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Biological test method for determining acute lethality of sediment to amphipods: chapter 4

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Section 4: Procedure for Testing Sediment

4.1 Sample Collection

Guidance on the collection of samples of marine or estuarine sediment for toxicity evaluations using marine or estuarine amphipods is given in Section 5.1 of Environment Canada (1992), and should be consulted beforehand. Environment Canada (1994) provides additional guidance on field sampling designs and appropriate techniques for sample collection; this guidance document should be referred to for further information.

Procedures and equipment used for sample collection (i.e., core, grab, dredge, or composite) will depend on the study objectives or regulatory requirements, and on the nature of the material being sampled. Samples of dredged material should be taken at all depths of interest. Samples of field-collected test or reference sediment, including those taken from or adjacent to ocean disposal sites, frequently represent the upper 2-cm depth. Sites for collecting samples of reference sediment should be sought where the geochemical properties of the sediment, including grain size characteristics, are similar to those at the site(s) where samples of test sediment are collected. Ideally, reference sediment should be collected from a site uninfluenced by the source(s) of contamination but within the general vicinity of the site(s) where samples of test sediment are taken. It is recommended that reference sediment from more than one site be collected to increase the likelihood of a good match with grain size and other physicochemical characteristics of the test sediments. Samples of control sediment are normally those taken at the site where test organisms are collected.

The number of stations to be sampled at a study site and the number of replicate samples per station will be specific to each study. This will involve, in most cases, a compromise between logistical and practical constraints (e.g., time and cost) and statistical considerations. Environment Canada (1994) should be consulted for guidance with respect to the sampling design, including the recommended minimum number of field replicates. Additional guidance on sampling is found in Environment Canada (1995) for disposal-at-sea applications. Applicants are encouraged to consult with their regional Environment Canada Ocean Disposal Office (see Appendices B and C for contact information), before sampling and testing.

Where practical and consistent with the study design and objectives, a minimum of five samples of sediment should be taken from each discrete sampling station and depth of interest. Where practical and appropriate (see Section 6), sample collection should also include ≥5 samples from each of one or more reference stations (i.e., sites where uncontaminated sediment, having physicochemical properties similar to that of the test sediments, can be found) within the vicinity. The objective of collecting replicate samples at each station is to allow for quantitative statistical comparisons within and among different stations (EC, 1994; 1998a). Each of these “true replicate” samples of sediment should be tested for its acute toxicity to amphipods, using a minimum of five test chambers per sample (i.e., laboratory replicates) (EC, 1992).

The collection of replicate samples at a given sampling station is often not necessary for certain dredging projects (EC, 1994). If the objective is to obtain a “cost-effective” assessment of sample toxicity within the project area, sampling as many stations as possible (subject to cost constraints) with a single sample from each station might be the best way to achieve this. In this instance, testing might be restricted to five laboratory replicates (i.e., 5 subsamples) per sample (and no replication of samples from each station), each of which is prepared in the laboratory (Section 4.3).

To sample sediment, a benthic grab (i.e., Smith-MacIntyre, Van Veen, PONAR) or core sampler should be used rather than a dredge, to minimize disruption of the sample. Care must be taken during sampling to minimize loss of fines. The same collection procedure should be used for all field sites sampled.

A per-sample volume of at least 5 to 7 L of whole sediment is frequently required (EC, 1994), although this will depend on the study objectives/design and on the nature of the physicochemical analyses to be performed. To obtain the required sample volume, it is frequently necessary to combine subsamples retrieved using the sampling device. Guidance provided in Environment Canada (1994) for compositing subsamples in the field should be followed.

4.2 Sample Labelling, Transport, and Storage

Instructions and guidance in Section 5.2 of Environment Canada (1992) pertaining to sample labelling, transport, and storage apply here, and should be reviewed and followed. Additional useful guidance in this respect is found in Environment Canada (1994) and USEPA (1994a).

