toxicity and bioaccumulation testingrpitt.eng.ua.edu/publications/booksandreports/stormwater...

APPENDIX D

Toxicity and Bioaccumulation Testing

CONTENTS

General Toxicity Testing Methods.................................................................................................710 Methods for Conducting Long-Term Sediment Toxicity Tests with Hyalella azteca..................710

Placement of Sediment into Test Chambers ........................................................................710 Acclimation...........................................................................................................................713 Placing Organisms in Test Chambers ..................................................................................713 Feeding .................................................................................................................................713 Monitoring a Test .................................................................................................................713 Ending a Test ........................................................................................................................714 Interpretation of Results .......................................................................................................716

Methods for Conducting Long-Term Sediment Toxicity Tests with Chironomus tentans...........718 Collection of Egg Cases.......................................................................................................719 Hatching of Eggs..................................................................................................................719 Placing Organisms in Test Chambers ..................................................................................720 Feeding .................................................................................................................................720 Dissolved Oxygen ................................................................................................................720 Monitoring Survival and Growth .........................................................................................721 Monitoring Emergence.........................................................................................................722 Ending a Test ........................................................................................................................722 Interpretation of Results .......................................................................................................723

In Situ Testing Using Confined Organisms ...................................................................................724 Toxicity Identification Evaluations ................................................................................................729 Toxicity — Microtox Screening Test ............................................................................................730

Scope and Application..........................................................................................................730 Summary of Method ............................................................................................................730 Sample Handling and Preservation ......................................................................................730 Interferences .........................................................................................................................730 Apparatus..............................................................................................................................731 Reagents................................................................................................................................731 Procedure ..............................................................................................................................731 Calculations ..........................................................................................................................732 Precision and Accuracy ........................................................................................................733 Health and Safety Information.............................................................................................733

References ......................................................................................................................................733

709

710 STORMWATER EFFECTS HANDBOOK

GENERAL TOXICITY TESTING METHODS

There are a large number of toxicity and bioaccumulation test methods that can be used in laboratory or field (in situ) settings. The strengths and weaknesses of the two settings were discussed in Chapter 6. The toxicity test methods most commonly used in North America are those required by the EPA and state environmental protection agencies, such as Pimephales promelas and Ceriodaphnia dubia for wastewater effluent testing. While these tests have been used successfully to evaluate stormwaters, there are also other options that may be acceptable to the regulatory authorities, since they have been found useful in the scientific peer-reviewed literature. In addition, there are many standardized test methods approved by Environment Canada (Table D.1) and ASTM (Table D.2) that are often quite similar to U.S. EPA procedures. Only a few examples are listed below to help familiarize the user with the procedures and associated quality assurance issues. The project manager should verify that the appropriate test methods are being used to meet any regulatory requirements. These tests should only be conducted by laboratories with documented experience in aquatic toxicology. Given the potential for sampling and method-related artifacts (Chapters 5 and 6), it is important that the project manager ensure that proper study design, sample collection, and testing protocols are adhered to. The categories of assessment tools that are useful in receiving water assessments are shown in Table D.3. The methods recommended for screening are listed on Tables D.4 through D.14.

METHODS FOR CONDUCTING LONG-TERM SEDIMENT TOXICITY TESTS WITH HYALELLA AZTECA

The EPA recently finalized methods for long-term chronic toxicity testing of sediments (EPA 2000). These methods have not been widely used but have been found to be more sensitive to sediment contaminants than the 10-day assays. In addition, they were found to have acceptable levels of variability based on interlaboratory variance studies. Since these assays require 42 days and longer to run, they are somewhat costly to perform. Conditions for evaluating sublethal endpoints in a sediment toxicity test with H. azteca are summarized in Table D.15. A general activity schedule is outlined in Table D.16.

The 42-day sediment toxicity test with H. azteca is conducted at 23°C with a 16L:8D photoperiod at an illuminance of about 500 to 1000 lux. Test chambers are 300-mL high-form lipless beakers containing 100 mL of sediment and 175 mL of overlying water. Amphipods in each test chamber are fed 1.0 mL of YCT daily. Each test chamber receives two volume additions/day of overlying water.

A total of 12 replicates, each containing ten 7- to 8-d-old amphipods are tested for each sample. For the total of 12 replicates, the assignment of beakers is as follows: 12 replicates are set up on Day –1, of which 4 replicates are used for 28-day growth and survival endpoints and 8 replicates for measurement of survival and reproduction on Day 35, and survival, reproduction, or growth on Day 42.

Placement of Sediment into Test Chambers

The day before the sediment test is started (Day –1), each sediment is thoroughly homogenized and added to the test chambers. Sediment is visually inspected to judge the degree of homogeneity.

Each test chamber will contain the same amount of sediment, determined by volume. Overlying water is added to the chambers on Day –1 in a manner that minimizes suspension of sediment. Renewal of overlying water is started on Day –1. A test begins when the organisms are added to the test chambers (Day 0). Hardness, alkalinity, and ammonia concentrations in the water above the sediment within a treatment should not vary by more than 50% during the test.

TOXICITY AND BIOACCUMULATION TESTING 711

Table D.1 Status Report — Environment Canada Biological Test Method Development Programa (Revised December 1999)

Test Method / Supporting Guidance Documents Status Publication Date Report Number

Universal Test Methods

1. Acute Lethality Test Using Rainbow Trout 2. Acute Lethality Test Using Threespine

Stickleback 3. Acute Lethality Test Using Daphnia spp. 4. Test of Reproduction and Survival Using the

Cladoceran Ceriodaphnia dubia 5. Test of Larval Growth and Survival Using

Fathead Minnows 6. Toxicity Test Using Luminescent Bacteria

(Photobacterium phosphoreum) 7. Growth Inhibition Test Using the Freshwater

Alga (Selenastrum capricornutum) 8. Acute Test for Sediment Toxicity Using

Marine or Estuarine Amphipods 9. Fertilization Assay with Echinoids (Sea

Urchins and Sand Dollars) 10. Early Life-Stage Toxicity Tests Using

Salmonid Fish (Rainbow Trout) – Second Edition

11. Survival and Growth in Sediment Using Freshwater Midge Larvae Chironomus tentans or riparius

12. Survival and Growth in Sediment Using the Freshwater Amphipod Hyalella azteca

13. Test for Measuring the Inhibition of Growth Using the Freshwater Macrophyte Lemna minor

14. Survival and Growth in Sediment Using Estuarine or Marine Polychaete Worms

Published July 1990 EPS 1/RM/9 Published July 1990 EPS 1/RM/10

Published July 1990 EPS 1/RM/11 Published February 1992 EPS 1/RM/21

Published February 1992 EPS 1/RM/22

Published October 1992 EPS 1/RM/24

Published November 1992 EPS 1/RM/25

Published December 1992 EPS 1/RM/26


Published July 1998 EPS 1/RM/28



Published March 1999 EPS 1/RM/37

Final draft Early 2001 — in preparation

Reference Methods

1. Reference Method for Determining Acute Published July 1990 EPS 1/RM/13 Lethality of Effluents to Rainbow Trout

2. Reference Method for Determining Acute Published July 1990 EPS 1/RM/14 Lethality of Effluents to Daphnia magna

3. Reference Method for Determining Acute Published 1999 EPS 1/RM/35 Lethality of Sediment to Estuarine or Marine Amphipods

Supporting Guidance Documents

1. Control of Toxicity Test Precision Using Reference Toxicants

2. Collection and Preparation of Sediment for Physicochemical Characterization and Biological Testing

3. Measurement of Toxicity Test Precision Using Control Sediments Spiked with a Reference Toxicant

4. Application and Interpretation of Single-Species Test Data in Environmental Toxicology

5. Statistics for the Determination of Toxicity Test Endpoints

Published August 1990 EPS 1/RM/12


Published September 1995 EPS 1/RM/30

Final version Spring 2000 EPS 1/RM/34 in preparation

Second draft Early 2001 — in preparation

a Documents available in French and English, copies of published documents can be obtained from EPS Publication Section, ETAD, Environment Canada, Fax: (819)953-7253) Tel: (819)953-5921.


Table D.2 ASTM Standards on Toxicity Testing

Std. No.

