EPA Report Collection
[ional Center for Environmental Information
Ill
19103
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U.S. EPA Region III
Regional Center for Environmental
Information
1650 Arch Street (3PM52)
Philadelphia, PA 19103
Standard Operating Procedures
for Conducting Acute and Chronic
Aquatic Toxicity Tests
with Eurytemora a/finis,
a Calanoid Copepod
October 1998
University of Maryland
Agricultural Experiment Station
Wye Research and Education Center
o
Regional Center tor Emironnu'lital Information
US EPA Region HI
16SOAichSl.
Philadelphia. PA 19W
Chesapeake Bay Program
410 Severn Avenue, Suite 109
Annapolis, Maryland 21403
1-800-YOUR-BAY
http://www.chesapeakebay.net/bayprogram
Printed by the U.S. Environmental Protection Agency for the Chesapeake Bay Program
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October 1998
Standard Operating Procedures
For Conducting Acute and Chronic
Aquatic Toxicity Tests
with Eurytemora affinis, a Calanoid Copepod
Michael C. Ziegenfuss
and
Lenwood W. Hall, Jr.
University of Maryland
Maryland Agricultural Experiment Station
Wye Research and Education Center
Queenstown, Maryland 21658
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FOREWORD
There are few estuarine toxicity test protocols currently available for
Chesapeake Bay resident aquatic species. The calanoid copepod, Eurytemora
affinis, was recommended as a test species for Standard Operating Procedures
development based on an extensive literature review and synthesis of data from 25
candidate species found in the Chesapeake Bay. This manual outlines standard
operating procedures for conducting acute and chronic toxicity tests with
Eurytemora. The U.S. Environmental Protection Agency and Maryland Department
of the Environment provided the funding for the development of this manual.
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TABLE OF CONTENTS
FOREWORD i
1.0 INTRODUCTION 1
2.0 DISTRIBUTION 1
3.0 LIFE CYCLE 2
4.0 TAXONOMY 2
5.0 TERMINOLOGY 4
6.0 SUMMARY OF TEST PROCEDURES 5
7.0 INTERFERENCES 5
8.0 HEALTH AND SAFETY 5
8.1 GENERAL PRECAUTIONS 5
8.2 SAFETY EQUIPMENT 6
8.3 GENERAL LABORATORY OPERATION 6
9.0 QUALITY ASSURANCE 6
9.1 INTRODUCTION 6
9.2 FACILITIES AND EQUIPMENT 7
9.3 TEST ORGANISMS 7
9.4 CULTURE AND DILUTION WATER 7
9.5 TEST SUBSTANCE HANDLING 7
9.6 TEST CONDITIONS 7
9.7 ANALYTICAL METHODS 8
9.8 CALIBRATION AND STANDARDIZATION 8
9.9 ACCEPTABILITY OF TOXICITY TEST RESULTS 8
9.10 REFERENCE TOXICANTS 8
9.11 RECORD KEEPING 8
10.0 APPARATUS, EQUIPMENT, AND MATERIALS 9
10.1 FACILITIES 9
10.2 CONSTRUCTION MATERIALS 9
1 0.3 MATERIALS FOR CULTURING AND TESTING 11
1 0.4 TEST CHAMBERS 12
10.5 CLEANING 12
11.0 CONTROL AND DILUTION WATER 14
12.0 TEST SUBSTANCE SAMPLING, RECEIVING, HANDLING,
AND STORAGE 14
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13.0 ORGANISM CULTURE PROCEDURES 14
13.1 INTRODUCTION 14
13.2 CULTURE WATER 16
13.3 CULTURE VESSELS 16
13.4 SALINITY, TEMPERATURE, AND PHOTOPERIOD 16
1 3.5 RENEWAL OF CULTURE WATER 17
13.6 FOOD AND FEEDING REGIME 17
13.7 PHYTOPLANKTON CULTURE 17
13.8 CULTURE RECORDS 18
14.0 TOXICITY TEST PROCEDURES 20
14.1 EXPERIMENTAL DESIGN 20
14.2 RANGE-FINDING TEST 20
14.3 DEFINITIVE TEST 20
14.4 TEST SOLUTIONS 22
14.5 OBTAINING NEONATES FOR TOXICITY TESTS 22
14.6 STARTING THE TEST 22
14.7 TEMPERATURE AND PHOTOPERIOD 23
14.8 FEEDING 23
14.9 PHYSICAL AND CHEMICAL ANALYSIS 25
14.10 TERMINATING THE TEST 25
15.0 DATA ANALYSIS 29
16.0 REFERENCES 31
APPENDICES 35
A Summary of Eurytemora affinis control survival during
acute and chronic toxicity testing over a four-year period.
B Summary of Eurytemora affinis sublethal endpoints
(reproduction and growth) evaluated under control
conditions from 8 day tests.
C Single laboratory precision of the 24 Eurytemora affinis
estuarine toxicity test at 1 5 ppt salinity with the reference
toxicant cadmium chloride.
D Relative sensitivity of Eurytemora affinis to selected toxic
substances.
in
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1.0 INTRODUCTION
Few standardized estuarine toxicity test protocols are currently available for
Chesapeake Bay resident aquatic species. The widespread availability of such
protocols would be an asset in the region's ambient toxicity testing, estuarine
criteria development (or other single chemical tests), and effluent toxicity testing
efforts which are currently limited in scope due to the availability of standardized
protocols for only a few test species. This lack of protocols may be preventing the
inclusion of locally important and ecologically key species, which could lead to
erroneous ambient toxicity assessments, development of criteria which are not
protective of the Chesapeake Bay aquatic biota, or inaccurate assessments of
potential harm of effluents to receiving water biota due to the use of species which
may be much more tolerant (or sensitive) to toxicants than species resident to the
receiving waters of concern.
The goal of the "Chesapeake Bay Basinwide Toxics Reduction Strategy" is to
reduce input of toxic substances to levels which do not result in toxic impact on
Bay living resources (Chesapeake Executive Council, 1989). Therefore, it is
necessary to assess accurately the effect of toxic substances on Bay biota.
Development of standardized toxicity testing protocols for key Chesapeake Bay
species will provide the mechanism for enhancing the reliability of toxicity
assessments and ultimately, the protection of Chesapeake Bay aquatic organisms
from impact of toxic substances.
Eurytemora affinis, a calanoid copepod, was selected for standard toxicity
testing protocol development subsequent to screening 25 resident Chesapeake Bay
species including fish, invertebrates, and plants (Ziegenfuss and Hall, 1993).
Eurytemora was selected because of its ecological importance as an essential
component in the trophic structure of the estuary, its relative practicability of
culturing in the laboratory for year-round availability, and its sensitivity to toxic
substances. The standard operating procedures described in this document provide
detailed procedures for culturing, holding, and toxicity testing of E. affinis.
2.0 DISTRIBUTION
Copepods of the genus Eurytemora are distributed worldwide depending on
suitable temperature and salinity. This species tolerates a wide salinity range (0-33
ppt), however the optimum range is 5-15 ppt (Jeffries, 1962). Eurytemora also
tolerate a broad temperature range (2.5-35 °C) with a resistance to extremes
correlated with salinity (Bradley, 1975). Its habitat in North America as described
by Wilson and Yeatman (1959) includes lakes, ponds, and fresh and brackish
waters of the Atlantic, Pacific and Gulf coasts. The east coast distribution extends
from Miramichi Estuary, Canada to the Florida Keys. The west coast range for £.
