Biological test method: fertilization assay using echinoids (sea urchins and sand dollars), chapter 4
Section 2: Test Organisms
The test must be carried out using one of the species listed below.
- Strongylocentrotus droebachiensis (O.F. Müller), the green sea urchin, a circumpolar species found on the Canadian Atlantic and Pacific coasts and across the Arctic Ocean to 80 °N.
- Strongylocentrotus purpuratus (Stimpson), called in this report the Pacific purple sea urchin (and commonly called the purple sea urchin), found on the Pacific coast of Canada and southwards to Baja California (Meinkoth, 1981).
- Dendraster excentricus (Eschscholtz), a sand dollar of the Pacific coast of Canada and southwards, called in this report the eccentric sand dollar, a standard common name (Meinkoth, 1981).
- Arbacia punctulata (Lamarck), called in this report Arbacia, although the common name of “Atlantic purple sea urchin” is sometimes used (Meinkoth, 1981). Found on the Atlantic coast of the United States from Cape Cod southerly into the Caribbean and Gulf of Mexico.
- Lytechinus pictus (Verrill), the white sea urchin, found from southern California to Panama.
The first four species can be collected on one or more Canadian coasts. All five species can be purchased from biological supply houses and shipped to the test laboratory.
All of these species have been listed as echinoids commonly used in the laboratory (NRC, 1981). Most of the species have been used frequently in toxicity tests (Appendix D). According to feedback provided by Canadian and US laboratories in response to a questionnaire circulated prior to the preparation of this second edition (see Section 1.1), the green sea urchin (S. droebachiensis) has been used the least of the five candidate test species included in the first edition of EPS 1/RM/27. This species is desirable from a Canadian perspective, however, since it is the only echinoid sp., listed herein, that is found in Pacific, Atlantic, and Arctic waters. In recent testing, it was confirmed that green sea urchin eggs are relatively large (i.e., 3 to 4 times larger than the other species listed herein), have highly visible fertilization membranes, and show good fertilization rates in uncontaminated seawater (Jackman, pers. comm., 2008) . These factors along with their presence on all three Canadian coasts have resulted in the green sea urchin being retained in this second edition of EPS 1/RM/27.
In general, toxicity results from fertilization assays using echinoids appear to be similar among species (Kobayashi, 1984; Nacci et al., 1986). There might, however, be differences in species sensitivity, depending on the toxicant being tested. For example, the eccentric sand dollar appears to be about 1.4 times more sensitive to sodium dodecyl sulphate than the Pacific purple sea urchin, and 1.7 times more sensitive to bleached sulphite mill effluent than the green sea urchin (NCASI, 1992). In a more recent study involving 3 species of sea urchins (Pacific purple, Arbacia, and white) and 1 species of sand dollar (eccentric), the white sea urchin and the eccentric sand dollar were found to be the most sensitive to specific samples of sediment pore water from Vancouver and Halifax Harbours, whereas Arbacia had the greatest sensitivity to ammonia. In the same study, the Pacific purple sea urchin and the eccentric sand dollar had the greatest sensitivity to copper (Jackman and Doe, 2004).
The common sand dollar, Echinarachnius parma (Lamarck), has not been used frequently in toxicity tests, and performed poorly in a multi-species interlaboratory evaluation of this echinoid fertilization assay (Miller et al., 1992). Accordingly, the common sand dollar is not presently recommended for the test until research proves suitable. The circumpolar distribution of the common sand dollar, including its frequent occurrence along the Atlantic coast of Canada southwards to Maryland (U.S.A.), support further research with this species. Adult common sand dollars were successfully used for month-long toxicity tests in Newfoundland by Osborne and Leeder (1989). The effect of growth-inhibiting chemicals and sediment contaminants on the early life stages of this sand dollar have been studied (Karnofsky and Simmel, 1963; Meador et al., 1990).
2.2 Life Stage, Size, and Source
Mature and gravid echinoids should be obtained to provide the gametes. Adult sizes range from about 3-cm diameter upwards for the various species (Table 1); a common size-range for specimens in the laboratory is 5 to 6 cm.
All adults used to provide gametes for a test should be derived from the same batch and source. The native species can be collected from clean-water coastal marine locations, some in shallow water at low tide, or by diving. All species can also be purchased from biological supply houses. Adult echinoids must be positively identified to species. Organisms that are purchased from a commercial supplier should be supplied with certification of the organisms’ species identification, and the taxonomic reference or name(s) of the taxonomic expert(s) consulted. After the initial taxonomic identification of each species provided by a given supplier, confirmation of the species of test organisms in a shipment can be conducted by the testing laboratory. All information needed to properly identify the adult echinoids transported to a testing laboratory must be provided with each shipment. Records accompanying each batch of test organisms must include, as a minimum: the quantity and source of test organisms in each shipment, supplier’s name, date of shipment, date of arrival at the testing laboratory, arrival condition, and species identification. To ensure that the echinoids’ health is maintained during transit, appropriate temperature, DO, and salinity conditions should be maintained as much as possible. Shipping containers should be insulated to minimize changes in temperature during transit. If the organisms cannot be delivered on the same day that they are shipped, the transport containers should be stored in such a way that the temperature of the echinoids is held as constant as possible. The temperature of the test organisms should be recorded upon departure from the supplier’s facility and upon arrival at the testing laboratory.
