WARNER-LAMBERT V ACTAVIS – DO WE HAVE AN EFFECTIVE SYSTEM FOR ENFORCING SECOND MEDICAL USE PATENTS IN THE UK? Why is this important? Increasingly, unmet medical needs are being met by "repurposing" of old medicines for new diseases. Before such medicines may be used to treat patients for the new disease, expensive research including clinical trials must usually be performed. One mechanism to incentivise this research is to grant innovator companies exclusivity for the new indications through second medical use patents. Thus, there is a public policy argument for granting second medical use patents, and to ensuring that these patents are enforceable. On the other hand, generic versions of innovative drugs should be allowed on to the
Reefcentral.ruReference: Biol. Bull. 204: 68 – 80. (February 2003)
2003 Marine Biological Laboratory
Collection and Culture Techniques for Gelatinous
KEVIN A. RASKOFF1,*, FREYA A. SOMMER2, WILLIAM M. HAMNER3, AND KATRINA M. CROSS4 1 Monterey Bay Aquarium Research Institute, Moss Landing, California 95039-9644; 2 Hopkins Marine Station, Pacific Grove, California 93950-3094; 3 University of California, Los Angeles, California 90095-1606; and 4 Monterey Bay Aquarium, Monterey, California 93940-1085 Gelatinous zooplankton are the least under- techniques have been developed that permit researchers and stood of all planktonic animal groups. This is partly due to aquarists to collect intact gelatinous animals at sea and to their fragility, which typically precludes the capture of maintain many of these alive in the laboratory. These new intact specimens with nets or trawls. Specialized tools and methods and technologies have allowed scientists to resolve techniques have been developed that allow researchers and the life cycles of many organisms whose hydroid and hy- aquarists to collect intact gelatinous animals at sea and to dromedusa stages were previously thought to be separate maintain many of these alive in the laboratory. This paper species, and to conduct a variety of experimental studies in summarizes the scientific literature on the capture, collec- the laboratory. In addition, aquarists are now able to rear tion, and culture of gelatinous zooplankton and incorporates and display many species of medusae and ctenophores for many unpublished methods developed at the Monterey Bay the first time in public aquariums. Paffenho¨fer and Harris Aquarium in the past 15 years.
(1979) and Strathmann (1987) provide good reviews ofmany gelatinous zooplankton culture studies, as well as a detailed review of culture methods for non-gelatinous or-ganisms. The general lack of information about gelatinous Gelatinous zooplankton is a generic term for transparent zooplankton is due not only to their extreme fragility, but and delicate planktonic animals with mesoglea-like internal also to a shift of emphasis in the discipline of biological tissues that aid in regulating buoyancy. These animals in-clude some radiolarians and foraminifera, as well as medu- oceanography that occurred more than 100 years ago. The sae, siphonophores, ctenophores, chaetognaths, pteropods, change led from a qualitative interest in the systematics and heteropods, appendicularians, salps, doliolids, and pyro- developmental biology of all zooplankton, to the present somes (e.g., Hamner et al., 1975). These taxonomic groups quantitative concern for the fisheries implications of certain are widely distributed in large numbers in all the world's components of the zooplankton as they relate to cycles of oceans, throughout the water column. They are the least energy and material in the sea.
understood of all planktonic animal groups. This is partly This dramatic shift of emphasis was advocated by Victor due to their fragility, which typically precludes the capture Hensen (1887). Ernst Haeckel and others used fine-meshed of intact specimens with nets or trawls. In fact, many plankton nets towed slowly at the surface from small boats, systematic descriptions of hydromedusae and siphono- or they carefully dipped individual animals from the sea phores during the past 200 years were based only on frag- surface by hand. In contrast, Hensen (1887) used large, ments of animals (e.g., Mayer, 1910; Russell, 1953). For- vertically hauled plankton nets from large ships to collect tunately, during the past 30 years, specialized tools and fish eggs and copepods. This procedure produced quantita-tive information on the distribution and abundance of fisheggs, copepods, and larval forms, but it seriously under- Received 24 April 2002; accepted 6 November 2002.
sampled and physically damaged the gelatinous fauna. Even though Haeckel believed that Hensen's approach to oceanic COLLECTION AND CULTURE OF GELATINOUS ZOOPLANKTON biology was flawed, Hensen's attempt to quantify plank- techniques developed at the Monterey Bay Aquarium dur- tonic ecology has prevailed, and the most recent manual on ing the past 15 years.
zooplankton methodology (Harris et al., 2000) is still pri-marily concerned with crustaceans and fish eggs.
Throughout the first half of the 20th century, systematists continued to collect individual gelatinous animals (e.g.,Kramp, 1965), but interest in the developmental biology of The primary goal of all collection methods for gelatinous these animals diminished as Haeckel's ideas about phylo- zooplankton is to minimize handling and damage.
genetic recapitulation through ontogeny lost favor. In the Surface collection. Many common species can be col- 1950s, neurophysiologists such as Pantin (1952), Bullock lected easily from surface waters by using a small boat or (1943), and Mackie (1960) began to investigate the neurol- while snorkeling. Ocean slicks, glassy patches on the ogy of the so-called lower invertebrates (anemones, medu- ocean's surface caused by a combination of wind and cur- sae, siphonophores, flatworms, and ctenophores), an effort rent (Haeckel's "animal roads"), are excellent sources of that required that these animals be collected carefully at sea epipelagic species (Alldredge and Hamner, 1980; Hamner and also maintained alive, if only briefly, in the laboratory.
