systems for bryozoans. (a) Short-term culture system using individual tanks. (b) Flow-through tank system. (c) Slow flow-through system. (d) Recirculating system. The option of a semirecirculating system is also shown with a red arrow. BF biological filter, F filter sock, H/C heating/cooling system, L light source, MF mechanical filter, Phyt. R phytoplankton room/area, pH pH meter, P pump. Colors: red = aeration system, blue = sea water inflow, brown: sea water outflow or waste water, yellow and light blue: water purification steps, green: phytoplankton as food

3.6 Spawning

• 1. Find out what kind of larvae is produced by the species you wish to spawn (see Note 28).

• 2. Keep colonies in seawater, undisturbed at a constant temperature in the dark for at least 24 h prior to spawning.

• 3. Induce spawning in marine bryozoans by sudden exposure to bright light; freshwater species will spawn overnight, but must be watched as the larvae can be very short-lived. The time until larval release can vary between 5 and 60 min depending on the species [19,20,21,22,23,24,25] (see Note 29).

• 4. Collect larvae by gentle pipetting and transfer to prepared aquarium set-up or experiment (see Note 30). Provide many different substrate options (see Note 28), as they may settle fairly randomly when competent.

3.7 Marking Colonies

Calcein is taken up during calcification and retained in the skeleton, so this technique is useful for growth or skeletal regeneration studies in calcified marine bryozoans (see Note 31).

• 1. Fill tank (leaving headspace) with 8 L Calcein in seawater solution (see Note 1).

• 2. Adjust the water temperature to the one of the tank where bryozoans have been living.

• 3. Add bubbler for aeration and circulation.

• 4. Gently place well-fed living bryozoans in Calcein tank, taking care that bubbles do not disturb or damage them (see Note 32).

• 5. Wrap tank with black plastic or keep in dark room for 8 to 24 h.

• 6. Gently rinse colonies in seawater at the same temperature and then place back in seawater culture tank.

• 7. When ready to measure growth rate, kill bryozoans in 5% bleach solution for 1 h.

• 8. Rinse in freshwater.

• 9. Dry gently in oven or under light at <60 °C.

• 10. Examine under fluorescence microscope at 495 nm wavelength.

• 11. Measure growth since marking date as distance from bright glowing band (Calcein mark) to growth edge.

3.8 Anesthetizing and Fixing for Histology

The process of relaxing the colony in such a way that the lophophores can be fixed in a protruded position needs to be controlled under the stereomicroscope, with minimal disturbance to the colony. A typical relaxation interval should not exceed 2–2.5 h.

• 1. Place the living colony in a container with 3/4 sea water and 1/4 MgCl₂ solution. The ratio of the colony to medium volume need not be large, indeed a smaller amount of medium is easier to replace and requires less relaxant.

• 2. After the polypides begin to protrude, gradually replace the medium: with a pipette gently withdraw a small (~1–3 mL) amount of the medium and slowly replace it with the same volume of MgCl₂ solution every 2–5 min, depending on the initial volume.

• 3. Check if the colony is relaxed, by giving the extended tentacles a gentle tap with a dissection needle or gently shake the container (see Note 33).

• 4. If the lophophores have lost sensitivity, wait an additional 10–15 min for MgCl₂ to further penetrate the polypides and immobilize the retractor muscles. It is common for a partly anesthetized polypide to lose sensitivity to touch, yet withdraw on contact with the fixative.

• 5. Briefly lift the whole colony and then return it back into the container. If the polypides remain everted, the colony is ready for fixation .

• 6. Fix specimens for paraffin-based histology in 4% buffered formalin at room temperature in a fume cupboard. The volume ratio of the material to medium should be no more than 1 to 10. Store at room temperature (see Note 34).

• 7. Fix specimens for immunocytochemistry using 4% PFA in buffer at room temperature for a minimum of 4 h, or at 4 °C overnight. The volume of the fixative should be 10 or more times larger than the volume of the sample. Remove colonies from the fixative within 10 days and store refrigerated in the same, but osmolarity-adjusted buffer.