Aquatic Toxicity Testing — Water

General E 1850-97 Guide for Selection of Resident Species as Test Organisms for Aquatic and Sediment Toxicity Tests E 1203-98 Practice for Using Brine Shrimp Nauplii as Food for Test Animals in Aquatic Toxicity E 1733-95 Guide for Use of Light in Laboratory Testing Phytoplankton D 3978-80 Practice for Algal Growth Potential Testing with Selenastrum capricornutum E1218-97a Guide for Conducting Static 96-hour Toxicity Testing with Microalgae E1913-97 Guide for Conducting Toxicity Tests with Bioluminescent Dinoflagellates Plant E 1841-96 Guide for Conducting Renewal Phytotoxicity Tests with Freshwater Emergent Macrophytes E 1498-92 (1998) Guide for Conducting Sexual Reproduction Tests with Seaweeds E 1415-91 (1998) Guide for Conducting Static Toxicity Tests with Lemna gibba G3 E 1913-97 Guide for Conducting Static, Axenic, 14-Day Phytotoxicity Tests in Test Tubes with the Submerged

Aquatic Macrophyte, Myriophyllum sibiricum Komarov Invertebrates E 1440-91 (1998) Guide for Acute Toxicity Test with the Rotifer Brachionus E 1562-94 Guide for Conducting Acute, Chronic, and Life Cycle Aquatic Toxicity Tests with Polychaetous

Annelids E 724-98 Guide for Conducting Static Acute Toxicity Tests Starting with Embryos of Four Species of

Saltwater Bivalve Molluscs E 1193-97 Guide for Conducting Daphnia magna Life Cycle Toxicity Tests E 1191-97 Guide for Conducting Life-Cycle Toxicity Tests with Saltwater Mysids E 1463-92 (1998) Guide for Conducting Static and Flow-Through Acute Toxicity Tests with Mysids from the West

Coast of the United States E 1295-89 (1995) Guide for Conducting Three-Brood, Renewal Toxicity Tests with Ceriodaphnia dubia E 1563-98 Guide for Conducting Static Acute Toxicity Tests with Echinoid Embryos Vertebrate E 729-96 Guide for Conducting Acute Toxicity Tests on Test Materials with Fishes, Macroinvertebrates, and

Amphibians E 1192-97 Guide for Conducting Acute Toxicity Tests on Aqueous Ambient Samples and Effluents with Fishes,

Macroinvertebrates, and Amphibians E 1241-98 Guide for Conducting Early Life-Stage Toxicity Tests with Fishes E 1439-98 Guide for Conducting the Frog Embryo Teratogenesis Assay-Xenopus (Fetax) General E 1022-94 Practice for Conducting Bioconcentration Tests with Fishes and Saltwater Bivalve Molluscs E 1242-97 Practice for Using Octanol-Water Partition Coefficient to Estimate Median Lethal Concentrations

for Fish Due to Narcosis Microcosm E 1366-96 Practice for Standardized Aquatic Microcosm; Freshwater Behavior E 1604-94 Guide for Behavioral Testing in Aquatic Toxicology E 1711-95 Guide for Measurement of Behavior during Fish Toxicity Tests E 1768-95 Guide for Ventilatory Behavioral Toxicology Testing of Freshwater Fishes

Aquatic Toxicity Testing — Sediment

General E 1391-94 Guide for Collection, Storage, Characterization, and Manipulation of Sediments for Toxicological

Testing E 1525-94a Guide for Designing Biological Tests with Sediments Marine Sediment Toxicity Tests E 1611-94 Guide for Conducting Sediment Toxicity Tests with Marine and Estuarine Polychaetous Annelids E 1367-99 Guide for Conducting 10-day Static Sediment Toxicity Tests with Marine and Setuarine Amphipods E 1688-97a Guide for Determination of the Bioaccumulation of Sediment-Associated Contaminates by Benthic

Invertebrates Freshwater Sediment Toxicity Tests E 1706-95b Test Methods for Measuring the Toxicity of Sediment-Associated Contaminants with Fresh Water

Invertebrates


Table D.3 Toxicity and Bioaccumulation Testing Categories

Site Type Assay Media Organisms (Examples)

Laboratory Acute/Screening Toxicity Low Flow P. promelas, C. dubia, Daphnia magna Short-term Chronic High Flow

Outfalls Acute or Chronic Sediments Hyalella azteca, Chironomus tentans,

Chironomus riparius Bioaccumulation Sediments Lumbriculus variegatus

Field Acute to Chronic Toxicity Low Flow P. promelas, C. dubia, D. magna, H. azteca, High Flow Gammarus, C. tentans, or C. riparius, bivalves Mixing Zones Sediment H. azteca, Gammarus, C. tentans or

C. riparius, P. promelas, D. magna, Bivalves Bioaccumulation Low Flow Lumbriculus variegatus, bivalves, fish

High Flow Mixing Zones Sediment

Bioaccumulation Surrogate Low Flow Semipermeable membrane devices High Flow Mixing Zones Interstitial water?

Acclimation

Test organisms are cultured and tested at the same temperature. Test organisms are cultured in the same water that is used in testing, as recommended by EPA (EPA 1994); therefore, no acclimation will be necessary.

Placing Organisms in Test Chambers

Amphipods are introduced into the overlying water below the air–water interface. Weight is measured on a subset of 20 amphipods used to start the test.

Feeding

For each beaker, 1.0 mL of YCT is added daily from Day 0 to Day 42. The amount of food added to the test chambers is kept to a minimum to avoid microbial growth and water fouling. If excess food collects on the sediment, a fungal or bacterial growth may develop on the sediment surface, in which case feeding is suspended for 1 or more days. A drop in dissolved oxygen below 2.5 mg/L during a test may indicate that the food added is not being consumed. Feeding is suspended for the amount of time necessary to increase the dissolved oxygen concentration. If feeding is suspended in one treatment, it should be suspended in all treatments. Detailed records of feeding rates and the appearance of the sediment surface are made daily.

Monitoring a Test

All chambers are checked daily and observations made to assess test organism behavior such as sediment avoidance. However, monitoring effects on burrowing activity of test organisms may be difficult because the test organisms are often not visible during the exposure. The operation of the exposure system is monitored daily.

Measurement of Overlying Water Quality Characteristics

Conductivity, hardness, alkalinity, and ammonia is measured in all treatments at the beginning and at the end of the sediment exposure portion of the test. Water quality characteristics are also


Table D.4 Recommended Toxicity Test Conditions and Test Acceptability Criteria for Ceriodaphnia dubia Screening and Definitive Acute Tests

Test Conditions Recommended

1. Test type: 2. Test duration: 3. Temperature: 4. Light quality: 5. Light intensity: 6. Photoperiod: 7. Test chamber size: 8. Test solution volume:

9. Renewal of test solutions: 10. Age of test organisms: 11. No. organisms per test chamber: 12. No. replicate chambers per concentration: 13. No. organisms per concentration: 14. Feeding regime:

15. Test chamber cleaning: 16. Test solution aeration: 17. Dilution water:

18. Test concentrations:

19. Dilution series:

20. Endpoint:

21. Sampling and sample holding requirements:

22. Sample volume required: 23. Test acceptability criterion:

Static non-renewal, static renewal or flow through 24, 48, or 96 h 20°C ± 1°C; or 25°C ± 1°C Ambient laboratory illumination 10–20 µE/m2/s (50–100 ft-c) (ambient laboratory levels) 16 h light, 8 h darkness 30 mL (minimum) 25 mL (minimum) – For whole sediment tests use 5 mL sediment, 20 mL water

Minimum, after 48 h Less than 24 h old Minimum, 5 for effluent and receiving water tests Minimum, 4 for effluent and receiving water tests Minimum, 20 for effluent and receiving water tests FeedYCT and Selenastrum while holding prior to the test; newly-released young should have food available a minimum of 2 h prior to use in a test; add 0.1 mL each of YCT and Selenastrum 2 h prior to test solution renewal at 48 h

Cleaning not required None Moderately hard synthetic water is prepared using MILLIPORE MILLI-Q® or equivalent deionized water and reagent grade chemicals or 20% DMW, receiving water, groundwater, or synthetic water, modified to reflect receiving water hardness

Effluents: minimum of five effluent concentrations and a control Receiving waters: 100% receiving water and a control Effluents: ≥ 0.5 dilution series Receiving Waters: None, or ≥ 0.5 dilution series Effluents: Mortality (LC50 or NOAEC) Receiving Waters: Mortality (significant difference from control)

Effluents and Receiving Waters: Grab or composite samples are used within 36 h of completion of the sampling period

1 L 90% or greater survival in controls

measured at the beginning and end of the reproductive phase (Day 29 to Day 42). Conductivity will be measured weekly and DO and pH three times/week

Dissolved oxygen is measured a minimum of three times/week and should be at a minimum of 2.5 mg/L. Aeration is used to maintain dissolved oxygen in the overlying water above 2.5 mg/L.

Temperature is measured at least daily in at least one test chamber from each treatment. The daily mean test temperature must be within 1°C of 23°C. The instantaneous temperature must always be within 3°C of 23°C.

Ending a Test

Endpoints monitored include 28-d survival and growth of amphipods and 35-day and 42-day survival, growth, and reproduction (number of young/female) of amphipods. Growth or reproduction of amphipods may be a more sensitive toxicity endpoint compared to survival.

On Day 28, four of the replicate beakers/sediment are sieved with a #40 mesh sieve (425-µm mesh; U.S. standard size sieve) to remove surviving amphipods for growth determinations. Growth


Table D.5 Recommended Toxicity Test Conditions and Test Acceptability Criteria for Daphnia pulex and D. magna Screening and Definitive Acute Tests



9. Renewal of test solutions: 10. Age of test organisms: 11. No. organisms per test chamber: 12. No. replicate chambers per concentration: 13. No. organisms per concentration: 14. Feeding regime:

15. Test chamber cleaning: 16. Test solution aeration: 17. Dilution water:



20. Endpoint:



Static non-renewal, static renewal or flow through 24, 48 or 96 h 20°C ± 1°C; or 25°C ± 1°C Ambient laboratory illumination 10–20 µE/m2/s (50–100 ft-c) (ambient laboratory levels) 16 h light, 8 h darkness 30 mL (minimum) 25 mL (minimum) — for whole sediment tests, use 10 mL, sediment, 40 mL water

Minimum, after 48 h Less than 24 h old Minimum, 5 for effluent and receiving water tests Minimum, 4 for effluent and receiving water tests Minimum, 20 for effluent and receiving water tests Feed YCT and Selenastrum while holding prior to the test; newly released young should have food available a minimum of 2 h prior to use in a test; add 0.1 mL each of YCT and Selenastrum 2 h prior to test solution renewal at 48 h

Cleaning not required None Moderately hard synthetic water prepared using MILLIPORE MILLI-Q or equivalent deionized water and reagent grade chemicals, or 20% DMW, receiving water, groundwater, or synthetic water, modified to reflect receiving water hardness

Effluents: minimum of five effluent concentrations and a control

Receiving Waters: 100% receiving water and a control Effluents: ≥ 0.5 dilution series Receiving Waters: None, or ≥ 0.5 dilution series Effluents: Mortality (LC50 or NOAEC) Receiving Waters: Mortality (significant difference from control)


1 L 90% or greater survival in controls

of amphipods are reported as weight. Dry weight of amphipods in each replicate are determined on Days 28 and 42. Dry weight of amphipods are determined by: (1) transferring rinsed amphipods to a preweighed aluminum pan; (2) drying these samples for 24 hours at 60°C; and (3) weighing the pan and dried amphipods on a balance to the nearest 0.01 mg. Average dry weight of individual amphipods in each replicate is calculated from these data.