and Schneider, 1986; Larson, 1991). Smooth-rimmed glass The first use of scuba to collect or view planktonic beakers and glass jars (perhaps attached to the end of a pole) animals at sea was in the 1960s by divers who collected are good collecting containers, since most cnidarian tenta- siphonophores in the Mediterranean (frontispiece in Totton, cles do not adhere to glass as readily as to plastic. Larger 1965) and by Ragulin (1969), who first viewed krill under- specimens can be collected in plastic buckets or in plasticbags. Some hardy species can be collected with dip nets or water in the Antarctic. Soon thereafter, use of scuba to small plankton nets. The smaller the mesh size the better, as investigate oceanic gelatinous animals became routine in larger meshes can cut into the soft gelatinous tissue. Knot- the epipelagic blue waters of the Gulf Stream (e.g., Gilmer, less mesh of broad, flat strands of soft material in the shape 1972; Hamner et al., 1975). Research submersibles in mid- of a gusseted bag is the most effective type of hand-held dip water extended these in situ observations of gelatinous net. It is important not only to minimize the stress and plankton from the upper 30 m of the sea to thousands of disturbance to the animal, but also to avoid introducing air meters below the surface (e.g., Larson et al., 1992; Robison bubbles into the body cavity; air bubbles are exceptionally et al., 1998; Raskoff, 2001, 2002).
difficult to remove, and if left within the animal either In situ observations of epipelagic and midwater animals produce tissue embolisms or cause the animal to rise to the stimulated a revival of attention to all aspects of the living surface, where exposure to air can damage it further.
biology of gelatinous zooplankton. We know today that Subsurface collection. Although many specimens can be gelatinous animals are an important component of marine collected in good condition at or just under the surface, ecosystems, with particular significance for fisheries man- others may already be damaged, particularly those garnered agement (e.g., Purcell, 1997; Purcell and Arai, 2001), an from surface convergences crowded with flotsam. Further- issue anticipated by Haeckel (1893). We now know that more, for the vast array of species not routinely found at the gelatinous organisms are often the dominant macrozoo- air-water interface, collectors may need to employ other plankton of oceanic ecosystems (e.g., Robison et al., 1998).
methods. Use of scuba is one possibility. When working in Recent studies with in situ techniques have shown that the disorienting, featureless, blue-water environment of the scyphomedusae, hydromedusae, siphonophores, and cteno- open sea, protocols for diving-safety must be followed (see phores are abundant, often quite large, and apt to play Hamner, 1975; Heine, 1986). Divers who are even slightly disproportionately important roles as top predators in their negatively buoyant can easily sink below safe depths. Also, food webs (see Mills, 2001; Purcell et al., 2001). Yet these divers who are not connected to one another and to the animals continue to be neglected in many syntheses of support boat often come to the surface away from the boat, where they can be difficult to see in a choppy sea. However, Our ignorance of gelatinous plankton biology is thus blue-water diving techniques, properly executed, provide a partly due to the history of oceanography, partly to inade- safe and effective way to collect specimens and to observe quate collection and observational technologies, and partly the behavior of undisturbed animals in their natural habitat.
to the fragility of many gelatinous taxa. Although it is now Many ethological discoveries in the last 20 years have been possible to capture most species of gelatinous animals in made using these techniques (e.g., Madin, 1974; Hamner, good condition, it is still difficult or impossible to keep 1985; Matsumoto and Harbison, 1993).
many of these taxa alive in laboratory aquaria for observa- For collection of specimens deeper than the limits of safe tion and experimentation. This paper summarizes the scien- scuba diving, or when diving conditions are not optimum, tific literature on the culture of gelatinous zooplankton and various nets have been deployed successfully from the incorporates many unpublished culturing and displaying surface. Midwater trawls, bottom trawls, and plankton nets K. A. RASKOFF ET AL. can all be effective in capturing delicate living specimens (Sameoto et al., 2000) if the nets are pulled slowly (⬍1.0km h⫺1) for a relatively short time, and if the cod end of the Once organisms have been collected, cultures can be net is large and without side windows, which generate started in a number of ways. The most common method is turbulence in the collecting well (Baker, 1963; Reeve, 1981; to facilitate natural spawning by grouping both sexes to- Childress and Thuesen, 1993). Thermally insulated cod gether in a small controlled space. Spawning can often be ends have also proved very successful in the capture of induced by crowding (some scyphozoans), by leaving ani- gelatinous organisms in good physiological condition (Chil- mals in the dark for several hours followed by periods of dress et al., 1978; Thuesen and Childress, 1994). The use of light (some hydrozoans), or by simply permitting the tem- research submersibles and remotely operated vehicles perature of the water to slowly rise over several hours (see (ROVs) has permitted gelatinous species to be collected Mills and Strathmann, 1987). In some cnidarian genera, from meso- and bathypelagic depths when the use of nets is such as Aurelia, females brood their planulae on their oral not an option (Youngbluth, 1984; Robison, 1993). These arms. It is often sufficient to place the brooding female in a vehicles may be equipped with large collection cylinders small volume of seawater and wait for a few hours for the open at both ends, permitting the vehicle pilot to slide the planulae to be released. Alternatively, larvae may be re- collecting container over a gelatinous animal by maneuver- moved from the edges of the oral arms with a pipette. A ing the entire vehicle and then gently closing the ends of the more labor intensive spawning technique involves in vitro sampler. Some species are so delicate that they have never fertilization. For this procedure, gonadal tissue from both survived even these samplers: descriptions of several deep- males and females are incubated together in a small volumeof water for several hours until the eggs are fertilized and sea species, such as Kiyohimea usagi (Matsumoto and Ro- larvae begin to develop. The sexing of zooplankton can be bison, 1992) and Lampocteis cruentiventer (Harbison et al., difficult, but the eggs can often be seen inside the female 2001), were based on in situ observations and photography.
reproductive tissue. The most accurate way to determine thesex of the specimens is to remove a small piece of the gonadand examine the tissue under a compound or dissecting Transport of specimens microscope to look for sperm and eggs. This will also helpdetermine if the specimen is mature.