• 8. Fix specimens for electron microscopy using GA or PFA-GA in buffer at room temperature.

• 9. Rinse specimens after fixing.

• 10. Store at +4 °C. Samples are best used within a month, but can be stored in the fixative for longer if needed (see Note 35).

• 11. Decalcify the specimen (if it is calcified) (see Note 36).

• 12. For subsequent processing stages please refer to methodology-specific protocols.

4 Notes

• 1. Concentrations of Calcein vary depending on organism and purpose of the study. Refer to [17] for review and recommendations.

• 2. Stained preparations can be observed under a microscope using the appropriate excitation filter for the stain used.

• 3. Store both the stock and resulting fixative refrigerated, use within days of preparation.

• 4. For marine species, adjust osmolarity with sucrose to be isotonic with local seawater. Since 1‰ = 30 mOsmol, convert target salinity, say 39‰, into osmolarity: 39*30 = 1170 mOsmol. Make up the fixative, measure its osmolarity and compare with sea water. The osmolarity of the fixative would be different depending on concentration of GA and the buffer used, so it is best to measure. Let us say the fixative has 974 mOsmol. Calculate the difference between it and the target, 1170–974 = 196 mOsmol. This is the amount that has to be added. Knowing that 0.342 g sucrose per 10 mL solution adds 100 mOsmol and knowing the volume of the fixative (in our case 50 mL) calculate the amount of sucrose: 0.342 * (196/100) * 5 = 3.35 g. Having a small difference of ~50 mOsmol between actual and target osmolarities is not a problem.

• 5. Bryozoans prefer the dark and are thus most easily found on the undersides of shells, on rocky overhangs, in submarine caves, on underwater vegetation and on the undersides of objects. In general they need hard substrate such as rocks or piers or bottles.

• 6. Freshwater bryozoans are not calcified, hard or rigid. They look like brown globs on logs, moss-like weeds or balls floating in the water. They are tricky: it is best to consult an expert for identification. In order to collect undisturbed colonies of the motile bryozoan Cristatella mucedo (Fig. 1), one can use a 3 mL plastic pipette with a cutoff tip. An opening (5–6 mm in diameter) should be large enough to take the colony in lengthwise, but small enough to provide enough suction. Other freshwater species live attached to the substrate and can be picked off (with or without substrate fragments) by hand, knife or forceps. Places to check: underwater vegetation and submerged surfaces (rotting wood, stones, plastic, cans, bottles) in streams, lakes, and ponds. In a fast-moving water, colonies are commonly located on the undersides of objects. Freshwater bryozoans are commonly identified using their reproductive resting stages: statoblasts. A very young colony may retain the valves of the original statoblast from which it “hatched.” Older colonies, especially near the end of the reproductive season, often contain numerous statoblasts in the body cavity. In nature, these near-microscopic propagules can be collected from the water’s edge, where they can form a brown “tideline,” or they might be found on floating items (e.g., wood, litter) or in foam. If you want to collect statoblasts particularly, tether a piece of polystyrene foam (Styrofoam) in the water and collect it a few days later, then rinse off the statoblasts—they adhere to it especially well. For microscopic examination, statoblasts should be opened and both valves placed in a drop of water on a glass slide. Coverslips are usually unnecessary as dry objectives (magnifications up to 40×) are sufficient.

• 7. For many purposes, it does not matter if specimens break as you collect them. Because bryozoans are modular, even a small fragment contains enough information for identification or genetic analysis. Many collectors choose to leave half the colony behind, as it will continue to grow after sampling.

• 8. Labeling specimens well is critical. We use this protocol : three-letters for general location (e.g., Otago Shelf would be OSH), number for exact location which will be detailed in log book, (e.g., 08), letters for bryozoan species if known (Cinctipora elegans would be CE) or random if not (XX). So, the sample label would read: OSH-08-CE. As ethanol dissolves ink, we use pencil on waterproof paper, and label everything at least twice (once in the container, once in the bag containing the containers from that location, and often writing on the container in permanent marker).