The spawning seasons listed in Table 1 show that in a given location, tests could be carried out for much of the year by collecting sea urchins and sand dollars at appropriate times.
The testing season could be lengthened by maintaining the adults at warm or cool temperatures to encourage early or late spawning. The green sea urchin is in spawning condition for only a few months in the spring (see Table 1), however Canadian laboratories might be able to obtain gametes of the green sea urchin over most or all of the year by such changes in holding conditions (Wells, 1982, 1984). The other alternative would be to purchase species that had a suitable spawning time, from another location. It should be realized that animals from different sources and climatic conditions might show variations in timing and length of spawning season, or in the optimum temperature for bringing about spawning. Sea urchins that are spawned early in the season can sometimes provide gametes again in a month or six weeks if fed a proper diet (Dinnel and Stober, 1985). These sea urchins should be held in a separate tank after the first spawning.
|Species||Spawning SeasonFootnote a||Maximum Diameter of Adult (cm)Footnote b||Holding Temperature in Laboratory (°C)Footnote c|
|Green Sea Urchin||generally April, but March to May at specific Canadian locations; a later cycle to June in the St. Lawrence estuary (January, June +)||8.3||12 ± 2, ≤15|
|Pacific Purple Sea Urchin||generally January to May, optimally January to March for feral animals; late October to April on California coast (December, June)||10||10 ± 2, ≤17|
|Eccentric Sand Dollar||May through summer to October (February to December)||9||13 ± 2, ≤17|
|Arbacia||June to August on Atlantic Coast; January to April on Gulf CoastFootnote d||5.1||17 ± 2, ≤22|
|White Sea Urchin||March through summer to November||♂2.8
|13 ± 2, ≥8 & ≤17|
Maturation should be checked before attempting to carry out a toxicity test with the gametes. Sperm and eggs obtained outside the main period of maturation can give poor fertilization rates and poor test results. Inspections for state of maturity require some experience on the part of the investigator, but can be assessed by spawning a sample of echinoids and examining the gametes (Section 4.2.1). Mature sperm are minute and quickly become very active in seawater. Mature eggs rapidly become spherical in seawater. Immature eggs have a clear spot in the cytoplasm. Some adults could be sacrificed to examine the gonads, and to obtain gametes directly instead of by forced spawning. In sea urchins, mature ovaries are coloured yellow to red depending on species, and testes are white.
Shipping extremely ripe or gravid individuals under stressful conditions (e.g., extreme temperature changes) might cause spawning or mortality during shipment or upon receipt. This can be avoided by having the animals acclimated to laboratory conditions, as much as possible, prior to being shipped. Adults should be shipped dry with cold packs to keep the temperature at 10 ± 2° C. The Pacific purple sea urchin can be shipped surrounded by algae or some other moist material. Shipping these organisms submerged in water might lead to oxygen depletion. The eccentric sand dollar, on the other hand, should be shipped in a small amount of chilled seawater. If adult echinoids spawn prematurely (i.e., during shipment or upon receipt in the laboratory), they can be separated by sex, and from those that have not spawned, and then housed in separate holding tanks.
Moving animals from one location to another marine location raises serious questions of introducing non-native species or transporting diseases and parasites. Any proposed procurement, shipment, or transfer of echinoids should be submitted for the approval of federal, provincial, or regional authorities. Provincial governments might require a permit to import organisms whether or not the species is native to the area, and movements of aquatic organisms might be controlled by a Federal-Provincial Introductions and Transplant Committee. Advice on contacting the committee or provincial authorities, and on sources of echinoids, can be obtained from the regional Environmental Protection Office (Appendix C). Application for a permit to bring in animals must be made to the above-mentioned committee, to the appropriate provincial agency, or to the Regional Director-General of the Department of Fisheries and Oceans (DFO), depending on procedures in place locally.
Testing laboratories might be required to establish and use a quarantine section within their facilities, where imported organisms or gametes can be isolated and all equipment and fluids that come in contact with the test organisms or gametes can be sterilized and disposed of according to provincial or federal regulations. Standard operating procedures detailing quarantine operations and procedures might also be required by provincial agencies or DFO.