Once collected, specimens can be safely transported to After a spawning event, it is necessary to examine the the rearing facility either in their collection containers or in water for larvae or fertilized eggs. Collection and handling larger jars or tubs. If the animals are transferred to a larger techniques for many larval taxa are summarized in Strath- container, it is important to minimize their exposure to air mann (1987). Planulae range in length from 100 to 1000 m and to avoid pouring them roughly from one container to the in hydrozoans, range from 100 to 400 m in scyphozoans, other. Each animal must be dipped gently out of the col- and up to 160 m in cubozoans (Martin and Koss, 2002).
lecting container with a transfer jar, which is then emptied Ctenophore larvae range in length from 280 to 1000 m by tipping it below the surface of the water in the transport (Baker and Reeve, 1974; F. Sommer, unpubl. data).
container. Several devices have been designed for the trans- Cnidarian planulae will typically settle and attach to the fer of individual gelatinous zooplankton (Acun˜a et al., substrate within a few days, often within hours if a suitable 1994; Sato et al., 1999). The water in the transport contain- substrate is available. Planulae will settle on many types of ers should be free of bubbles and have the same temperature substrate (Brewer, 1984). Glass or plastic microscope slides and salinity as the water in the collecting vessel; if not, or cover slips are often used due to the ease of post- small volumes of water should be exchanged between them settlement manipulation. Some species may preferentially slowly, over the course of perhaps an hour, to permit tem- settle on substrates that have been "conditioned" by several perature and osmotic adjustment by the animals. Transport days' immersion in seawater to accumulate a light microbial vessels can then be put into an insulated box or cooler with film (Brewer, 1984; Schmahl, 1985). Several chemicals cold or hot packs as needed for the duration of the trip. The (TPA, DAG, Cs⫹, Li⫹, NH ⫹) have been shown to posi- water can be saturated with oxygen before transport, but this tively affect larval settlement (Siefker et al., 2000). Larvae is more crucial for large animals, those being shipped in typically settle to the bottom of the chamber and often are warm water, or those kept in the shipping container for a thigmotactic, tending to settle at the edges (Brewer, 1976; long time. All air must be removed from the container Orlov, 1996). Some species (Aurelia aurita, Cyanea capil- before sealing because even small air bubbles can damage lata, Ptychogena lactea) are also light sensitive and will gelatinous specimens. With an appropriately low ratio of settle under opaque objects such as small rocks or shell biomass to volume of water (⬍1:2), the animals often (Custance, 1964; Brewer, 1978, 1984; Raskoff, unpubl.
survive trips of 18 h or more. For small medusae and polyp data), while others (Clava multicornis) are positively pho- cultures, air-permeable plastic fish bags are very effective.
totactic (Orlov, 1996). There is evidence that some species COLLECTION AND CULTURE OF GELATINOUS ZOOPLANKTON settle preferentially in areas with a high density of conspe-cifics (Keen, 1987). Some larvae may also settle at theair-water interface, attaching upside down onto the surfacefilm (Pagliara et al., 2000). These can be dislodged bygently disturbing the surface tension with a drop of water,whereupon the polyps drop to the bottom and reattach to abenthic substrate. Additionally, planulae will attach to afloating substrate that is gently placed on the water surface.
Larvae induced to settle on microscope slides can be raisedoff the bottom of the culture chamber after they have startedto reproduce asexually, and inverted so the polyps hangupside down. This facilitates their feeding and allows theirwastes to fall to the bottom of the tank, reducing fouling.
With consistent feeding and a debris-free environment, healthy polyps will generally grow and produce juvenilemedusae. However, several treatments can be used to initi-ate or speed up the process. Scyphozoan polyps typicallyproduce juvenile medusae by the process of strobilation,which can be induced in various ways. These include brieftemperature increases of ⬇5.0 °C, prolonged (4 – 6 weeks)reduction of water temperature by ⬇5-10 °C followed by a Figure 1.
Flow-though culture tanks and grow-out facility. Tank sizes return to normal temperatures over a few days, and changes should allow for free, unrestricted feeding and movements of specimens.
in the amount of feeding (Abe and Hisada, 1969; Calder, Mesh sizes should be smaller than the smallest dimension of the organism.