• 9. After collection , guard against transport-related damage: overheating, oxygen deprivation and mechanical damage associated with shaking. Overheating can be avoided with the use of cold packs and thermo-isolating boxes. Over short travel times, oxygen depletion is not significant if the density of animals is low, so we recommend filling the containers almost to the brim with water. This makes the colonies more resistant to shaking and mechanical damage. For long travel times, however, leave some air above the water and take extra care not to shake the containers.

• 10. Bryozoans can be dredged from the seafloor or collected from rocks or macroalgae.

• 11. When working in shelf depths from a sea-going vessel, a dredge is a quick way to get a huge volume of sediment on deck. Run the dredge for a very short time (2 min). If the location turns out to be full of bryozoans, then deploy the grab sampler to get discrete, less damaged samples. In the subtidal (water depths of 5–15 m), you can work from a smaller vessel and deploy a small hand-held grab or grapnel to collect macroalgae (on which the bryozoans may be living). Throw the grapnel far overboard, holding it parallel to the water. If collecting in the area with strong currents, throw the grapnel upstream. When the grapnel hits the bottom, release some additional rope (~5–10 m), then move the boat, dragging the grapnel along the bottom for about 2 min. Collected macroalgae can be roughly sorted and the epibiont-rich fragments placed into buckets with sea water for future processing.

• 12. Picking bryozoans from a heap of dredged material on a heaving boat while the deck hose is running is a messy job. Full waterproofs are needed.

• 13. Fill shallow trays with cool seawater and put the sampled specimens in there, grouping by apparent species. If hands are sensitive, use rubber gloves.

• 14. It is sometimes useful to collect 1–2 L of unsorted dredged material as a record of the sediment type and other taxa present. This can be transported dry, in seawater, or in ethanol. Ethanol will reduce the smell as the living material rots, and allows for future molecular analysis.

• 15. Morphologically similar colonies might be classified by colour (orange, purple), texture (soft, fluffy), or growth habit (e.g., branching, net-like). They may belong to multiple species, but often are the same family or genus.

• 16. Ethanol dissolves pen, so be sure to use pencil. It works to precut long thin (3–4 mm) strips of waterproof paper. Then write each location number (e.g., SE4) and place it straight into the container. Easier to handle than tiny rectangles of paper.

• 17. Avoid touching the mounting stub or anything that will go into the SEM, because grease from fingers will diminish the vacuum. If you do touch it, wipe with ethanol and start over.

• 18. When mounting bryozoans on the carbon tape, be gentle; they will break easily. Orient them to show zooids which are undamaged and variable, and try to include heterozooids, especially avicularia and gonozooids/ovicells. In the case of statoblasts, make sure some are upside down and some are right side up. For small specimens, it is possible to fit more than one species on a single stub. Use silver paint to close any gaps between the bryozoan and the tape—you are aiming to let electrons conduct.

• 19. If you have an Environmental SEM, coating is not required, and very low voltages can be used. Check with technician what best practice is for a fairly conductive substance like calcium carbonate.

• 20. You will need to develop the best protocol for your own SEM. We recommend voltage of 12–15 kV and a working distance in the range of 12–20 mm.

• 21. Orient photos so that the growing edge of the branch/colony is “up.” Usually the orifice is nearer the top of the zooid. If there are interesting details (such as a tooth or process in the orifice, or an elaborate pore), zoom in and get a photo of it, often at about 300×. Sometimes you might want a photo of the ultrastructure of the surface or of a broken part, that would be at about 1000×.

• 22. For marine species, the most practical approach is to work at a dedicated marine laboratory, using available, locally common species, along with a locally obtained seawater supply pumped ashore into the facility and into a dedicated flow-through or semiclosed recirculating tank system. Use of a continuous supply of “natural” seawater alleviates some of the practical difficulties associated with food culturing and maintaining appropriate environmental parameters. However, this method comes with some risks—such as introducing predators or competitors into tanks, and increased vulnerability to episodic perturbations, such as heatwaves or storms, which can rapidly change in-tank water quality by changing the temperature or smothering colonies with fine sediments if seawater is piped directly into the system. Potentially, colony recruitment and culturing can be done in situ, using artificial settlement substrates attached to the seafloor or suspended beneath artificial structures such as buoys and wharves. Exclusion cages can be used to reduce mechanical damage and predation of colonies, although these may be largely ineffective against bryozoan micropredators. A hybrid strategy is to settle larvae in the natural environment, then transfer the developing colonies, along with their substrates, to a culture system. If research objectives require tightly controlled conditions, or the absence of contaminating organisms, a closed aquarium system will be required. However, provision of artificial or sterilized natural seawater and a cultured microalgal diet adds a significant level of time investment to a project. Furthermore, some bryozoans have been shown to develop different and often-unusual colony morphology when kept in highly controlled laboratory conditions and fed microalgal monocultures [26].