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affinis includes Vancouver, British Columbia to San Francisco Bay. Eurytemora is a
year-round resident dominating the zooplankton community in winter and early
spring in Chesapeake Bay (Tepper, 1986). It is confined to lower saline upper
reaches of Bay tributaries during the warm summer months.
3.0 LIFE CYCLE
Eurytemora affinis is obligate sexual in its reproduction (Figure 1 from
Gurney, 1931). The antennas and modified fifth legs of the male are used to clasp
the female. During clasping, the male transfers sperm to the female in
spermatophores usually with the aid of the legs. Sperm is stored in a ventral area
of the female genital segment that serves as a seminal receptacle. Fertilization
occurs as the eggs leave the female reproductive tract. Fertilized eggs are retained
by the female in ovisacs containing 5 to 40 eggs located ventrally on the genital
segment. Development of the eggs is temperature-dependant and can range from
> 12 days at 5 °C to approximately 1 day at 25 °C. Growth and time to sexual
maturity are temperature and food-dependent. During development, the molting
process includes six nauplii stages and six copepodid instars, the last of which is
the adult. Generation time may vary from about 105 days at 2 °C to nine days at
23.5 °C (Katona, 1970).
4.0 TAXONOMY
The following section describes anatomical features of female and male
Eurytemora affinis (Mori, 1964). The female metasome has five segments with the
fifth segment expanded to form wing-like structures. The urosome has three
segments and is symmetrical. The anal segment and caudal rami are densely
covered with spine-like hairs on the dorsal side. Fifth legs are symmetrical and the
terminal segment is small, globular and contains two spines. One spine is long and
located on the terminal segment and the second spine is short and located on the
outer distal corner. The average length of females is about 1.42 mm.
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Figure 1. Diagrams of male and female E. affinis. A - male showing urosome with
five segments; B - right first antenna of male (right antenna has asymmetric apering
from proximal to distal end unlike left antenna of male and both first antennae of
female); C - female (ovisac of gravid female is carried on ventral sice of 3
segmented urosome); D - dorsal view of female showing wing-like structures on
last segment of metasome.
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The first right anterior antenna of the male is modified into a grasping organ.
The metasome has five segments with the last segment rounded posteriorly. The
urosome has five segments and is symmetrical. The anal segment has many spine-
like hairs. Caudal rami also have a row of fine hairs located on their inner border.
Rami setae are well developed. Fifth legs are asymmetrical with the right leg being
larger. The right leg has four segments with the terminal segment being fairly large
at the proximal end and tapering to form a blunt point. The left leg has four
segments with the terminal segment being heavier but shorter than that of the right
leg. It ends in a complicated structure. Males are usually smaller than females,
with an average length of 1.15 mm.
5.0 TERMINOLOGY
Lethality (mortality) is the most often measured endpoint in toxicity tests.
Experimentally, the effect on 50% of a group of test organisms is the most
reproducible and easily determined measure of toxicity. Forty-eight and/or 96 hours
are short exposure durations that provide a rapid measure of toxicity. The measure
of acute toxicity is the LC50 (median lethal concentration) expressed as a 48-h or
96-h value. The LC50 is the statistically derived best estimate of the sample
concentration that is lethal to 50% of the test organisms at the end of the
exposure duration.
Chronic toxicity tests for E. affinis require a longer exposure period, 8-d at
25 °C, spanning the post-hatch (naupliar) lifestage to sexual maturity. Measures of
chronic toxicity include mortality and sublethal effects such as growth and
reproduction or fecundity. The chronic toxicity value is derived by calculating the
geometric mean of the No Observed Effect Concentration (NOEC) and the Lowest
Observed Effect Concentration (LOEC).
Acute and chronic tests can be conducted by at least three methods. Static
tests are conducted by placing test solution and organisms in chambers within test
beakers for the duration of the test without renewal of test solution. Static
renewal tests are conducted by periodically (usually every 24-h) replenishing the
test solution with fresh solution of the same composition. Flow-through tests are
conducted by continuously passing fresh test solution over the organisms for the
duration of the exposure. A metering system (serial diluter or fluid metering pump)
controls the flow of sample and dilution water.
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6.0 SUMMARY OF TEST PROCEDURES
Separate groups of nauplii (less than 24-hours old) are exposed to various
treatments of test solution for a specific period of time. A control treatment is
used to provide a measure of acceptability of the test by providing information
relative to health or quality of test organisms, the suitability of dilution water,
testing conditions (temperature, light regime, etc.) and handling procedures.
Control water consists of contaminant-free estuarine water adjusted to a desired
salinity matching the test solution. Test treatments consist of undiluted sample or
a series of dilutions (or concentrations) in geometric progression.
The acute toxicity test is 48 or 96 hours in duration and provides mortality
data at the end of the exposure period. The chronic test has an 8-day exposure
period (at 25 °C) at which time mortality is determined. In addition to lethality,
reproduction (fecundity) and growth (maturation) are evaluated.
7.0 INTERFERENCES
Toxic substances may be introduced by contaminants in dilution water and
testing apparatus. Improper collection and handling of test sample solutions may
also adversely affect test results. Adverse effects of low dissolved oxygen and
extreme of pH may mask the presence of toxic substances. Pathogenic and/or
predatory organisms in the dilution water or test water may affect test organism
survival. Inadvertent introduction of copepods from test water (ambient estuarine
water) into test chambers will also confound test results. Food (phytoplankton)
added during the test, i.e., renewal of ambient water may sequester toxic
substances and also affect test results.
8.0 HEALTH AND SAFETY
8.1 General Precautions
Conducting toxicity tests may involve differing levels of risk. Effluents are
accepted for testing only with a copy of the chemical composition shown in the
NPDES discharge permit (preferably prior to delivery of sample). This allows
adequate safety precautions to be taken to prevent injury. Personnel conducting
tests protect themselves by taking all safety precautions necessary to avoid
inhalation or absorption of toxic substances through the skin and to prevent
asphyxiation due to lack of oxygen or presence of volatile noxious substances.
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8.2 Safety Equipment
Personnel use safety equipment, as required, such as disposable rubber
gloves, lab coats and/or aprons, respirators, and safety glasses. Laboratory safety
equipment includes a proper ventilation system, first aid kits, fire extinguishers and
blanket, and an emergency eye wash and shower unit.
8.3 General Laboratory Operation
Work with samples containing suspected toxic substances is performed in
compliance with accepted rules pertaining to the handling of hazardous materials.
Toxicity tests with volatile compounds are conducted under a ventilation hood.
Because the chemical composition and toxicity of samples are usually poorly
understood, samples are considered potential health hazards and exposure to them
is minimized.
Toxicity testing and organism culture maintenance are conducted in separate
defined areas of the laboratory. Containers used in the laboratory are always
labeled to indicate their contents and prevent contamination. The laboratory is
generally kept clean and orderly to promote safety and reliable test results.