On Day 28, the remaining eight beakers/sediment are sieved and the surviving amphipods in each sediment beaker are placed in 300-mL water-only beakers containing 150 to 275 mL of overlying water and a 5 × 5 cm piece of Nitex screen or 3M fiber mat. Each water-only beaker receives 1.0 mL of YCT stock solution and about two volume additions of water daily.

Reproduction of amphipods is measured on Day 35 and Day 42 in the water-only beakers by removing and counting the adults and young in each beaker. On Day 35, the adults are then returned to the same water-only beakers. Adult amphipods surviving on Day 42 are preserved in sugar formalin. The number of adult females is determined by simply counting the adult males (mature male amphipods will have an enlarged second gnathopod) and assuming all other adults are females.


Table D.6 RecommendedToxicityTest Conditions and Test Acceptability Criteria for the Fathead Minnow (Pimephales promelas) Screening and Definitive Acute Tests



9. Renewal of test solutions: 10. Age of test organisms: 11. No. organisms per test chamber: 12. No. replicate chambers per concentration:

13. No. organisms per concentration:

14. Feeding regime:

15. Test chamber cleaning: 16. Test solution aeration:

17. Dilution water:



20. Endpoint:



Static non-renewal, static renewal or flow through 24, 48 or 96 h 20°C ± 1°C or 25°C ± 1°C Ambient laboratory illumination 10–20 µE/m2/s (50–100 ft-c) (ambient laboratory levels) 16 h light, 8 h darkness 250 mL (minimum) 200 mL (minimum) — for whole sediment tests, use 150 mL sediment, 600 mL water

Minimum, after 48 h 1–14 days: 24 h range in age Minimum, 10 for effluent test Minimum, 2 for effluent tests Minimum, 4 for receiving water tests Minimum, 20 for effluents tests Minimum, 40 for receiving waters tests Artemia nauplii are made available while holding prior to the test; add 0.2 mL Artemia nauplii concentrate 2 h prior to test solution renewal at 48 h

Cleaning not required None, unless DO concentration falls below 40% saturation; rate should not exceed 100 bubbles/min

Moderately hard synthetic water prepared using MILLIPORE MILLI -Q or equivalent deionized water and reagent grade chemicals or 20% DMW, receiving water, groundwater, or synthetic water, modified to reflect receiving water and hardness




2 L for effluents and receiving waters 90% or greater survival in controls

The number of females is used to determine number of young/female/beaker from Day 28 to Day 42. Growth will also be measured for these adult amphipods.

Interpretation of Results

Endpoints measured in the 42-day H. azteca test include survival (Days 28, 35, and 42), growth (Days 28 and 42), and reproduction (number of young/female produced from Days 28 to 42). Reproduction is often more variable than growth. Some investigators have shown growth provides unique information that can help discriminate toxic effects of exposure to contaminants in sediment, while others have not seen differences from survival information.

On rare occasions, test organism responses in control sediments may exhibit responses which are less than reference or test sediments. This may be due to the poor nutritional content of the control sediment or other unknown physicochemical factors. Currently, there are no standard control sediments which can be strongly recommended for chronic toxicity testing due to a lack of testing


Table D.7 Recommended Toxicity Test Conditions and Test Acceptability Criteria for the Rainbow Trout (Oncorhynchus mykiss) and Brook Trout (Salvelinus fontinalis) Screening and Definitive Acute Tests


1. Test type: 2. Test duration: 3. Temperature: 4. Light quality: 5. Light intensity: 6. Photoperiod:

7. Test chamber size:

8. Test solution volume:

9. Renewal of test solutions: 10. Age of test organisms:

11. No. organisms per test chamber: 12. No. replicate chambers per concentration:

13. No. organisms per concentration:

14. Feeding regime: 15. Test chamber cleaning: 16. Test solution aeration:

17. Dilution water:



20. Endpoint:


22. Sample volume required:

23. Test acceptability criterion:

Static non-renewal, static-renewal or flow-through 24, 48 or 96 h 12 ± 2°C Ambient laboratory illumination 10–20 µE/m2/s (50–100 ft-c) (ambient laboratory levels) 16 h light, 8 h darkness. Light intensity should be raised gradually over a 15 min period at the beginning of the photoperiod, and lowered gradually at the end of the photoperiod, using a dimmer switch or other suitable control device.

5 L (minimum) (test chamber should be covered to prevent fish from jumping out)

4 L (minimum) — for whole sediment tests, use 80 mL sediment, and 320 mL water

Minimum, after 48 h Rainbow Trout: 5–30 days, “24 h (after yolk sac absorption to 30 days)

Brook Trout: 30–60 days Minimum, 10 for effluent and receiving water tests Minimum, 2 for effluent tests Minimum, 4 for receiving water tests Minimum, 20 for effluent tests Minimum 40 for receiving water tests Feeding not required Cleaning not required None, unless DO concentration falls below 6.0 mg/L; rate should not exceed 100 bubbles/min

Moderately hard synthetic water is prepared using MILLIPORE MILLI-Q or equivalent deionized water and reagent grade chemicals or 20% DMW, receiving water, groundwater, or synthetic water, modified to reflect receiving water hardness




20 L for effluents 40 L for receiving waters 90% or greater survival in controls

and research. Should poor responses be observed in a control sediment, a secondary control or reference sediment may be substituted for comparisons of significance. This will not invalidate the test, but simply adds some degree of uncertainty in the determination of ecological significance.

Recently, the U.S. EPA conducted interlaboratory variance testing with the 42-day H. azteca assay. In these tests, the draft standard methods were used. The minimum detectable differences for amphipod survival at 28 and 42 days ranged from 8 to 12% in moderately contaminated sediments. Minimum detectable differences for reproductive endpoints were higher, as expected, ranging from 19 to 25%.


Table D.8 Recommended Toxicity Test Conditions and Test Acceptability Criteria for the Ceriodaphnia dubia Survival and Reproduction Test


1. Test type: 2. Test duration:

3. Temperature: 4. Light quality: 5. Light intensity: 6. Photoperiod: 7. Test chamber size: 8. Test solution volume:

9. Renewal of test solutions: 10. Age of test organisms: 11. No. neonates per test chamber: 12. No. replicate test chambers per concentration: 13. No. neonates per concentration: 14. Feeding regime:

15. Test solution aeration: 16. Dilution water:


18. Dilution factor:

19. Endpoints: 20. Sampling and sample holding requirements:

21. Sample volume required: 22. Test acceptability criteria:

Static renewal Until 60% of control females have three broods (maximum test duration 8 days)

25 ± 1°C Ambient laboratory illumination 10–20 µE/m2/s (50–100 ft-c) (ambient laboratory levels) 16 h light, 8 h darkness. 30 mL (minimum) 15 mL (minimum) — for whole sediment assays, use 5 mL sediments and 20 mL water

Daily Less than 24 h and all released within an 8 h period 1 10 10 Feed 0.1 mL each of YCT and 0.1 mL of algal suspension per test chamber daily

None Uncontaminated source of receiving water or other natural water, synthetic water prepared using MILLIPORE MILLI-Q or equivalent deionized water and reagent grade chemicals or DMW

Effluents: Minimum of five effluent concentrations and a control

Receiving water: 100% receiving water or minimum of five concentrations and a control

Effluents ≥ 0.5 Receiving waters: None or ≥ 0.5 Survival and reproduction For on-site tests, samples collected daily and used within 24 h of the time they are removed from the sampling device. For off-site tests, a minimum of three samples collected on days one, three, and five with a maximum holding time of 36 h before first use

1 L 80% or greater survival and an average of 15 or more young per surviving female in the control solutions; 60% of surviving control organisms must produce three broods

METHODS FOR CONDUCTING LONG-TERM SEDIMENT TOXICITY TESTS WITH CHIRONOMUS TENTANS

Conditions for conducting a long-term sediment toxicity test with C. tentans are summarized in Table D.17. A general activity schedule is outlined in Table D.18.

The long-term sediment toxicity test with C. tentans is conducted at 23°C with a 16L:8D photoperiod at an illuminance of about 500 to 1000 lux. Test chambers, sediment addition, water renewal, and water quality monitoring are as described above for H. azteca.

A total of 16 replicates, each containing 12, <24-hour-old larvae are tested for each sample. For the total of 16 replicates, the assignment of beakers is as follows: initially, 12 replicates are set up on Day –1, of which 4 replicates are used for 20-day growth and survival endpoints and 8 replicates for determination of emergence and reproduction. It is typical for males to begin emerging 4 to 7 days before females. Midges in each test chamber are fed 1.5 mL of a 4-g/L Tetrafin™

suspension daily. Endpoints monitored include 20-day survival and ash-free dry weight, emergence; and time to death (adults). Reproduction and egg hatchability are not assessed.