• 23. The most tolerant and logistically feasible bryozoan species come from near-shore or intertidal environments, or from freshwater habitats. It is useful to conduct a pilot culturing study using a variety of locally obtained species to determine which taxa are best-suited for your system. When collecting colonies, make a note of the conditions in the microenvironments in which they occur, to help inform system design. Among the fast-growing encrusting genera are Watersipora, Electra, and Einhornia. Cryptosula pallasiana is a moderately well-studied, cosmopolitan and intertidal cheilostome [10] able to be cultured in the laboratory [27]. Ctenostomes amenable to culture include victorellids and Amathia [28, 29]. Among the cyclostomes, short-lived, fast-growing “weedy” species, such as Tubulipora, Crisia, and Filicrisia, are probably the most amenable to laboratory experimentation [30]. Heavily calcified marine taxa and deep-water species have proven to be very challenging to culture in the laboratory. Lack of knowledge regarding their natural environment (e.g., aquatic chemistry, food source) means that aquarium conditions can only be an educated guess. In some cases, culturing the specific phytoplankton species found in their natural environment might be the tipping point between failure and success. Freshwater bryozoans are abundant but have few genera. Cristatella, Plumatella, Fredericella, Lophopus, and Pectinatella in particular are among the most ubiquitous and easy to find taxa; and techniques for their culture have been refined [18]. Cristatella mucedo and Fredericella sultana have potential as a model species for WBR research, as they are common, easy to grow, and have fully sequenced genomes [31, 32].

• 24. Numerous commercial tank systems are available, and it is relatively easy to make custom systems. Tanks do not need to be large, but as a general rule, larger volumes (~20–50 L) tend to be more stable in terms of temperature and water quality and, for the same biomass of colonies, will require less frequent feeding at a given microalgal cell concentration if pulse feeding is used. Various low-volume rearing apparatus have been successfully developed [15], although these designs are relatively sophisticated. Tanks and other components in contact with the water should be nonmetallic and should not contain natural rubber, as both can be toxic to bryozoans. It is good practice to “condition” immersed system parts, especially new plastics, by placing them under running (sea)water for 24 h; this removes soluble residues and encourages the establishment of biofilms. Recirculating, semirecirculating and flow-through tanks can be used for bryozoan culture. These options have different advantages and drawbacks. Flow-through tanks are preferable if using natural seawater as the food source. Cultured food, however, can also be used in the case of a slow flow-through system. Recirculating or semirecirculating systems are ideal if precise control of water quality conditions is needed. Depending on the objectives, a very simple system can be effective: for example, plastic buckets can be used as tanks, and manual water changes every ~24 h can work for short-term studies. A good general principle for culturing bryozoans is to replicate natural conditions as much as possible. Many marine and freshwater bryozoans prefer low-light levels and shaded environments, growing best on underhanging substrates. If culturing is taking place in a well-lit room, a light cover over tanks should be considered. Note that exposure to high-intensity light can induce spawning in some species (see Subheading 3.4). Water movement is another consideration. Linear or oscillating currents can be generated using various methods, including submersible pumps, wavemakers, aerators and mechanical stirrers. Some species grow well in still water, although a small amount of water movement is beneficial to ensure that food particles do not sink to the bottom. In our experience care should be taken to ensure microbubbles are not introduced into the system during aeration, as these can adhere to colonies, disrupting normal function. Similarly, high current speeds can be damaging. For many bryozoan species, management of water temperature is important. Nearshore marine and freshwater bryozoans tend to be eurythermal. For stenothermal taxa, a responsive temperature controller heating and/or cooling system should be used. Tanks can also be housed in a controlled-temperature environment.

