Biological test method for toxicity tests using early life stages of rainbow trout: chapter 3

3.1 Facilities and Materials
3.2 Lighting
3.3 Test Apparatus
3.4 Control/Dilution Water

3.1 Facilities and Materials

The test should be conducted in a facility isolated from general laboratory disturbances. If a separate room is unavailable, the test area should be surrounded with an opaque curtain (e.g., black plastic) to minimize stress to embryos, alevins, or swim-up fry during testing. Dust and fumes should be minimized within the facilities.

The test facility must be able to maintain the daily mean temperature of all test solutions at 14 ± 1.0°C (see Section 4.3.3). This might require in-line heating and/or cooling of the control/dilution water, a temperature-and photoperiod-controlled wet laboratory, or various types of equipment such as portable water-cooling and/or heating units.

The laboratory must have instruments for measuring the basic water quality variables (temperature, conductivity, dissolved oxygen, pH), and should be prepared to undertake prompt and accurate analysis of other variables such as hardness, alkalinity, ammonia, and residual chlorine.

Any construction materials or equipment that might contact the test substance, test solutions, or control/dilution water, must not contain any substances that can be leached into the sample, solutions, or water at concentrations that could cause toxic effects. Materials such as borosilicate glass (e.g., Pyrex^TM), stainless steel, porcelain, nylon, high-density polystyrene, or perfluorocarbon plastics (Teflon^TM), should be used. Other nontoxic plastics, such as polypropylene or polyethylene, may be used but their re-use should be only after careful and thorough cleaning (e.g., using a phosphate-free detergent wash followed by an acid soak and several rinses with deionized water) to minimize the possible release of sorbed toxicants during a subsequent test. For tests with chemicals or chemical products (see Section 5), glass is the recommended material for containers and apparatus which contact the test solutions before or during the test.

3.2 Lighting

The test should be conducted in the dark until one week after the embryos have hatched.^Footnote 3 For the remainder of the test, subdued lighting should be used. Light intensity at the water surface should be within the range of 100 to 500 lux. Depending upon test requirements and intent, lighting might be provided by overhead full-spectrum fluorescent fixtures.^Footnote 4 The photoperiod should normally be a constant sequence of 16 ± 1 h of light and 8 ± 1 h of darkness. A 15- to 30-minute transition period between light and dark is recommended.^Footnote 5

3.3 Test Apparatus

In 1996, the test apparatus illustrated in Environment Canada (1992a), which had been used in a number of early life-stage toxicity tests with salmonid embryos or alevins (McLeay and Gordon, 1980; Martens et al., 1980; Hodson et al., 1991), was re-designed. The modified design was tested and proved effective (Yee et al., 1996). The incubation unit and associated apparatus now recommended (Figure 3) is easily and inexpensively constructed, and enables gentle aeration and/or continuous circulation of test solutions past incubating embryos or alevins and easy renewal of solutions with minimal disturbance. In addition, the unit allows the embryos or alevins to remain bathed in the test solution during the solution renewal process. This incubation unit is made from an 800-mL (or larger) Tri-Pour^TM plastic beaker having slightly tapered sides. Unlike the earlier design which required replacing the bottom of the beaker with a plastic mesh floor (EC, 1992a), the bottom of the beaker is left intact, and a series of horizontal slits are cut in the sides, near the bottom, to allow the circulation of test solutions within the beaker (Yee et al., 1996; Figure 3A). A circular hole is drilled in the centre of the bottom of the beaker, and a removable "pressure-fit" 5-cm long standpipe, cut from a standard-supply 10-mL disposable polystyrene volumetric pipette, is inserted through the hole.

The incubation unit can be easily suspended in a test chamber by inserting it through a 10-cm diameter hole cut in the cover of the chamber (e.g., the white plastic lid of a 4-L white plastic pail). A second hole of smaller diameter (~6 cm; Yee et al., 1996) can be cut in the lid to enable monitoring of water quality during the test (Section 4.3). The volume of solution held in each test chamber should not be chosen arbitrarily but should be considered in light of the requirements for amounts of new test solution to be supplied daily (Section 4.3.2), throughout the exposure period involving embryos or alevins. The daily (or more frequent) renewal of each test solution is achieved either by siphoning out ~80% of the old solution and replacing it immediately with a fresh (new) solution prepared to the same strength (i.e., static-renewal test), or by the continuous addition of fresh solution to the the test chamber (i.e., a flow-through test, see Section 4.3.2).