Containers for transporting and storing samples must be new or thoroughly cleaned, and rinsed with clean water. Environment Canada (1994) should be consulted for guidance in selecting suitable containers. Each sample container should be filled completely, to exclude air. Immediately after filling, each sample container must be sealed and labelled or coded. Labelling and accompanying records made at this time must include at least a code which can be used to identify the sample or subsample. A cross-referenced record, which might or might not accompany the sample or subsample, must be made by the field personnel identifying the sample type (e.g., grab, core, composite), source, precise location (e.g., water body, latitude, longitude, depth), replicate number, and date of collection. This record should also include the name and signature of the sampler(s). Sediment sample collectors should also keep records describing:

Upon collection, warm (>7°C) samples should be cooled to between 1 and 7°C with regular ice or frozen gel packs, and kept cool (4 ± 3°C) in darkness throughout transport (EC, 1994). As necessary, gel packs, regular ice, or other means of refrigeration should be used to assure that sample temperatures range within 1 to 7°C during transit.

Upon arrival at the laboratory, the sample temperature and date of receipt must be recorded. Samples to be stored for future use must be held in airtight containers and in darkness at 4 ± 2°C (EC, 1992; 1994). Any air headspace in the storage container should be purged with nitrogen gas, before capping tightly (EC, 1994). Samples must not freeze or partially freeze during transport or storage, and must not be allowed to dry (EC, 1992; 1994). It is recommended that samples of sediment or similar particulate material be tested as soon as possible after collection. The sediment toxicity test should begin within two weeks of sampling, and preferably within one week; the test must start no later than six weeks after sample collection.

4.3 Sample Manipulation and Characterization

Samples of field-collected test sediment and reference sediment must not be wet-sieved. Large debris or large indigenous macro-organisms should be removed using forceps or a gloved hand. If a sample contains a large number of indigenous macro-organisms which cannot be removed using forceps or a gloved hand, the sample may be press-sieved (not washed) through one or more suitably sized (e.g., 1 or 2 mm) mesh stainless steel screens. Any pore water that has separated from the sample during shipment and storage must be mixed back into the sediment. To achieve a homogeneous sample, either mix it in its transfer/storage container, or transfer it to a clean mixing container. The sample should normally be stirred using a nontoxic device (e.g., stainless steel spoon or spatula), until its texture and colour are homogeneous (EC, 1992). Alternatively, a mechanical method (USEPA, 1994a; EC, 1994) may be used to homogenize the sample. For each sample included in a test, mixing conditions including duration and temperature must be as similar as possible. If there is concern about the effectiveness of sample mixing, subsamples of the sediment should be taken after mixing, and analyzed separately to determine homogeneity.

The portion of control sediment obtained from the amphipod collection site for use in the toxicity test, and for particle size and chemical analysis, must be previously wet-sieved through a 0.5-mm stainless steel screen to remove small amphipods and other organisms. Procedures described in Section 3.4 of Environment Canada (1992) should be followed. Sieved control sediment should be stored as described in the previous section (4.2) until used.

Immediately following sample mixing, subsamples of test material required for the toxicity test and for physicochemical analyses must be removed and placed in labelled test chambers, and in the labelled containers required for storage of samples for subsequent physicochemical analyses. Any remaining portions of the homogenized sample that might be required for additional toxicity tests using amphipods or other test organisms should also be transferred at this time to labelled containers. All subsamples to be stored should be held in sealed containers with no air space, and must be stored in darkness at 4 ± 2°C until used or analyzed. Just before it is analyzed or used in the toxicity test, each subsample must be thoroughly re-mixed to ensure that it is homogeneous.

Each sample (including all samples of control and reference sediment) must be characterized by analyzing subsamples for at least the following (EC, 1992; USEPA, 1994a): for whole sediment -- percent very coarse-grained sediment (i.e., particles >1.0 mm), percent sand (>0.063 to 2.0 mm), percent silt (>0.004 to 0.063 mm), percent clay (<0.004 mm), percent water content, and total organic carbon content; for pore water -- salinity, pH, and ammonia (total and un-ionized). Other analyses could include: total inorganic carbon, total volatile solids, biochemical oxygen demand, chemical oxygen demand, cation exchange capacity, acid volatile sulphides, metals, synthetic organic compounds, oil and grease, petroleum hydrocarbons, and porewater analyses for various physicochemical characteristics such as hydrogen sulphide. Recommended procedures for collecting pore water are described in Environment Canada (1994) and should be followed here. For disposal-at-sea applications, minimum information requirements are explained in Environment Canada (1995).