Guidance on safe practices when collecting samples and conducting toxicity tests is
frequently obtained from the permittee, in the case of discharge effluents, and
general industrial safety manuals including U.S. EPA (1977) and Waters and
Jameson (1984).
9.0 QUALITY ASSURANCE
9.1 Introduction
The following quality assurance (QA) section is adapted from U.S. EPA
(1 991 a). Quality assurance practices for conducting toxicity tests with Eurytemora
should address all aspects that affect the integrity of the final data, such as: (1)
test substance sampling, handling, and storage; (2) quality of dilution water; (3)
condition of test organisms; (4) condition and operation of laboratory equipment;
(5) test conditions; (6) instrument calibration; (7) replication; (8) use of reference
toxicants; (9) recording data and observations; and (10) data evaluation. For more
information on quality assurance and good laboratory practices related to toxicity
testing see: FDA (1978), U.S. EPA (1975, 1979a, 1980a, 1980b, 1991b),
Dewoskin (1984), and Taylor (1987).
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9.2 Facilities and Equipment
Separate culture and toxicity test areas are necessary to avoid possible cross
contamination which could result in the loss of cultures. The laboratory should be
equipped with a ventilation system to prevent recirculation of contaminated air from
testing areas, sample preparation and storage areas, and chemical analysis areas.
In addition to space and ventilation requirements, temperature control equipment
must be capable of maintaining test temperatures with minimal variation,
programmable lighting is required to simulate day-night conditions, and an oil-free
mechanical air supply is needed for both toxicity testing and culture areas.
9.3 Test Organisms
Test organisms must be identified to species. The organisms used in toxicity
testing experiments must appear healthy, vigorous, and have low mortality in
cultures, during holding, and in test controls. Copepods should be cultured at the
same (within 3 ppt) salinity as the test salinity. All copepods in a test should be
the same lifestage (less than 24-hours). In most instances, tests are initiated with
nauplii (24-hours old or less) because this lifestage is generally considered to be
most sensitive.
9.4 Culture and Dilution Water
Water used for culturing and testing purposes should be from the same
source. The water should be tested for toxic contaminants (metals and organics) at
least once per year.
9.5 Test Substance Handling
Procedures for sampling, receiving, handling, and storage should conform to
the conditions described in section 12, TEST SUBSTANCE SAMPLING, RECEIVING,
HANDLING, AND STORAGE.
9.6 Test Conditions
The temperature of the test solution should be measured by placing a
thermometer or probe directly into the test solution or a surrogate beaker containing
the same volume of solution as the test beaker. Dissolved oxygen concentration,
pH, and salinity must be measured in the actual test solutions. Test condition
parameters should be measured at least initially and at the end of the exposure
duration.
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9.7 Analytical Methods
All routine chemical and physical analysis (T, DO, Sal, pH, etc.) of culture
and dilution water, and test solutions are performed as outlined in U.S.EPA (1979a,
b). Reagent containers, chemical stock solutions, and working solutions are dated
when received from the supplier and shelf life is not exceeded.
9.8 Calibration and Standardization
Instruments used for routine chemical and physical parameter measurements
are calibrated prior to use according to the instrument manufacturer's procedures.
9.9 Acceptability of Toxicity Test Results
Control survival data from 96-h and 8-d tests (and a few 48 h tests)
conducted during a four-year period are presented in Appendix A. The mean
survival for the 96 h and 8 d tests ranged from 82 - 86 %. Based on these data,
control survival (mean %) for E. affinis should equal or exceed 80% in acute or
chronic tests. A summary of the reproduction and growth endpoints from control
conditions are presented in Appendix B. Due to the variability with these
endpoints, an acceptable value cannot be recommended. Within test comparisons
of control and test conditions are used to determine statistical differences.
9.10 Reference Toxicants
Reference toxicants are used to establish the validity of toxicity data
generated from toxicity tests. The reference toxicants provide information
regarding the relative health and sensitivity of the copepods used in toxicity testing.
Several toxicants, CdCI2 (cadmium chloride), CuS04 (copper sulfate), and SDS
(sodium dodecyl sulfate), are available for use. Data generated in our laboratory
with cadmium chloride are presented in Appendix C. A control chart (Figure 2) is
prepared for each reference toxicant and LC50 values are plotted to determine if
results are within expected limits (U.S. EPA, 1991 a). If the LC50 does not fall in
the expected range, then the sensitivity of the copepods are
suspect. The sensitivity of E. affinis to various other toxic chemicals is presented
in Appendix D.
9.11 Record Keeping
Proper record keeping is very important. Bound notebooks are used to
maintain detailed records of culture maintenance, equipment maintenance and
calibration, receipt and storage of test solutions, test conditions employed, and
results. Annotations are made in ink to prevent the loss of information. All data
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from the toxicity tests are kept in either bound notebooks or bioassay data sheets.
10.0 APPARATUS, EQUIPMENT, AND MATERIALS
10.1 Facilities
The bioassay laboratory should consist of separate and defined toxicity
testing and organism culture areas. The laboratory should be equipped with an oil-
free air supply, programmable lighting for day-night simulation, and controlled
temperature. Biological incubators can be used for conducting toxicological tests to
increase precision in temperature. A water treatment system, such as Millipore
Milli-Q, Super-Q, or equivalent, is required to deliver contaminant-free freshwater.
Water supply lines should be constructed from PVC or other non-toxic plastic.
10.2 Construction Materials
Glass, polycarbonate, Nitex screen, and 100% clear silicon adhesive are
used for construction of equipment that comes in contact with copepods, culture
water, and test solutions.
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Figure 2. Control chart and calculations for reference toxicant tests
UPPER CONTROL LIMIT (X +20)
CENTRAL TENDENCY
LC50
LOWER CONTROL LIMIT (X-2o)
1 1 T~
I
0 5 10 15 20
TOXICITY TEST WITH REFERENCE TOXICANTS
In
(n-l)
X. = Successive LC50 s from toxicity tests
n = Number of tests
X =MeanLC50
a = Standard deviation
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10.3 Materials For Culturing and Testing
Eurytemora affinis can be obtained from cultures at the University of
Maryland's Wye Research and Education Center, Queenstown, Maryland, or the
University of Maryland's Chesapeake Biological Laboratory, Solomons, Maryland.
A detailed description of culturing and testing material is presented below:
• Algal cultures, Isochrysis galbana, Thalassiosira pseudonana and T.
f/uviati/is, can be obtained from commercial sources.
• Glass aquaria, 10-18 I capacity, are used for culturing copepods.
• Glass Fernbach flasks, 2.8 I capacity, are used for culturing algae for
feeding copepods.
• Non-toxic plastic buckets, 7 I capacity, are used for mixing culture and
dilution water to the desired salinity.
• Airline tubing, airstones, disposable sterile plugged 1.0 ml pipettes,
and gang valves are required for aeration of cultures.
• Polycarbonate carboys, 20 I capacity, are used for holding sterile
culture water.
• Autoclave or filter apparatus are used for sterilizing water for algal
cultures.
• Nitex screens (mesh sizes 202 and 53/;m, 250ml), polycarbonate jars,
and wash bottles are used for sorting and containing copepods.
• Borosilicate glass wide-bore pipettes with fire-polished ends and bulbs
are used for transferring copepods.
• Glass beakers, 150 - 250ml capacity, are used as test vessels during
toxicity tests.