• 25. Bryozoan colonies can be fed simply by the provision of natural seawater in flow-through systems, or by adding cultured food (usually microalgae) to tanks. For freshwater culturing, Wood [18] describes use of a closed mixed culture system using a tank containing well-fed goldfish and a rich microbiota; food-rich water from this reservoir is supplied to bryozoan culture tanks via an airlift pump. Marine bryozoan culture requires cultured microalgae, using commercially obtained strains developed for the aquaculture industry. Conveniently their nutritional content is usually well-documented. Many marine laboratories maintain a dedicated phytoplankton culture room, and requests for culture of specific strains should be made 1–2 months in advance to allow time for culturing. Common microalgal genera used for feeding bryozoans include Dunaliella, Rhodomonas, Tetraselmis, and Pavlova. It is important to supply appropriate cell concentrations of the cultured microalgae to the bryozoan tank, taking into account the dilution volume of the tank itself. Feeding can be done by manual daily additions of cultured cells, or ideally, by a drip feed system, which can be applied both to closed and flow-through systems. Microalgal monocultures are commonly used for experimental studies of growth and feeding , but mixed microalgal diets may be more appropriate for some studies. It should be noted that abrupt changes in diet from one monoculture to another is reported to induce colony-wide polypide regeneration in some taxa [15]. Optimal cell concentrations can be found in the literature for some commonly studied species, such as Electra pilosa [33] or Cryptosula pallasiana [27]. To calculate the volume of cultured algae required for feeding , cell counts can be made using a cavity slide or an automated cell counter.

• 26. Bryozoan colonies are delicate, and must avoid direct contact with hard surfaces other than the attachment substrate. If colonies are grown on slides or plates they can be suspended vertically or upside-down in the water column. Wood [18] recommends use of inverted petri dishes as substrates for freshwater bryozoans. An attachment system, such as a frame, rails, Velcro, or even Lego, makes it easy to remove and replace colonies. Cyanoacrylate glue is a safe adhesive for bryozoans; aquarium silicone works as a strong and flexible glass adhesive, but requires curing and conditioning before use. If tank aeration is used, ensure that colonies are placed away from the bubble stream, and away from the strong flows generated by submersible pumps and inlets.

• 27. Cleaning of bryozoan culture tanks and the colonies themselves is necessary, especially in closed systems. Excess food and fecal pellets from colonies can contribute to rapid build-up of bacteria, which can inhibit normal function in some bryozoans [15]. Cleaning may also reduce the buildup of ciliates, which can often be problematic in marine culture systems. Regular siphoning of detritus from the tank floor is recommended, as well as regular water changes in closed systems. Cleaning of bryozoan colonies requires care and is best done in response to observed detritus clinging to the colony (some bryozoans appear quite adept at cleaning themselves). Common practice is to use an ultrafine artist’s brush or a dissecting needle to gently sweep away sticking detritus. An alternative is to employ gentle puffs from a disposable plastic pipette that has had the tip cut off to enlarge the opening. In both cases great care should be taken to avoid damaging the colonies, and it is prudent to do some “test cleans,” followed by examination of colonies several hours later to ensure they are undamaged (e.g., feeding normally). Some bryozoans, such as free-walled cyclostomes, are prone to having their membranous body walls scraped off, and individual brush hairs can easily enter zooidal tubes, damaging terminal membranes and polypides. Water filtration can reduce the buildup of mobile detritus in a closed or semirecirculating system; however, doing so also removes food particles. One method is to run a timer-activated power filter once a day for a short period (~1 h) before feeding or turn off the water filtration system for a short period of time (~2 h) while feeding .

• 28. Most gymnolaemates, and all stenolaemates, produce lecithotrophic (nonfeeding) coronate larvae, but other species produce planktotrophic (feeding ) cyphonautes [34]. Refer to [19] to identify the two larva types. A pilot study might be required prior to spawning the species of interest, in order to understand the type of larvae produced and potential settlement intervals.