1974; Cargo, 1975). Other inducers found to have somesuccess are changes in illumination level and pH, increases brush. Razor blades or narrow-tipped utility knives are in salinity, and treatment with various chemicals (iodine, helpful for scraping polyps off smooth, flat surfaces such as thyroxine, etc.) (Spangenberg, 1971; Olmon and Webb, glass slides. Once removed, polyps are placed into separate tanks and allowed to resettle. Many species reattach quickly When the polypoid phase begins to release juvenile me- when simply resting on the bottom of a dish; others may dusae, it is helpful to remove them from the culture chamber take longer. One method for raising these polyps off the and place them into a rearing tank as soon as possible.
bottom to facilitate feeding is to tie a tight loop of small- Young ephyrae and hydromedusae can be injured or eaten gauge monofilament line around, or slip a small rubber band by other members of the polyp colony. The young medusae over, a glass microscope slide, and then insert the base of will often be swept out the outflow of the polyp culture tank the polyp under the line on the flat portion of the microscope (Fig. 1) and into the grow out tank, but transporting them slide (Groat et al., 1980; F. Boero, Universita di Lecce, via a large-diameter pipette is preferable because it reduces Italy, pers. comm.). The tension of the monofilament line the stress on the juvenile medusae.
holds the polyp next to the surface of the slide without The medusae of most species can be placed directly into cutting through the stalk of the polyp. The microscope slides flow-through or aerated rearing tanks after release (Fig. 1), can then be inverted, allowing the polyp's tentacles to hang although some species such as Pelagia colorata and Ae- freely. After several days to weeks, the polyp will attach to quorea victoria respond better if first placed into small the slide, and the monofilament can be cut and removed.
dishes with still, filtered water for several days to weeks.
Asexual reproductive bodies, such as cysts and frustules, Juvenile medusae may need to be transferred into several can also be removed from the original tank to seed a grow-out tanks of increasing sizes and decreasing conspe- replicate culture. Cysts can be removed by scraping, and the cific densities throughout their development, depending on damage caused to the capsule of the cyst sometimes stim- the species and size of the medusae (Spangenberg, 1965).
ulates excystment and subsequent growth of the polyp, as Once a polyp culture has been started, it is often neces- can changes in temperature (Brewer and Feingold, 1991).
sary to propagate the polyps in additional culture containers.
Swimming frustules are produced in some species (hydroid Propagated polyps can be used to set up replicate cultures example: Craspedacusta; scyphozoan example: certain rhi- for experimentation, for transfer to other researchers or zostomes such as Cassiopeia and Mastigias), and these can aquarium facilities, or as backup in case of problems. Both be pipetted into a dish where they will settle and develop hydrozoan and scyphozoan polyps can be removed from into polyps. After settlement, the dish can be transferred to substrates by gently scraping with a small instrument, such a flow-through tank.
as a plastic toothpick or a trimmed, hard-bristled paint The use of antibiotics to aid in the culture of gelatinous K. A. RASKOFF ET AL. organisms has not had much study. Strathmann (1987) lists available for capture only briefly. These foods must be several antibiotics and fungicides that might help fight in- removed or they rot and promote growth of fouling organ- fections. The antibiotic tetracycline has been used to treat isms. Live Aurelia and other medusae are a good and bacterial infections on large scyphomedusae. After being sometimes necessary dietary supplement for many medu- placed in a 20-ppm bath for 2 h a day, 5 days in a row (B.
sivorous jellyfish, including Pelagia, Cyanea, Chrysaora, Upton, Monterey Bay Aquatium, pers. comm.), the infected Phacellophora, and Aequorea. Smaller stages of these me- medusae improved markedly. This technique shows great dusivores can be fed Aurelia ephyrae, finely diced adult promise for treating the common "bell rot" encountered medusae, or small hydromedusae. Small, newly released with many large medusae.
hydromedusae, such as Aequorea, Eutonina, and Bougain-villea, are especially important in the diet of Pelagia colo- rata ephyrae, which are difficult to raise on Artemia alone(Sommer, 1993). Wild-caught plankton also offer an impor- Among the types of food that can be used to feed gelat- tant dietary supplement to gelatinous zooplankton in cul- inous zooplankton are Artemia nauplii, krill, chopped squid ture. Live copepods are desirable for tentaculate cteno- and fish tissue, medusae, wild plankton (copepods, etc.), phores and scyphomedusae. Recent research has pointed out rotifers, trochophore larvae, agar-based foods, algae, bi- the importance of utilizing natural prey whenever possible.
valve hepatopancreas, and "grow-lights" for those species The reason that at least two species of naturally biolumi- of medusae with zooxanthellae. Artemia nauplii are the nescent medusae do not produce light when reared in the most common food items used in culture of polyps and laboratory is a dietary deficiency of the luciferin coelentera- medusae and provide the backbone of most species' diets in zine (Haddock et al., 2001). Thus, even seemingly healthy laboratory conditions. Most species can be fed Artemia cultured animals may not receive all of their nutritional daily, but some very small polyps may have difficulty needs from convenient laboratory prey, and alternative or capturing and ingesting prey of this size (about 400 m).
supplemental foods should be tried routinely.
Tentaculate ctenophores thrive on Artemia, but non-tentacu- Small and newly metamorphosed animals can be difficult late beroid ctenophores need gelatinous prey. The lack of to feed due to their diminutive size. Various live single- appropriate food items is a major stumbling block for the celled algae, such as Tetraselmis spp., Isochrysis galbana, culture and study of many gelatinous taxa. For example, the and Nannochloropsis spp., can be valuable food sources for natural food of many pteropods is other species of ptero- small polyps, as well as for filter-feeding salps and doliolids pods, which are difficult to culture in the laboratory; there- (Paffenho¨fer, 1970, 1973; Heron, 1972; Paffenho¨fer and fore, even if the animals themselves can be successfully Harris, 1979). Rotifers (⬇100 –200 m), such as Brachio- maintained in tanks, providing them with adequate nutrition nus plicatilis, and oyster trochophores (⬇50 m) are in the over long periods of time is a challenge (Conover and Lalli, right size range for capture and consumption by polyps, which may be unable to consume the much larger Artemia Hatching times and water temperatures vary between the nauplii. Rotifers can be fed on the above algae as well. All different species and strains of Artemia, so recommenda- of the above prey items are commercially available. Rotifers tions provided by the supplier should be consulted. After and algae are easily cultured in tanks similar to those used hatching, the nauplii should be fed for a day or so with a for Artemia, and the trochophores can be purchased frozen.
food supplement (Super Selco, Algamac, algae, yeast, etc.).