• Polycarbonate test chambers (Section 10.4) and plastic-covered
paperclips are used during testing.
• A dissecting microscope is required for counting copepods and a
compound microscope is used for counting algae.
• A Spencer improved Neubauer corpuscle counting chamber, or
equivalent, is needed for determining algal densities.
• Refractometer, pH meter, dissolved oxygen meter, and thermometer
are required for measuring routine physical and chemical parameters in
culture water and test solutions.
• Adjustable pipettes, 0.2, 1.0, and 5.0ml with disposable tips are used
for mixing test solutions, and adding reagents to algal growth medium.
• Waterproof markers are used for labeling containers in the laboratory.
• Reagents needed for algal growth medium, as described by Guillard
(1975), include NaNO3, NaH2PO4, Na2Si03, Na2 EDTA, FeCI3, CuSO4,
ZnS04, CoCI2, MnCI2, NaMo04, Thiamine-HCI, Biotin, and vitamin B12.
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• Synthetic seasalts or hypersaline brine are required for salinity
adjustments of culture water and test solutions.
• Reagents needed for routine physical and chemical water quality
parameters include pH calibration buffers (4, 7, and 10) and electrode
filling and storage solutions, dissolved oxygen probe membranes and
filling solution, and salinity standards for refractometer calibration.
• Laboratory glassware required for preparation of standard, chemical
stocks, test solutions and dilutions include beakers (1 50 ml - 2.0 I),
volumetric flasks, and graduated cylinders.
• Reference toxicant solutions can be prepared in-house by obtaining
reagents (see section 9.10) from a commercial supplier.
• Neutral buffered formalin (10%) or ethanol is used to preserve adult
copepods at the end of a chronic test.
• Scintillation vials (20ml) are used to contain preserved copepods at the
termination of a chronic test.
10.4 Test Chambers
Test chambers are constructed from rigid polycarbonate tube (6.35cm height
x 3.8cm diameter). Silicon adhesive is used in minimal amounts to attach 53//m
mesh Nitex screen to the bottom of the chamber. Subsequently, all test chambers
are leached for 24 hours in contaminant-free freshwater prior to use in toxicity
experiments. Test chambers are suspended within 150 - 250 ml glass beakers
with plastic-covered paper clips (Figure 3.)
10.5 Cleaning
All glassware used to prepare stock solutions and test solutions, and contain
organisms during toxicity tests are cleaned before use according to procedures
outlined by the U.S. EPA (1985) and ASTM (1980). Briefly, glassware is washed
with detergent and rinsed with tap water, 10% nitric or hydrochloric acid,
deionized water, pesticide-free acetone, followed by a minimum of three rinses with
de-ionized water. Polycarbonate chambers are cleaned following similar procedures
except that methanol is substituted for acetone.
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Figure 3. Beaker and test chamber used for toxicity tests.
150 ml Beaker
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11.0 CONTROL AND DILUTION WATER
Natural estuarine water is preferred over synthetic seawater for use as
culture, control, and dilution water. Natural estuarine water contains necessary
trace metals, biogenic colloids, and some of the microbial components necessary
for growth, survival, and reproduction of estuarine organisms. Salinity adjustments
are frequently necessary to match the salinity of receiving waters during effluent
testing, standardizing salinities for ambient toxicity testing, or determining the
effect of salinity on single or multiple chemicals. The salinity of the natural water
can be increased with hypersaline brine (100 ppt) or a good-quality commercial
seasalt such as HW Marine Mix (Hawaiian Marine Imports, Inc., Houston, TX). De-
ionized water is used to reduce the salinity of natural water. The estuarine water
should be filtered (0.4um) or autoclaved prior to use to eliminate the introduction of
predators, feral copepods, and undesirable phytoplankton into the cultures and test
conditions.
12.0 TEST SUBSTANCE SAMPLING, RECEIVING, HANDLING, AND STORAGE
See U.S. EPA (1991 a) section 8 for a detailed discussion of effluent and
receiving water sampling and handling. The decision to collect grab or composite
samples is based on the objectives of the study. Collection of composite samples
should not exceed 24-h so that dilution of toxicity spikes is minimized. Sample
containers should be constructed from non-toxic plastic or glass which has been
appropriately cleaned (Section 10.5). Samples should be shipped on ice and stored
at 4 C to minimize microbial action, chemical reaction, and volatilization. A chain
of custody form must accompany all samples (Figure 4). Sample holding time, the
time elapsed between the collection of the sample and the initiation of the toxicity
test, should not exceed 36 hours. Health and safety issues addressed in Section
8.3 should be followed when handling the samples.
13.0 ORGANISM CULTURE PROCEDURES
13.1 Introduction
Techniques for rearing Eurytemora affinis in the laboratory have been
described by Heinle (1969), Katona (1970), and Hall et al. (1991). Brood stock are
most easily obtained with plankton nets in the spring when Eurytemora dominate
the zooplankton community in Chesapeake Bay. During summer months,
Eurytemora can be obtained in lower saline water in tributaries of the Bay.
Continuous cultures of Eurytemora affinis require minimal space, water, and
equipment. This species is relatively easy to culture and requires minimal
maintenance.
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FIGURE 4. CHAIN OF CUSTODY FORM FOR RECEIVING AND HANDLINGTEST SUBJECTS
Project/Facility Name
Permit No.
Sample I.D. No.
Outfall No.
Collected by
Sample Type:
Grab Collected
Grab Composite
Time Date
/ /
Time Date
/ /
Time Date
Total No. Grab Samples:
Sampling Interval:
Automatic Composite
Collected
Sample Cooled
from:
to:
Time Date
/ /
Time Date
During Collection Yes No
During Delivery Yes No
Upon Receipt Yes No
Name_
Name_
Name
On-site Physical Chemistry Measurement
Conducted by:
Date/Time:
PH ~
Dissolved Oxygen, mg/l
Temperature, °C
Salinity, ppt
Conductivity, /c/mhos
Comments
Sample Possession
From:
From:
Name, Date, Time
Name Date, Time
To:
To:
Name, Date, Time
Name, Date, Time
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In addition to copepod cultures,monocultures of microalgaa must be
maintained as a source of food for the copepods. Laboratory cultures provide year-
round availability of copepods for toxicity testing. The generation time of
Eurytemora is short (approximately 8-d at 25 °C), therefore high density cultures
can be achieved in relatively little time. Established cultures contain all
developmental stages (egg-adult). The number of animals in each culture is
maintained at approximately 100 per liter of culture medium.
13.2 Culture Water
Natural estuarine water is preferred over synthetic saltwater for culturing
Eurytemora. Natural estuarine water used for the maintenance of stock cultures
contains essential components necessary for healthy copepods, however natural
water may also contain unwanted microalgae and predators. Estuarine water
should be filtered (0.4//m) or autoclaved prior to use to eliminate chances of biotic
contamination. Salinity adjustments can be made with deionized water, hypersaline
brine, or a good quality synthetic sea salt such as H-W Marinemix (Hawaiian Marine
Imports, Inc., Houston, TX).