Table D.9 Recommended Toxicity Test Conditions and Test Acceptability Criteria for the Fathead Minnow (Pimephales promelas) Larval Survival and Growth Test



9. Renewal of test solutions: 10. Age of test organisms:

11. No. larvae per test chamber: 12. No. replicate test chambers per concentration: 13. No. larvae per concentration: 14. Source of food: 15. Feeding regime:

16. Test chamber cleaning: 17. Test solution aeration:

18. Dilution water:



21. Endpoints: 22. Sampling and sample handling requirements:

23. Sample volume required: 24. Test acceptability criteria:

Static renewal 7 days 25 ± 1°C Ambient laboratory illumination 10–20 µE/m2/s (50–100 ft-c) (ambient laboratory levels) 16 h light, 8 h darkness. 500 mL (minimum) 250 mL (minimum) — for whole sediment tests use 50 mL sediment and 200 mL water

Daily Newly hatched larvae less than 24 h old. If shipped, not more than 48 h old, 24 h range in age

15 (minimum of 10) 4 (minimum of 3) 60 (minimum of 30) Newly hatched Artemia nauplii (less than 24 h old) Feed 0.1 mL newly hatched (less than 24 h old) brine shrimp nauplii three times daily at 4 h intervals or, as a minimum 0.15 mL twice daily, 6 h between feedings (at the beginning of the work day following renewal). Sufficient larvae are added to provide an excess. Larvae are not fed during the final 12 h of the test

Siphon daily, immediately before test solution renewal None, unless DO concentration falls below 4.0 mg/L. Rate should not exceed 100 bubbles/min. Uncontaminated source of receiving water or other natural water, synthetic water prepared using MILLIPORE MILLI-Q or equivalent deionized water and reagent grade chemicals or DMW

Effluents: Minimum of five effluent concentration and a control


Effluents: ≥ 0.5 Receiving waters: none or ≥ 0.5 Survival and growth (weight) For on-site tests, samples are collected daily, and used within 24 h of the time they are removed from the sampling device. For off-site tests, a minimum of three samples are collected on days one, three, and five with a maximum holding time of 36 h before first use

2.5 L/day 80% or greater survival in controls: average dry weight per surviving organism in control chambers equals or exceeds 0.25 mg

Collection of Egg Cases

Egg cases are obtained from adult midges held in a sex ratio of 1:3 male:female. Adults are collected 4 days before starting a test. The day after collection of adults, 6 to 8 of the larger “C”shaped egg cases are transferred to a petri dish with culture water and incubated (at 23°C). Hatching typically begins around 48 hours and larvae typically leave the egg case 24 hours after the first hatch.

Hatching of Eggs

Hatching of eggs should be complete by about 72 hours. Hatched larvae remain with the egg case for about 24 hours and appear to use the gelatinous component of the egg case as an initial


Table D.10 Toxicity Test Conditions and Test Acceptability Criteria for the Fathead Minnow (Pimephales promelas) Embryo-Larval Survival and Teratogenicity Test



9. Renewal of test solutions: 10. Age of test organisms: 11. No. embryos per test chamber: 12. No. replicate test chambers per concentration: 13. No. embryos per concentration: 14. Feeding regime: 15. Test solution aeration: 16. Dilution water:



19. Endpoint: 20. Sampling and sample handling requirements:

21. Sample volume required:

22. Test acceptability criteria:

Static renewal 7 days 25 ± 1°C Ambient laboratory illumination 10–20 µE/m2/s (50–100 ft-c) (ambient laboratory levels) 16 h light, 8 h darkness. 150–500 mL 70 mL (minimum) — for whole sediment tests, use 50 mL sediment and 200 mL water

Daily Less than 36 h old embryos (maximum 48 h if shipped) 15 (minimum of 10) 4 (minimum of 3) 60 (minimum of 30) Feeding not required None, unless DO concentration falls below 4 mg/L Uncontaminated source of receiving water or other natural water, synthetic water prepared using MILLIPORE MILLI-Q or equivalent deionized water and reagent grade chemicals or DMW

Effluents: Minimum of five effluent concentration and a control


Effluents: ≥ 0.5 Receiving waters: none or ≥ 0.5 Combined mortality (dead and deformed organisms) For on-site tests, samples are collected daily, and used within 24 h of the time they are removed from the sampling device. For off-site tests, a minimum of three samples are collected on days one, three, and five with a maximum holding time of 36 h before first use

1.5 to 2.5 L/day depending on the volume of test solutions used

80% or greater survival in controls

source of food. After the first 24-hour period with larvae hatched, egg cases are transferred from the incubation petri dish to another dish with clean test water. The action of transferring the egg case stimulates the remaining larvae to leave the egg case within a few hours. These are the larvae that are used to start the test.

Placing Organisms in Test Chambers

To start the test, larvae are collected with a Pasteur pipette from the bottom of the incubation dish with the aid of a dissecting microscope. Test organisms are pipetted directly into overlying water. Larvae are transferred to exposure chambers within 4 hours of emerging from the egg case.

Feeding

Each beaker received a daily addition of 1.5 mL of Tetrafin (4 mg/mL dry solids). Feeding is curtailed under circumstances described in the amphipod methods.

Dissolved Oxygen

Routine chemistries on Day 0 should be taken before organisms are placed in the test beakers. Excursions of DO as low as 1.5 mg/L did not seem to have an effect on midge survival and


Table D.11 Toxicity Test Conditions and Test Acceptability Criteria for the Algal (Selenastrum capricornutum) Growth Test


1. Test type: 2. Test duration: 3. Temperature: 4. Light quality: 5. Light intensity: 6. Photoperiod: 7. Test chamber size: 8. Test solution volume: 9. Renewal of test solutions:

10. Age of test organisms: 11. Initial cell density in test chamber:

Static non-renewal 48–96 h 25 ± 1°C “Cool white” fluorescent lighting 86 ± 8.6 µE/m2/s (400 ± 40 ft-c or 4306 lux) Continuous illumination 125 or 250 mL 50 or 100 mL None 4 to 7 days 10,000 cells/mL

12. No. replicate chambers per sample: 4 (minimum or 3) 13. Shaking rate: 100 rmp continuous, or twice daily by hand 14. Test solution aeration: None 15. Dilution water: Algal stock culture medium, enriched uncontaminated source of

receiving or other natural water, synthetic water prepared using MILLIPORE MILLI-Q or equivalent deionized water and reagent grade chemicals, or DMW without EDTA or enriched surface water

16. Test concentrations: Effluents: Minimum of five effluent concentrations and a control Receiving water: 100% receiving water or minimum of five concentrations and a control

17. Test dilution factor: Effluents: ≥ 0.5 Receiving waters: None or ≥ 0.5

18. Endpoint: Growth (cell counts, chlorophyll fluorescence, absorbance, biomass)

19. Sample requirements: For on-site tests, one sample collected at test initiation, and used within 24 h of the time being removed from the sampling device. For off-site tests, holding time must not exceed 36 h

20. Sample volume required: 1 or 2 L depending on test volume 21. Test acceptability criteria: 1 × 106 cells/mL with EDTA or 2 × 105 cells/mL without EDTA in

the controls: Variability of controls should not exceed 20%

development (P.K. Sibley, University of Guelph, Guelph, Ontario, personal communication). Based on these findings, periodic depressions of DO below 2.5 mg/L (but not below 1.5 mg/L) are not likely to adversely affect test results, and thus should not be a reason to discard test data. Nonetheless, tests should be managed toward a goal of DO > 2.5 mg/L to ensure satisfactory performance. If the DO level of the water falls below 2.5 mg/L for any one treatment, aeration is conducted in all replicates for the duration of the test.

Monitoring Survival and Growth

At 20 days, four of the initial 12 replicates are selected for use in growth and survival measurements. Using a #40 sieve (425-µm mesh) to remove larvae from sediment, C. tentans is collected. Surviving larvae are kept separated by replicate for weight measurements; if pupae are recovered, these organisms are included in survival data but not included in the growth data.

The AFDW of midges is determined for the growth endpoint. All living larvae per replicate are combined and dried to a constant weight (e.g., 60°C for 24 hours). All weigh boats are ashed before use to eliminate weighing errors due to the pan oxidizing during ashing. The sample is brought to room temperature in a desiccator and weighed to the nearest 0.01 mg to obtain mean weights per surviving organism per replicate. The dried larvae in the pan are then ashed at 550°C for 2 hours. The pan with the ashed larvae is then reweighed and the tissue mass of the larvae is determined as the difference between the weight of the dried larvae plus pan and the weight of the ashed larvae plus pan.