Another food that has been used with some success is By enriching the content of protein and free amino acids in agar-enriched medium. Homogenized food items mentioned the nauplii acid (Helland et al., 2000), these supplements above, as well as amino acids, lipids, and protein sources, contribute to the subsequent growth and health of the ani- can be mixed into heated agar and, when cooled, a gel is mals to which the nauplii are fed. The nauplius must have a formed. This gel can be cut into small pieces and fed by mouth (2nd instar stage) before it can ingest the enrichment hand to polyps, medusae, and beroid ctenophores. This is medium, which must be dispersed (emulsified, aerated, or labor intensive but useful for some species that are other- otherwise kept in the water column) so that the nauplii can wise difficult to feed. Another common feeding technique eat it. The "shells" of the Artemia cysts can be removed to uses bivalve hepatopancreatic tissue, finely chopped and reduce fouling and increase hatching efficiency. Several cleaned in successive changes of seawater. These pieces are methods of cyst decapsulation are available on the Internet.
then hand-fed to individual polyps. Common intertidal Decapsulated cysts can be kept for extended periods, refrig- copepods can be cultured in shallow pans as a food source.
erated in water, until they are needed.
Several species of medusae depend on the photosynthetic Krill, squid, and other large or fleshy prey can be cut or products of zooxanthellae for nutrition. In addition to a homogenized to an appropriate size and fed to many polyps, normal diet of prey, these species require a strong light with medusae, and heteropods. A disadvantage is that these food an appropriate action spectrum for photosynthesis by the items quickly sink to the bottom of the tank and thus are zooxanthellae. The type and power of the light can be COLLECTION AND CULTURE OF GELATINOUS ZOOPLANKTON variable depending on tank size and depth. For example, at above (Fig. 1B). The addition of an air line close to the the Monterey Bay Aquarium, the scyphozoans Mastigias screen in any catch or grow-out tank will cause bubbles to papua and Cassiopeia xamachana have been reared for rise along the screen, and these will create a gentle upward several months in tanks with metal halide and actinic or current that encourages juvenile medusae to stay up in the daylight fluorescent lamps.
water column and off the screen.
Pelagic stages. Tanks for pelagic animals offer unique challenges, but the aim is to mimic a natural environment asclosely as possible. The vast majority of gelatinous zoo- Benthic stages. With careful cleaning and frequent water plankton are pelagic, and their tanks must minimize contact changes, benthic hydroids and polyps can be kept in simple between the animal and all tank surfaces. That being said, jars and dishes (e.g., Miglietta et al., 2000). However, when many gelatinous taxa have been maintained or cultured in dealing with large cultures, or when flowing water is desired the laboratory in nothing more than jars or aquaria of still for efficient feeding, more complex facilities are needed.
water in temperature-controlled environments. Radiolarians Rees and Russell (1937) designed the first successful large- (Sugiyama and Anderson, 1997) and foraminifera (Hemle- scale culture system for cnidarian polyps. This consisted of ben and Kitazato, 1995) have been kept for extended peri- rows of glass beakers that held the polyps, and vertical ods in small jars and culture dishes. Reeve (1970) and microscope slides attached to a rocker arm driven by an Reeve and Walter (1972) raised chaetognaths in 30-l automatic pipette washer. This moved the slides gently aquaria with daily water changes. Conover and Lalli (1972) forward and back at the top of the beaker, keeping the water kept the pteropod Clione limacina "indefinitely" in small stirred and aerated, and the food in suspension. This type of dishes and beakers with filtered water. Baker and Reeve system has also been used to raise a variety of larvae (1974) and Martindale (1987) raised the ctenophore Mne- (Strathmann, 1987). The water in the beakers was changed miopsis mccradyi in 30-l aquaria with gentle aeration, but and the beakers were cleaned regularly. A better arrange- had very low survival. Hirota (1972) used large jars for the ment for polyp cultivation uses flowing seawater, and the culture of the ctenophore Pleurobrachia bachei. Heron culture tank therefore requires an incoming water line and (1972) raised the salp Thalia democratica in small tanks an exit drain. Rectangular clear plastic boxes of various with lids that prevented the salps from encountering the sizes (pet cages available from most pet stores) make ideal air-water interface. Many researchers continue to raise small culture and grow-out tanks. Plastic containers can be easily hydromedusae, ctenophores, and other gelatinous organisms modified and are inexpensive, but any small tank can suf- in dishes, small-volume culture plates, and jars of various fice. Depending on the purpose of the tank, the drainage can sizes (Rees, 1979; Mills et al., 2000).
flow into another tank to collect newly released medusae, or The standard pelagic tank designs used today are all the drainage can be screened off with mesh (Fig. 1). If the variations of the planktonkreisel designed originally by exit drain is to be screened, the screen mesh must be smaller Greve (1968, 1970, 1975), which was modified and re- than the smallest medusae that will be released (mesh sizes designed for shipboard use by Hamner (1990) and for public of 120 –500 m are commonly used). In addition, the sur- display by the Monterey Bay Aquarium (Sommer, 1992, face area of the exit screen must be maximized so that the 1993). Paffenho¨fer (1970) described a rotating culture ap- drain pressure at any one point is low enough to prevent the paratus used very successfully for copepods, appendicular- medusae from being trapped against the screen. Screens are ians, and doliolids, which has been modified to various typically put across one entire side of the tank, several degrees (e.g., Sato et al., 2001; Gibson and Paffenho¨fer, centimeters from the drain.