Analyses for particle size distribution and porewater salinity must be undertaken as soon as possible after sample collection, to confirm that the values for these characteristics are within the application limits for the intended species of test organism (see Section 2.6 as well as Appendices D for R. abronius, E for E. washingtonianus, F for E. estuarius, and G for A. virginiana). Analyses for porewater pH, salinity, and ammonia must be undertaken within 24 h of the start of the test and should be initiated at the beginning of the test, to determine the initial concentrations of total and un-ionized ammonia to which test organisms were exposed at the start of the test. Ammonia analyses must be conducted using a recognized and standardized procedure (for example, APHA et al., 1995; Standard Methods). Calculations of concentrations of un-ionized ammonia must be based on the test temperature and on the porewater pH and salinity of the sample (Trussell, 1972; Bower and Bidwell, 1978; USEPA, 1985).

4.4 Test Water

Test water must be the same as that used to acclimate the test organisms (see Section 2.4). This may be reconstituted seawater or an uncontaminated supply of natural seawater. Natural or reconstituted seawater may be adjusted to the required salinity (i.e., that to which the amphipods have been acclimated; see Section 2.4) by the addition of dry ocean salts or brine (if too brackish), or distilled water (if too saline). Guidance provided in Environment Canada (1992; Section 2.5.4) for preparing and storing test water should be followed.

Test water must be adjusted to the required test temperature (i.e., 15 ± 2°C for R. abronius, E. washingtonianus, or E. estuarius; 10 ± 2°C for A. virginiana) and salinity before use, and its dissolved oxygen concentration must be 90 to 100% of the air-saturation value for that temperature and salinity. As necessary, the required volume of water should be aerated vigorously (using oil-free compressed air passed through one or more air stones) immediately before use, and its dissolved oxygen content checked to confirm that 90 to 100% saturation has been achieved.

4.5 Test Conditions

4.6 Criteria for a Valid Test

4.7 Beginning the Test

Details for preparing for and starting the test are provided in Section 4.1 of Environment Canada (1992); instructions therein should be followed when undertaking this reference method.

Each test chamber placed within the test facility must be clearly coded or labelled to enable sample identification. The date and time when the test is started must be recorded, either directly on the labels or on separate data sheets specific to the test. The test chambers should be positioned for ease of observation and taking measurements. A minimum of five replicates per treatment, including at least five samples or subsamples of control sediment, should be included in each test (see Section 4.1). Each set of replicate treatments should be positioned randomly within the test facility.

On the day preceding the start of the test (i.e., Day -1) each sample of test sediment to be evaluated should be homogenized (Section 4.3). Thereafter, a 175-mL aliquot of each sample or subsample must be added to a separate test chamber. The aliquot should be smoothed to form a layer approximately 2-cm deep on the bottom of the test chamber, either by tapping the side of the test chamber against the side of the hand or by smoothing the sample with a clean plastic or stainless steel spatula. Highly contaminated sediment should be added to test chambers in a certified fume hood. Following sample addition, test water (see Section 4.4) should be added without disturbing the sample (see EC, 1992; Section 4.1), to a standard height on the test chamber. The (identical) volume of water added to each test chamber should approach the 950-mL mark (i.e., the combined volume of sediment and overlying water in the chamber at the start of test), but allow space for the transfer of test organisms (in a small volume of test water) the next day (i.e., Day 0). Each test chamber should then be covered, placed within the temperature-controlled test facility, and the overlying water aerated gently. Sediment in test chambers must not be stirred with the overlying water or otherwise disturbed, at any time before (i.e., Day -1) or during the test.