13.3 Culture Vessels
Copepods are cultured in 18-L glass aquaria. Larger or smaller culture
volumes can be maintained to accommodate individual laboratory needs or
constraints. A minimum of two cultures are normally maintained to provide
adequate numbers of copepods for testing purposes and provide back-up security in
the event that one culture "crashes". Cultures are kept covered to prevent
accidental contamination from spills and also to minimize evaporation which results
in an increase in culture salinity. Minimal aeration is supplied using 3/16" airline
connected to a 1.0 mL glass pipett.
13.4 Salinity, Temperature, and Photoperiod
Eurytemora affinis tolerate a wide salinity and temperature range. Katona
(1 970) demonstrated the ability of Eurytemora to reproduce in temperatures
ranging from 2 to 23 °C and salinities from 5 to 33 ppt. The duration of copepod
development is affected by temperature-salinity combinations, as observed by
Nagarai (1 992); however, temperature has a greater influence on development rate.
Increasing temperature (at constant salinity) accelerates developmental rate but
reduces longevity. For toxicity testing purposes, cultures are maintained at 25 °C
and 15 ppt salinity. By culturing at 25 °C and 15 ppt, copepods are acclimated to
a standard test temperature and salinity representative of the estuarine
environment. Salinity adjustment can be achieved by using a rate increase or
16
-------
decrease of 2-3 ppt/day. Cultures are protected from direct light with screens or
tinted covers. Artificial lighting is controlled to provide a photoperiod of 16 hours
light: 8 hours dark.
1 3.5 Renewal of Culture Water
Culture water is replenished at least once per month for 18L cultures to
remove biological wastes (dead algal cells and copepod fecal pellets) as well as
adjust the density of copepods. Culture volume is reduced by 75% and renewed
with fresh, filtered or autoclaved, salinity adjusted estuarine water. Water and
copepods are removed from the cultures using a slow siphon. For 18L cultures,
four to five liters of old culture water and copepods are collected and the remaining
water is discarded. Culture aquaria are wiped clean with fresh water (without
detergent) and dried. Fresh culture water and food are added followed by addition
of the previously collected copepods.
13.6 Food and Feeding Regime
Copepods store little energy reserves, therefore food supply has a significant
influence on generation time, reproduction, and mortality (Gaedke, 1990).
Eurytemora feed on phytoplankton, microzooplankton, and detritus. Phytoplankton
is directly or indirectly the most important food source. A two-species microalgal
diet provides adequate nutrition for sustained healthy cultures of Eurytemora aff/nis.
Stock cultures are fed on a diet of Isochrysis galbana (Tahitian) and a species of
centric diatom such as Tha/assiosira fluviati/is or T. pseudonana. A 50/50 (vol/vol)
mixture of the two phytoplankton, each in log-phase growth, is added to copepod
cultures three times per week. Algal density within the copepod cultures is
maintained at approximately 104 cells/ml Isochrysis and 103 cells/mL Thalassiosira.
13.7 Phytoplankton Culture
The algae are cultured using aseptic techniques in order to exclude biotic
contaminants. Monocultures of Isochrysis and Thalassiosira are grown in batch
culture in 2.8L Fernbach flasks. Algal cultures are maintained in a temperature
controlled room (20-25 °C), with fluorescent lighting (16:8 L:D photoperiod), and
vigorous aeration via a sterile cotton-plugged glass pipette. New cultures are
initiated every week with a 250ml_ inoculum, from the previous week's culture,
added to 2L of f/2 enrichment medium (Guillard, 1975). The algal culture medium
consists of autoclaved (filtered and salinity adjusted) estuarine water with nutrients,
trace metals, and vitamins added (Table 1).
17
-------
13.8 Culture Records
Details on the culture history, daily culture maintenance operations, and
phytoplankton cultures are recorded in a laboratory notebook.
18
-------
Table 1. Algal culture medium (modified f/2) from Guillard (1975).
Stock Solutions
NaNO3
NaH2PO4
NH4CI
Na2SiO3
Major Elements
% (W/V)
7.5
0.5
2.65
3.0
(Required for diatom)
% (W/V) n grams brought to a volume of 100 mis with de-ionized water.
Use 1.0 ml per liter of estuarine water to obtain f/2 medium
Trace Elements
Primary Stock Solutions %(W/V)
CuSO4 5H20 0.98
CoCI2 6H2O 1.05
Na2MoO4 2H2O 0.63
ZnS04 7H20 2.2
MnCI2 4H2O 18.0
Working Stock Solution
Dissolve 3.15g FeCI3 and 4.36g Na2EDTA in approximately 900 ml de-ionized
water. Add 1.0 ml of each trace element primary stock and bring to 1.0 L. Use
1.0 ml_ of the working stock for each liter of f/2 medium.
Vitamins
Primary Stock Solutions
Biotin 0.1 mg/ml
B12 1.0 mg/ml
Working Stock Solution
Dissolve 20 mg thiamine HCI in 100 ml de-ionized water and add 1.0 ml biotin
primary stock and 0.1 ml B12 primary stock.
Add 0.5 ml vitamin working stock to each liter f/2 medium
19
-------
14.0 TOXICITY TEST PROCEDURES
14.1 Experimental Design
Eurytemora bioassays are initiated with copepods of the same developmental
stage (within 12-hours in age). Acute tests are started with nauplii or copepedids
and chronic tests are started with nauplii (~ 24-hours old). Four replicates for each
test condition (control and various sample dilutions) are recommended to better
distinguish differences in biological effects between treatments.Each replicate
contains 12-16 organisms which are held in chambers suspended in the test
solution. Tests are conducted as static, static-renewal, or flow-through. Test
organisms are fed daily with a 50/50 algal mixture. Physical and chemical
parameters are measured initially and with each renewal of test solution. At the
end of the exposure duration, the test solution is lowered (removed by slow siphon)
in each replicate and survival is evaluated. Surviving copepods from chronic tests
are preserved in formalin. Subsequently, growth (maturation) and fecundity are
evaluated. A summary of test conditions is listed in Table 2.
14.2 Range-Finding Test
When the toxicity of an effluent, chemical, or environmental sample is
unknown, a range-finding bioassay is performed to determine the concentration
series to be used in a definitive test. Groups of copepods are exposed for 24-h to
at least three dilutions (1, 10, and 50%, and a control) for effluents.
Concentrations should span four orders of magnitude (1, 10, 100, 1000 x/L) for
single or multiple chemicals.
14.3 Definitive Test
Toxicity tests usually consist of one control treatment and a series of
toxicant dilutions (i.e., 100, 56, 32, 18, 10, and 5.6%). The dilution series is
determined by the range-find test. Each dilution, except for the highest
concentration and the control, is at least 50% of the next higher one. The control
treatment consists of the same dilution water, conditions, procedures, and
copepods as are used in the other treatments. For a valid test, average control
survival must equal or exceed 80% in acute and chronic tests.
20
-------
Table 2. Recommended Test Conditions for Eurytemora affinis.