2000). Ward (1974) described some simple aquarium sys- A simple way to set up a "medusa factory" using this tems for maintenance of ctenophores and jellyfish. Dawson technique is to clip a beaker or dish containing polyps to an (2000) devised a horizontal mesocosm that stratified by edge of the culture chamber, suspending it slightly above various salinity layers and may hold promise for species that the water level of the tank. Incoming water runs into the require complex water masses for development. The plank- polyp beaker and spills over the side into the tank (Sommer, tonkreisel design, however, has proved to be the most 1993). In this manner, newly produced medusae are washed useful, and it has been modified over the years into several out of the polyp beaker into the catch tank, where they will designs that offer more complex flow patterns and easier be safe from capture by the polyps (Fig. 1A). Alternatively, access to the inside of the tank and to the animals (Sommer, the polyps can be kept at the bottom of a tank without a 1992, 1993). Despite these alterations, the basic principles screened-off outflow. As the medusae are produced, they of the planktonkreisel remain unchanged.
tend to swim up; eventually most will go out the outflow. A The main chamber of the tanks is circular, with curved second catch tank with a screened-off outflow is placed sides and bottom and a flat back and front (Fig. 2). The below to collect the juvenile medusae (Utter, 2001). This water inlets and drains are designed to keep organisms from catch tank can be of similar design to the tanks described coming in contact with the screen that shields the drain.
K. A. RASKOFF ET AL. lid, which allows animals to be put into or removed from thekreisel without danger of being sucked down the drain. Alarger lid allows for easier access into the tank for cleaningand manipulation of the specimens (Sommer, 1993). Forscientific purposes, a matte black back plate allows for sidelighting of transparent plankton, achieving dark-field illu-mination (Hamner, 1990). For display aquaria, a mattetranslucent blue-and-white acrylic back, illuminated frombehind with fluorescent lamps, can be used to create theappearance of a lifelike blue-water environment. Spotlightsfrom the sides of the tank are used to illuminate animals fordisplay or photographic purposes. Strong lights do not ap-pear to bother many gelatinous species, which typicallyhave limited visual equipment. Most gelatinous organismscan do well in planktonkreisels (see Tables 1 and 2 for asummary). Plans of a planktonkreisel developed by KimReisenbichler at the Monterey Bay Aquarium ResearchInstitute (Fig. 2) are available for download at http:// Figure 2.
Contemporary planktonkreisel design showing separated inlet/outlet chamber and tank access lid. Detailed plans of this tank are Another variation on the planktonkreisel design is the available online at http://www.mbari.org/midwater/tank/tank.htm.
stretch kreisel, or Langmuir kreisel (Fig. 3). The tank hastwo inlet/outlet chambers that are located on each side of a Water flows from the inlet chamber and jets in a laminar rectangular tank, sending flow upward. The dimensions of flow across the lower side of a fine-mesh screen, which the rectangular tank (still with circular ends) must be about separates the main tank from the drain outflow. In this way twice as wide as tall, permitting the formation of two gyres, any specimen that drifts near the outflow screen will be one of which rotates clockwise and the other counterclock- pushed away by the incoming water. The placement of a wise. The top of the tank is open and the flows meet in the few parallel layers of polycarbonate double-wall sheet, middle, where they are joined by water added from a commonly used as greenhouse siding, into the space be- horizontally positioned perforated tube, creating convergent tween the inlet chamber and the main tank will force the currents that descend down the center of the tank. The two inlet water to enter with a smooth laminar flow. Modifica- opposing circular flows result in downwelling at the center tions to the planktonkreisels made by the Monterey Bay of the tank and upwelling at either end. This design works Aquarium include the construction of a separate outflow and well with species that tend to swim actively into a current Selected culture techniques for medusae commonly used for display Temperature (°C) Aurelia aurita Artemia, Krill Aurelia labiata Artemia, Krill Artemia, Juvenile Aurelia, Krill Pelagia colorata Artemia, Juvenile Aurelia, Krill Artemia, Juvenile Aurelia, Krill Artemia, Lighting Mastigias papua Artemia, Lighting Aequorea victoria Artemia, Rotifers (hydroids), Juvenile Aurelia, Eutonina Eutonina indicans Artemia, Rotifers Wild freshwater plankton, Frozen Daphnia Tima formosa Artemia, Rotifers Data summarized from Sommer (1992, 1993) for the Monterey Bay Aquarium.
* K ⫽ Kreisel; PK ⫽ Pseudokreisel; SK ⫽ Stretch kreisel; RF ⫽ Reverse flow; HP ⫽ Horizontal pseudokreisel; RT ⫽ Rectangular tank.
COLLECTION AND CULTURE OF GELATINOUS ZOOPLANKTON Selected culture techniques of non-cnidarian gelatinous zooplankton Temperature (°C) Wild-caught zooplankton Wild-caught copepods Wild-caught zooplankton Wild-caught zooplankton Baker and Reeve, 1974 Beroe spp.