The test chamber should be adapted to accommodate either static-renewal or flow-through conditions, depending on the requirements and objectives of the test.^Footnote 6 For static-renewal tests (Section 4.3.2), the experimental design shown in Figure 3B is recommended. Using this setup, filtered, oil-free air is bubbled through a suitable length (e.g., 60 cm) of 0.5-mm ID polyethylene capillary tubing, which is inserted in the centre standpipe protruding through the bottom of the incubation unit (Figure 3B).^Footnote 7 This aeration system, which can also be used if necessary for flow-through tests, provides a continuous current of aerated water past the embryos or alevins.

For flow-through tests (see Section 4.3.2), the experimental design shown in Figure 3C is recommended. This setup may be used with or without aeration.^Footnote 6^,^Footnote 8 A flow of fresh solution passes continuously over the embryos or alevins as it drains from the test chamber (Figure 3C). Using the flow-through design shown in Figure 3C, the incubation unit is suspended around a suitable length of standpipe drain that is secured through both the centre of the incubation unit and the bottom of the test chamber. A plastic mesh insert is placed in the top of the standpipe to prevent loss of embryos or alevins.

Each test chamber, whether it contains one or more incubation units suspended in it, is one replicate. For each test concentration or control, there must be at least three separate test chambers providing true replication, adequate for calculation of experimental error (see Sections 4.1 and 4.5). If hypotheses testing is to be applied, such as estimating NOEC/LOEC, four separate test chambers should be used; that is the minimum number required if it were necessary to use nonparametric statistical tests.

Other apparatus (e.g., Canaria et al., 1996) may also be used as test chambers, provided that the objectives of the test and criteria for validity are achieved (Section 4.6). However, use of apparatus illustrated in Figure 3 is recommended for incubating embryos and alevins, and if used will provide a greater degree of standardization of conditions during incubation.

Figure 3 Recommended Design for Static-renewal or Flow-through Setups for Incubating Embryos or Alevins in Test Solutions

Recommended design for static-renewal of flow-through setups for incubating embryos or alevins in test solutions.

Exploded view of modified incubation unit and associated apparatus, showing horizontal side slits.
Incubation unit suspended in a test chamber, showing aeration and upwelling current.
Incubation unit suspended in a test chamber with no aeration, showing standpipe drain and current created by solution flow.

Long description for Figure 3

This figure is composed of three parts. Part A is an exploded view of the modified incubation unit and the associated apparatus. There are three primary components in this apparatus. One component is an 800 mL beaker to serve as the incubation unit. This beaker has six horizontal slits in its side and a hole cut in the bottom to accommodate a 5 cm section of pipette. The final component pictured in part A is for cover of a test chamber (e.g. a 4 L plastic pail) with two holes cut into it. The holes are 10 cm and 6 cm in diameter to accommodate the incubation unit and solution exchange respectively.

Part B is an illustration of a static-renewal setup. The pipette section has been inserted into the bottom of the incubation unit and the unit has been lowered into the test chamber. The chamber is filled with test solution until the solution reaches approximately the mid-point of the incubation unit. Flow of the test solution through the incubation unit is generated by suspending a 0.5 mm airline through the top of the incubation unit such that it terminates in the middle of the pipette (roughly in line with the bottom of the incubation unit). Air is then passed through this narrow tube and the rising bubbles pull test solution up through the bottom of the incubation unit and out the horizontal side slits.

Part C illustrates a flow-through setup. This setup is similar to that described in part B but with the addition of a mesh insert to the top of the pipette and a waste tray under the test chamber. The pipette now extends down through the bottom of the test chamber and the mesh insert is added to the top of the pipette to prevent the loss of alevins or embryos. In this setup, test solution is continually added through the aforementioned hole cut into the lid of the test chamber. This constant influx of solution flows in through the horizontal slits in the incubation unit and drains through the pipette section. Test solution flowing out through the pipette section is collected by the waste tray.

