£EPA
United States
Environmental Protection
Agency
Office of
Research and Development
Washington.-DC 20460
EPA/600'4-90/027
September 1991
Methods for Measuring the
Acute Toxkity of
Effluents and Receiving
Waters to Freshwater and
Marine Organisms
Fourth Edition
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EPA-600/4-90-027
SEPTEMBER 1991
METHODS FOR MEASURING THE ACUTE TOXICITY
OF EFFLUENTS AND RECEIVING WATERS
TO FRESHWATER AND MARINE ORGANISMS
(FOURTH EDITION)
Edited by
Cornelius I. Weber
Floor
Chicago, II 60604-^0
ENVIRONMENTAL MONITORING SYSTEMS LABORATORY - CINCINNATI
OFFICE OF RESEARCH AND DEVELOPMENT
U.S. ENVIRONMENTAL PROTECTION AGENCY
CINCINNATI, OHIO 45268
OO) Printed on Recycled Paper
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DISCLAIMER
This report has been reviewed by the Environmental Monitoring Systems
Laboratory - Cincinnati (EMSL-Cincinnati), U. S. Environmental Protection
Agency (USEPA), and approved for publication. The mention of trade names
or commercial products does not constitute endorsement or recommendation
for use.
11
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FOREWORD
Environmental measurements are required to determine the chemical and
biological quality of drinking water, surface waters, ground waters,
waste waters, sediments, sludges, and solid waste. The Environmental
Monitoring Systems Laboratory - Cincinnati (EMSL-Cincinnati) conducts
research to:
o Develop and evaluate methods to identify and measure the
concentration of chemical pollutants.
o Identify and quantitate the occurrence of viruses, bacteria,
other human pathogens, and indicator organisms.
o Perform ecological measurements and measure the toxicity of
pollutants to representative species of aquatic organisms and
determine the effects of pollution on communities of indigenous
freshwater, estuarine, and marine organisms, including the
phytoplankton, zooplankton, periphyton, macrophyton,
macroinvertebrates, and fish.
o Develop and operate a quality assurance program to support
achievement of data quality objectives for environmental
measurements.
The Federal Water Pollution Control Act Amendments of 1972 (PL
92-500), the Clean Water Act (CWA) of 1977 (PL 95-217), and the Water
Quality Act of 1987 (PL 100-4) explicitly state that it is the national
policy that the discharge of toxic substances in toxic amounts be
prohibited. The detection of acutely toxic effluents, therefore, plays
an important role in identifying and controlling toxic discharges to
surface waters. This manual is the fourth edition of the acute toxicity
test manual for effluents, first published by EMSL-Cincinnati in January,
1978. It provides updated methods for estimating the acute toxicity of
effluents to freshwater, estuarine, and marine organisms for use by the
U.S. Environmental Protection Agency (USEPA) regional programs, the state
programs, and the National Pollutant Discharge Elimination System (NPDES)
permittees.
Thomas A. Clark
Director
Environmental Monitoring Systems
Laboratory - Cincinnati
m
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PREFACE
This manual represents the fourth edition of the general purpose effluent
acute toxicity test manual initially published by EMSL-Cincinnati in January,
1978. The current edition reflects changes recommended by the Toxicity
Assessment Subcommittee of the EMSL-Cincinnati Biological Advisory Committee,
USEPA headquarters program offices and regions, state and interstate water
pollution control programs, environmental protection groups, trade
associations, major industries, consulting firms, academic institutions
engaged in aquatic toxicology research, and other interested parties in the
private sector.
The membership of the Toxicity Assessment Subcommittee, EMSL-Cincinnati
Biological Advisory Committee is as follows:
William Peltier, Subcommittee Chairman,
Environmental Services Division, Region 4
Peter Nolan, Environmental Services Division, Region 1
Steve Ward, Environmental Services Division, Region 2
Ronald Preston, Environmental Services Division, Region 3
Charles Steiner, Environmental Services Division, Region 5
Evan Hornig, Environmental Services Division, Region 6
Terry Hollister, Environmental Services Division, Region 6
Michael Tucker, Environmental Services Division, Region 7
Loys Parrish, Environmental Services Division, Region 8
Peter Husby, Environmental Services Division, Region 9
Joseph Cummins, Environmental Services Division, Region 10
Bruce Binkley, National Enforcement Investigations Center, Denver
Wesley Kinney, Environmental Monitoring Systems Laboratory - Las Vegas
George Morrison, Environmental Research Laboratory - Narragansett
Douglas Middaugh, Environmental Research Laboratory - Gulf Breeze
Teresa Norberg-King, Environmental Research Laboratory - Duluth
Donald Klemm, Environmental Monitoring Systems Laboratory - Cincinnati
Philip Lewis, Environmental Monitoring Systems Laboratory - Cincinnati
Richard Swartz, Environmental Research Laboratory - Newport
Margarete Heber, Health and Ecological Criteria Division, Office of Science
and Technology (OST), Office of Water (OW)
Chris Zarba, Health and Ecological Criteria Division, OST,OW
Bruce Newton, Assessment and Watershed Protection Division, Office
of Wetlands, Oceans, and Watersheds, OW
Dan Rieder, Hazard Evaluation Division, Office of Pesticide Programs
Jerry Smrchek, Health and Environmental Review Division, Office of
Toxic Substances
Gail Hansen, Office of Solid Waste
Royal Nadeau, Emergency Response Team, Edison, NJ
James M. Lazorchak, Ph.D.
Chairman, Biological Advisory Committee
IV
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ABSTRACT
This manual describes methods for measuring the acute toxicity of
effluents to freshwater, estuarine, and marine macroinvertebrates and
fish. The methods include single and multiple concentration static
non-renewal, static-renewal, and flow-through toxicity tests for
effluents and receiving waters. Also included are guidelines on
laboratory safety; quality assurance; facilities and equipment; test
species selection and handling; dilution water; effluent and receiving
water sample collection, preservation, shipping, and holding; test
conditions; toxicity test data analysis; report preparation; organism
culturing; and dilutor and mobile laboratory construction.
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CONTENTS
Foreword iii
Preface iv
Abstract v
Figures ix
Tables x
Acknowledgments xiii
1. Introduction 1
2. Types of Tests 2
3. Health and Safety 5
General Precautions 5
Safety Equipment 5
General Laboratory and Field Operations 6
Disease Prevention 6
Safety Manuals 6
4. Quality Assurance 7
Introduction 7
Facilities, Equipment, and Test Chambers 7
Test Organisms 8
Laboratory Water used for Culturing and
Test Dilution Water 8
Effluent Sampling and Sample Handling 8
Test Conditions 8
Quality of Test Organisms 8
Food Quality 9
Acceptability of Acute Toxicity Test Results .... 9
Analytical Methods 10
Calibration and Standardization 10
Replication and Test Sensitivity 10
Variability in Toxicity Test Results 10
Demonstrating Acceptable Laboratory Performance ... 11
Documenting Ongoing Laboratory Performance 11
Reference Toxicants 12
Record Keeping 12
5. Facilities and Equipment 20
General Requirements 20
Cleaning Test Chambers and Laboratory Apparatus ... 21
Apparatus and Equipment for Culturing and
Toxicity Tests 22
Reagents and Consumable Materials 23
Test Organisms 25
6. Test Organisms 26
Test Species 26
Sources of Test Organisms 27
Life Stage 28
Laboratory Culturing 28
vi
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Holding and Handling of Test Organisms 28
Transportation to the Test Site 29
Test Organism Disposal 31
7. Dilution Water 32
Types of Dilution Water 32
Standard Synthetic Dilution Water 32
Use of Receiving Water as Dilution Water 34
Use of Tap Water as Dilution Water 38
Dilution Water Holding 38
8. Effluent and Receiving Water Sampling and
Sample Handling 39
Effluent Sampling 39
Effluent Sample Types 39
Effluent Sampling Recommendations 40
Receiving Water Sampling 41
Effluent and Receiving Water Sample Handling,
Preservation, and Shipping 41
Sample Receiving 42
Persistence of Effluent Toxicity During Sample
Shipping and Holding 42
9. Acute Toxicity Test Procedures 44
Preparation of Effluent and Receiving Water Samples
for Toxicity Tests 44
Preliminary Toxicity Range-finding Tests 46
Multi-concentration (Definitive) Effluent
Toxicity Tests 46
Receiving Water Tests 47
Static Tests 47
Flow-through Tests 48
Number of Test Organisms 49
Replicate Test Chambers 49
Loading of Test Organisms 51
Illumination 51
Feeding 51
Test Temperature 52
Stress 52
Dissolved Oxygen Concentration 52
Test Duration 55
Acceptability of Test Results 55
Summary of Test Conditions for the Principal
Test Organisms 55
10. Test Data 70
Biological Data 70
Chemical and Physical Data 70
11. Acute Toxicity Data Analysis 74
Introduction 74
Determination of the LC50 from Definitive, Multi-
Effluent-Concentration, Acute Toxicity Tests ... 75
Graphical Method 77
VII
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Spearman-Karber Method 80
Trimmed Spearman-Karber Method 82
Probit Method 86
Determination of No-Observed-Adverse-Effect
Concentration (NOAEC) from Multi-Concentration Tests,
and Determination of Pass or Fail (Pass/Fail) for
Single-Concentration (Paired) Tests 91
General Procedure 91
Single Concentration Test 100
Multi-Concentration Test 106
12. Report Preparation 117
Cited References 119
Bibliography 124
Appendices 130
A. Distribution, Life Cycle, Taxonomy, and Culture
and Holding Methods 131
Daphnid, Cen'odaphm'a dubia 131