• 29. Most marine bryozoans are light-induced spawners, [20,21,22, 34, 35] while spawning in freshwater bryozoans is dark-induced instead of light-induced [23]. Sheet-encrusting marine species such as Membranipora membranicea can be prepared for spawning with the “one-zooid-row preparation.” It includes cutting portions of colonies with a scalpel blade into strips one-zooid wide and several zooids long [20, 36]. This method provides biological replicates (clones, ramets) for experiments. The “one-zooid strips” can then be placed in a small petri dish with seawater for spawning and observation. When choosing encrusting colonies growing on algae such as M. membranacea, it is advisable to choose algae which do not secret large amount of mucus when they are cut [20, 36]. If the “one-zooid-row preparation” is not possible, different colonies or colony parts can be kept in separate containers with the chosen substrate for settlement such as acetate sheets, glass slides, a chicken egg-shell membrane previously soaked for 24 h in seawater, other adult non–sexually reproductive bryozoan species (preferably dead) or any other calcified substrate (e.g., bivalve shells) [19, 37, 38]. Similar principles apply for freshwater species; each colony can be placed in a separate container or petri dish until spawning [23, 24]. For both marine and freshwater species, regular check-ups are required (every ~2 h) to ensure whether settlement has occurred. Spawning and settlement intervals can also be recorded with a microscope camera, either by video recordings or photo time frames [36]. Settlement intervals vary, but planktotrophic cyphonautes larvae produced by the marine ctenostome species Amathia gracilis may swim up to 10 h before settlement [37].

• 30. Larvae of marine bryozoans are phototactic, and therefore a light source can be used to lure the larvae so they can be easily collected using a pipette. Care should be taken so that larvae do not swim to the air–water interface, where they can be trapped, therefore leading to larval mortality. In order to prevent that, the water–air interface can be taped over with black tape or tinfoil, all around the glass jar or aquarium. Apply the light source laterally below the tape line, which makes it possible to concentrate and collect the swimming larvae.

• 31. Calcein binds to CaCO₃ so this method generally works best on well-calcified species. Calcein concentration should be 50–150 mg/L. Higher concentrations work in a shorter time, but have a higher chance of being toxic [17].

• 32. Calcein marking can be done in situ—surround bryozoans with a plastic bag, sealed against the substrate or around them, then inject with concentrated Calcein to achieve necessary concentration.

• 33. If partially anesthetized polypides retract, they are often unable to evert again and the process needs to be started over with a different colony. Bryozoan species vary significantly in their sensitivity. While some are relatively hard to disturb, others may prove extremely skittish. In the latter case one may consider doing relaxation after hours or on a weekend. Both marine and freshwater species can be successfully relaxed in the regular lab settings, although a temperature-controlled room may be a better option for deep-water species. If done at room temperature, container may need to be cooled locally with a freezer pack. Have a reserve pack ready for long relaxations.

• 34. Freshwater species are more difficult to anesthetize. MgCl₂ can be used but is less effective for these bryozoans since solution isotonic to freshwater has very few Mg²⁺ ions, and increasing concentration causes osmotic shock. Instead, use one of the other two substances which work better: menthol and chloral hydrate solutions. Keep in mind that freshwater bryozoans are even more easily damaged by overexposure. Living tissues begin to macerate: the epidermal layer on the tentacle loses cohesion and the cells slough off. A polypide with visibly narrowed or uneven tentacle surfaces is too damaged to fix. A typical relaxation should not exceed 1 h. There are two methods using menthol for relaxation. Menthol is not easily soluble in water, so one may prepare a solution beforehand and add it gradually, or else place small menthol crystals on the water surface of the culture container. It is important to keep the container covered, because menthol is an irritant and evaporates easily. The lid should be transparent to allow microscopic observation and cover the container in such a way as to be removed without shaking the colony. A large upturned Petri dish is usually good for this purpose. Use of a benchtop extraction hood during relaxation with menthol is recommended. The same procedure of gradually adding the relaxant applies for chloral hydrate solution. Chloral hydrate: 20 g C₂H₃Cl₃O₂ in 1 L H₂O, prepared 1 month before use to saturate properly.