3.4 Control/Dilution Water

Depending on the test substance and intent (Sections 5 to 7), the control/dilution water may be: "uncontaminated" ground or surface water from a river or lake; reconstituted water of a desired pH and hardness (e.g., simulating that of the receiving water); a sample of receiving water collected upstream of the source of contamination, or adjacent to the source but removed from it; or dechlorinated municipal water.^Footnote 9

The water supply should previously have been demonstrated to consistently and reliably support good survival, health, and growth of the test species. Monitoring and assessment of variables such as residual chlorine (if municipal water is used), pH, hardness, alkalinity, total organic carbon, conductivity, suspended solids, dissolved oxygen, total dissolved gases, chemical oxygen demand, temperature, ammonia nitrogen, nitrite, metals, and pesticides, should be performed as frequently as necessary to document water quality (e.g., monthly or more frequently if deleterious changes in water quality are suspected or found) (see Table 1). Conditions for the collection, transport, and storage of samples of receiving water, if used as control/dilution water, should be as described in Section 6.2.

If surface water is used as the control/dilution water, it should be filtered and/or sterilized. A conventional sand filter or commercial in-line filter would be suitable. Small quantities might be filtered through a fine-mesh net (≤60 µm). Ultraviolet sterilization is recommended to reduce the possibility of introducing pathogens into the laboratory and fish-holding system. The control/dilution water must be adjusted to the required test temperature before use (see Section 4.3.3). The total gas pressure (TGP) of this water should not exceed 100%.^Footnote 10

Additionally, its dissolved oxygen concentration (DO) should be 90 to 100% of the air-saturation value, before its use. If necessary, the control/dilution water should be aerated vigorously (oil-free compressed air passed through air stones) immediately before use, and a check made to confirm that a dissolved oxygen concentration of 90 to 100% saturation has been achieved.

Table 1 Recommended Quality for Control/Dilution Water ^Footnote a
Variable	Recommended Limits for Exposure
pH	6.5 to 8.5 (7.5 to 8.0 is desirable)
Hardness	15 to 150 mg CaCO₃/L
Alkalinity	20 to 200 mg CaCO₃/L
Aluminum	<5 µg/L (pH ≤6.5) <0.1 mg/L (pH >6.5)
Ammonia (un-ionized)	<5 µg/L (preferably not detectable)
Cadmium	<0.3 µg/L (in soft water) <0.5 to 0.75 µg/L (in hard water)
Chlorine	<2 µg/L
Copper	<6 µg/L (in soft water) <30 µg/L (in hard water)
Dissolved carbon dioxide	0.03 to 15 mg/L
Dissolved oxygen	90 to 100% of saturation
Hydrogen cyanide	<10 µg/L
Hydrogen sulphide	<2 µg/L (preferably not detectable)
Iron	<0.3 mg/L
Lead	<1 µg/L (in soft water) <2 µg/L (in hard water)
Mercury	<0.05 µg/L
Nitrite	<60 µg/L (preferably not detectable)
Nitrogen (dissolved gas)	<100 to 103% (max. partial pressure) <103 % (total gas pressure)
Selenium	<10 µg/L
Total suspended solids	<3 mg/L during incubation <2 5 m g/L during larval and fry stages
Zinc	< 0.03 mg/L (in soft water) < 0.3 mg/L (in hard water)

Footnotes

Footnote a

For salmonid species (from Klontz et al., 1979; CCREM, 1987; and Gordon et al., 1987). Soft water is defined here as 60 mg/L total hardness as calcium carbonate (CaCO₃). This table is intended as a general guide for water quality. Local water conditions, particularly variations of hardness, alkalinity, and dissolved organic matter, can reduce or increase the threshold for metal toxicity. Available studies of metal toxicity in local water should be consulted.

Other important variables, such as total organic carbon, chemical oxygen demand, and pesticide residues in the control/dilution water, should be monitored and their potential effects on toxicity should be evaluated.

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Footnotes

Footnote 3

Dim incandescent lighting (i.e., <200 lux) may be used for short periods during observations and maintenance (R. Watts, personal communication, Pacific Environmental Science Centre, North Vancouver, BC). Alternatively, the use of red ("dark room") incandescent lighting is recommended (G. van Aggelen, personal communication, Pacific Environmental Science Centre, North Vancouver, BC).