Daphnids, Daphnia pulex and D. magna 148
Mysid, Mysidopsis bahia 169
Brine Shrimp, Artemia salina 189
Fathead Minnow, Pimephales promelas 198
Rainbow Trout, Oncorhynchus mykiss
and Brook Trout, Salvelinus fontinalis 217
Sheepshead minnow, Cyprinodon variegatus 227
Silversides: Inland Silverside, Mem'dia
beryllina, Atlantic Silverside, M. mem'dia,
and Tidewater Silverside, N. pem'nsulae , 246
B. Supplemental List of Acute Toxicity Test Species 263
C. Dilutor Systems 266
Solenoid and Vacuum Siphon Dilutor Systems 266
Solenoid System Equipment List 270
Vacuum System Equipment List 273
Dilutor Control Panel Equipment List 278
D. Plans for Mobile Toxicity Test Labortory 279
Tandem-axle Trailer 279
Fifth-wheel Trailer 282
E. Check Lists and Information Sheets 283
Bioassay Field Equipment List 281
Information Check List for On-site Industrial and
Municipal Waste Toxicity Tests 285
Daily Events Log 291
Dilutor Calibration Form 292
Daily Dilutor Calibration Check 293
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FIGURES
Number Page
1. Control (cusum) charts 19
2. Approximate times required to replace water in
test chambers in flow-through tests 50
3. Rawson's nomograph for obtaining oxygen saturation
values in freshwater at different temperatures at
sea level 53
4. Data sheet for effluent toxicity tests 72
5. Check list on back of effluent toxicity data sheet 73
6. Flowchart for determination of the LC50 for multi-
concentration acute toxicity tests 76
7. Plotted data and fitted line for Graphical Method,
using all-or-nothing data 79
8. Example of input for computer program for Trimmed
Spearman-Karber Method 84
9. Example of output from computer program for Trimmed
Spearman-Karber Method 85
10. Example of input for computer program for Probit Method ... 88
11. Example of output for computer program for Probit Method ... 89
12. Plot of adjusted probits and predicted regression line .... 90
13. Flowchart for analysis of single-effluent-concentration
test data 92
14. Flowchart for analysis of multi-effluent-concentration
test data 93
15. Plot of mean survival proportion data in Table 27 108
IX
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TABLES
Number Page
1. Intra-laboratory precision of LCSOs from static acute
toxicity tests with aquatic organisms, using reference
toxicants , 14
2. Intra- and inter-laboratory precision of 48-h acute toxicity
tests with Daphm'a magna, using a standard effluent 15
3. Inter-laboratory precision of acute toxicity tests with
aquatic organisms, using reference toxicants 16
4. Inter!aboratory study of acute toxicity test precision, 1990:
summary of responses using KC1 as the reference toxicant ... 17
5. National interlaboratory study of acute toxicity test
precision, 1991: summary of responses using reference
toxicants 18
6. Preparation of synthetic freshwater using reagent
grade chemicals 35
7. Preparation of synthetic freshwater using mineral water ... 35
8. Preparation of synthetic seawater using reagent
grade chemicals 36
9. Percent unionized NH3 in aqueous ammonia solutions:
temperature 15-26°C and pH 6.0-8.9 45
10. Oxygen solubility (mg/L) in water at equilibrium with air
at 760 mm Hg (after Richards and Corwin, 1956) 54
11. Summary of test conditions and test acceptability criteria
for Ceriodaphm'a dubia acute toxicity tests with
effluents and receiving waters 56
12. Summary of test conditions and test acceptability criteria
for Daphm'a pulex and D. magna acute toxicity tests
with effluents and receiving waters 58
13. Summary of test conditions and test acceptability criteria
for fathead minnow, Pimephales prome/as, acute toxicity
tests with effluents and receiving waters 60
14. Summary of test conditions and test acceptability criteria
for rainbow trout, Oncorhynchus mykiss, and brook trout,
SalveTinus fontinalis, acute toxicity tests with effluents
and receiving waters 62
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TABLES (CONT.)
Number Page
15. Summary of test conditions and test acceptability criteria
for mysid, Mysidopsis bahia, acute toxicity tests with
effluents and receiving waters 64
16. Summary of test conditions and test acceptability
criteria for sheepshead minnow, Cyprinodon
variegatus, acute toxicity tests with effluents and
receiving waters 66
17. Summary of test conditions and test acceptability
criteria for silverside, Mem'dia beryllina, M. menidia,
and M. pem'nsulae, acute toxicity tests with effluents
and receiving waters 68
18. Mortality data (number of dead organisms) from acute
toxicity tests used in examples of LC50 determinations
(20 organisms in the control and all test concentrations) . . 78
19. Coefficients for the Shapiro-Milk's test 95
20. Quantiles of the Shapiro-Milk's test statistic 99
21. Critical values for Milcoxon's rank sum test five
percent critical level 103
22. Data from an acute single-concentration toxicity
test with Cen'odaphm'a 103
23. Example of Shapiro-Milk's test: centered observations .... 104
24. Example of Shapiro-Milk's test: ordered observations .... 104
25. Example of Shapiro-Milk's test: Table of coefficients
and differences 104
26. Example of Milcoxon's rank sum test: Assigning ranks
to the control and 100% effluent concentrations 105
27. Fathead minnow survival data 107
28. Centered observations for Shapiro-Milk's example 107
29. Ordered centered observations for the
Shapiro-Milk's example 110
xi
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TABLES (CONT.)
Number Page
30. Coefficients and differences for the
Shapiro-Wilk's example 110
31. Anova table 112
32. Anova table for Dunnett's Procedure example 113
33. Calculated T values 114
34. Dunnett's "T" values 115
Xll
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ACKNOWLEDGMENTS
The principal authors of this document are Cornelius Weber, Florence Fulk,
Donald Klemm, Philip Lewis, and James Lazorchak, Environmental Monitoring
Systems Laboratory - Cincinnati; William Peltier, Environmental Services
Division, Region 4; Stephen Ward, Environmental Services Division, Region 2;
Margarete Heber, Office of Science and Technology, Office of Water; Teresa
Norberg-King, Environmental Research Laboratory, Duluth; George Morrison and
David Bengtson, Environmental Research Laboratory, Narragansett; Douglas
Middaugh, Environmental Research Laboratory, Gulf Breeze; Mark Smith and
Quentin Pickering, Technology Applications, Inc. Cincinnati; and Cathy Poore,
Computer Sciences Corporation, Cincinnati. Contributors to specific sections
of this manual are listed below.
1. Sections 1-10 and 12: General Guidelines
Margarete Heber
James Lazorchak
Teresa Norberg-King
George Morrison
William Peltier
Cornelius Weber
2. Section 11: Data Analysis
Florence Fulk
Cathy Poore
3. Appendices
Appendix A.I
Appendix A.2
Appendix A.3
Appendix A.4
Appendix A.5
Appendix A.6
Appendix A.7
Appendix A.8
Appendix B:
Appendix C:
Appendix D:
Appendix E:
and David Bengtson
, Quentin Pickering,
Philip Lewis and James Lazorchak
Philip Lewis and James Lazorchak
Stephen Ward
Philip Lewis
Donald Klemm
Donald Klemm
Donald Klemm
Douglas Middaugh and Donald Klemm
Margarete Heber
William Peltier
William Peltier
William Peltier
and Mark Smith
xm
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ACKNOWLEDGMENTS (CONT.)
Review comments from the following persons are gratefully acknowledged:
Barbara Albrecht, Environmental Research Laboratory, U.S. Environmental
Protection Agency, Gulf Breeze, Florida
Robert Burm, Environmental Services Division, U.S. Environmental Protection
Agency, Denver, Colorado
Randy Crawford, Missouri Department of Natural Resources, Jefferson City,
Missouri
Geri Cripe, Environmental Research Laboratory, U.S. Environmental Protection
Agency, Gulf Breeze, Florida
Philip Crocker, Environmental Services Division, U.S. Environmental Protection
Agency, Dallas, Texas
Joseph Cummins, Environmental Services Division, U.S. Environmental Protection
Agency, Seattle, Washington
Robert Donaghy, Environmental Services Division, U.S. Environmental Protection
Agency, Wheeling, West Virginia
Lee Dunbar, Connecticut Department of Environmental Protection, Hartford,
Connecticut
William Gidley, Nebraska Department of Environmental Control, Lincoln,
Nebraska
James Green, Environmental Services Division, U.S. Environmental Protection
Agency, Wheeling, West Virginia
Steve Haslouer, Kansas Department of Health and Environment, Topeka, Kansas
Thorn Haze, Connecticut Department of Environmental Protection, Hartford,
Connecticut
Michael Henebry, Illinois Environmental Protection Agency, Springfield,
Illinois
Terry Hollister, Environmental Services Division, U.S. Environmental
Protection Agency, Houston, Texas
Jack Kennedy, University of Iowa Hygienic Laboratory, Iowa City, Iowa
Alfred Korndorfer, New Jersey Department of Environmental Protection, Trenton,
New Jersey
Robert Masnado, Wisconsin Department of Natural Resources, Madison, Wisconsin
Ann McGinley, Texas Water Commission, Austin, Texas
Mary Moffett, Environmental Services Division, U.S. Environmental Protection
Agency, Kansas City, Kansas
Michael Morton, Environmental Services Division, U.S. Environmental Protection
Agency, Dallas, Texas
Peter Nolan, Environmental Services Division, U.S. Environmental Protection
Agency, Lexington, Massachusetts
Loys Parrish, Environmental Services Division, U.S. Environmental Protection
Agency, Denver, Colorado
Glen Rodriguez, Environmental Services Division, U.S. Environmental Protection
Agency, Denver, Colorado
Janice Smithson, West Virginia Division of Natural Resources, Charleston, West
Virginia
xiv
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ACKNOWLEDGMENTS (CONT.)