• 35. 4% Formalin and Bouin’s solution (150 mL filtered saturated solution of picric acid, 50 mL formalin, 10 mL glacial acetic acid, use within a few days of preparing) both work well for fixation of both marine and freshwater bryozoans for paraffin based histological sectioning. Samples can be stored in formalin for several months, but prolonged storage in Bouin’s solution is not recommended, as it will dissolve skeletal carbonate. Histological handbooks such as [39] provide more background and details on specific fixatives and processing methods. Formalin-fixed material can also be processed for aceto-orcein staining . For immunocytochemical studies with the use of confocal laser scanning microscope, Triton X-100 is a common permeabilization agent, with goat or bovine serum albumin as common blocking solutions. Primary antibodies successfully used with bryozoans include rabbit anti-serotonin and mouse anti-acetylated α-tubulin; secondary antibodies include goat and donkey anti-rabbit as well as goat and donkey anti-mouse (e.g., [40]). For SEM examination of larvae, use 2.5% GA fixative, osmolarity-adjusted and buffered with 0.2 M Milloning’s phosphate buffer (pH 7.4) for 1 h at 20 °C. Freshwater and ctenostome bryozoans may require stronger concentrations. Postfix the animals in 2% osmium tetroxide and 1.25% sodium bicarbonate (pH = 7.2 with 1 N HCL immediately before use) at 20 °C, for 1–2 h, then follow standard protocols for rinsing, dehydration, critical-point drying and coating [37]. Fixation and processing for visualization of ovaries, oocytes and nuclei requires some very specialized stains. For DNA-specific fluorochrome Bisbenzimide H333342 use samples fixed with buffered 4% formalin, stain with Bisbenzimide H333342 (10 pg/mL) for 5 min or more at room temperature. Rinse three times with filtered seawater [20]. The specimens can be stored at 4 °C until imaging. For aceto-orcein staining , specimens can be fixed either directly with 3:1 methanol–acetic acid for 30 min, or in two stages. For the latter method, first fix with 4% formalin with 0.2 M PBS for 20 min and rinse thoroughly with any phosphate buffer, then postfix with 3:1 methanol–acetic acid for 60 min. Use 45% solution of aceto-orcein to stain the samples for 30 min. Rinse with 20% acetic acid [20, 34].

• 36. Most marine bryozoan taxa are biomineralized, with skeletons comprising CaCO₃ in the form of calcite and/or aragonite. For histological and EM sectioning purposes, the skeleton must be fully removed. Failure to properly decalcify can lead to wholesale destruction of the expensive diamond knives used for ultramicrotomy. Decalcification can have harsh effects on delicate bryozoan tissues, and lead to mechanical damage and extraction of cell contents, so gentler, longer-period protocols generally work best for heavily calcified taxa if excellent-quality sections are required. Immersion of samples in decalcification solution in the absence of CaCO₃ is especially damaging to tissues, so close monitoring of progress is recommended. Typically, a weak acid or a chelating agent is used to remove the skeleton. Popular acids for decalcification include ascorbic acid and formic acid, and these are used in diluted form (~2–4%), in a seawater-isotonic solution; this process can take several weeks and requires regular changes of solution. Calcium chelation is a highly regarded method, especially for TEM , and the preferred agent is EDTA (in 5–20% range). As decalcification protocols are required for marine bryozoans only, decalcifying solutions should done in a seawater isosmotic environment, ideally similar to that used for fixation and washing stages, so buffers should be used during this procedure, for example, PBS/cacodylate buffer. An osmometer is useful, and the osmolarity of local seawater should be used as a target. In most cases decalcification will be undertaken after the initial fixation , but can be done either before or after postfixation, in the case of TEM . Following decalcification, samples should be washed several times in seawater-isotonic buffer. As a general rule, further processing of the decalcified material is best done by hand, rather than with a tissue processor. For the most delicate specimens, embedding in low- or ultralow temperature agarose before decalcification dramatically improves overall sample integrity.