2.3 Holding and Acclimating Adults in the Laboratory
Groups of male and female echinoids are held in tanks and used to provide gametes when required for a test. There is no particular limitation on time that the adults may, or must be kept in the laboratory before providing gametes. All five test species have been successfully maintained in spawning condition in the laboratory for extended periods of time (i.e., 3 months to 1 year), lthough some species (i.e., Arbacia and the white sea urchin) are reportedly more easily maintained than others. For the Pacific purple sea urchin and the eccentric sand dollar, there are varying reports on the ease with which these species can be held in the laboratory for extended periods of time. Many Canadian laboratories have resorted to purchasing these two test species from a commercial supplier when tests are requested, and spawning adults on the day of, or within a few days after arrival at the laboratory (i.e., without a thorough acclimation). Problems associated with holding these test species for extended periods of time include: adults spontaneously spawn prior to testing; adults spawn for only a short period of time after arrival at the laboratory; and difficulty keeping them healthy or alive for extended periods of time (>3 months). As a result, this second edition of EPS 1/RM/27 includes an option for “holding adults for immediate use”, where gametes may be collected within a short period of time (≤ 3 days) after the adults are received at the laboratory.Footnote 1
For adults that are to be spawned and gametes tested within a 3-day period after the adults arrive at the testing laboratory, confirmation should be obtained from the supplier that the adults are mature and that the eggs are viable prior to shipping. The temperature at which the test organisms are shipped should be maintained at or near the required test conditions, since there is little if any time for acclimation upon arrival at the testing laboratory. Even with “holding for immediate use”, the adults should be moved to laboratory holding conditions as gradually as possible, so that the stress of the rapid changes in holding conditions for the sexually-mature adults does not influence the test results for their gametes (i.e., test validity criteria are met, see Section 4.5.1). Gradual exposure of the adult echinoids to the testing laboratory’s control/dilution water is recommended in all cases, but especially in instances where there is a marked difference in quality (i.e., temperature, salinity, pH) from that to which they were previously acclimated. This gradual exposure should minimize any stress on the animals caused by different water quality characteristics. For adults that are to be spawned for testing on the same day that they arrive at the laboratory, a minimum holding period of three hours is required to allow for observation of the general health of the adults and to move the adults from their shipping conditions (i.e., temperature and water) to testing conditions. For adults shipped in water, a useful procedure for moving adults from their shipping water to control/dilution water prior to spawning is to hold them for 1-2 h in a 50:50 mixture of shipping water:control/dilution water, then for 1-2 h in a 25:75 mixture of shipping water:control/dilution water, followed by a final 1-2 h in 100% control/dilution water before they are spawned. Another useful procedure is to siphon off 20 to 30% of the shipping water every 1 to 2 hours and replace it with laboratory control/dilution water until at least three exchanges have been made. Adults that are shipped “dry” (i.e., wrapped in moist paper towel or seaweed), do not have to be placed in control/dilution water prior to spawning, however they must be held for a 3-hour observation period, prior to spawning, and any adjustment of their temperature (i.e., air temperature) to the test temperature should be made as gradually as possible, if necessary. The shift of adults from shipping conditions to test conditions should be started as soon as possible after the sexually-mature adult echinoids arrive at the testing facility.
Some Canadian and US laboratories have had good success in holding both the eccentric sand dollar and the Pacific purple sea urchin in the laboratory for extended periods of time. Success in holding both of these species might be due to the use of fairly simple systems with lots of clean natural seawater flowing continuously through the holding tanks.Footnote 2 For the Pacific purple sea urchin, these conditions might include temperatures below 15 °C, high DO, pH > 8, good water flow, prompt removal of fecal material, and organisms being held in the dark (Bay and Greenstein, pers. comm., 2008; and Buday, pers. comm., 2008). Problems experienced by other laboratories could be due to a combination of adverse conditions during transport, poor exchange rate of seawater in the holding tanks, and prolonged exposure to artificial sea salts of a source and mixture that is foreign to the echinoids.
Both Arbacia and the white sea urchin have been successfully and easily held in the laboratory for extended periods of time (> 1 year), and spawned repeatedly (every 4 to 6 weeks) throughout the year when maintained under the right conditions (Doe and Jackman, pers. comm., 2008; Jonczyk and Holtze, pers. comm., 2008; and Carr, Nipper, and Biedenbach, pers. comm., 2008). They can be sexed and housed in separate aquaria to facilitate the quick selection of the appropriate numbers of males and females required for testing. Most laboratories report very little mortality with either of these two species during acclimation and holding. Both Arbacia and the white sea urchin are easily acclimated and maintained in closed, recirculating, temperature-controlled aquariums.
For adult echinoids that are going to be held in the laboratory for extended periods of time (i.e., > 3 days), it is desirable to provide a gradual acclimation and a minimum holding time of 3 or 4 days, at the test temperature, salinity, and in the water to be used for controls and dilution, prior to gamete collection. Acclimation should be started as soon as possible, upon arrival of the adults at the testing facility. The need for appropriate procedures for “holding for immediate use” or gradual acclimation and satisfactory long-term holding conditions, is dependent on the requirement for the delivery of viable gametes that meet the needs and validity criteria (see Section 4.5.1) of the test.