The overlying water in each test chamber must be aerated continuously once the water is added (i.e., Days -1 through Day 10); except perhaps during the brief period when test organisms are added, and when observations and measurements are made during the test. Compressed air, previously filtered and free of oil, should be bubbled through a glass or plastic pipette and attached plastic tubing (aquarium supply). The tip of the pipette should be suspended 2 to 4 cm above the surface of the sediment layer. Air flow to each test chamber must be gentle (e.g., 2 to 3 bubbles/s), and should not disturb the surface of the sediment. The rate of air flow should be adjusted as required to maintain a dissolved oxygen concentration in the overlying water of at least 90% saturation (EC, 1992; USEPA, 1994a).

Instructions in Section 4.1 of Environment Canada (1992) should be followed when adding amphipods to test chambers the next day (i.e., Day 0). Test organisms must be sieved from their acclimation chamber(s) (see Sections 2.4 and 2.5, herein) on that day, and 20 amphipods randomly added to each test chamber. Any animal that appears atypical or that is dropped or injured during the sieving and transfer process must not be used. Following the addition of test organisms, the water level in the test chamber must then be brought up to the 950-mL mark, after which the test chamber is covered and aeration of the overlying water is resumed after one hour at a gentle rate.

Within the first hour of the test, each test chamber must be examined to see if the amphipods have buried into the sediment. With the exception of A. virginiana (see EC, 1992; footnote 18), animals that do not bury within one hour must be replaced with those from the same sieved population, unless they are observed to repeatedly burrow into the sample and immediately emerge in an apparent avoidance response, or unless there is an obvious difference between the control and the test sediments. This would indicate a contaminant-related response, in which instance animals in any treatment would not be replaced. Amphipods displaying an avoidance behaviour during the initial hour of the test must not be replaced, i.e., they are to comprise the 20 test animals in the test chamber. Observations of apparent avoidance responses must be recorded.

4.8 Test Measurements and Observations

Test measurements must be made in at least one test chamber representing each treatment. The temperature of the overlying water must be measured at the beginning of the test and thereafter at least three times per week (e.g., Mondays, Wednesdays, Fridays) on non-consecutive days until test completion. More frequent (i.e., daily) measurements of temperature are recommended. Additionally, it is recommended that the temperature of any water bath used, and/or of the air in a temperature-controlled room or chamber used for the test, be recorded continuously.

For at least one test chamber representing each treatment, the concentration of dissolved oxygen in the overlying water must be measured at the beginning of the test, and thereafter at least three times/week (e.g., Mondays, Wednesdays, Fridays) on non-consecutive days until test completion. More frequent (e.g., daily; USEPA, 1994a) measurements are recommended and should be performed for sediments having a high oxygen demand that depresses the dissolved oxygen of the overlying water below 90% saturation. A probe and calibrated dissolved oxygen (DO) meter is recommended for these measurements. The probe must be inspected carefully after each reading to ensure that organisms are not adhered to it, and must be rinsed in deionized or distilled water between samples to minimize cross-contamination. The position of the tip of the pipette in each test chamber and the rate of aeration should be checked frequently and routinely (e.g., daily) throughout the test, and adjustments made if necessary to maintain a gentle (e.g., 2 to 3 bubbles/s) rate of aeration.

If at any time during the test the air flow to one or more test chambers is observed to have stopped, the dissolved oxygen concentration in the overlying water must be measured and then the air flow re-established at a gentle rate. Any DO readings that have fallen below 60% saturation (USEPA, 1994a) must be included in the test-specific report (Section 7.1.6), and must be considered when interpreting the test results (Section 6.2).