Charles Steiner, Environmental Services Division, U.S. Environmental
Protection Agency, Chicago, Illinois
Donald Thurston, New Hampshire Department of Environmental Services, Concord,
New Hampshire
Michael Tucker, Environmental Services Division, U.S. Environmental Protection
Agency, Kansas City, Kansas
Bruce Walker, Michigan Department of Natural Resources, Lansing, Michigan
Audrey Weber, Virginia State Water Quality Control Board, Richmond, Virginia
Charles Webster, Ohio Environmental Protection Agency, Columbus, Ohio
Many, very useful public comments on the third edition of the acute
toxicity test methods (EPA/600/4-85/013) were received in response to the
proposed rule, published in the Federal Register, December 4, 1989 [FR
54(231):50216-50224], regarding the Agency's intent to include the acute
toxicity tests in Table IA, 40 CFR Part 136. These comments were considered
in the preparation of the fourth edition of the manual, and are included in
the Public Docket for the rulemaking, located at room 2904, EPA Headquarters,
Washington, DC.
xv
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SECTION 1
INTRODUCTION
1.1 This manual describes acute toxicity tests for use in the National
Pollutant Discharge Elimination System (NPDES) Permits Program to
identify effluents and receiving waters containing toxic materials in
acutely toxic concentrations. The methods included in this manual are
referenced in Table IA, 40 CFR Part 136 regulations and, therefore,
constitute approved methods for acute toxicity tests. They are also
suitable for determining the toxicity of specific compounds contained in
discharges. The tests may be conducted in a central laboratory or
on-site, by the regulatory agency or the permittee.
1.2 The data are used for NPDES permits development and to determine
compliance with permit toxicity limits. Data can also be used to predict
potential acute and chronic toxicity in the receiving water, based on the
LC50 and appropriate dilution, application, and persistence factors. The
tests are performed as a part of self-monitoring permit requirements,
compliance biomonitoring inspections, toxics sampling inspections, and
special investigations. Data from acute toxicity tests performed as part
of permit requirements are evaluated during compliance evaluation
inspections and performance audit inspections.
1.3 Modifications of these tests are also used in toxicity reduction
evaluations and toxicity identification evaluations to identify the toxic
components of an effluent, to aid in the development and implementation
of toxicity reduction plans, and to compare and control the effectiveness
of various treatment technologies for a given type of industry,
irrespective of the receiving water (USEPA, 1988a,1988b, 1989a, 1989b,
1991a).
1.4. This methods manual serves as a companion to the short-term chronic
toxicity test methods manuals for freshwater and marine organisms (USEPA,
1991b, 1991c).
1.5 Guidance for the implementation of toxicity tests in the NPDES
program is provided in the Technical Support Document for Water Quality-
Based Toxics Control (USEPA, 1991d).
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SECTION 2
TYPES OF TESTS
2.1 The selection of the test type will depend on the NPDES permit
requirements, the objectives of the test, the available resources, the
requirements of the test organisms, and effluent characteristics such as
fluctuations in effluent toxicity.
2.2 Effluent acute toxicity is generally measured using a multi-
concentration, or definitive test, consisting of a control and a minimum
of five effluent concentrations. The tests are designed to provide dose-
response information, expressed as the percent effluent concentration
that is lethal to 50% of the test organisms (LC50) within the prescribed
period of time (24-96 h), or the highest effluent concentration in which
survival is not statistically significantly different from the control.
2.3 Use of pass/fail tests consisting of a single effluent concentration
(e.g., the receiving water concentration or RWC) and a control is not
recommended. If the NPDES permit has a whole effluent toxicity limit for
acute toxicity at the RWC, it is prudent to use that permit limit as the
midpoint of a series of five effluent concentrations. This will ensure
that there is sufficient information on the dose-response relationship.
For example, the effluent concentrations utilized in a test may be:
(1) 100% effluent, (2) (RWC + 100)/2, (3) RWC, (4) RWC/2, and (5) RWC/4.
More specifically, if the RWC = 50%, the effluent concentrations used in
the toxicity test would be 100%, 75%, 50%, 25%, and 12.5%.
2.4 Receiving (ambient) water toxicity tests commonly employ two
treatments, a control and the undiluted receiving water, but may also
consist of a series of receiving water dilutions.
2.5 A negative result from an acute toxicity test does not preclude the
presence of chronic toxicity. Also, because of the potential temporal
variability in the toxicity of effluents, a negative test result with a
particular sample does not preclude the possibility that samples
collected at some other time might exhibit acute (or chronic) toxicity.
2.6 The frequency with which acute toxicity tests are conducted under a
given NPDES permit is determined by the regulatory agency on the basis of
factors such as the variability and degree of toxicity of the waste,
production schedules, and process changes.
2.7 Tests may be static (static non-renewal or static renewal), or flow-
through.
2.7.1 Static Tests
2.7.1.1 Static non-renewal tests - The test organisms are exposed to the
same test solution for the duration of the test.
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2.7.1.2 Static-renewal tests - The test organisms are exposed to a fresh
solution of the same concentration of sample every 24 h or other
prescribed interval, either by transferring the test organisms from one
test chamber to another, or by replacing all or a portion of solution in
the test chambers.
2.7.2 Flow-through Tests
2.7.2.1 Two types of flow-through tests are in common use: (1) sample is
pumped continuously from the sampling point directly to the dilutor
system; and (2) grab or composite samples are collected periodically,
placed in a tank adjacent to the test laboratory, and pumped continuously
from the tank to the dilutor system. The flow-through method employing
continuous sampling is the preferred method for on-site tests. Because of
the large volume (often 400 L/day) of effluent normally required for
flow-through tests, it is generally considered too costly and impractical
to conduct these tests off-site at a central laboratory.
2.8 Advantages and disadvantages of the types of tests are as follows:
2.8.1 Static non-renewal tests:
2.8.1.1 Advantages:
1. Simple and inexpensive
2. Very cost effective in determining compliance with permit
conditions.
3. Limited resources (space, manpower, equipment) required; would
permit staff to perform many more tests in the same amount of
time.
4. Smaller volume of effluent required than for static renewal or
flow-through tests.
2.8.1.2 Disadvantages:
1. Dissolved oxygen (DO) depletion may result from high chemical
oxygen demand (COD), biological oxygen demand (BOD), or
metabolic wastes.
2. Possible loss of toxicants through volatilization and/or
adsorption to the exposure vessels.
3. Generally less sensitive than static renewal or flow-through
tests, because the toxic substances may degrade or be adsorbed,
thereby reducing the apparent toxicity. Also, there is less
chance of detecting slugs of toxic wastes, or other temporal
variations in waste properties.
2.8.2 Static-renewal, acute toxicity tests:
2.8.2.1 Advantages:
1. Reduced possibility of dissolved oxygen (DO) depletion from high
chemical oxygen demand (COD) and/or biological oxygen demand
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(BOD), or ill effects from metabolic wastes from organisms in
the test solutions.
2. Reduced possibility of loss of toxicants through volatilization
and/or adsorption to the exposure vessels.
3. Test organisms that rapidly deplete energy reserves are fed when
the test solutions are renewed, and are maintained in a
healthier state.
2.8.2.2 Disadvantages:
1. Require greater volume of effluent than non-renewal tests.
2. Generally less sensitive than flow-through tests, because the
toxic substances may degrade or be adsorbed, thereby reducing
the apparent toxicity. Also, there is less chance of detecting
slugs of toxic wastes, or other temporal variations in waste
properties.
2.8.3 Flow-through tests:
2.8.3.1 Advantages:
1. Provide a more representative evaluation of the acute toxicity
of the source, especially if sample is pumped continuously
directly from the source and its toxicity varies with time.
2. DO concentrations are more easily maintained in the test
chambers.
3. A higher loading factor (biomass) may be used.
4. The possibility of loss of toxicant due to volatilization,
adsorption, degradation, and uptake is reduced.
2.8.3.2 Disadvantages:
1. Large volumes of sample and dilution water are required.
2. Test equipment is more complex and expensive, and requires more
maintenance and attention.
3. More space is required to conduct tests.
4. Because of the resources required, it would be very difficult to
perform multiple or overlapping sequential tests.
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SECTION 3
HEALTH AND SAFETY
3.1 GENERAL PRECAUTIONS
3.1.1 Development and maintenance of an effective health and safety
program in the laboratory requires an ongoing commitment by laboratory
management, and includes (1) the appointment of a laboratory health and
safety officer with the responsibility and authority to develop and
maintain a safety program, (2) the preparation of a formal, written,
health and safety plan, which is provided to each laboratory staff
member, (3) an ongoing training program on laboratory safety, and (4)
regularly scheduled, documented, safety inspections.
3.1.2 Collection and use of effluents in toxicity tests may involve
significant risks to personal safety and health. Personnel collecting
effluent samples and conducting toxicity tests should take all safety
precautions necessary for the prevention of bodily injury and illness
which might result from ingestion or invasion of infectious agents,
inhalation or absorption of corrosive or toxic substances through skin
contact, and asphyxiation due to lack of oxygen or presence of noxious
gases.