Table D.12 Toxicity Test Conditions and Test Acceptability Criteria for the Amphipod (Hyalella azteca) Survival Test


1. Test type: Whole sediment toxicity test with renewal of overlying water 2. Test duration: 10 d 3. Temperature: 23 ± 1°C 4. Light quality: Wide-spectrum fluorescent lights 5. Illuminance: About 100 to 1000 lux 6. Photoperiod: 16 h light, 8 h dark 7. Test chamber size: 300 mL high-form lipless beaker 8. Sediment volume: 100 mL 9. Overlying water volume: 175 mL

10. Renewal of overlying water: 2 volumes additions/d; continuous or intermittent (e.g., 1 volume addition every 12 h)

11. Age of test organisms: 7 to 14 d old at the start of the test (1 to 2 d range in age) 12. No. organisms per test chamber: 10 13. No. replicate chambers per treatment: Depends on the objective of the test. Eight replicates are

recommended for routine testing 14. Feeding regime: YCT food, fed 1.0 mL daily (1800 mg/L stock) to each test

chamber 15. Test solution aeration: None, unless DO in overlying water falls below 2.5 mg/L 16. Overlying water: Culture water, well water, surface water, site water, or

reconstituted water 17. Test chamber cleaning: If screens become clogged during a test, gently brush the

outside of the screen 18. Overlying water quality: Hardness, alkalinity, conductivity, pH, and ammonia at the

beginning and end of a test; temperature and dissolved oxygen daily

19. Endpoint: Survival and growth 20. Test acceptability criterion: Minimum mean control survival of 80% and measurable growth

of test organisms in the control sediment

Monitoring Emergence

Emergence traps are placed on the reproductive replicates on Day 20 (emergence traps for the auxiliary beakers are added at the corresponding 20-day time interval for those replicates. At 23°C, emergence in control sediments typically begins on or about Day 23 and continues for about 2 weeks. However, in contaminated sediments, the emergence period may be extended by weeks.

Two categories are recorded for emergence: complete emergence and partial emergence. Complete emergence occurs when an organism has shed the pupal exuviae completely and escapes the surface tension of the water. If complete emergence has occurred but the adult has not escaped the surface tension of the water, the adult will die within 24 hours. Therefore, 24 hours will elapse before this death is recorded. Partial emergence occurs when an adult has only partially shed the pupal exuvia. These adults will also die, an event which can be recorded after 24 hours.

Between Day 23 and the end of the test, emergence of males and females, pupal and adult mortality, and time to death for adults is recorded daily for the reproductive replicates.

Ending a Test

The point at which the life cycle test is ended depends upon the sediments being evaluated. In clean sediments, the test typically requires 40 to 50 days from initial setup to completion if all possible measurement endpoints are evaluated. However, test duration will increase in the presence of environmental stressors that act to reduce growth and delay emergence. Where a strong gradient of sediment contamination exists, emergence patterns between treatments will likely become asynchronous, in which case each treatment needs to be ended separately. For this reason, emergence is used as a guide to decide when to end a test. Testing will be terminated with completion of emergence.


Table D.13 Toxicity Test Conditions and Test Acceptability Criteria for the Midge (Chironomus tentans) Survival Test


1. Test type: Whole sediment toxicity test with renewal of overlying water 2. Test duration: 10 d 3. Temperature: 23 ± 1°C 4. Light quality: Wide-spectrum fluorescent lights 5. Illuminance: About 100 to 1000 lux 6. Photoperiod: 16 h light, 8 h dark 7. Test chamber size: 300 mL high-form lipless beaker 8. Sediment volume: 100 mL 9. Overlying water volume: 175 mL

10. Renewal of overlying water: 2 volumes additions/d; continuous or intermittent (e.g., 1 volume addition every 12 h)

11. Age of test organisms: Second to third instar larvae (about 10 d old larvae; all organisms must be third instar or younger with at least 50% of the organisms at third instar)

12. No. organisms per test chamber: 10 13. No. replicate chambers per treatment: Depends on the objective of the test. Eight replicates are

recommended for routine testing 14. Feeding regime: Tetrafin goldfish food, fed 1.5 ml daily to each test chamber

(1.5 mL contains 6.0 mg of dry solids) 15. Test solution aeration: None, unless DO in overlying water falls below 2.5 mg/L 16. Overlying water: Culture water, well water, surface water, site water, or




19. Endpoint: Survival and growth (ash-free dry weight, AFDW) 20. Test acceptability criterion: Minimum mean control survival must be 70%, with minimum

mean weight/surviving control organisms of 0.48 mg AFDW

For treatments in which emergence has occurred, the treatment (not the entire test) is ended when no further emergence is recorded over a period of 7 days (the 7-day criterion). At this time, all beakers of the treatment are sieved through a #40 mesh screen (425 µm) to recover remaining larvae, pupae, or pupal casts. When no emergence is recorded in a treatment at any time during the test, that treatment can be ended once emergence in the control sediment has ended using the 7-day criterion.

Interpretation of Results

Endpoints measured in the C. tentans test include survival, growth, and emergence. On rare occasions, test organisms in control sediments may exhibit responses which are less than reference or test sediments. This may be due to the poor nutritional content of the control sediment or other unknown physicochemical factors. Currently, there are no standard control sediments that can be strongly recommended for chronic toxicity testing due to a lack of testing and research. Should poor responses be observed in a control sediment, a secondary control or reference sediment may be substituted for comparisons of significance. This will not invalidate the test, but simply adds a degree of uncertainty to the determination of ecological significance.

Recently, the U.S. EPA conducted interlaboratory variance testing with the chronic C. tentans assay. In these tests, the draft standard methods were used. The minimum detectable differences have not been calculated at this time, but will be available in the near future to provide a point of comparison for the test assays. It is expected that the minimum detectable difference for 28-day survival and emergence endpoints will be in the 15 to 30% range.


Table D.14 Toxicity Test Conditions and Test Acceptability Criteria for the Oligochaete (Lumbriculus variegatus) Survival Test


1. Test type: Whole sediment toxicity test with renewal of overlying water 2. Test duration: 23 d 3. Temperature: 23 ± 1°C 4. Light quality: Wide-spectrum fluorescent lights 5. Illuminance: About 100 to 1000 lux 6. Photoperiod: 16 h light, 8 h dark 7. Test chamber size: 4 to 6 L aquaria with stainless steel screens or glass stand pipes 8. Sediment volume: 1 L or more depending on TOC 9. Overlying water volume: 1 L or more depending on TOC

10. Renewal of overlying water: 2 volumes additions/d; continuous or intermittent (e.g. 1 volume addition every 12 h)

11. Age of test organisms: Adults 12. No. organisms per test chamber: Ratio of total organic carbon in sediment to organism dry weight

should be no less than 50:1. Minimum of 1 g/ replicate, preferably 5 g/replicate

13. No. replicate chambers per treatment: Depends on the objective of the test. Five replicates are recommended for routine testing

14. Feeding regime: None 15. Test solution aeration: None, unless DO in overlying water falls below 2.5 mg/L 16. Overlying water: Culture water, well water, surface water, site water, or




19. Endpoint: Bioaccumulation 20. Test acceptability criterion: Performance-based criteria specifications

Four test species will be evaluated in situ in exposure chambers. The exposure chambers are constructed on plastic core tubes of ~3-in diameter and 4-in length. Two windows are cut on opposite sides of the chamber and covered with nylon mesh. The mesh size varies with the experimental treatment, ranging from 10- to 1000-µm openings. For high flow testing, only water column chambers will be exposed. One duplicate set of chambers will have reduced mesh size openings to allow determinations of flow and suspended solids effects. Chambers are placed in the stream, either in the overlying water or partially buried in the sediment, with exposures varying with the treatment. Organisms are slowly acclimated to site water temperatures and then added to each test chamber (10 organisms/chamber). The age of the organisms, handling, and culturing follow U.S. EPA toxicity test methods for short-term chronic toxicity testing. For bioaccumulation testing, additional organisms are placed to provide enough tissue mass. For the oligochaete assay, 5 g of tissue are used in each chamber. Chambers are placed in the stream in replicates of four and secured with netting and steel stakes. At Days 2 and 10, chambers will be retrieved and organisms enumerated within 2 hours of collection. Test endpoints are shown in Table D.20.

The effects of water quality during high flow events will be measured at all test sites. This will involve exposures using chambers with small and large mesh sizes to vary the organism exposure to suspended solids. Exposures will be for 48 hours and include D. magna, H. azteca, and C. tentans. Testing will only be conducted when organisms can be exposed to a significant first flush event.

IN SITU TESTING USING CONFINED ORGANISMS

There are many reasons for evaluating toxicity and bioaccumulation in situ, such as those shown in Table D.19 and discussed in Section 6. Numerous assessments of stormwater quality have found


Table D.15 Test Conditions for Conducting a 42-day Sediment Toxicity Test with Hyalella azteca

Parameter Conditions

1. Test type: Whole-sediment toxicity test with renewal of overlying water 2. Temperature: 23 ± 1°C 3. Light quality: Wide-spectrum fluorescent lights 4. Illuminance: About 500 to 1000 lux 5. Photoperiod: 16L:8D 6. Test chamber: 300-mL high-form lipless beaker 7. Sediment volume: 100 mL 8. Overlying water volume: 175 mL in the sediment exposure from Day 0 to Day 28 (175

to 275 mL in the water-only exposure from Day 28 to Day 42) 9. Renewal of overlying water: 2 volume additions/d; continuous or intermittent (e.g., one

volume addition every 12 h) 10. Age of organisms: 7- to 8-d old at the start of the test 11. Number of organisms/chamber: 10 12. Number of replicate chambers/treatment: 12 (4 for 28-d survival and growth and 8 for 35- and 42-d

survival, growth, and reproduction). Reproduction is more variable than growth or survival; hence, more replicates might be needed to establish statistical differences among treatments

13. Feeding: YCT food, fed 1.0 mL (1800 mg/L stock) daily to each test chamber

14. Aeration: None, unless dissolved oxygen in overlying water drops below 2.5 mg/L

15. Overlying water: Culture water, well water, surface water, or site water. Use of reconstituted water is not recommended

16. Test chamber cleaning: If screens become clogged during a test; gently brush the outside of the screen

17. Overlying water quality: Hardness, alkalinity, conductivity, and ammonia at the beginning and end of a sediment exposure (Day 0 and 28). Temperature daily. Conductivity weekly. Dissolved oxygen (DO) and pH three times/ week. Concentrations of DO should be measured more often if DO drops more than 1 mg/L since the previous measurement.