The salinity and pH of the overlying water must be measured at the beginning and end of the test in at least one test chamber representing each treatment. Additionally, ammonia concentrations in the overlying water must be measured (total ammonia; see for example APHA et al., 1995) and calculated (un-ionized ammonia; Trussell, 1972; Bower and Bidwell, 1978; USEPA, 1985) at the beginning and end of the test in at least one test chamber representing each treatment. Salinity and pH may be measured using probes and calibrated meters. Ammonia may be measured using an ion-specific electrode or by extracting an aliquot of the overlying water for this analysis. As with DO measurements, any probe inserted in a test chamber must be inspected carefully immediately after each reading, and rinsed in deionized or distilled water between samples. For measurements of ammonia requiring sample aliquots, samples of overlying water must be taken just before the addition of test organisms and upon completion of the test. On each occasion, no more than 10% of the volume of the overlying water in a test chamber should be removed for this purpose. A pipette should be used carefully to remove water from a depth of about 1 to 2 cm above the sediment surface. The pipette should be checked to ensure that no amphipods are removed during water sample collection.

Each test chamber must be examined frequently and routinely during the test (i.e., at least three times per week on non-consecutive days, and preferably daily) to note if amphipods have emerged from the sediment, or if they are swimming in the overlying water or floating on its surface. The number of animals seen swimming in the water, floating on its surface, moving on the surface of the sediment, or emerged from the sediment but apparently dead, should be noted and recorded during each observation period. Amphipods caught in the surface film should be gently pushed down into the water using a glass rod or pipette. Animals that appear to be dead should not be removed.

4.9 Ending the Test

The test is terminated after 10 days of exposure. At that time, the final set of observations of numbers of amphipods seen floating on the surface of the overlying water, swimming in it, moving on the surface of the sediment, or emerged from the sediment but apparently dead, must be made and recorded. Just before sieving the contents of a test chamber, all live and apparently dead amphipods in the water column or on the surface of the sediment should be pipetted from the test chamber.

Individuals that are completely inactive but not obviously dead (e.g., not decomposing) should be held in test water within a petri dish or other suitable container, and examined closely at this time using a low-power microscope or handheld magnifying glass. These individuals should be prodded gently with a sharp point to confirm that they show no sign of life (such as a pleopod twitch). Any animals that fail to show signs of life before and after prodding must be counted as dead.

A consistent amount of time should be taken to sieve the contents of each test chamber for recovery of live or dead organisms. To ensure that the procedure used to recover amphipods is adequate, it is recommended that the laboratory personnel responsible for sieving the contents of test chambers demonstrate that they are able to retrieve an average of at least 90% of the organisms from control sediment. For example, test organisms could be added to control sediment and recovery could be determined after one hour (USEPA, 1994a; Tomasovic et al., 1995).

The contents of each test chamber must be sieved through a 1.0-mm (or smaller) mesh screen to remove all remaining test organisms, and to determine if they are dead or alive. Test water adjusted to the salinity and temperature of that in the test chambers should be used for sieving. Material retained on the screen should be washed into a sorting tray using clean test water. A small portion of the material should be sorted through at a time, removing amphipods as they are found (USEPA, 1994a). Amphipods that are inactive but are not obviously dead should be examined closely as described previously, and counted as dead if they fail to show signs of life. Animals that are missing are presumed to have died and are counted as dead organisms in the calculations (Section 4.10).

4.10 Test Endpoints and Calculations

The biological endpoint for this 10-day solid-phase sediment toxicity test is percent survival. The mean (± SD) percentage of amphipods that survived the 10-day exposure is calculated, for each treatment (i.e., each set of replicates representing a test sediment). This calculation is typically based on 100 organisms per treatment (i.e., 20 amphipods exposed to each of five replicate samples or subsamples; see Sections 4.1 and 4.7).

To enable this calculation, numbers of amphipods found to be dead, missing, or alive in each test chamber at Day 10 are determined and recorded (Section 4.9). Missing individuals are assumed to have died and disintegrated during the test, and must be included in the count of number dead per chamber. The mean (± SD) percent survival for the replicate groups within a given treatment is then calculated, for each treatment. Thereafter, the mean (± SD) value for percent survival determined for each treatment is compared against that for the reference sediment or, as necessary, against the mean percent survival for the control sediment (see Section 6 for guidance).

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