3.1.3 Prior to sample collection and laboratory work, personnel must
determine that all required safety equipment and materials have been
obtained and are in good condition.
3.1.4 Guidelines for the handling and disposal of hazardous materials
must be strictly followed.
3.2 SAFETY EQUIPMENT
3.2.1 Personal Safety Gear
3.2.1.1 Personnel must use safety equipment, as required, such as rubber
aprons, laboratory coats, respirators, gloves, safety glasses, hard hats,
and safety shoes.
3.2.2 Laboratory Safety Equipment
3.2.2.1 Each laboratory (including mobile laboratories) must be provided
with safety equipment such as first aid kits, fire extinguishers, fire
blankets, emergency showers, and eye fountains.
3.2.2.2 Mobile laboratories should be equipped with a telephone to
enable personnel to summon help in case of emergency.
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3.3 GENERAL LABORATORY AND FIELD OPERATIONS
3.3.1 Guidance in Material Safety Data Sheets should be followed for
reagents and other chemicals purchased from supply houses.
3.3.2 Work with effluents must be performed in compliance with accepted
rules pertaining to the handling of hazardous materials (see Safety
Manuals, Paragraph 3.5). Personnel collecting samples and performing
toxicity tests should not work alone.
3.3.3 Because the chemical composition of effluents is usually only
poorly known, they must be considered as potential health hazards, and
exposure to them should be minimized. Fume and canopy hoods over the
test areas must be used whenever necessary.
3.3.4 It is advisable to cleanse exposed parts of the body immediately
after collecting effluent samples.
3.3.5 All containers must be adequately labeled to indicate their
contents.
3.3.6 Strong acids and volatile organic solvents employed in glassware
cleaning must be used in a fume hood or under an exhaust canopy over the
work area.
3.3.7 Good housekeeping contributes to safety and reliable results.
3.3.8 Electrical equipment or extension cords not bearing the approval
of Underwriter Laboratories must not be used. Ground-fault interrupters
must be installed in all "wet" laboratories where electrical equipment is
used.
3.3.9 Mobile laboratories must be properly grounded to protect against
electrical shock.
3.4 DISEASE PREVENTION
3.4.1 Personnel handling samples which are known or suspected to contain
human wastes should be immunized against hepatitis B, tetanus, typhoid
fever, and polio.
3.5 SAFETY MANUALS
3.5.1 For further guidance on safe practices when collecting effluent
samples and conducting toxicity tests, check with the permittee and
consult general industrial safety manuals, including USEPA (1986) and
Walters and Jameson (1984).
-------
SECTION 4
QUALITY ASSURANCE1
4.1 INTRODUCTION
4.1.1 Development and maintenance of a toxicity test laboratory quality
assurance (QA) program requires an ongoing commitment by laboratory
management, and includes the following: (1) appointment of a laboratory
quality assurance officer with the responsibility and authority to develop and
maintain a QA program, (2) preparation of a quality assurance plan with data
quality objectives, (3) preparation of written descriptions of laboratory
standard operating procedures (SOP's) for test organism culturing, toxicity
testing, instrument calibration, sample chain-of-custody, laboratory sample
tracking system, etc. and (4) provision of adequate, qualified technical staff
and suitable space and equipment to assure reliable data.
4.1.2 QA practices within an aquatic toxicology laboratory must address all
activities that affect the quality of the final effluent toxicity data, such
as: (1) effluent sampling and handling; (2) the source and condition of the
test organisms; (3) condition and operation of equipment; (4) test conditions;
(5) instrument calibration; (6) replication; (7) use of reference toxicants;
(8) record keeping; and (9) data evaluation.
4.1.3 Quality control practices, on the other hand, consists of the more
focused, routine, day-to-day activities carried out within the scope of the
overall QA program. For more detailed discussion of quality assurance, and
general guidance on good laboratory practices related to toxicity testing,
see: FDA, 1978; USEPA, 1975, 1979a, 1980a, 1980b, 1991b; DeWoskin, 1984; and
Taylor, 1987.
4.2 FACILITIES, EQUIPMENT, AND TEST CHAMBERS
4.2.1 Separate test organism culturing and toxicity testing areas should be
provided to avoid possible loss of cultures due to cross-contamination.
Ventilation systems should be designed and operated to prevent recirculation
or leakage of air from chemical analysis laboratories or sample storage and
preparation areas into test organism culturing or toxicity testing areas, and
from toxicity test laboratories and sample preparation areas into culture
rooms.
4.2.2 Laboratory and toxicity test temperature control equipment must be
adequate to maintain recommended test water temperatures. Recommended
materials must be used in the fabrication of the test equipment which comes in
contact with the effluent (see Section 5, Facilities and Equipment).
'Adapted from USEPA, 1989c.
-------
4.3 TEST ORGANISMS
4.3.1 The test organisms used in the procedures described in this manual are
listed in Section 6. The organisms should appear healthy, behave normally,
feed well, and have low mortality in cultures, during holding, and in test
controls. Test organisms should be positively identified to species.
4.4 LABORATORY WATER USED FOR CULTURING AND TEST DILUTION WATER
4.4.1 The quality of water used for test organism culturing and for dilution
water used in toxicity tests is extremely important. Water for these two uses
should come from the same source. The dilution water used in effluent
toxicity tests will depend in part on the objectives of the study and
logistical constraints, as discussed in detail in Section 7. Types of water
treatment facilities are discussed in Section 5. Water supplies used for
culturing and test dilution water should be analyzed quarterly, as a minimum,
for toxic metals and organics. The concentration of the metals, Al, As, Cr,
Co, Cu, Fe, Pb, Ni, Zn, expressed as total metal, should not exceed 1 ug/L
each, and Cd, Hg, and Ag, expressed as total metal, should not exceed 100 ng/L
each. Total organochlorine pesticides plus PCBs should be less than 50 ng/L
(APHA, 1989). Individual pesticide concentrations should not exceed the
limits set in the USEPA National Water Quality Guidelines.
4.5 EFFLUENT SAMPLING AND SAMPLE HANDLING
4.5.1 Sample holding times and temperatures must conform to conditions
described in Section 8, Effluent Sampling and Sample Handling.
4.6 TEST CONDITIONS
4.6.1 The temperature of test solutions must be measured by placing the
thermometer or probe directly into the test solutions, or by placing the
thermometer in equivalent volumes of water in surrogate vessels positioned at
appropriate locations among the test vessels. Temperature should be recorded
continuously in at least one vessel during the duration of each test. Test
solution temperatures must be maintained within the limits specified for each
test. DO concentration and pH in test chambers should be checked daily
throughout the test period, as prescribed in Section 9.
4.7 QUALITY OF TEST ORGANISMS
4.7.1 The quality of test organisms obtained from an outside source must be
verified by conducting a reference toxicant test concurrently with the
effluent toxicity test. The supplier should provide data with the shipment
describing the history of the sensitivity of organisms from the same source
culture, determined in monthly tests using a suitable reference toxicant.
4.7.2 The supplier should also certify the species identification of the test
organisms, and provide the taxonomic reference (citation and page) or name(s)
of the taxonomic expert(s) consulted.
8
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4.7.3 If the laboratory performing toxicity tests maintains its own stock
cultures, the sensitivity of the offspring should be determined in a toxicity
test performed with a reference toxicant at least once each month (see
Paragraph 4.15). If preferred, this reference toxicant test may be performed
concurrently with each effluent toxicity test. However, if a given species of
test organism produced by inhouse cultures is used only monthly, or less
frequently, in effluent toxicity tests, a reference toxicant test must be
performed concurrently with effluent toxicity test (See Section 6 for further
information on test organisms).
4.8 FOOD QUALITY
4.8.1 The nutritional quality of the food used in culturing and testing fish
and invertebrates is an important factor in the quality of the toxicity test
data. This is especially true for the Cen'odaphm'a diet and the unsaturated
fatty acid content of Artetm'a nauplii used in mysid culturing. Suitable trout
chow, Artemia, and other foods must be obtained as described in this manual.
4.8.2 Problems with the nutritional suitability of the food will be reflected
in the survival, growth, and reproduction of the test organisms in cultures
and toxicity tests. If a batch of food is suspected to be defective, the
performance of organisms fed with the new food can be compared with the
performance of organisms fed with a food of known quality in side-by-side
tests. If the food is used for culturing, its suitability should be
determined using a short-term chronic test which will determine the affect of
food quality on growth or reproduction of each of the relevant test species in
culture, using four replicates with each food source. Where applicable, foods
used only in acute toxicity tests can be compared with a food of known quality
in side-by-side, multi-concentration acute tests, using the reference toxicant
regularly employed in the laboratory QA program.
4.8.3 New batches of food used in culturing and testing should also be
analyzed for toxic organics and metals. If the concentration of total organic
chlorine exceeds 0.15 ug/g wet weight, or the total concentration of
organochlorine pesticides plus PCBs exceeds 0.30 ug/g wet weight, or toxic
metals exceed 20 ug/g wet weight, the food should not be used (for analytical
methods see USEPA, 1979b, 1982).
4.9 ACCEPTABILITY OF ACUTE TOXICITY TEST RESULTS
4.9.1 For the test results to be acceptable, control survival must equal or
exceed 90%.
4.9.2 An individual test may be conditionally acceptable if temperature, DO,
and other specified conditions fall outside specifications, depending on the
degree of the departure and the objectives of the tests (see test condition
summaries). The acceptability of the test will depend on the experience and
professional judgment of the laboratory analyst and the reviewing staff of the
regulatory authority. Any deviation from test specifications must be noted
when reporting data from a test.