Ctenophores, gelatin Beroe gracilis Beroe cucumis Clione limacina Wild-caught pteropods Conover and Lalli, 1972 Cliopsis krohni Sagitta hispida Reeve, 1970; Reeve and Walter, 1972 Pelagic Tunicates Oikopleura dioica Cultivated phytoplankton Paffenho¨fer, 1973 Cultivated phytoplankton Paffenho¨fer and Harris, 1979 nd ⫽ no data.
* K ⫽ Kreisel; PK ⫽ Pseudokreisel; RF ⫽ Reverse flow; RJ ⫽ Rotating jars.
(such as Chrysaora fuscescens), since they will tend tocongregate in the center of the tank, away from the walls(Tables 1 and 2).
Any rectangular tank can be modified into a "pseudo- kreisel," but care must be taken to ensure that the height andwidth of the tank are about equal, or the water in the tankwill not be able to rotate in a perfect circle and will createareas within the tank of limited flow where the animals mayaccumulate and contact the sides. Rectangular tanks aremodified by glueing a screen across the upper corner at anangle of about 30°– 40° from vertical in front of the over-flow (Fig. 4). Water enters the tank through a perforatedtube positioned so that the flow sweeps across the screendown towards the bottom of the tank. It is important that thetube is positioned so that the flow is parallel to the screenand covers the entire screen so that specimens are sweptaway rather than drawn against it. Curved plastic or vinylinserts are glued with silicone into the bottom corners toround them into a more circular shape. Friction-fitting stiffscreens can also be used to round the corners, although thisoption makes the tank more difficult to clean and maintainthan one with solid corners.
Several water quality issues are important for the suc- cessful culture and rearing of gelatinous organisms. Tem- Figure 3.
Stretch kreisel design showing the two rotating Langmuir perature and salinity must be kept within a range appropri- cells set up by the placement of the side and downwelling inlets.
K. A. RASKOFF ET AL. Figure 4.
Pseudokreisel design made from a standard tank. Bottom corners are filled in with silicone and solid pieces of plastic or vinyl. Outflow is separated from the tank by the inlet and screen.
ate for the species being reared. The water must be gently lift and collect debris. Kitchen basters work well for relatively clean and filtered, especially if the animals are to removing larger items because of their large reservoir vol- be used for any display purpose. Small particles in the water ume and wide bore. Siphons are best constructed from will quickly clog the outflow screens. Filtering the water small-bore acrylic tubes with flexible plastic tubing at- with 20-m pleated cartridge filters is usually sufficient; tached, so that the tubing may be pinched to stop flow if an however, some cultures that are very sensitive to biological animal gets too close to the suctioning tip. Additionally, fouling (such as many hydroid species) may need additional siphoning the "waste water" into a temporary container filtration to the 3 m level. Although air bubbles can be allows for the retrieval of any specimen that might inadver- helpful in the culture of many small gelatinous animals by tently be removed. To protect the insides of the tanks from increasing water circulation, they can be detrimental to scratches, it is helpful to dip the end of the acrylic tube into larger adult sizes (⬎3 cm). The bubbles can be ingested and liquid plastic, available from most hardware stores; alterna- collect in the gut and radial canals of medusae and cteno- tively, a small ring of Nalgene tubing may be placed on the phores, causing the animals to become positively buoyant, end of the siphon tube. Floating layers of lipid-rich mate- disrupting their normal swimming and feeding behaviors. A rials can be removed by skimming with small jars or more serious problem is that these bubbles will slowly work beakers or fine-meshed nets, or by absorbing the material themselves through the mesoglea, which can lead to infec- onto paper towels floated on the surface of the water. The tion. A degassing system for the water may be needed if the sides of the tanks can be cleaned by wiping with brushes incoming water tends to be supersaturated. A degassing (firm paint brushes work well) or non-abrasive pads. For tower in which the water trickles down through small plastic larger tanks (⬎75 l), painting or scrub pads can be covered balls or other material serves to degas the water before it with nonabrasive nylon mesh fabric and attached to poles enters the tank. Deep-sea animals may be sensitive to the for cleaning hard-to-reach areas of the tanks. The wood or high oxygen concentrations of surface waters. Reducing the metal handles of these scrubbers can be covered in plastic oxygen concentration in tank water by bubbling nitrogen tubing to reduce the adherence of tentacles. Flow to the gas has been used in the past with some success, although it tanks can also be temporarily shut off and the animals does not appear to be critical for most deep-sea species.
allowed to collect on the bottom of the tank during cleaning.