18. Test duration: 42 d 19. Endpoints: 28-d survival and growth; 35- and 42-d survival, growth,

reproduction, and number of adult males and females on Day 42

20. Test acceptability: Minimum mean control survival of 80% on Day 28

the following study design example useful. The typical assessment will be an upstream–downstream evaluation of an outfall with an additional reference site. The assessment must include both low and high flow periods to separate the role of stormwater and nonpoint source runoff from low flow conditions that may include point sources and groundwater upwelling inputs. For in situ toxicity and/or bioaccumulation tasks, a variety of exposure periods can be used, depending on several issues, such as species resilience, meteorological conditions, concern over acute vs. chronic effects, and available resources (longer assessments are more expensive). A great challenge in any stormwater assessment is detecting chronic toxicity effects. The literature has documented (see Chapter 6) that delayed effects may occur days to weeks after pulse exposures to pesticides or metals. This is obviously difficult to determine in routine receiving water assessments. However, given the reality that chronic toxicity may be occurring, it is important to try and assess effects for as long a period as possible. Some test species, such as the cladocerans C. dubia and D. magna and early life stages of the fathead minnow P. promelas, do not survive well within typical in situ chambers for more than 4 days. The benthic macroinvertebrates, such as the amphipods H. azteca and Gammarus, midge C. tentans, and bivalves, can be exposed for periods of over a week (Brooker 2000). Fish may also be exposed for longer periods, but often require routine feeding. When determining bioaccumulation potential, the oligochaete worm L. variegatus is recommended. It accumulates nonpolar organic chemicals relatively quickly, so exposures as short as 4 days are acceptable.


Table D.16 General Activity Schedule for Conducting a 42-d Sediment Toxicity Test with Hyalella azteca

Day Activity

Pre-Test

–7 Remove adults and isolate <24-h-old amphipods (if procedures outlined in Section 12.3.4 are followed).

–8 Separate known-age amphipods from the cultures and place in holding chambers. Begin preparing food for the test. The <24-h amphipods are fed 10 mL of YCT (1800 mg/L stock solution) and 10 mL of Selenastrum capricornutum (about 3.0 x 107 cells/mL) on the first day of isolation and 5 mL of both YCT and S. capricornutum on the 3rd and 5th d after isolation.

–6 to –2 Feed and observe isolated amphipods, monitor water quality (e.g., temperature and dissolved oxygen).

–1 Feed and observe isolated amphipods, monitor water quality. Add sediment into each test chamber, place chambers into exposure system, and start renewing overlying water.

Sediment Test

0 Measure total water quality (pH, temperature, dissolved oxygen, hardness, alkalinity, conductivity, ammonia). Transfer ten 7- to 8-day-old amphipods into each test chamber. Release organisms under the surface of the water. Add 1.0 mL of YCT (1800 mg/L stock) into each test chamber. Archive 80 amphipods for dry weight determination. Observe behavior of test organisms.

1 to 27 Add 1.0 mL of YCT to each test beaker. Measure temperature daily, conductivity weekly, and dissolved oxygen (DO) and pH three times/week. Observe behavior of test organisms.

28 Measure temperature, dissolved oxygen, pH, hardness, alkalinity, conductivity and ammonia. End the sediment-exposure portion of the test by collecting the amphipods with a #40 mesh sieve (425-µm mesh; U.S. standard size sieve). Use four replicates for growth measurements: count survivors and preserve organisms in sugar formalin for growth measurements. Eight replicates for reproduction measurements: Place survivors in individual replicate water-only beakers and add 1.0 mL of YCT to each test beaker/d and 2 volume additions/d of overlying water.

Reproduction Phase

29 to 35 Feed daily. Measure temperature daily, conductivity weekly, DO and pH three times a week. Measure hardness and alkalinity weekly. Observe behavior of test organisms.

35 Record the number of surviving adults and remove offspring. Return adults to their original individual beakers and add food.

36 to 41 Feed daily. Measure temperature daily, conductivity weekly, DO and pH three times a week. Measure hardness and alkalinity weekly. Observe behavior of test organisms.

41 Same as Day 1. Measure total water quality (pH, temperature, dissolved oxygen, hardness, alkalinity, conductivity, ammonia).

42 Record the number of surviving adults and offspring. Surviving adult amphipods on Day 42 are preserved in sugar formalin solution. The number of adult males in each beaker is determined from this archived sample. This information is used to calculate the number of young produced per female per replicate from Day 28 to Day 42.

A routine assessment of in situ toxicity and bioaccumulation requires that organisms be deployed during low flow conditions; once when the entire exposure period is at baseflow and a second time that captures a high flow event. The organisms at baseflow should be exposed for a period of time greater than or equal to the period of the high flow exposure period (usually 2 to 4 days). Another useful design is to deploy a large number of replicates on Day 0 and then subsample every 2 days for an extended period (such as 14 days). Between one and four species can be evaluated simultaneously, depending on available resources. Often two species are used in each test chamber (as described below). The in situ chambers are constructed of clear core sampling tubes (cellulose acetate butyrate) cut to a length of approximately 15 cm. Polyethylene closures cap each end. Two rectangular windows (~85% of the core surface area) are usually covered with 80 µm Nitex® mesh and silicon glued opposite each other. The mesh size varies with the experimental treatment, ranging from 10 to 1000 µm openings. For high flow testing, only water column chambers need be exposed. Duplicate sets of chambers having small vs. large mesh size openings (e.g., 10 vs. 250 µm) allow determinations of flow and suspended solids effects. The source of toxicity/bioaccumulation can also be measured as originating from sediments or overlying water by varying the chamber posi-


Table D.17 Test Conditions for Conducting a Long-Term Sediment ToxicityTest with Chironomus tentans

Parameter Conditions

1. Test type: Whole-sediment toxicity test with renewal of overlying water 2. Temperature: 23 ± 1°C 3. Light quality: Wide-spectrum fluorescent lights 4. Illuminance: About 500 to 1000 lux 5. Photoperiod: 16L:8D 6. Test chamber: 300-mL high-form lipless beaker 7. Sediment volume: 100 mL 8. Overlying water volume: 175 mL 9. Renewal of overlying water: 2 volume additions/d; continuous or intermittent (e.g.,

one volume addition every 12 h) 10. Age of organisms: <24-hour-old larvae 11. Number of organisms/chamber: 10 12. Number of replicate chambers/treatment: 16 13. Feeding: Tetrafin goldfish food, fed 1.5 mL daily to each test chamber

(1.5 mL contains 6.0 mg of dry solids); starting Day –1 14. Aeration: None, unless dissolved oxygen in overlying water drops

below 2.5 mg/L 15. Overlying water: Culture water, well water, surface water, site water, or

reconstituted water 16. Test chamber cleaning: If screens become clogged during a test; gently brush the

outside of the screen 17. Overlying water quality: Hardness, alkalinity, conductivity, and ammonia at the

beginning and end of a test. Temperature daily (ideally continuously). Dissolved oxygen (DO) and pH three times/week. Concentrations of DO should be measured more often if DO has declined by more than 1 mg/L since previous measurement.

18. Test duration: About 40 to 50 d; each treatment is ended separately when no additional emergence has been recorded for seven consecutive days. When no emergence is recorded from a treatment, termination of that treatment should be based on the control sediment using this 7-d criterion.

19. Endpoints: 20-d survival and AFDW; female and male emergence, adult mortality

20. Test acceptability: Minimum average size of C. tentans in the control sediment at 20 d must be at least 0.6 mg/surviving organism as dry weights or 0.48 mg/surviving organism as AFDW. Emergence should be ≥ 50%. Time to death after emergence is <6.5 d for males and <5.1 d for females.

tioning and design. Prior to chamber deployment, 10 of each organism (H. azteca, C. tentans, and D. magna) were gently added to 50-mL test tubes of culture water for ease of transport to field locations (one test tube contained one species only). Transportation of organisms to field sites by this method has proven to minimize handling and travel-related stressors (Chappie and Burton 1997). Upon acclimation, in situ chambers capped on one end were immersed into the river, allowing water to fill the chamber by infiltration through the mesh, and test organisms were slowly delivered from the test tubes into the open end and the chambers then capped. Before placement into in situ baskets, chambers were held below the water surface and purged of all internal air. Chambers exposed to the sediment interface are secured under wire baskets (see Figure 6.161) and placed with the mesh windows against the sediment. Quadruplicate chambers exposed to overlying waters are secured on top of the wire baskets. The baskets were weighted down with bricks and anchored to the stream bed with rebar. Organisms are acclimated to site water temperatures slowly (1 to 2 degree/hour) and then added to each test chamber (10 organisms/chamber). For example, C. tentans and H. azteca were placed together in replicate chambers for a total of 20 organisms per chamber. Ground-up laboratory paper toweling is provided as a substrate to reduce stress on these benthic species. Test water for laboratory controls should be the organism culture water. These controls


Table D.18 General Activity Schedule for Conducting a Long-Term Sediment Toxicity Test with Chironomus tentans

Day Activity

–4 Start reproduction flask with cultured adults (1:3 male:female ratio). For example for 15 to 25 egg cases, 10 males and 30 females are typically collected. Egg cases typically range from 600 to 1500 egg/case.