1. Temperature1:
2. Lighting:
3. Photoperiod:
4. Size of Test Vessel:
5. Volume of Test Solution;
6. Age of Test Copepods:
7. No. of Copepods per Test
Vessel:
8. No. of Replicates per
Concentration:
9. Feeding Regime:
10. Aeration:
11. Dilution Water
12. Test Duration
13. Effect Measured
20 or 25 ± 2 °C
100-1 50 fc
16 L:8 D
150 ml Beaker
100ml
= 24 hours
12-16
Daily Algal Mixture
104 cells/ml for I. galbana
103 cells/ml for T. Fluviatilis
None, unless DO concentration
falls below 40% saturation
Autoclaved Natural Estuarine
Water (Salinity Adjusted)
Acute Test - 48 or 96 hours
Chronic Test - 8 days
Mortality, Fecundity, Maturation
25 °C should be used for chronic test.
21
-------
14.4 Test Solutions
The volume of test solution required depends on the dilution series,
frequency of renewals, and duration of exposure. Usually, a one liter sample
provides enough volume to start a bioassay with six dilutions (100 to 5.6%), four
replicates per treatment, and 100 ml per replicate. Physical and chemical
parameters are measured with the remainder of the sample. For each renewal
(50%), an additional 500 ml are required. Tests should begin as soon as possible,
preferably within 24 hours after sample collection. Single chemical stock solutions
should be prepared within 24-48 hours of initiating the test. If the persistence of
toxicity is not known, the maximum holding time of a sample should not exceed 72
hours. Samples are stored at 4 °C. Immediately prior to starting a bioassay, the
sample is warmed to the test temperature (20 or 25 °C) by placing the sample in a
warm water bath.
14.5 Obtaining Neonates For Toxicity Tests
Approximately 24 hours prior to starting a bioassay, adult copepods are
removed from stock cultures. A 202//m Nitex mesh sieve is passed through the
culture numerous times to capture adults. The sieve is removed from the culture
(with adult copepods retained) and gently rinsed with dilution water to collect
adults in a 1-1 beaker containing approximately 750 mL of dilution water. A small
volume (10-20 ml) of the two-species algal mixture is added. The beaker is
covered and placed out of direct light, without aeration, and at the desired test
temperature (20-25 °C).
Within 24 hrs from the time the adult copepods were isolated, nauplii are
collected by slowly siphoning the volume of water and copepods from the 1 -L
beaker through a 202 //m Nitex mesh sieve into another 1-1 beaker. The bottom of
the beaker is not disturbed while siphoning to reduce the amount of algal cells and
fecal pellets in the nauplii collection. The adults are trapped on the mesh while the
neonates pass through the mesh. Entrapped adults are rinsed back into stock
cultures. Nauplii are then concentrated in a 250 ml polycarbonate jar in which the
bottom has been removed and replaced with a 53 //m Nitex mesh.
14.6 Starting the Test
Test beakers (150 ml) are labeled (salinity, concentration, replicate number)
and test chambers are suspended in the beakers with plastic coated paperclips.
Test solution (100 ml) is added to each beaker. Nauplii are transferred to test
chambers by drawing an aliquot of water and nauplii into a fire-polished, wide bore,
glass pipette and counting the number of nauplii in the pipette under a dissecting
microscope (15 x magnification). The nauplii are gently expelled into the test
22
-------
chamber and the number of nauplii within the corresponding test chamber are
recorded on data sheets (Figure 5). Several small aliquots are often necessary to
introduce 12-16 nauplii to each chamber. The concentration of nauplii in the 250
ml polycarbonate jar is adjusted so that an aliquot contains approximately five
organisms. If nauplii are too concentrated, counts under the microscope are
difficult and less accurate. If nauplii are not concentrated, too many aliquots are
needed and the test solution is diluted.
14.7 Temperature and Photoperiod
Test beakers containing organisms are placed randomly in a biological
incubator to control temperature (20 or 25 ± 2 °C) and lighting (100-150 fc). A
photoperiod of 1 6 hour light and 8 hour dark is utilized. Beakers are covered to
reduce cross contamination and evaporation.
14.8 Feeding
Eurytemora are fed during acute or chronic testing because copepods store
little energy reserves. Test organisms are fed daily after renewals with an equal
volume mixture of Isochrysis and Thalassiosira. Algal density in test beakers is
approximately the same as in the Eurytemora stock cultures (104 Isochrysis and 103
Thalassiosira cells/ml). Cell densities in subsamples from the algal cultures are
measured with a Spencer improved Neubauer corpuscle counting chamber
(hemacytometer). Algal density is adjusted so that 1.0 ml or less of the algal
mixture is added to each test beaker. The volume of algae mixture added is
minimized to avoid diluting the test condition. If algal density is low, the algae is
centrifuged and the cells re-suspended in an appropriate volume of the dilution
water.
23
-------
Figure 5. Sample data sheet for Eurytemora affinis test survival results.
Test ID:
Toxicant:
Performed by:
Date sample collected:
Salinity:
Date test began:
Date test ended:
Dilution water used:
Conc./% eff. Rep. N0 Nt % Survival Mean ± S.D.
1. A
B
C
D
2. A
B
C
D
3. A
B
C
D
4. A
B
C
D
5. A
B
C
D
6. A
B
C
D
7. A
B
C
D
24
-------
14.9 Physical and Chemical Analysis
The temperature, dissolved oxygen concentration, pH, and salinity are
measured in the dilution water and each set of test solutions and recorded (Figure
6). These parameters are measured daily prior to and following renewals by
collecting a composite sample from each test condition. The sample is obtained by
slowly siphoning test solution from each test beaker (outside the test chamber) into
a 100 mL plastic cup. Instruments are calibrated immediately prior to use with
fresh buffers and standards.
Aeration during the test may alter the results and should be used only as a
last resort to maintain the required DO. Aeration can reduce toxicity of test
solutions by stripping them of highly volatile toxic substances, or increase its
toxicity by altering pH. However, the DO in the test solution must not be permitted
to fall below 4.0 mg/l. If aeration is necessary, air is bubbled through a 1.0 ml
pipette at the rate of 100 bubbles/min.
14.10 Terminating the Test
An acute test is terminated after 48 or 96-hours of exposure. A chronic test
is terminated after 8-days of exposure. Survival is evaluated by reducing the
volume of test solution in each test beaker (using a slow siphon) to approximately
10 ml. The remaining test solution and copepods are removed from the test
chamber with a wide-bore glass pipette, examined under a dissecting microscope,
and the number of live copepods and corresponding test beaker are recorded.
After all the solution in the test chamber has been removed and surviving
copepods counted, the chambers are finally examined under the microscope for the
presence of any remaining copepods entrapped on the chamber mesh. If copepods
are present, the mesh is rinsed with a squeeze bottle and the remaining copepods
are pipetted from the chamber.
Surviving copepods are preserved for maturation and fecundity evaluation in
chronic tests. Copepods from each test beaker are preserved in separate 20 mL
scintillation vials containing 2.0 mL of 10% buffered formalin. Preserved copepods
are examined under a dissecting microscope and categorized as: gravid female;
non-gravid female; male; and immature (Figure 7). A gravid female is an adult
carrying eggs in an external ovisac. A non-gravid female has the adult female
characteristic of wing-like structures on the last segment of the metasome without
an external ovisac carried under the urosome. An adult male has the identifying
characteristic of a modified right first antenna. Immature E. affinis are smaller than
either adult males or females and lack the adult characteristics of either a modified
right first antenna or wing-like structures on the last segment of the metasome.