Echinoids must be handled with care and should not be subjected to sudden shocks or changes in holding conditions. In particular, large changes in temperature or hydrostatic pressure might stimulate spawning at a time that is not desired by the investigator (Dinnel and Stober, 1985). Some laboratories that use natural seawater without fine filtration have noticed mass spawning of sea urchins occurring at times of plankton blooms, and the phenomenon has been observed in Canadian waters (Starr, 1990; Starr et al., 1990). In addition, spawning by individual animals might induce others to spawn, so such animals should be isolated immediately upon detection, to prevent mass spawning.
For adult echinoids that are going to be maintained in the laboratory for an extended period of time (i.e., > 3 days), the recommended conditions for holding echinoids, outlined in the following Sections 2.3.2 to 2.3.10 and summarized in Tables 1 and 2, are intended to allow some degree of flexibility within a laboratory, while at the same time standardizing those elements which, if uncontrolled, might affect the health of animals or viability of their gametes. Where applicable, guidance on “holding for immediate use” (i.e., spawning of adults within 3 days of arrival at the laboratory) is also provided. Recommended conditions have been drawn, in general, from Appendix D, and guidance derived from the feedback provided by US and Canadian laboratories responding to a questionnaire (see Section 1.1). Further details and rationale are given in some of the publications included in Appendix D, and in the References, particularly ASTM (1990), USEPA (1988, 1994, 1995, 2002), NCASI (1991), and papers of Dinnel and colleagues listed in the References and in the Bibliography of Appendix E. In addition, the website developed at Stanford University, might also provide useful details related to sea urchin care and embryology.
2.3.2 Holding Containers
Adults may be held in aquaria, troughs, or tanks made of nontoxic materials such as glass, stainless steel, porcelain, fibreglass-reinforced polyester, perfluorocarbon plastics (TeflonTM), acrylic, polyethylene, or polypropylene. Tanks containing about 50 to 150 L of water, and fitted with a standpipe drain, are most commonly used. The holding tanks should be located away from any major physical disturbances and preferably in a location separate from that used for testing. To help avoid undesired mass spawning, adults should be held in groups of 20 or fewer animals.
For sea urchins, the water depth should be ≥20 cm. For sand dollars, trays are frequently used, for example, 1 × 2 m with a water depth of 10 cm. There should be 2 to 3 cm of sediment or sand, rich in detritus including settled algal cells, on the bottom of containers used for sand dollars.
The lighting conditions for holding echinoids currently used by various Canadian and US laboratories are quite varied and include: ambient laboratory light levels (100 - 500 lux), with a 16-h light : 8-h dark photoperiod; natural outside light and seasonal photoperiod (i.e., tanks are outside); or complete darkness. For sea urchins, the strength of lighting and photoperiod do not seem to be of major importance, and a low intensity of normal laboratory lighting is most common. For sand dollars, overhead fluorescent lighting at the equivalent of bright office lighting encourages algal growth on the sediment, which might result in desirable nutritional self-sufficiency for the tray of sand dollars.
For adult echinoids that are to be held for a prolonged period (i.e., > 3 days) prior to collecting gametes for use in a test, ambient laboratory lighting (100 - 500 lux) and a 16-h light : 8-h dark photoperiod are recommended. For tanks that are maintained outside, normal daylight and a seasonal photoperiod are recommended.
For the Pacific purple sea urchin, some laboratories have reported that cultures have been maintained successfully in complete darkness for extended periods of time. Constant darkness might disrupt some of the seasonal patterns of the animals that provide them a cue to spawn, so that they will maintain ripe gonads for a longer period of time (Bay, pers. comm., 2008). In instances where adults are transferred to a testing facility for “same-day” collection of gametes or collection of gametes within an ensuing period of 3 days or less, lighting conditions representing those to be used in the test are recommended.
The water in containers holding adults should be renewed continuously or periodically to prevent a buildup of metabolic wastes. The water may be either an uncontaminated supply of natural seawater or reconstituted seawater (also known as artificial seawater), made up to a desired salinity according to Environment Canada’s recommended procedure (EC, 2001). Any commercially-available sea salts (e.g., Instant OceanTM, Ocean Pure Sea SaltTM, Red Sea SaltTM) or appropriate mixture of reagent-grade salts (e.g., modified GP2; see Bidwell and Spotte, 1985 or Table 2 in USEPA, 1994 or USEPA, 1995), used to prepare the reconstituted water, should have previously been shown to consistently and reliably support good survival and health of echinoids. The water supply should be monitored and assessed as frequently as required to document its quality. Temperature, salinity, dissolved oxygen, pH, and the volume of flow to each tank should be measured, preferably daily. Assessment of other variables such as total dissolved gases, ammonia, nitrogen, nitrite, metals, pesticides, suspended solids, and total organic carbon, should be performed as frequently as necessary to document water quality.