Also, tanks can be cleaned just after the animals have been fed, when tentacles are typically retracted and less apt to Throughout the course of feeding and rearing, tanks ac- become ensnared (C. Widmer, Monterey Bay Aquarium, cumulate debris that should be removed regularly. The use per. comm.). Screens in the tanks collect debris quickly and of pipettes, small brushes, basters, and siphons for removing need to be scrubbed and cleaned at regular intervals. When larger debris, including waste, uneaten Artemia, and other screens become clogged, organisms are more likely to stick food items, will help keep the tanks clean and discourage to them, possibly with fatal results.
fouling growth. Pipettes of any size and type can be used to Even with proper cleaning and filtration, biofouling in COLLECTION AND CULTURE OF GELATINOUS ZOOPLANKTON culture and rearing tanks can become a serious problem. In Through the use of culture methodologies, laboratory- some cases of diatom and algae fouling, reducing the light based experimentation on salps and larvaceans has begun to that shines on the tank can help reduce growth, but typi- address important ecological questions about the role these cally, scrubbing the tanks eventually becomes necessary.
animals play in the nutrient cycling of the oceans and their When diatom, hydroid, or other fouling organisms cannot impact on the ecosystem. These organisms have some of the be satisfactorily removed by any of the means discussed fastest generation times and largest nutrient turnovers in the previously, bleaching is necessary. This can be especially world, and their fecal pellets and associated "marine snow" useful on the screens, pumps, and waterlines, which can be are important sources of carbon transport into the deep sea very difficult to clean by other means. The entire tank (e.g., Alldredge, 1972; Silver et al., 1998).
system may need to be bleached every 1– 6 months, depend- Recent laboratory studies have shown that some species ing on the size and fouling rate. During bleaching, the of medusa have chemically-regulated feeding behaviors occupants of the tank must be removed and transferred to a (Arai, 1991, 1997; Tamburri et al., 2000), with several holding facility. The longer the tanks and lines are allowed different chemical stimuli controlling the feeding and swim- to bleach, the more complete the fouling kill will be. Over- ming of both hydrozoan and scyphozoan medusae. Tank- night is preferred, but bleaching for even an hour kills most based studies on the vertical migration of medusae (Mackie fouling organisms. As a rule of thumb, 1 l of standard et al., 1981; Mills, 1983) and on their swimming and 3%– 6% sodium hypochlorite (NaOCl) bleach will treat feeding behaviors (e.g., Costello and Colin, 1995; Suchman about 200 l of water (⬇1 gallon bleach/800 gallons of and Sullivan, 2000) have provided much information on the water), but this amount can be increased or decreased de- physiological and behavioral components of medusa loco- pending on the severity of the fouling and the time available motion as it relates to prey selection and capture.
to let the tank bleach. The water level in the tank should be The interactions between gelatinous zooplankton and hu- dropped so that there is no overflow when the bleach is mans are increasing, whether from envenomation (Burnett, added. If the tanks have self-contained pumps, these should 2001); blooms that clog power plant intakes (Masilamoni et be run at a high flow rate to mix the bleach and flush it into al., 2000); interactions, both positive and negative, with the pump housings.
fisheries (Mutlu et al., 1994; Mutlu, 1999; Mills, 2001; To complete the process, the bleach must be neutralized.
Purcell and Arai, 2001); or the general increase in gelati- This can be accomplished by adding about 60 g of sodium nous zooplankton populations in perturbed or eutrophic thiosulfate (Na S O ) per liter of bleach used (⬇1 cup/ environments (Mills, 1995, 2001; Arai, 2001). The oppor- gallon). The sodium thiosulfate crystals may be dissolved in tunities for scientific studies of gelatinous zooplankton are a bucket of water prior to adding to the tank. When the color vast and largely untouched. We hope researchers can use of the water in the aquarium changes from yellow-green to some of the techniques presented here to expand the re- clear, sufficient thiosulfate has been added for neutraliza- search being done on these important but poorly understood tion. Allow the thiosulfate several minutes to run through marine organisms.
the entire tank and pumps. The treated water is then drainedfrom the tank and discarded. While draining, thoroughly The public's fascination with and appreciation of gelati- rinse out the tank with freshwater. Stubborn growth can be nous zooplankton is growing rapidly. What were once con- removed at this time by scrubbing. After all debris and sidered nasty animals that might sting or otherwise disturb treated water is removed, begin to refill the tank with beachgoers are now a major attraction in public aquaria all seawater, minimizing turbulence and bubbles during the over the globe. The time and money spent by the aquarium refilling since bubbles will stick to the walls of the tank and industry to provide compelling exhibits on gelatinous zoo- will have to be removed before gelatinous animals are plankton is a testament to their appeal. Over 3.4 million people visited the Monterey Bay Aquarium during the tem-porary "Planet of the Jellies" exhibit in 1992 and 1993(Powell, 2001; J. Tomulonis, Monterey Bay Aquarium, pers comm.). Jellyfish and ctenophores were given permanent The use of the techniques described herein for the cap- starring roles in the Outer Bay Wing, and in a new tempo- ture, culture, and rearing of gelatinous zooplankton has rary exhibit, "Jellies: Living Art." Aquarists in the United allowed researchers to address many important biological States and elsewhere are responsible for many of the tech- issues. Historically, these contributions were limited pri- niques discussed in this paper. Aquariums around the world marily to the disciplines of systematics, developmental bi- provide the bulk of the layperson's information on gelati- ology, and evolution. More recently, new advances in our nous zooplankton, and we hope that the rising public ap- understanding of behavior, physiology, ecology, and ocean- preciation of these important and beautiful animals may ographic processes from the sea surface to the abyssal lead to increased financial and societal support for their depths have also been possible.
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IJMCM Original Article Autumn 2015, Vol 4, No 4 Expression Pattern of Neuronal Markers in PB-MSCs Treated by Growth Factors Noggin, bFGF and EGF Zahra Fazeli1#, Sayyed Mohammad Hossein Ghaderian1,2#. Masoumeh Rajabibazl3, Siamak Salami3, Nader Vazifeh Shiran4, Mir Davood Omrani1,2∗ 1. Department of Medical Genetics, Faculty of Medicine, Shahid Beheshti University of Medical Sciences,