-------
4.10 ANALYTICAL METHODS
4.10.1 All routine chemical and physical analyses for culture and dilution
water, food, and test solutions, must include established quality assurance
practices outlined in Agency methods manuals (USEPA, 1979a,b).
4.10.2 Reagent containers should be dated when received from the supplier,
and the shelf life should not be exceeded. Also, working solutions should be
dated when prepared, and the recommended shelf life should be observed.
4.11 CALIBRATION AND STANDARDIZATION
4.11.1 Instruments used for routine measurements of chemical and physical
parameters such as pH, DO, temperature, conductivity, salinity, alkalinity,
and hardness, must be calibrated and standardized prior to use each day
according to the instrument manufacturer's procedures as indicated in the
general section on quality assurance (see EPA Methods 150.1, 360.1, 170.1, and
120.1, USEPA, 1979b). Calibration data are recorded in a permanent log.
4.11.2 Wet chemical methods used to measure hardness, alkalinity, and total
residual chlorine, must be standardized prior to use each day according to the
procedures for those specific EPA methods (see EPA Methods 130.2 and 310.1,
USEPA, 1979b).
4.12 REPLICATION AND TEST SENSITIVITY
4.12.1 The sensitivity of toxicity tests will depend in part on the number of
replicates per concentration, the probability level selected, and the type of
statistical analysis. If the variability remains constant, the sensitivity of
the test will increase as the number of replicates is increased. The minimum
recommended number of replicates varies with the objectives of the test and
the statistical method used for analysis of the data.
4.13 VARIABILITY IN TOXICITY TEST RESULTS
4.13.1 Factors which can affect test success and precision include: the
experience and skill of the laboratory analyst; test organism age, condition,
and sensitivity; dilution water quality; temperature control; and the quality
and quantity of food provided. The results will depend upon the species used
and the strain or source of the test organisms, and test conditions such as
temperature, DO, food, and water quality. The repeatability or precision of
toxicity tests is also a function of the number of test organisms used at each
toxicant concentration. Jensen (1972) discussed the relationship between
sample size (numbers of fish) and the standard error of the test, and
considered 20 fish per concentration as optimum for Probit Analysis.
4.13.2 Test precision can be estimated by using the same strain of organisms
under the same test conditions, and employing a known toxicant, such as a
reference toxicant. The single-laboratory (intra-laboratory) and
multi-laboratory (inter-laboratory) precision of acute toxicity tests with
several common test species and reference toxicants are listed in
Tables 1-4. Intra- and inter-laboratory precision are described by the
10
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mean, standard deviation, and relative standard deviation (percent coefficient
of variation, or CV) of the calculated endpoints from the replicated tests.
4.13.3 Intra-laboratory precision data from 268 acute toxicity tests with
four species and five reference toxicants are listed in Tables 1 and 2. The
precision, expressed as CV%, ranged from 9% to 120%. More recent CV values
reported by Jop et al. (1986), Dorn and Rogers (1989), Hall et al. (1989), and
Cowgill et al. (1990), fell in a somewhat lower range (8% to 41%).
4.13.4 Inter-laboratory precision of acute toxicity tests from 253 reference
toxicant tests with seven species, are listed in Tables 2, 3, 4, and 5
(expressed as CV%), ranged from 12% to 167%.
4.13.5 No clear pattern of differences were noted in the intra- or inter-
laboratory test precision with the species listed, although the test results
with some toxicants, such as cadmium, appear to more variable than those with
other reference toxicants.
4.13.6 Additional information on toxicity test precision is provided in the
Technical Support Document for Water Quality-Based Toxics Control (see pp. 3-
4, and 11-15, USEPA 1991d).
4.14 DEMONSTRATING ACCEPTABLE LABORATORY PERFORMANCE
4.14.1 It is a laboratory's responsibility to demonstrate its ability to
obtain consistent, precise results with reference toxicants before it performs
toxicity tests with effluents for permit compliance purposes. To meet this
requirement, the intra-laboratory precision, expressed as percent coefficient
of variation (CV%), of each type of test to be used in a laboratory should be
determined by performing five or more tests with different batches of test
organisms, using the same reference toxicant, at the same concentrations, with
the same test conditions (i.e., the same test duration, type of dilution
water, age of test organisms, feeding, etc.), and same data analysis methods.
A reference toxicant concentration series (0.5 or higher) should be selected
that will consistently provide partial mortalities at two or more
concentrations.
4.15 DOCUMENTING ONGOING LABORATORY PERFORMANCE
4.15.1 Satisfactory laboratory performance on an ongoing basis is
demonstrated by conducting at least one acceptable test per month with a
reference toxicant for each toxicity test method commonly used in the
laboratory. The requirement for acute toxicity data can be satisfied by
determining the 48-h LC50 from a routine, monthly, short-term chronic test in
which one or more higher (acutely toxic) concentrations of reference toxicant
have been added. For a given test method, successive tests must be performed
with the same reference toxicant, at the same concentrations, in the same type
of dilution water, using the same data analysis method.
4.15.2 A control chart is prepared for each reference-toxicant-organism
combination, and successive test endpoints (LCSOs) are plotted and examined to
11
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determine if the results are within prescribed limits (Figure 1A). In this
technique, a running plot is maintained for the endpoints (Xs) from successive
tests with a given reference-toxicant-organism combination. The type of
control chart illustrated (USEPA, 1979a) is used to evaluate the cumulative
trend of the statistics from a series of such tests. For point estimation
techniques, the cumulative mean (X) and upper and lower control limits of two
standard deviations (+ 2S) are re-calculated with each successive test result.
Precision may vary with the test species, reference toxicant, and type of
test. Examples of control charts for fathead minnow and Cen'odaphm'a acute
toxicity tests are illustrated in Figures IB and 1C.
4.15.3 Outliers, which are values falling outside the upper and lower control
limits, and trends of increasing or decreasing sensitivity are readily
identified. If the toxicity value from a given test falls well outside the
"expected" range, the sensitivity of the organisms and the credibility of the
test results are suspect. In this case, the test procedure should be examined
for defects and should be repeated with a different batch of test organisms.
4.15.4 Performance should improve with experience, and the control limits
should gradually narrow (see Figure Ib), as the statistics stabilize.
However, control limits of + 2S, by definition, will be exceeded 5% of the
time, regardless of how well a laboratory performs. For this reason, good
laboratories which develop very narrow control limits may be penalized if a
test result which falls just outside the control limits is rejected de facto.
The width of the control limits should be considered in determining if data
which exceed control limits should be rejected.
4.16 REFERENCE TOXICANTS
4.16.1 Reference toxicants such as NaCl, KC1, cadmium (CdCl2), copper (CuS04),
SDS (sodium dodecyl sulfate), and K2Cr207, are suitable for use in the NPDES
and other Agency programs requiring aquatic toxicity tests. EMSL-Cincinnati
plans to release EPA-certified solutions of cadmium and copper, with
accompanying toxicity data for the recommended test species, for use as
reference toxicants in FY-92, through cooperative research and development
agreements with commercial suppliers, and will continue to develop additional
reference toxicants for future release. Interested parties can determine the
availability of "EPA Certified" reference toxicants by checking the EPA-
Cincinnati electronic bulletin board, using a modem to access the following
telephone numbers: FTS 684-7610, or Comm. 513-569-7610. Standard reference
materials also can be obtained from commercial supply houses, or can be
prepared inhouse using reagent grade chemicals. The regulatory agency should
be consulted before reference toxicant(s) are selected and used.
4.17 RECORD KEEPING
4.17.1 Proper record keeping is important. A complete file should be
maintained for each individual toxicity test or group of tests on closely
related samples. This file should contain a record of the sample chain-of-
custody; a copy of the sample log sheet; the original bench sheets for the
test organism responses during the toxicity test(s); chemical analysis data on
the sample(s); detailed records of the test organisms used in the test(s),
12
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such as species, source, age, date of receipt, and other pertinent information
relating to their history and health; information on the calibration of
equipment and instruments; test conditions employed; and results of reference
toxicant tests. Laboratory data should be recorded on a real-time basis to
prevent the loss of information or inadvertent introduction of errors into the
record. Original data sheets should be signed and dated by the laboratory
personnel performing the tests.
4.17.2 The regulatory authority should retain records pertaining to discharge
permits. Permittees are required to retain records pertaining to permit
applications and compliance for a minimum of 3 years [40 CFR 122.41(j)(2)].
13
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TABLE 1. INTRA-LABORATORY PRECISION OF LC50S FROM STATIC ACUTE
TOXICITY TESTS WITH AQUATIC ORGANISMS USING REFERENCE
TOXICANTS
TEST ORGANISM
Pimephales promelas
Daphnia magna
Daphnia magna
Daphnia magna
Daphnia magna
Daphnia pulex
Daphnia pulex
Daphnia pulex
Daphnia pulex
Mysidopsis bah i a
REFERENCE TOXICANT'1
(96
(24
(24
(48
(48
(24
(24
(48
(48
(96
h, 21°C)^
h, 20°CK
h, 26°C)7
h, 20°C),
h, 26°cr
h, 20°C),
h, 26°o;
h, 20°C)?
h, 26°C>*
h, 25°C)5
N
9
8
10
10
9
9
10
10
9
SDS
LC50
8.6
20.9
12.9
13.5
10.8
18.4
13.9
12.6
10.2
CV(%)
20
28
48
29
33
23
25
32
36
N
12
10
9
10
9
9
9
9
8
NAPCP
LC50
0.14
0.69
0.67
0.42
0.48
0.64
0.62
0.48
0.47
CV(%)
40
14
25
21
23
15
25
16
32
N
9
11
9
9
8
5
10
10
6
13
CD
LC50
0.15
0.121
0.026
0.038
0.009
0.147
0.063
0.042
0.006
0.346
CV(%)
120
49
77
58
35
30
45
45
14
9
Precision expressed as percent coefficient of variation, where
CV% = (standard deviation X 100)/mean.