25
-------
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15.0 DATA ANALYSIS
The data are tabulated and summarized. The proportion of surviving
copepods, females that are gravid/total females, and copepods that have reached
maturity/total surviving copepods, in each replicate is calculated. The endpoints of
toxicity are based on the reduction in survival, proportion of gravid females, and
proportion of copepods reaching maturity. Previously, reproduction success was
evaluated by determining the number of neonates produced by each gravid female
24 hours after terminating the test. This method was abandoned because females
are capable of producing more than one brood after a single mating and no
significant effects were observed in three years of testing.
The LC50 and 95% confidence interval is calculated by the Spearman-Karber
Method, the Trimmed Spearman-Karber Method and the Probit Method. The
Spearman-Karber Method (Finney, 1978) requires 0% mortality in the lowest test
concentration (not including control) and 100% mortality in the highest test
concentration, and at least one condition with partial mortality. The Trimmed
Spearman-Karber Method (Hamilton et al., 1977) requires mortalities bracket 50%
and at least one concentration with partial mortality. The Probit Method (Finney,
1978) requires mortalities bracket 50% and at least two concentrations with partial
mortality.
Determination of the no-observed-effect-concentration (NOEC) and the
lowest-observed-effect-concentration (LOEC) for multi-concentration tests is
accomplished with hypothesis testing. Survival, reproduction, and maturation data
from each concentration are compared to the control values. The NOEC is the
lowest concentration that is not significantly different than the control value. The
LOEC is the lowest concentration that is statistically different (p <0.05) than the
control value. Proportions are transformed by the arc-sine-square-root
transformation. This transformation is commonly used on proportionality data to
stabilize variance and satisfy the normality requirement for parametric tests. After
data are transformed, normality and homogeneity of variance is determined with
the Shapiro-Wilks Test and Bartlett's Test, respectively. If the data meet the
normality and homogeneity of variance assumptions, Dunnett's Procedure
(parametric test) is used to determine significant differences. If, however, the data
fail to satisfy the assumptions, Steel's Many-one Rank Test (nonparametric test) is
used to determine the NOEC and LOEC. If unequal numbers of replicates occur
among the test concentrations, the parametric analysis alternative is the t-test with
a Bonferroni adjustment, and the nonparametric analysis alternative is the Wilcoxon
Rank Sum Test with the Bonferroni adjustment. A chronic value is determined by
calculating the geometric mean of the NOEC and LOEC values. Recently,
environmental managers have been utilizing the IC25, the concentration which
causes a 25% reduction in survival, growth or reproduction, as a measure of
29
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chronic toxicity (see EPA software - ICPIN Program).
Statistical procedures and calculations for the interpretation of toxicity test
data are described in greater detail in U.S.EPA (1991 a). Computer programs for
analyzing toxicity test data are available by contacting:
Center for Water Quality Monitoring
U.S. Environmental Protection Agency
Environmental Research Laboratory
College Station Road, Athens, Georgia 30613
Telephone: 404-546-3123
and
Fish Physiology and Toxicology Laboratory
Department of Zoology and Physiology
University of Wyoming
Laramie, Wyoming 82071
Telephone: 307-745-8504
30
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16.0 REFERENCES
Allan, S.J. and R.E. Daniels. 1982. Life table evaluation of chronic exposure of
Eurytemora affinis (Copepoda) to Kepone. Mar. Biol, 66:179-184.
American Society for Testing and Materials (ASTM). 1980. Standard Practice for
Conducting Toxicity Tests with Fishes, Macroinvertebrates and Amphibians. ASTM
E 729-80, Philadelphia, PA. 25 pp.
Bradley, B.P. 1975. The anomalous influence of salinity on temperature tolerances
of summer and water populations of the copepod, Eurytemora affinis. Biol. Bull.
148:26-34.
Bushong, S.J., L.W. Hall Jr., W.E. Johnson, W.S. Hall and M.C. Ziegenfuss. 1987.
Acute and chronic toxicity of tributyltin to selected Chesapeake Bay fish and
invertebrates. Final Report, Johns Hopkins University Applied Physics Laboratory,
Shady Side, MD.
Chesapeake Executive Council. 1989. Chesapeake Bay Basinwide Toxics
Reduction Strategy. Annapolis, MD.
Daniels, R.E. and J.D. Allan. 1981. Life table evaluation of chronic exposure to a
pesticide. Can. J. Fish. Aquat. Sci. 38:485-494.
DeWoskin, R.S. 1984. Good laboratory practice regulations: comparison.
Research Triangle Institute, Research Triangle Park, NC. 63pp.
Federal Drug Administration. 1978. Good laboratory practices for non-chemical
laboratory studies. Part 58. Federal Register 43(247):6001 3-60020, December 22,
1978.
Finney, D.J. 1978. Statistical method in biological assay. 3rd edition. Charles
Griffin and Co. Ltd., London. 508 pp.
Gaedke, V. 1 990. Population dynamics of the calanoid copepods, Eurytemora
affinis and Acartia tonsa in the Ems-Dollart Estuary: A numerical simulation. Arch.
Hydrobiol. 118:185-226.
Guillard, R.R.L. 1975. Culture of Phytoplankton for Feeding Marine Invertebrates.
In: Culture of Marine Invertebrate Animals. W.L. Smith and M.H. Chanley (eds.), pp.
29-60. Pleum Publishing, New York, NY.
Gurney, R. 1931. Eurytemora affinis. In: British Freshwater Copepoda. R. Gurney
31
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(ed.), pp. 202-214. Ray Society, London, England.
Hall, L.W. Jr., M.C. Ziegenfuss, R.D. Anderson, T.D. Spittler and H.C. Leichtweis.
1994. Influence of salinity on atrazine toxicity to a Chesapeake Bay copepod
(Eurytemora affinis) and fish (Cyprinodon variegatus). Estuaries 17:181-186.
Hall, L.W. Jr., M.C. Ziegenfuss, S.A. Fischer, R.W. Alden, E. Deaver, J. Gooch and
N. Debert-Hastings. 1991. A pilot study for ambient toxicity testing in
Chesapeake Bay. Vol. 1, Year 1 Report. U.S. EPA Chesapeake Bay Program
Office, Annapolis, MD.
Hamilton, M.A., R.C. Russo, and R.V. Thurston. 1977. Trimmed Spearman-Karber
method for estimating median lethal concentrations. Environ. Sci. Tech., 11:714-
719.
Heinle, D.R. 1969. Culture of calanoid copepods in synthetic seawater. J. Fish.
Res. Bd. Can. 26:150-153.
Jeffries, H.P. 1962. Succession of two Acartia species in estuaries. Limnol.
Oceanogr. 7:354-364.
Katona, S.K. 1970. Growth characteristics of the copepods, Eurytemora affinis
and E. herdmani in laboratory cultures. Helgolandes wiss. Meesesunters 20:373-
384.
Mori, T. 1964. The pelagic Copepoda from the neighboring waters of Japan. 150
pp., 80 pi.
Nagaraj, M. 1982. Combined effects of temperature and salinity on the
development of the copepod, Eurytemora affinis. Aquaculture 103:65-71.
Sullivan, B.K., E. Buskey, D.C. Milles and P.J. Ritacco. 1983. Effects of copper
and cadmium on growth, swimming and predator avoidance in Eurytemora affinis
(Copepoda). Mar. Biol. 77:299-306.