–3 Collect egg cases (a minimum of 6 to 8) and incubate at 23°C. –2 Check egg cases for viability and development. –1 1. Check egg cases for hatch and development.

2. Add 100 mL of homogenized test sediment to each replicate beaker and place in corresponding treatment holding tank. After sediment has settled for at least 1 h, add 1.5 mL Tetrafin slurry (4g/L solution) to each beaker. Overlying water renewal begins at this time.

0 1. Transfer all egg cases to a crystallizing dish containing control water. Discard larvae that have already left the egg cases in the incubation dishes. Add 1.5 mL food to each test beaker with sediment before the larvae are added. Add 12 larvae to each replicate beaker (beakers are chosen by random block assignment). Let beakers sit (outside the test system) for 1 h following addition of the larvae. After this period, gently immerse all beakers into their respective treatment holding tanks.

2. Measure temperature, pH, hardness, alkalinity, dissolved oxygen, conductivity and ammonia at start of test.

1–End On a daily basis, add 1.5 mL food to each beaker. Measure temperature daily. Measure the pH and dissolved oxygen three times a week during the test. If the DO has declined more than 1 mg/L since previous reading, increase frequency of DO measurements and aerate if DO continues to be less than 2.5 mg/L. Measure hardness, alkalinity, conductivity, ammonia weekly.

6 For auxiliary male production, start reproduction flask with culture adults (e.g., 10 males and 30 females; 1:3 male to female ratio).

7–10 Set up schedule for auxiliary male beakers (4 replicates/treatment) same as that described above for Day –3 to Day 0.

19 In preparation for weight determinations, ash weigh-pans at 550°C for 2 h. Note that the weigh boats should be ashed before use to eliminate weighing errors due to the pan oxidizing during ashing of samples.

20 Randomly select four replicates from each treatment and sieve the sediment to recover larvae for growth and survival determinations. Pool all living larvae per replicate and dry the sample to a constant weight (e.g., 60°C for 24 h). Install emergence traps on each reproductive replicate beakers.

21 The sample with dried larvae is brought to room temperature in a desiccator and weighed to the nearest 0.01 mg. The dried larvae in the pan are then ashed at 550oC for 2 h. The pan with the ashed larvae is then reweighed and the tissue mass of the larvae determined as the difference between the weight of the dried larvae plus pan and the weight of the ashed larvae plus pan.

23–End On a daily basis, record emergence of males and females, pupal, and adult mortality, and time to death for previously collected adults.

33–End Transfer males emerging from the auxiliary male replicates to individual inverted petri dishes. The auxiliary males are used for mating with females from corresponding treatments from which most of the males had already emerged or in which no males emerged.

40–End After 7 d of no recorded emergence in a given treatment, end the treatment by sieving the sediment to recover larvae, pupae, or pupal exuviae. When no emergence occurs in a test treatment, that treatment can be ended once emergence in the control sediment has ended using the 7-d criterion.

are typically maintained in a hotel room during field assessments. The age of the organisms, handling, and culturing follow U.S. EPA toxicity test methods for short-term chronic toxicity testing. For bioaccumulation testing, additional organisms are placed to provide enough tissue mass. For the oligochaete assay, 1 to 5 g of tissue (equal to approximately 1:10 animal wet wt:sediment organic carbon) is used in each chamber, depending on analytical requirements. After exposures of 1 to 30 days depending on species and objectives, chambers were gently lifted out of the river and placed into coolers of site water and returned to the field laboratory for enumeration. Upon arrival at the lab, chambers were checked for damage, the outsides rinsed, then individually emptied into crystallizing dishes and the survivors of each species enumerated and logged. Typical measurement endpoints are shown in Table D.20.


Table D.19 In Situ Stressor and Sediment Toxicity Tasks and Outcomes

Task Rationale and Outcome

1. Sediment toxicity: H. azteca, C. tentans

2. In situ toxicity and uptake: D. magna, H. azteca, C. tentans, L. variegatus

3. In situ partitioning of exposure and D. magna, H. azteca

4. In situ assessment of bioaccumulation and transport potential: SPMDs and peepers.

Laboratory measure of sediment chronic toxicity. Trigger for comprehensive sediment toxicity survey. Determine the potential for adverse effects on benthic organisms.

Realistic field exposures to water, suspended solids, and sediments. Determine low and high flow responses. Relate to storm flow and food web modeling. Assess the potential for, and source of, adverse effects on the ecosystem.

In field exposures, determine and rank primary stressors: flow and stressors: turbidity, photoinduced toxicity, ammonia, metals, nonpolar organics, overlying water, pore water. Relate to transport and food web modeling. Assess the contribution and source of various stressors that produce adverse effects.

In field exposures, determine presence and potential for uptake of nonpolar organics through time with SPMDs in surficial waters and pore waters. Assess the presence and transport of contaminants through time with peepers.Target side channel seepage to support transport and food web modeling.

Table D.20 In situ Toxicity and Bioaccumulation Measurement Endpoints

Test Organism Endpoints

Daphnia magna Survival (2 d) Hyalella azteca Survival (2, 7 d) Tissue concentration (7 d) Chironomus tentans Survival (2, 7 d), growth (7 d), tissue concentration (7 d) Lumbriculus variegatus Tissue concentration (7 d)

The effects of water quality during high flow events should be measured at all test sites. Physicochemical water quality parameters are measured as often as is practical. Preferably, continuous measures of flow and general water quality parameters are made using a data sonde-type instrument. At a minimum, however, measures are made at test initiation, then again at test termination at each field site for each of the following: temperature (°C), dissolved oxygen (mg/L), pH, hardness (mg/L CaCO3), alkalinity (mg/L CaCO3), turbidity (NTU), conductivity (µmhos), and flow. Samples for other potentially useful parameters, such as ammonia, pathogen indicators, BOD, and nutrients, are also collected.

Organisms sampled for tissue analyses are allowed to depurate in culture for several hours. Following that time, organisms are counted, weighed, and frozen. Tissue analyses should be conducted by a laboratory capable of low detection limits with small quantities of tissues.

TOXICITY IDENTIFICATION EVALUATIONS

The toxicity identification evaluation (TIE) is a process by which effluent or pore water samples are fractionated into various classes of contaminants and then tested for toxicity. This allows one to characterize which class of contaminants is primarily responsible for toxicity (EPA 1991a,b). These groups of contaminants include: pH-sensitive and volatile compounds (such as ammonia), metals, oxidant/reductants, and nonpolar organics. Toxicity is determined by exposing C. dubia for 24 hours to the various treatment fractionations and then measuring survival. A TIE was conducted following modified draft EPA guidelines for TIEs of sediments (EPA 1991b). Pore water aliquots were used for initial toxicity tests (within 24 hours of sample receipt), baseline ambient pore water, pH adjusted with aeration, pH adjusted with filtration, pH adjusted with C18 filtration, sodium thiosulfate addition, and EDTA addition fractions. If toxicity is removed in any fraction, subsequent chemical analyses will be conducted to confirm the removal of compounds which may be contributing to pore water toxicity. These manipulations and data interpretation can be quite involved and should only be conducted by a laboratory with documented experience.


TOXICITY — MICROTOX SCREENING TEST

Scope and Application

This test measures the reduction of light output at a specific time during the run by bacteria exposed to a water sample. This light output is compared to that of a control sample to calculate relative toxicity. The Microtox Screening Procedure has a range of relative toxicities between 0 and 100% (light output reduction, as compared to the control).

Summary of Method

The Microtox Screening Procedure uses a bioluminescent marine bacteria, Photobacterium phosphoreum, to measure the toxicity of a sample relative to a control sample at three times during the 25-min run. At each of the three reading times, the light output of each sample and each control is measured on a chart recorder and recorded as the height of the peak light output on a scale of 0 to 100.

P. phosphoreum emit light as a by-product of respiration. If a sample contains one or more components that interfere with respiration, then the bacteria’s light output is reduced proportional to the amount of interference with respiration, or toxicity. The light output reduction is proportional to the toxicity of the sample. The relative toxicity of a sample to the control can then be calculated. These relative toxicities can be compared to toxicity test results using standard reagents specified by this procedure.

For samples that are calculated to be more than 50% toxic, an EC50 concentration is calculated. The EC50 concentration is the fraction of sample, using the Microtox diluent as the dilution solution, that causes a light output from the sample that is 50% of the light output of the control. It is also called the 50% effective concentration.

Sample Handling and Preservation

Glass sample containers must be clean and free of soap residues, and stoppers and lids must not be made of cork. Detergents, cork, and other materials may add chemicals to the sample and may add to its toxicity.

Tap water and distilled water are fatal to the bacteria due to high levels of chlorine. Sample storage containers must be rinsed with deionized or ultra-pure water prior to use, with ultra-pure water being preferable.

Samples should be analyzed soon after arrival at the laboratory. Until they are analyzed, samples should be stored at 4°C. Stored samples may be kept up to 1 week in the refrigerator. Freshwater samples should not be salted until the samples are ready to be analyzed, as salt–metal complexes seem to readily form, reducing the toxicity of the sample. Salted samples can only be stored for approximately 15 to 30 min.