SDS = Sodium dodecyl (lauryl) sulfate; NAPCP = Sodium pentachlorophenate; CD = Cadmium;
N = Number of tests; toxicant concentration in mg/L.
Pimephales promelas tests were performed in soft, synthetic freshwater; total hardness,
40-48 mg/L as CaCOj, by J. Dryer, Aquatic Biology Section, EMSL-Cincinnati.
/
Daphnia data from Lewis and Horning, 1991. Tests with D. magna used hard reconstituted
water (total hardness, 180-200 mg/L as CaCOj); tests with D. pulex used moderately-hard
reconstituted water (total hardness, 80-100 mg/L as CaCOj).
Mysid tests were performed in 25 ppt salinity, natural seawater. Data were provided by
Steve Ward, Environmental Services Division, U.S. Environmental Protection Agency,
Edison, New Jersey. Personal communication, November 14, 1990.
14
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TABLE 2. INTRA- AND INTER-LABORATORY PRECISION OF ACUTE
TOXICITY TESTS WITH DAPHNIA MAGNA, USING A
STANDARD EFFLUENT1'2
INTER-LABORATORY PRECISION: INTRA-LABORATORY
LABORATORY LCSOs FROM REPLICATE TESTS PRECISION
24 H 48 H
INDUSTRY
1 14.4 4.2
11.4 4.9
2 13.9 6.8
16.6 6.1
13.7 6.1 6.4
3 11.7 3.5
17.4 7.1
GOVERNMENT
1 14.0 4.4
10.0 4.4
10.8 4.1 4.0
2 13.2 4.5
14.1 4.5
3 11.6 4.2
COMMERCIAL
1
2
3
20.1
20.1
8.9
12.3
14.8
25.4
26.4
N 20
MEAN 15.0
SD 4.75
CV% 31.6
4.9
4.7
3.7
5.6
9.0
9.1
8.6
20
5.52
1.75
31.6
—
—
3.0
3
4.47
1.75
39.1
From Table 2, p. 191, Grothe and Kimerle, 1985. Tests performed at
20°C + 2°C; dilution water hardness, 100 mg/L as CaCO^; dilution water
alkalinity, 76 mg/L as CaCOj; effluent hardness, approx. 1000 mg/L as
CaCO-,; effluent alkalinity, 310 mg/L as CaCO,; effluent dilutions -
56%, 32%, 18%, 10%, 5.6%, 3.1%, 1.7%.
LC50 expressed in percent effluent.
Intra-laboratory precision expressed as the weighted mean CV(%).
15
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TABLE 3. INTER-LABORATORY PRECISION OF ACUTE TOXICITY
TESTS WITH AQUATIC ORGANISMS, USING REFERENCE
TOXICANTS1
TEST ORGANISM
REFERENCE TOXICANT
SILVER
N LC50 CV<%)'
ENDOSULFAN
LC50 CV(%)
1. Pimephales promelas (96 h, 22°C)
96-h static test (Meas) 10 14.0 53
96-h flow-through test (Meas) 9 7.49 40
2. Oncorhyncus mykiss (96 h, 12°C)
96-h static test (Meas) 10 34.5 88
96-h flow-through test (Meas) 9 11.5 33
Daphnia magna (48 h, 20 C)
48-h static (Meas)
12 10.6 166
4. Mvsidopsis bahia (96 h, 22 C)
96-h static test (Norn) 6 210 27
96-h flow-through test (Norn) 6 251 22
96-h flow-through test (Meas) 6 192 58
5. Cyprinodon variegatus (96 h, 22°C)
96-h static test (Norn) 4 1122 35
96-h flow-through test (Norn) 5 1573 50
96-h flow-through test (Meas) 5 1216 50
12 2.03 38
12 0.96 46
12 1.15 50
12 0.40 42
11 328
51
5 0.84 62
6 1.02 58
5 0.94 167
6 2.41 37
6 1.69 46
6 0.81 46
Data for Pimephales promelas (fathead minnow), Oncorhvncus mykiss
(rainbow trout), and Daphnia magna were taken from USEPA, 1983.
Data for, Hysidopsis bahia, and Cyprinodon varieqatus (sheepshead
minnow) were taken from USEPA, 1981. Six laboratories
participated in each study. Test salinity was 28°/oo.
LCSOs expressed in ug/L.
In the studies with the freshwater organisms, the water hardness for
five of the six laboratories ranged between 36 and 75 mg/L. However,
the water hardness for the sixth laboratory was 255 mg/L, resulting in
LC50 values for silver more than an order of magnitude larger
than for the other five. These values were rejected in calculating
the CV%. The mean weights of test fish were from 0.05-0.26 g for fathead
minnows, and 0.22-1.32 g for rainbow trout. Daphnia were < 24-h old.
In studies with the marine organisms, only one LC50 (presumably the
combined LC50 from duplicate tests) was reported for each toxicity
test. LCSOs for flow-through tests with Mysidopsis bahia and Cyprinodon
varieqatus were calculated two different ways -- (1) on the basis of
the nominal toxicant concentrations (Norn), and (2) on the basis of
measured (Meas) toxicant concentrations. Test organism age was
< 2 days for Mvsidopsis bahia, and 28 days for Cyprinodon varieqatus.
The salinity of test solutions was 28°/oo.
N, the total number of LC50 values used in calculating the CV(%) varied
with organism and toxicant because some data were rejected due to
water hardness, lack of concentration measurements, and/or because some
of the LCSOs were not calculable.
2CV% = Percent coefficient of variation = (standard deviation x 100)/mean.
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TABLE 4. INTERLABORATORY STUDY OF ACUTE TOXICITY TEST PRECISION, 1990:
SUMMARY OF RESPONSES USING KCL AS THE REFERENCE TOXICANT1
TEST TYPE
TEST PRECISION (CV%)2
NO. LABS GRAPH3 STAT4
SUBMITTING METHOD METHOD TOTAL5
VALID DATA N LC50 CV% N LC50 CV% N LC50 CV%
Pimephales prome I as
Pimephales prome I as
Ceriodaphnia dubia
Mysidopsis bahia
(96 h, 22°C)6 17
(24 h, 25°C)7 6
(48 h, 25°C)7 11
(96 h, 22°C)8 14
6 944 28.8 13 832 27.8 17 864 29.6
6 832 11.5 6 832 11.5
11 256 53.1 11 264 48.5
7 292 32.9 11 250 36.0 14 268 37.3
Inter-laboratory study of toxicity test precision conducted in 1990 by the Environmental Monitoring
Systems Laboratory - Cincinnati, U.S. Environmental Protection Agency, Cincinnati, Ohio 45268, in
cooperation with the states of New Jersey and North Carolina, and the Office of Water Enforcement
and Permits, U.S. Environmental Protection Agency, Washington, DC.
Percent coefficient of variation = (standard deviation X 100)/mean. Calculated for LC50 from acute
tests. LCSOs expressed as mg/L KCl added to the dilution water.
LC50 estimated by the Graphical Method.
LC50 estimated by Probit, Litchfield-WiIcoxon, or Trimmed Spearman-Karber method.
LC50 usually reported for only one method of analysis for each test. Where more than one LC50 was
reported for a test, the lowest value was used to calculate the statistics for "Total."
Data from the New Jersey Department of Environmental Protection: static daily-renewal tests, using
moderately-hard synthetic freshwater.
Data from North Carolina certified laboratories: static non-renewal tests, using moderately-hard
reconstituted freshwater.
Data from the New Jersey Department of Environmental Protection: static daily-renewal tests, using
25 ppt salinity, FORTY FATHOMS synthetic seawater.
17
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TABLE 5. NATIONAL INTERLABORATORY STUDY OF ACUTE TOXICITY
TEST PRECISION, 1991: SUMMARY OF RESPONSES USING
REFERENCE TOXICANTS
TEST TYPE
Pimephales p_
Ceriodaphnia
rome I as
dubia
Mysidopsis bahia
Menidia
beryllina
(48
(48
(48
(48
h,
h.
h,
h.
25°(
25°(
25°(
25°(
,}3
,j3
,}5
,}5
NO. LABS
SUBMITTING
DATA
203
171
61
39
LC50
CV%2
8964
432
532
164
4
4
6
28
39
30
42
.6
.8
.1
.2
1
performed in 1991 by the Environmental Monitoring Systems Laboratory -
Cincinnati, U.S. Environmental Protection Agency, Cincinnati, Ohio 45268.
Participants included Federal, state, and private laboratories engaged in
NPDES permit compliance monitoring. LCSOs were estimated by the graphical
or Spearman-Karber method.
Percent coefficient of variation = (standard deviation X 100)/mean.
Static non-renewal tests, using moderately-hard synthetic freshwater
(total hardness = 80-100 mg/L as CaCOj).
Expressed as mg KCl added per liter of dilution water.
Static non-renewal tests, using 30 ppt modified GP2 artificial seawater.
Expressed as ug Cu added per liter of dilution water.