Taylor, J.K. 1987. Quality assurance of chemical measurements. Lewis
Publishers, Inc., Chelsea, Ml.
Tepper, B. 1986. Genetic correlations in natural populations of the copepod,
Eurytemora affinis. Ph.D. Thesis, University of Maryland, College Park, MD.
United States Environmental Protection Agency (U.S. EPA). 1975. Methods for
acute toxicity tests with fish, macroinvertebrates and amphibians. Environmental
32
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Research Laboratory, U.S. Environmental Protection Agency, Duluth, MN.
United States Environmental Protection Agency (U.S. EPA). 1977. Occupational
health and safety manual. Office of Planning and Management. U.S.
Environmental Protection Agency, Washington, DC.
United States Environmental Protection Agency (U.S. EPA). 1979a. Handbook for
analytical quality assurance in water and wastewater laboratories. U.S.
Environmental Protection Agency, Environmental Monitoring and Support
Laboratory, EPA/600/4-79-019. Cincinnati, OH.
United States Environmental Protection Agency (U.S. EPA). 1979b. Methods for
the chemical analysis of water and wastes. Environmental monitoring and support
laboratory. U.S. Environmental Protection Agency, EPA/600/4-79-020, Cincinnati,
OH.
United States Environmental Protection Agency (U.S. EPA). 1980a. Proposed
good laboratory practice guidelines for toxicity testing. Paragraph 163.60-6.
Federal Register 45:26377-26382, April, 18, 1980.
United States Environmental Protection Agency (U.S. EPA). 1980b. Physical,
chemical, persistence and ecological effects testing; good laboratory practice
standards (proposed rule). 40 CFR 772. Federal Register 45:77353-77365,
November 21, 1980.
United States Environmental Protection Agency (U.S. EPA). 1985. Methods for
Measuring the Acute Toxicity of Effluents to Freshwater and Marine Organisms.
W.H. Peltier and C.I. Weber (eds.). EPA/600-4-85/013, Washington, DC.
United States Environmental Protection Agency (U.S. EPA). 1991 a. Methods for
Measuring the Acute Toxicity of Effluents and Receiving Waters to Freshwater and
Marine Organisms. C.I. Weber (ed.). U.S. Environmental Protection Agency,
EPA/600/4-9-027. Cincinnati, OH.
United States Environmental Protection Agency (U.S. EPA). 1991b. Technical
support document for water quality-based toxics control. Office, Water, U.S.
Environmental Protection Agency, EPA/505/2-90-001. Washington, DC.
Waters, D.B. and C.W. Jameson. 1984. Health and safety or toxicity testing.
Butterworth Publishers, Woburn, MA.
Wilson, M.S. and H.C. Yeatman. 1959. Free-living Copepoda, ]n: Freshwater
Biology. W.T. Edmondson (ed.), pp. 795-861. John Wiley and Sons, New York,
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NY.
Ziegenfuss, M.C. and L.W. Hall, Jr. 1993. Screening of candidate species for
development of standard operating procedures for aquatic toxicity testing with
resident Chesapeake Bay biota. Report. Maryland Department of the Environment,
Baltimore, MD.
34
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APPENDIX A
SUMMARY OF EURYTEMORA AFFINIS CONTROL SURVIVAL DURING ACUTE AND
CHRONIC TOXICITY TESTING OVER A FOUR-YEAR PERIOD.
Date
Aug 1990
Sept 1990
Aug 1990
Dec 1990
Dec 1990
Dec 1990
Aug 1991
Aug 1991
Sept 1991
Apr 1992
Apr 1992
Apr 1992
Apr 1992
Apr 1992
June 1992
June 1992
July 1992
July 1992
Nov 1992
Oct 1992
Dec 1992
Dec 1992
Apr 1993
Oct 1993
Oct 1993
Oct 1993
Oct 1993
Nov 1993
Dec 1993
Jan 1994
Feb 1994
Mar 1994
Mar 1994
Mar 1994
June 1994
June 1994
June 1994
Sept 1994
Oct 1994
Control
Survival %
100
92
69
45
70
92
78
90
93
77
40
95
96
68
97
94
78
88
86
19
98
86
81
53
88
92
58
97
90
83
90
90
78
75
92
97
80
82
93
Duration
8-d
8-d
48-h
8-d
8-d
8-d
48-h
8-d
8-d
96-h
96-h
96-h
96-h
96-h
96-h
96-h
96-h
96-h
48-h
8-d
8-d
8-d
8-d
96-h
96-h
96-h
96-h
96-h
96-h
8-d
8-d
8-d
8-d
8-d
96-h
96-h
96-h
48-h
8-d
Water Source
Synthetic
Synthetic
Synthetic
Synthetic
Estuarine
Estuarine
Synthetic
Synthetic
Synthetic
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Salinity
15
15
15
15
15
15
15
15
15
5
15
25
25
5
15
25
5
15
15
15
5
15
15
15
25
5
25
15
25
5
5
5
15
25
5
15
25
15
15
Temp
PPt°C
25
25
25
25
25
25
25
25
25
20
20
20
20
20
20
20
20
20
25
25
25
25
25
25
25
25
25
25
25
25
25
25
25
25
25
25
25
25
25
Mean % control survival: 96-h = 82,
8-d = 86
35
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APPENDIX B
SUMMARY OF E. AFFIN1S CHRONIC ENDPOINTS REPRODUCTION (FECUNDITY)
AND GROWTH (MATURATION) EVALUATED UNDER CONTROL CONDITIONS
FROM 8 DAY TESTS.
Date Control Water Salinity Temperature
Source
Gravid Immatures ppt °C
Females,? Mean %
Mean %
Jan 1994 82 7 Estuarine 5 25
Feb 1994 83 3 Estuarine 5 25
Mar 1994 89 3 Estuarine 5 25
Mar 1994 6 37 Estuarine 15 25
Mar 1994 13 40 Estuarine 25 25
Oct 1994 97 7 Estuarine 15 25
36
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APPENDIX C
SINGLE LABORATORY PRECISION OF THE EURYTEMORA AFFINIS ESTUARINE
TOXICITY TEST AT 15 PPT SALINITY WITH THE REFERENCE TOXICANT
CADMIUM CHLORIDE.
Test
48-h LC50 Cd CI2
95% Conf. Interval
1.
2.
3.
98
113
143
Mean:
Standard Deviation:
Coeff. Variation:
67-144
72-177
111 - 184
118
22.9
19.4%
37
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APPENDIX D
RELATIVE SENSITIVITY OF EURYTEMORA AFFINIS TO SELECTED TOXIC
SUBSTANCES.
Copper 96-h LC50 = 30 ^g/L (Adult) (Sullivan et al., 1983)
Dieldrin 48-h LC50 = 23 ^glL (Napulii) (Daniels & Allan, 1981)
TBT 48-h LC50 = 1.4 ^glL (Subadult) (Bushong et al., 1988)
Kepone 48-h LC50 = 40 /^g/L (Nauplii) (Allan & Daniels, 1982)
Atrazine 96-h LC50 = 0.5-13.2 mg/L (Nauplii) (Hall et al., 1994)
38
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