As a general guideline for the optimal maintenance of high-quality water, the flow rate of seawater in “once-through” systems should provide 5 to 10 L/d or more for each organism held, and have a flow that equals the tank volume in 6 to 12 h. For static holding tanks, a similar and acceptable exchange rate would be replacement of most of the water on a daily basis. There is no apparent consensus for optimal amounts of water and exchange times in the existing methods (Appendix D), nor among Canadian and US laboratories responding to a questionnaire in a recent survey (see Section 1.1).Footnote 3 Most methods do not specify the flow, and the few that do, range from a high rate of hundreds of litres per animal per day, with an inflow equalling the tank volume in a few minutes, to lower rates which equal the tank volume in about 5 h. NCASI (1991, 1992) uses seawater flows similar to those recommended here, with 7 to 14 L/d per sand dollar and flow that equals the tank volume in 1.3 to 2.7 hours.
|Source of Adults||collected from clean-water areas or purchased from supply houses|
|Water||uncontaminated natural seawater or reconstituted (artificial) seawater; flow-through or semi-static (e.g., once every 24 h) replacement; average salinity from 28 to 34 g/kg, and individual measurements not outside 25 to 36 g/kg; rate of salinity change ≤5 g/kg/d for adults to be held for >3 d; as a general guideline, volume of flow should provide 5 to 10 L/d for each animal and equal the volume of tank in 6 to 12 h|
|Temperature||from 10 to 17 °C depending on species, somewhat lower or higher to delay or speed spawning, see Table 1; rate of temperature change ≤5 °C/d for adults to be held for >3 d|
|Oxygen/aeration||dissolved oxygen 80 to 100% saturation; maintained by aeration with filtered, oil-free air if necessary|
|pH||within the range 7.5 to 8.5, in normal circumstances 8.0 ± 0.2|
|Water quality||monitor temperature, salinity, dissolved oxygen, pH, and flow to each holding tank, preferably daily|
|Lighting||normal laboratory lighting at low intensity, 16-h light : 8-h dark; normal daylight, seasonal photoperiod; or complete dark. Lighting conditions not considered critical|
|Feeding||for sea urchins; kelp, other macroalga, or romaine lettuce, spinach and carrots; for sand dollars; provide sediment with detritus and alga, use lighting to encourage growth of algae, and if necessary add cultured alga or algal paste|
|Cleaning||removal of old alga, fecal material, and debris, daily or as required, unless intended as food|
|Disease/mortality||monitor mortality daily; for adults held >3 d, mortality should be ≤2%/d averaged over 7 d preceding collection of gametes, and cumulative mortality over the same 7-d period must be ≤20%; for adults held ≤3 d, cumulative mortality must be ≤20%; remove diseased or moribund animals; groups of diseased animals should be discarded|
“Less-than-optimal” exchange rates and loading densities may be used provided that the criterion for survival in holding tanks (see Section 2.3.10) as well as those for test validity (see Section 4.5.1) are achieved. The regular monitoring and documentation of water quality variables in holding tanks (ammonia and nitrite in particular) is highly recommended, and values should be compared to the recommended target values to ensure that metabolic wastes do not reach harmful levels in the holding tanks. Target values, recommended for the protection of aquatic organisms, are ≤0.02 mg/L of un-ionized ammonia and ≤0.06 mg/L of nitrite (CCREM, 1987).Footnote 4
The average salinity of the holding water should be 28 to 34 g/kg, preferably 30 to 32 g/kg. Extreme salinity values must not be less than 25 or should not be more than 36 g/kg during holding of echinoids.Footnote 5 For organisms that are to be held in the laboratory for extended periods of time, the rate of any salinity adjustment should be ≤3 g/kg/day and must be ≤5 g/kg/day. Certain situations (e.g., adults spawned for testing on the same day they arrive in the laboratory), however, might require a daily shift of more than 5 g/kg/day. Many laboratories have reported that for tests initiated on the same day or the day after adults arrive in the laboratory, rapid changes in salinity of 6 - 8 g/kg have no effect on the gametes of the test organisms. Therefore, for adults “held for immediate use” (e.g., spawning adults for test purposes within 3 days of arrival at the laboratory), salinity adjustments should be made as gradually as possible. However, a daily shift of >5 g/kg may be made if the criteria for test validity can be met (see Section 4.5.1), and the sensitivity of the gametes in reference toxicant tests is not affected (see Section 4.6).
There are reportedly some species-specific differences in salinity tolerance among the echinoids listed herein. During a recent survey (see Section 1.1), some Canadian and US laboratories reported that the Pacific purple sea urchin and the white sea urchin thrive better at higher salinities (34-35 g/kg), however other laboratories reported being able to maintain these species at salinities of 28 to 30 g/kg without problems. For the white sea urchin, salinities <28 g/kg might result in stressed animals and high mortality. Some laboratories reported that the eccentric sand dollar has been found to be slightly more sensitive to large salinity changes, and to salinities lower than 32 g/kg.