18
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LT3
CJ
UPPER CONTROL LIMIT(X+ 2S)
CENTRALTENDENCY
LOWER CONTROL LI MIT (X - 2S)
05 10 15 20
TOXICITY TEST WITH REFERENCE TOXICANTS
O
12
10
O 9
U)
3 •
i i i _. i i t i i
246 8101214161820222426283032343638404244464850
3.0
O
§2.0
O
1.5
2 4 6 8 10 12 14 16 18 20 22 24 26 28 30 32 34 36 38 40
Cumulative Test Number
Figure 1. Control (cusum) charts: A, General case; B and C, 48-h acute
tests with sodium chloride. (B) Fathead minnow (Pimephales
promelas), and (C) Ceriodaphnia dubia, with the individual LCBOs
(triangles), cumulative LC50 means (dotted line), and upper and
lower control limits of two standard deviations (squares).
(Provided by the Environmental Services Division, U.S.
Environmental Protection Agency, Kansas City, Kansas).
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SECTION 5
FACILITIES AND EQUIPMENT
5.1 GENERAL REQUIREMENTS
5.1.1 Effluent toxicity tests may be performed in a fixed or mobile
laboratory. Facilities should include equipment for rearing and/or holding
organisms.
5.1.2 The facilities must be well ventilated and free of toxic fumes. Sample
preparation, culturing, and toxicity testing areas should be separated to
avoid cross contamination of cultures or toxicity test solutions with toxic
fumes. Laboratory ventilation systems should be checked to ensure that return
air from chemistry laboratories and/or sample handling areas is not circulated
to test organism culture rooms or toxicity test rooms, or that air from
toxicity test rooms does not contaminate culture areas. Air pressure
differentials between such rooms should not result in a net flow of
potentially contaminated air to sensitive areas through open or loosely-
fitting doors.
5.1.3 Control of test solution temperature can best be achieved using
circulating water baths, heat exchangers, or environmental chambers.
Photoperiod can be controlled using automatic timers in the laboratory or
environmental chambers.
5.1.4 Water used for rearing, holding, and testing organisms may be diluted
mineral water, reconstituted synthetic water, ground water, surface water, or
dechlorinated tap water. Dechlorination can be accomplished by carbon
filtration, laboratory water conditioning units, or the use of sodium
thiosulfate. After dechlorination, total residual chlorine should be
non-detectable. Sodium thiosulfate may be toxic to the test organisms, and if
used for dechlorination, paired controls with and without sodium thiosulfate
should be incorporated in effluent toxicity tests.
5.1.4.1 A deionizing system providing 18 mega-ohm, laboratory grade water
should be provided with sufficient capacity for laboratory needs. If large
quantities of high quality deionized water are needed, it may be advisable to
supply the laboratory grade water deionizer with preconditioned water from a
Culligen", Continental , or equivalent, mixed-bed water treatment system.
5.1.5 Air used for aeration must be free of oil and fumes. Oil-free air pumps
should be used where possible. Particulates can be removed from the air using
BALSTONR Grade BX or equivalent filters (Balston, Inc., Lexington,
Massachusetts), and oil and other organic vapors can be removed using
activated carbon filters (BALSTONR, C-l filter, or equivalent).
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5.1.6 During rearing, holding, and testing, test organisms should be shielded
from external disturbances such as rapidly changing light conditions
(especially salmonids) and pedestrian traffic.
5.1.7 Materials used for exposure chambers, tubing, etc., that come in
contact with the effluent and dilution water should be carefully chosen.
Tempered glass and perfluorocarbon plastics (TEFLONR) should be used whenever
possible to minimize sorption and leaching of toxic substances, and may be
reused after cleaning. Containers made of plastics, such as polyethylene,
polypropylene, polyvinyl chloride, TYGONR, etc., may be used to ship, store,
and transfer effluents and receiving waters, but they should not be reused
unless absolutely necessary, because they could carry over adsorbed toxicants
from one test to another. However, these containers may be repeatedly reused
for storing uncontaminated waters such as deionized or laboratory-prepared
dilution waters and receiving waters. Glass or disposable polystyrene
containers can be used as test chambers. The use of large (> 20 L) glass
carboys is discouraged for safety reasons.
5.1.8 New plastic products should be tested for toxicity before general use
by exposing organisms to them under ordinary test conditions.
5.1.9 Equipment which cannot be discarded after each use because of cost,
must be decontaminated according to the cleaning procedures listed below.
Fiberglass, in addition to the previously mentioned materials, can be used for
holding and dilution water storage tanks, and in the water delivery system.
All material should be flushed or rinsed thoroughly with dilution water before
using in the test.
5.1.10 Copper, galvanized material, rubber, brass, and lead must not come in
contact with holding or dilution water, or with effluent samples and test
solutions. Some materials, such as neoprene rubber (commonly used for
stoppers), may be toxic and should be tested before use.
5.1.11 Silicone adhesive used to construct glass test chambers absorbs some
organochlorine and organophosphorus pesticides, which are difficult to remove.
Therefore, as little of the adhesive as possible should be in contact with
water. Extra beads of adhesive inside the containers should be removed.
5.2 CLEANING TEST CHAMBERS AND LABORATORY APPARATUS
5.2.1 New plasticware used for effluent or dilution water collection or
organism test chambers does not require thorough cleaning before use. It is
sufficient to rinse new sample containers once with sample before use. New,
disposable, plastic test chambers may have to be rinsed with dilution water
before use. New glassware, however, must be soaked overnight in 10% acid (see
below).
5.2.2 All non-disposable sample containers, test vessels, tanks, and other
equipment that has come in contact with effluent must be washed after use in
the manner described below to remove surface contaminants.
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1. Soak 15 min in tap water, and scrub with detergent, or clean in an
automatic dishwasher.
2. Rinse twice with tap water.
3. Carefully rinse once with fresh, dilute (10%, V:V) hydrochloric or
nitric acid to remove scale, metals, and bases. To prepare a 10%
solution of acid, add 10 ml of concentrated acid to 90 ml of deionized
water.
4. Rinse twice with deionized water.
5. Rinse once with full-strength, pesticide-grade acetone to remove
organic compounds (use a fume hood or canopy).
6. Rinse three times with deionized water.
5.2.3 All test chambers and equipment should be thoroughly rinsed with the
dilution water immediately prior to use in each test.
5.3 APPARATUS AND EQUIPMENT FOR CULTURIN6 AND TOXICITY TESTS
5.3.1 Culture units -- see Appendix. It is preferable to obtain test
organisms from in-house culture units. If it is not feasible to maintain
cultures in-house, test organisms can be obtained from commercial sources, and
should be shipped to the laboratory in well oxygenated water in insulated
containers to minimize excursions in water temperature during shipment.
The temperature of the water in the shipping containers should be measured on
arrival, to determine if the organisms were subjected to obvious undue thermal
stress.
5.3.2 Samplers -- automatic samplers, preferably with sample cooling
capability, that can collect a 24-h composite sample of 2 L or more.
5.3.3 Sample containers -- for sample shipment and storage (see Section 8,
Effluent and Receiving Water Sampling and Sample Handling).
5.3.4 Environmental chamber or equivalent facility with temperature control
(20°C or 25°C)
5.3.5 Water purification system -- MILLIPORE MILLI-QR, SUPER-QR or
equivalent. Depending on the quantity of high grade water needed, a first-
stage pre-conditioner deionizer, such as a CULLIGENR or CONTINENTAL System,
or equivalent, may be needed to provide feed water to the high-purity system.
5.3.6 Balance -- analytical, capable of accurately weighing to 0.0001 g.
5.3.7 Reference weights, Class S -- for documenting the performance of the
analytical balance(s). The balance(s) should be checked with reference
weights which are at the upper and lower ends of the range of the weighings
made when the balance is used. A balance should be checked at the beginning
of each series of weighings, periodically (such as every tenth weight) during
a long series of weighings, and after the last weight of a series is taken.
5.3.8 Test chambers -- borosilicate glass or non-toxic disposable plastic
test chambers are suitable. Test chamber volumes are indicated in the method
summaries. To avoid potential contamination from the air and excessive
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evaporation of test solutions during the test, the chambers should be covered
with safety glass plates or sheet plastic, 6 mm (1/4 in) thick.
5.3.9 Volumetric flasks and graduated cylinders -- Class A, borosilicate
glass or non-toxic plastic labware, 10-1000 mL for making test solutions.
5.3.10 Volumetric pipets -- Class A, 1-100 ml.
5.3.11 Serological pipets -- 1-10 ml, graduated.
5.3.12 Pipet bulbs and fillers -- PROPIPETR, or equivalent.
5.3.13 Droppers, and glass tubing with fire polished edges, 4 mm ID -- for
transferring test organisms.
5.3.14 Wash bottles -- for rinsing small glassware and instrument electrodes
and probes.
5.3.15 Glass or electronic thermometers -- for measuring water temperature.
5.3.16 Bulb-thermograph or electronic-chart type thermometers -- for
continuously recording temperature.
5.3.17 National Bureau of Standards Certified thermometer (see USEPA Method
170.1, USEPA 1979b).
5.3.18 pH, DO, and specific conductivity meters -- for routine physical and
chemical measurements. Unless the test is being conducted to specifically
measure the effect of one of the above parameters, a portable, field-grade
instrument is acceptable.
5.3.19 Refractometer -- for measuring effluent, receiving, and test solution
salinity.
5.3.20 Amperometric titrator -- for measuring total residual chlorine.
5.4 REAGENTS AND CONSUMABLE MATERIALS
5.4.1 Reagent water -- defined as MILLIPORE MILLI-QR or equivalent water (see
paragraph 5.3.5 above).