Interferences

Samples having pH values outside the range of 6.3 to 7.8 may be toxic to the bacteria. Normally, the pH of the sample is not adjusted because pH may be the parameter causing toxicity in a natural environment. Color and turbidity will interfere with, and probably reduce, the amount of emitted light leaving the cuvette and reaching the photomultiplier. Organic matter may provide a second food source for the bacteria and may result in a sample whose relative toxicity is calculated to be less than zero.


Apparatus

• Microtox 2055 Analyzer • 500 µL pipettor (with disposable tips) • 10 µL pipettor (with disposable tips) • Glass cuvettes (disposable)

Reagents

• Microtox bacterial reagent • Microtox reconstitution solution • Microtox diluent • Microtox osmotic adjusting solution • Reagent grade sodium chloride

Procedure

Sampling, Sample Preparation

Note: The older Microtox 2055 instrument has space in its incubator for 15 cuvettes. We label these positions with letters for each of the three rows (A, B, and C) and label the five columns with numbers (1 to 5), giving each position a letter and number, such as A1 for the first position and C5 for the last position. For a normal run, three of the cuvettes (A1, B1, and C1) are reserved for the control solution. One of the remaining 12 cuvettes is reserved for the standard solution whose concentration is approximately the predetermined ZnSO4·7H2O EC50 concentration. The remaining 11 cuvettes contain the samples to be tested using this screening procedure.

1. Rinse clean 40-mL sample vials, vial caps, and Teflon septa with ultra-pure water. 2. Mix the sample by inverting the container several times. 3. Pour 10 mL of sample into the vial. 4. Add 0.2 g NaCl (reagent grade) to the vial. 5. Mix the sample and salt by inverting the vial until the salt is completely dissolved.

Preparation of Apparatus

1. Discard the cuvettes remaining in the incubator and pre-cool slots from any prior run (used cuvettes are normally left in the incubator to reduce condensation problems).

2. Put new cuvettes into the 15 slots in the incubator and one in the pre-cool slot. 3. Pipette 1.0 mL of diluent into the cuvettes in positions A1, B1, and C1. 4. Pipette 1.0 mL of reconstitution solution into a cuvette in the pre-cool position. 5. Pipette 1.0 mL of each sample (already adjusted for salinity, as specified above) into separate

cuvettes in positions A2 through A5, B2 through B5, or C2 through C5. 6. Set the timer for 5 min to allow for temperature stabilization of the reconstitution solution. 7. Get a vial of the Microtox reagent bacteria out of the freezer. (Must be stored in a freezer at no

warmer than –20°C.) 8. Tap the reagent vial on the countertop gently several times to break up the contents. 9. After the 5 min temperature stabilization period has expired, open the vial.

10. Quickly, pour the reconstitution solution in the pre-cool slot into the reagent vial. 11. Swirl the contents to mix (all solid reagent should go into solution). 12. Pour the reagent solution back into the pre-cool cuvette. 13. Mix the reagent solution approximately 20 times with a 500 µL pipette. 14. Set the timer for 15 min.

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Analysis of Samples

1. Pipette 10 µL of reagent solution into each cuvette in the following order: A1, B1, C1, A2 through A5, B2 through B5, and C2 through C5. Do not immerse the pipette tip in the solutions.

2. Gently mix each cuvette’s contents 20 times with a 500 µL pipette. Mix the cuvettes in the same order in which reagent solution was added. Use a single pipette tip for the three controls, but a new tip for each sample and the standard.

3. Push in the “HV” and “HV Check” buttons on the front of the Microtox analyzer. The panel on the front should read between –700 and –800.

4. Push in the “HV Check” button (so it toggles back out) and push in the “Sensitivity X10” and “Run” buttons.

5. Turn on the strip chart recorder. 6. Zero the chart recorder using the knob located on the right side of the machine. 7. Make sure the speed setting is for 1 in/min. 8. Make sure the pen is touching the recorder paper by putting the pen arm down. 9. Place the cuvette in A1 into the turret and close the turret to get a reading on A1.

10. After the reading is obtained, remove the cuvette from the turret. 11. Read the cuvettes in B1 and C1 also to determine which of the three has the largest reading. Place

that cuvette back in the turret and close. 12. Adjust the chart reading to between 90 and 100 using the Scan knob on the front of the analyzer.

If display reads “1” (not “001”), change the sensitivity setting to “Sensitivity X1.” 13. Open the turret and check the zero point again on the chart recorder. Adjust as necessary. 14. Close the turret. 15. Set the timer for 5 min. 16. When the timer rings, read the samples in the following order: A1, B1, C1, A1 through A5,

B1 through B5, C1 through C5, A1, B1, and C1. 17. Place the control cuvette (A1, B1, or C1) which has the highest reading in the turret and close. 18. Set the timer for 10 min. 19. When the timer rings, read the samples in the following order: A1, B1, C1, A1 through A5,

B1 through B5, C1 through C5, A1, B1 and C1. 20. Place the control cuvette (A1, B1, or C1) which has the highest reading in the turret and close. 21. Set the timer for 10 min. 22. When the timer rings, read the samples in the following order: A1, B1, C1, A1 through A5,

B1 through B5, C1 through C5, A1, B1 and C1. 23. Shut off the chart recorder and cap the pen. 24. Return the C1 cuvette to the incubator and close the turret. 25. Push in the “HV” and “Turret” buttons on the front of the analyzer (toggle them off). 26. If, at the end of the test, the light output of any sample is less than half of the light output of the

controls, the EC50 concentration of that sample must be found. This is done by rerunning the Microtox test using three to four dilutions of that sample (including one at 100% strength). The previously prepared (salted) sample cannot be used either to create the dilutions or as the 100% strength sample.

Calculations

At each of the three times that a sample is read, each of the three control samples is read three times. The results of these nine analyses are averaged and their standard deviation and coefficient of variation calculated. If the coefficient of variation for the control samples at any time in the run is greater than 0.05 (5%), the run is rejected.

Relative toxicity is calculated as follows:

%Reduction [at time t] = Control – Sample- × 100%

Control


where Control = average peak height of the control samples at t Sample = peak height of sample at t

Precision and Accuracy

The Microtox Analyzer is calibrated using solutions of either zinc sulfate or phenol. A standard solution of approximately 10 mg/L zinc sulfate or of approximately 50 mg/L phenol is made. Four dilutions of the standard solution, with three replicates of each dilution, are used in place of the 12 samples in the normal Microtox screening procedure. The four dilutions should bracket the expected EC50 concentration of the standard solution. However, instead of using sodium chloride to adjust the ionic strength of the sample, the Microtox osmotic adjusting solution (MOAS) should be used. The amount of MOAS used should be 10% of the volume of the standard.

During each run, one of the 12 sample positions is occupied by the standard solution at the EC50 concentration. If the relative toxicity of the standard sample is outside the range of 45 to 55%, the run is rejected and repeated with freshly made standard solution. If the EC50 on the repeat again falls outside the range of 45 to 55%, the calibration is repeated. If the calibrated EC50 is significantly higher than the previous calibrations on that box of reagent, then a new box of reagent is opened and the calibration screening procedure is performed on one of the reagents in that box.

Extensive work has been done to establish the precision and accuracy of this procedure. Please refer to A. Ayyoubi, Physical Treatment of Urban Stormwater Runoff Toxicants, pp. 11–23.

Health and Safety Information

Refer to the MSDSs for information regarding the use of the reagents in this procedure. None of the reagents and materials has OSHA PEL(s), AGGIH TLV(s), or other limits. Oral

rat LD50 data have not been established for any of the reagents supplied by Microtox. Sodium chloride, which is one of the reagents and is a component of most of the reagents

supplied by Microtox, has an LD50 of 3000 mg/kg. The sodium chloride, either as a reagent or as a component of the other reagents, may cause eye irritation, and ingestion of large quantities may cause vomiting, diarrhea, and dehydration.

No special storage requirements are needed beyond keeping the freeze-dried bacteria culture in a freezer. Reagents are not considered to be a fire or explosion hazard (water may be used to extinguish a fire), and have no hazardous decomposition products. The reagents are stable under ordinary conditions of use and storage. Spilled reagent, whether reacted or not, may be cleaned up by adsorption with paper towels, and excess fluid may be flushed down a regular sewer drain.

REFERENCES

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Ayyoubi, A. Physical Treatment of Urban Stormwater Runoff Toxicants, Master’s thesis. University of Alabama at Birmingham, Birmingham, AL. 1993.

Brooker, J. Use of the Asian Clam and Mayflies as in Situ Toxicity Indicators. M.S. thesis. Wright State University, Dayton, OH. 2000.

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Burton, G.A., Jr. Assessing freshwater sediment toxicity. Environ. Toxicol. Chem., 10: 1585–1627. 1991. Burton G.A., C.G. Ingersoll, L.C. Burnett, M. Henry, M.L. Hinman, S.J. Klaine, P.F. Landrum, P. Ross, and

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Burton G.A., T.J. Norberg-King, C.G. Ingersoll, G.T. Ankley, P.V. Winger, J. Kubitz, J.M. Lazorchak, M.E. Smith, I.E. Greer, F.J. Dwyer, D.J. Call, K.E. Day, P. Kennedy, and M. Stinson. Interlaboratory study of precision: Hyalella azteca and Chironomus tentans freshwater sediment toxicity assay. Environ. Toxicol. Chem., 15: 1335–1343, 1996b.

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EPA. Methods for Aquatic Toxicity Identification Evaluations. Phase 1, Toxicity Characterization Procedures, 2nd edition. EPA/600/6-91/003. Office of Research and Development, Washington, D.C. 1991a.

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