Water entering the containers should not be supersaturated with gases, as might occur if the water were warmed. If that is a valid concern, total gas pressure in the water should be checked frequently (Bouck, 1982). Remedial measures must be taken (e.g., use of aeration columns or vigorous aeration in an open reservoir) if dissolved gases exceed 100% saturation.
If reconstituted (artificial) seawater is to be used as dilution and control water (see Section 4.1.1 and Section 5.3), and if adults are going to be held for >3 days after arrival in the laboratory, adults should be acclimated to that water for at least three days immediately before they are forced to spawn. Holding in reconstituted seawater or in limited seawater supply might require filtration and recirculation of water, or its periodic renewal in static systems; ammonia and nitrite should then be measured frequently to check that they do not reach harmful levels.
If reconstituted (artificial) seawater is to be used, it must be made up to the desired salinity by adding hypersaline brine (HSB) and/or commercially-available dry ocean salts or reagent-grade salts to the appropriate quantity of a suitable fresh water (see EC, 2001 for guidance). The HSB should have a salinity of 90 ± 1 g/kg. Any reconstituted water prepared by the direct addition of dry salts must be aerated vigorously for a minimum of 24 h before being used (EC, 2001), however, longer periods of aging (i.e., ≥3 days) with aeration are recommended.Footnote 6
The use of HSB derived from an uncontaminated, source of high quality (and preferably high salinity) natural seawater is recommended (EC, 2001), however, artificial hypersaline brine may also be prepared using commercially available dry ocean salts (e.g., Instant OceanTM ) or reagent-grade salts (i.e., “modified GP2;” see Bidwell and Spotte, 1985 or Table 2 in USEPA, 1994 or USEPA, 1995). Any artificial HSB which is prepared using commercial sea salt or reagent-grade salts must be filtered (≤1 µm), and then aerated vigorously for a minimum of 24 h before use, however longer periods of aging (i.e., ≥3 days) with aeration are recommended.Footnote 6 HSB derived from natural seawater should be filtered (≤1 µm) and may be used immediately for salinity adjustment. Unused portions of prepared natural or artificial HSB should be capped and stored in the dark at 4 ± 2 °C until used (EC, 2001). Additionally, testing laboratories should obtain the “best quality” of commercial sea salts (e.g., Forty FathomsTM Toxicity Test Grade) available from the supplier. The suitability and consistency of any new products or batches should be evaluated for their ability to meet the test-specific validity criteria (see Section 4.5.1), before artificial seawater is used to prepare HSB or control/dilution water (EC, 2001), since some investigators feel that specific types and/or batches of sea salt might produce low fertilization rates in controls, produce unwanted toxic effects, and/or sequester test substances. If ocean salts are used to prepare HSB or control/dilution water, the suitability and consistency of these salts should also be verified by testing.
Hypersaline brine derived from natural seawater may be prepared by concentrating seawater (natural or, less desirably, artificial) by freezing or evaporation. The seawater should be filtered to at least 10 µm before placing it into the freezer or the evaporation chamber. Once prepared, its salinity should be 90 ± 1 g/kg (EC, 2001). If prepared by freezing, freeze at -10 to -20 °C for ≥6 h, and collect the HSB under the ice when it reaches a salinity of 90 ± 1 g/kg. If prepared by evaporation, heat the seawater in a non-corrosive, non-toxic container at ≤40 °C while aerating it, until the desired salinity (i.e., 90 ± 1 g/kg) is achieved (USEPA, 1994, 1995; EC, 2001). Regardless of which technique is used (i.e., freezing or evaporation), the salinity of the brine should be monitored during its preparation, and must not exceed 100 g/kg. HSB may be added to natural seawater, fresh water, distilled water, deionized water, or test samples, to increase the salinity to the level desired for testing. Guidance in Environment Canada (2001) should be followed when preparing, aging, and storing HSB. If HSB with a salinity of 90 g/kg is used to prepare control/dilution water with a salinity of 30 g/kg (see Sections 3.4 and 4.1.1), the maximum concentration of effluent (or other freshwater sample) that could be tested would be 67%.Footnote 7 If, however sample salinity is adjusted by the direct addition of dry salt to the freshwater sample, toxicity can be determined at 100% test concentration.
Sources of water used for preparing reconstituted seawater may be deionized water, distilled water, an uncontaminated supply of groundwater or surface water, or dechlorinated municipal drinking water. If municipal or natural freshwater sources are used, this water should also be chemically assessed as appropriate to document its quality, for example the items listed at the beginning of this Section (2.3.4).