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Footnote 4

Full-spectrum fluorescent or equivalent lamps, supplemented with natural outdoor illumination if desired, are suitable for simulating the visible range of natural light. However, full-spectrum lights do not emit ultraviolet (UV-B) radiation at intensities approaching that of natural illumination, and the toxicity of certain effluents and chemicals can be altered markedly by photolysis reactions caused by UV-B radiation. For certain tests (e.g., those concerned with photoactivation or photodegradation of toxic substances due to ultraviolet radiation), lights with particular spectral qualities may be selected (e.g., high-pressure mercury arc lamps). ASTM (1996) provides useful guidance in this regard.

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Footnote 5

A "dawn/dusk" transition period is recommended since abrupt changes in intensity startle and stress fish. Automated dimmer control systems are available for dimming and brightening the intensity of fluorescent lights, although they are costly. Alternatively, a secondary incandescent light source, regulated by time clock and automated rheostat, may be used to provide the transition period.

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Footnote 6

With many types of test substances, static tests with 12- or 24-h renewal of test solutions, when done properly, can be as sensitive and accurate as flow-through tests (Sprague, 1973). Static-renewal tests might also be desirable or necessary when the degradation products of the test substance are of concern. High chemical or biochemical oxygen demand, volatility, or instability of certain substances might necessitate the use of a flow-through test. A flow-through test is also necessary if embryos or alevins are to be incubated in test solutions without aeration, in order to provide a continuous exchange of solution within the incubation unit.

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Footnote 7

A simple and practical design for aerating several incubation units simultaneously can be quickly assembled by inserting three or four lengths of 0.5 mm ID capillary tubing into one length (e.g., 40 cm) of standard aquarium airline tubing. Clear silicone sealant can be used to fuse the juncture and prevent air leaks (Yee et al., 1996). Other apparatus, such as tubing with Eppendorf tips attached, is also suitable for generating small bubbles and thus for providing gentle aeration through a narrow-bore aperture. Disposable Pasteur pipettes or other air-delivery apparatus used routinely for acute lethality tests with fish (e.g., EC, 1990b) may be used to aerate test solutions with volumes 6 L.

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Footnote 8

Aeration might strip volatile chemicals from solution or increase their rates of oxidation and degradation to other substances. Gentle aeration (Section 4.3.4) might be desirable or even necessary during a flow-through test, to maintain adequate levels of dissolved oxygen when the chemical or biochemical oxygen demand of the test substance is particularly high. If aeration is to be provided using a flow-through setup (e.g., Figure 3C), the end of the airline tubing should be placed in the test solution outside the standpipe drain.

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Footnote 9

If municipal drinking water is to be used for culturing fish and as control/dilution water, effective dechlorination must rid the water of any harmful concentration of chlorine. It is difficult to remove the last traces of residual chlorine and chlorinated organic substances, which might be toxic to developing fish. Vigorous aeration of the water followed by the use of activated carbon (bone charcoal) filters can be applied to strip out part of the volatile chlorine gas, and subsequent ultraviolet radiation (Armstrong and Scott, 1974) can be used for removing most of the residual chloramine and other chlorinated organic compounds. Aging the water in aerated holding tanks might also help. The addition of thiosulphate or other chemicals to dilution water to remove residual chlorine is not recommended, as such chemical(s) could alter sample toxicity. The target value for total residual chlorine, recommended for the protection of freshwater aquatic life, is 0.002 mg/L (CCREM, 1987). Anything greater than 0.002 mg/L might risk interaction of chlorine toxicity with whatever was being tested (Brungs, 1973; NAS/NAE, 1974). In addition to measurements of chlorine, monitoring of egg production and fish survival can provide evidence of satisfactory water.

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Footnote 10

Water entering the test chamber should not be supersaturated with gases (i.e., TGP 100%). In situations where gas supersaturation within the water supply is a valid concern (e.g., groundwater source, without subsequent gas stripping), or the control/dilution water is either actively or passively heated to accommodate a specified test temperature, total gas pressure within water supplies should be checked frequently (Bouck, 1982). Remedial measures should be taken (e.g., use of aeration columns or vigorous aeration in an open reservoir) if TGP exceeds 100% saturation. It is not a simple matter to completely remove supersaturation, and frequent checking should be done if the problem is known or suspected to exist.

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Date modified:: 2014-08-26

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