If municipal drinking water is to be used for preparing reconstituted seawater, effective dechlorination must rid the water of any harmful concentration of chlorine. The target value for total residual chlorine in water used for holding, control tests or dilution, is ≤0.002 mg/L (CCREM, 1987). Available chlorine as low as 0.05 mg/L is a potent spermicide for echinoids (Muchmore and Epel, 1973). Vigorous aeration of the water might strip out volatile chlorine gas. The use of activated carbon (bone charcoal) filters and subsequent ultraviolet radiation (Armstrong and Scott, 1974) is recommended for removing residual chloramine and other chlorinated organic compounds.Footnote 8
Echinoids may be held at the optimum temperature ranges identified herein (Table 1), or, if desired, at normal seasonal temperatures (i.e., using the temperature of the incoming natural seawater supplied to the laboratory). A pre-spawning optimum temperature of 12 ± 2°C for the green sea urchin, 10 ± 2°C for the Pacific purple sea urchin, 13 ± 2°C for the eccentric sand dollar, 17 ± 2°C for Arbacia, and 13 ± 2°C for the white sea urchin, should be maintained.Footnote 9
Groups of adults may be held for delayed spawning at temperatures that are lower than the seasonal norm for their habitat, the exact values varying with the species and the desired degree of delay. Laboratories responding to a questionnaire, however, indicated that they had little experience with holding adults at lower temperatures since it is typically not necessary to manipulate test organisms at lower temperatures to delay spawning. Arbacia and white urchins can be held in the laboratory for extended periods of time and therefore it is unnecessary to hold them at lower temperatures to delay spawning, however Arbacia has been reported to feed and regenerate faster if they are held at 15 °C after spawning. Similarly, temperature may be raised to encourage early development of gametes. Excessively high temperatures should be avoided in order to prevent spontaneous spawning and stress. Recommended upper limits are 15 °C for green sea urchins, 17 °C for Pacific purple urchins and eccentric sand dollars, 22 °C for Arbacia, and 17 °C for white sea urchins, a species which should not be held at less than 8 °C.
Gradual acclimation to test temperature before gamete collection is advised, even if the gametes are to be collected on the day of, or the day after, the spawning adults are received in the laboratory. Water temperatures should be changed to the desired value at a rate of ≤5 °C per day. Certain situations (e.g., adults spawned for testing on the same day they arrive in the laboratory), however might require a daily shift of more than 5 °C. For rapid temperature adjustment of adults held ≤ 3 days prior to spawning, the procedure described in Section 2.3.1 for mixing culture water (or shipping water) and control/dilution water should be used.Footnote 10
2.3.6 Dissolved Oxygen
The dissolved oxygen (DO) content of the water within holding containers should be 80 to 100% of air saturation. If necessary to achieve that, mild aeration of the water should be carried out using filtered, oil-free compressed air. Such aeration through a commercial aquarium airstone also assists in mixing the water. Overly vigorous aeration should be avoided.
The pH of water used for holding adults should normally be in the range 8.0 ± 0.2, and must be within limits of 7.5 to 8.5.Footnote 11 The average pH of ocean waters is 8.1 (Thurman, 1975) and seawater has a strong buffering capacity. Coastal waters have a lower salinity than the open ocean, however, and some variation occurs from runoff of fresh water. Uncontaminated seawater is normally within the range of 7.5 to 8.5, whether it is brackish or full-strength, although the extremes of that range would be unusual. Existing methods for toxicity tests with echinoids do not give recommendations for the pH of water used to hold adults (Appendix D). Most laboratories surveyed (see Section 1.1) indicated that the pH range defined herein was not problematic for holding, acclimating, or spawning the echinoid species included in this biological test method. It has been reported, however, that the Pacific purple sea urchin has difficulty acclimating to waters with pH > 8.1 and that high mortality might be experienced if adults are to be held at a pH of >8.1 for longer than a few days (Carr, Nipper, and Biedenbach, pers. comm., 2008).
Adult echinoids that are spawned for testing within 3 days of arrival at the laboratory do not require feeding.
Sea urchins that are held in the laboratory for an extended period of time (i.e., > 3 days) should be fed with kelp or macroalga (Laminaria, Nereocystis, Macrocystis, Egregia, Hedophyllum) or, alternatively, with romaine lettuce. During a recent survey (see Section 1.1), Canadian and US laboratories provided feedback on their experience with the relevance of diet on adult survival, health and spawning success. Most laboratories feed urchins leafy greens (romaine lettuce, spinach, macroalgae), supplemented with carrots and/or algal pellets. One laboratory reported that the nutritional state of the adult sea urchins can affect the sensitivity of their gametes to contaminants and that romaine lettuce alone did not provide adequate nutrition. Researchers at that laboratory found that romaine lettuce supplemented with leaf spinach and carrots offered a better response (Carr, Nipper, and Biedenbach, pers. comm., 2008). Another response to the survey indicated that carrots were an essential component of food provided and enabled the long-term holding of adults with viable gametes (Agius, pers. comm., 2008). Food should be added frequently enough (weekly, daily) that it is always available to the urchins (i.e., ad libitum), and old or decomposing food should be removed. Restricted food supply would likely limit success in holding animals in good spawning condition (Bay and Greenstein, pers. comm., 2008). Sea urchins have been held in the laboratory for years using macroalgae. The brown alga Fucus has been recommended as food (EVS, 1989) and also recommended against use (Dinnel et al., 1987). The green sea urchin in Newfoundland eats Fucus and other brown alga such as Alaria esculenta as a major component of diet (Himmelman and Steele, 1971). The apparent feeding preference of the sea urchins being held should guide the investigator on use of Fucus and other potential food.
Sand dollars normally ingest particles selectively from the bottom and make use of the organic detritus available to them, including microalgae. For this reason, the natural and uncontaminated sediment used on the bottom of containers holding sand dollars should contain such detritus, and especially, settled plankton. Sand dollars have been said to require microalgae such as diatoms on the surfaces of sediment particles, and sufficient lighting can encourage growth of such algae on the sediment, increasing the success of long-term holding of the animals. Algae from a culture, or as an algal paste should be added to the sediment, if necessary (ASTM, 1990).
There are alternatives for feeding sand dollars which might sometimes be useful. Shredded eel grass (Zostera sp.) or even spinach could be added weekly, so that the animals can feed on the detritus (EVS, 1989). Flaked fish food may be used as a supplement (NCASI, 1991). However, any decomposing food in the tanks should be removed.
2.3.9 Cleaning the Holding Containers
Holding containers should be cleaned by scrubbing and rinsing before introducing a new batch of adults. Disinfectants may be used if it is desired to minimize the transmission of disease. Suitable disinfectants include those containing chlorinated or iodophore compounds or n-alkyl dimethyl benzyl ammonium chloride (e.g., CometTM, OvidineTM, ArgentyneTM, RoccalTM). Disinfectants are toxic to aquatic animals, and traces could carry over on the tanks and affect the echinoids. If disinfection is used, each container must be thoroughly rinsed with the water used for holding.
When holding adults, the containers should be kept reasonably clean. Old macroalga should be removed from urchin tanks, daily or as required. Periodic siphon-cleaning can be used in containers holding sea urchins, and also in sand dollar containers for removing light detritus, fecal pellets, or replacing the sediment. Shell fragments could be left in tanks with sea urchins, since healthy urchins commonly cover themselves with such fragments.
2.3.10 Disease and Mortality
Adult mortality should be low if organisms are acclimated properly to laboratory conditions. Occasionally, laboratories experience some adult mortality in the first week or two after the organisms arrive at the laboratory, or when adults are shipped with very ripe gonads and there is spontaneous spawning. Laboratories have also reported increased mortality in organisms collected and spawned late in their spawning season (i.e., September or later for the eccentric sand dollar and April or later for the Pacific purple sea urchin). Additionally, some species and/or batches demonstrate a high rate of mortality after spawning.
Adults should be inspected upon arrival at the laboratory and thereafter, daily, for signs of disease. Dead individuals should be removed immediately. In groups of animals which are held in the laboratory for an extended period of time (i.e., >3 days) before their gametes are collected for use in a test, mortality should not exceed 2% per day, averaged over the seven days preceding collection of gametes. The cumulative mortality over the same 7-day period must not exceed 20%. If a number of organisms from a given batch die after spawning is induced for testing purposes, those individuals may be excluded in the calculations of daily/weekly mortality. Adults spawned for use in a test may be separated from the remainder of the batch and may be excluded from mortality calculations, unless they are to be used for testing again.
For adults that are to be spawned for testing ≤3 days of arrival at the laboratory, the cumulative mortality data for the 7-day period prior to shipment should be obtained for the batch of organisms shipped from the supplier, and should not exceed 20%. No adults are to be used for same-day gamete collection (i.e., testing on the day that adults arrive at the laboratory), if their cumulative mortality rate exceeds 20% upon their receipt at the laboratory. This same criterion for mortality applies in instances where adults are held briefly (i.e., up to 3 days) before spawning (i.e., the cumulative mortality rate upon receipt and for the ≤3 days before spawning must be <20%).
For those groups of adults with a high mortality rate (i.e., exceeding any of the criteria described herein), surviving echinoids should be either discarded or held for an extended period until the mortality rate is acceptably low. Discard also, any moribund animals, sea urchins with significant loss of spines, and sand dollars with patches of fungus. Moribund sea urchins can usually be distinguished by lack of activity of the tube feet, inability to right themselves when turned over, and in particular by lack of adhesion to the substrate. Moribund sand dollars are usually distinguished by external appearance and activity. Such individuals often show patchy or overall pale colour as the epidermis degenerates, and do not rebury themselves. There is only weak activity of tube feet upon close inspection (magnifying glass or dissecting microscope), coupled with limpness of spines and pedicellaria (small pincer-bearing appendages among the tube feet). Dead sand dollars develop a coating of slime and often turn black.
Treatment of diseased adults with chemicals should not be attempted; it is strongly recommended that groups of animals showing a high incidence of disease be discarded.
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