5.4.2 Effluent, dilution water, and receiving water -- see Section 7,
Dilution Water, and Section 8, Effluent and Receiving Water Sampling and
Sample Handling.
5.4.3 Reagents for hardness and alkalinity tests (see USEPA Methods 130.2 and
310.1, USEPA 1979b).
5.4.4 Standard pH buffers 4, 7, and 10 (or as per instructions of instrument
manufacturer) for instrument calibration (see USEPA Method 150.1, USEPA
19795).
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5.4.5 Specific conductivity and salinity standards (see USEPA Method 120.1,
USEPA 1979b).
5.4.6 Laboratory quality control check samples and standards for the above
chemistry methods.
5.4.7 Reference toxicant solutions (see Section 4, Quality Assurance).
5.4.8 Membranes and filling solutions for dissolved oxygen probe (see USEPA
Method 360.1, USEPA 1979b), or reagents for modified Winkler analysis.
5.4.9 Sources of Food for Cultures and Toxicity Tests.
5.4.9.1 All food should be tested for nutritional suitability, and chemically
analyzed for organic chlorine, PCBs, and toxic metals (see Section 4).
5.4.9.2 Brine Shrimp (Artemia) -- see Appendix A.
1. Brine Shrimp (Artemia) Cysts.
There are many commercial sources of brine shrimp cysts. Sources
include: Aquarium Products, 180L Penrod Ct., Glen Burnie, Maryland
21061); San Francisco Bay Brand, 8239 Enterprise Dr., Newark,
California, 94560 (Phone: 415-792-7200); Argent Aquaculture, 8702 152nd
Ave, N.E., Redmond, Washington, 98052 (206-885-3777)(Argentina brine
shrimp eggs, Grade 1, Gold Label); and Jungle, Inc. Additional sources
are listed in the section on Artemia culture in Appendix A. The
quality of the cysts may vary from one batch to another, and the cysts
in each new batch (can or lot) should be evaluated for nutritional
suitability and chemical contamination. The nutritional suitability
(see Leger et al., 1985, 1986) of each new batch is checked against
known suitable reference cysts by performing a side-by-side growth
and/or reproduction tests using the "new" and "reference" cysts. If
the results of tests for nutritional suitability or chemical
contamination do not meet standards, the Artemia should not be used.
2. Frozen Adult Brine Shrimp
Frozen adult brine shrimp are available from San Francisco Bay Brand,
8239 Enterprise Dr., Newark, California, 94560 (415-792-7200).
5.4.9.3 Trout Chow
5.4.9.3.1 Starter or No. 1 pellets, prepared according to current U.S. Fish
and Wildlife Service specifications, are available from: Zeigler Bros., Inc.,
P.O. Box 95, Gardners, Pennsylvania, 17324 (717-780-9009); Glencoe Mills, 1011
Elliott, Glencoe, Minnesota, 55336 (612-864-3181); and Murray Elevators, 118
West 4800 South, Murray, Utah 84107 (800-521-9092). (The flake food,
TETRAMINR or BIORILR, can be used regularly as a substitute for trout chow in
preparing food for daphnids, and can be used as a short-term substitute for
trout chow in feeding fathead minnows.)
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5.4.9.4 Dried, Powdered Leaves (CEROPHYLLR)
5.4.9.4.1 Dried, powdered, cereal leaves are available from Sigma Chemical
Company, P.O. Box 14508, St. Louis, Missouri, 63178, (800-325-3010); or
as CEROPHYLL*, from Ward's Natural Science Establishment, Inc., P.O. Box
92912, Rochester, New York, 14692-9012, (716-359-2502). Dried, powdered,
alfalfa leaves obtained from health food stores have been found to be a
satisfactory substitute for cereal leaves.
5.4.9.5 Yeast
5.4.9.5.1 Packaged dry yeast, such as Fleischmann's, or equivalent, can be
purchased at the local grocery store.
5.4.9.6 Flake Food
5.4.9.6.1 The flake foods, TETRAMIN* and BIORILR, are available at most pet
supply shops.
5.5 TEST ORGANISMS
5.5.1 Test organisms are obtained from inhouse cultures or commercial
suppliers (see Section 6).
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SECTION 6
TEST ORGANISMS
6.1 TEST SPECIES
6.1.1 The species used in characterizing the acute toxicity of effluents
and/or receiving waters will depend on the requirements of the regulatory
authority and the objectives of the test. It is essential that good
quality test organisms be readily available throughout the year from
inhouse or commercial sources to meet NPDES monitoring requirements. The
organisms used in toxicity tests must be identified to species. If there
is any doubt as to the identity of the test organisms, they should be
sent to a taxonomic expert for examination.
6.1.2 Toxicity test conditions and culture methods are provided in this
manual for the following principal test organisms:
Freshwater Organisms:
1. Ceriodaphm'a dubia (daphnid) (Table 11).
2. Daphm'a pulex and D. magna (daphnids) (Table 12).
3. Pimephales promelas (fathead minnow) (Table 13).
4. Oncorhynchus mykiss (rainbow trout) and Salvelinus
fontinalis (brook trout) (Table 14).
Estuarine and Marine Organisms:
1. Mysidopsis bahia (mysid) (Table 15).
2. Cyprinodon variegatus (sheepshead minnow) (Table 16).
3. Menidia beryl Una (inland silverside),
M. mem'dia (Atlantic silverside),
and M. peninsulae (tidewater silverside)(Table 17).
6.1.3 The test species listed in Section 6.1.2 are the recommended acute
toxicity test organisms. They are easily cultured in the laboratory, are
sensitive to a variety of pollutants, and are generally available
throughout the year from commercial sources. Summaries of test
conditions for these species are provided in Tables 11-17. Guidelines for
culturing and/or holding the organisms are provided in Appendix A.
6.1.4 Additional species may be suitable for toxicity tests in the NPDES
Program. A list of alternative acute toxicity test species and minimal
testing requirements (i.e., temperature, salinity, and life stage) for
these species are provided in Appendix B. It is important to note that
these species may not be as easily cultured or tested as the species on
the list in 6.1.2, and may not be available from commercial sources.
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6.1.5 Some states have developed culturing and testing methods for
indigenous species that may be as sensitive or more sensitive than the
species recommended in 6.1.2. However, EPA allows the use of indigenous
species only where state regulations require their use or prohibit
importation of the species in 6.1.2. Where state regulations prohibit
importation or use of the recommended test species, permission must be
requested from the appropriate state agency prior to their use.
6.1.6 Where states have developed culturing and testing methods for
indigenous species other than those recommended in this manual, data
comparing the sensitivity of the substitute species and one or more of
the recommended species must be obtained in side-by-side toxicity tests
with reference toxicants and/or effluents, to ensure that the species
selected are at least as sensitive as the recommended species. These
data must be submitted to the permitting authority (State or Region) as
required by 40 CFR 136.4. EPA acknowledges that reference toxicants
prepared from pure chemicals may not always be representative of
effluents. However, because of the observed and/or potential variability
in the quality and toxicity of effluents, it is not possible to specify a
representative effluent.
6.1.7 Guidance for the selection of test organisms where the salinity of
the effluent and/or receiving water requires special consideration is
provided in the Technical Support Document for Water Quality-Based Toxics
Control (USEPA, 1991d).
1. Where the salinity of the receiving water is < l°/oo, freshwater
organisms are used regardless of the salinity of the effluent.
2. Where the salinity of the receiving water is > l°/°o, the choice of
organisms depends on state water quality standards and/or permit
requirements.
6.2 SOURCES OF TEST ORGANISMS
6.2.1 Inhouse Cultures
6.2.1.1 Inhouse cultures should be established wherever it is cost
effective. If inhouse cultures cannot be maintained, test organisms
should be purchased from experienced commercial suppliers.
6.2.2 Commercial Suppliers
6.2.2.1 All of the principal test organisms listed in Paragraph 6.1.2
are available from commercial suppliers.
6.2.3 Feral Organisms
6.2.3.1 The use of test organisms taken from the receiving water has
strong appeal, and would seem to be the logical approach. However, it is
impractical for the following reasons:
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1. Sensitive organisms may not be present in the receiving water
because of previous exposure to the effluent or other pollutants.
2. It is often difficult to collect organisms of the required age and
quality from the receiving water;
3. Most states require collection permits, which may be difficult to
obtain. Therefore, it is usually more cost effective to culture the
organisms in the laboratory or obtain them from private, state, or
Federal sources. Fish such as fathead minnows, sheepshead minnows,
and silversides, and invertebrates such as daphnids and mysids, are
easily reared in the laboratory or purchased.
4. The required QA/QC records, such as the single laboratory precision
data, would not be available.
5. Since it is mandatory that the identity of test organisms is known
to the species level, it would be necessary to examine each organism
caught in the wild to confirm its identity, which would usually be
impractical or, at the least, very stressful to the organisms.
6. Test organisms obtained from the wild must be observed in the
laboratory for a minimum of one week prior to use, to assure that
they are free of signs of parasitic or bacterial infections and
other adverse effects. Fish captured by electroshocking must not be
used in toxicity testing.
6.2.3.2 Guidelines for collection of feral organisms are provided in
USEPA, 1973, 1990a.
6.2.4 Regardless of their source, test organisms should be carefully
observed to ensure that they are free of signs of stress and disease, and
in good physical condition. Some species of test organisms, such as
trout, can be obtained from stocks certified as "disease-free."
6.3 LIFE STAGE
6.3.1 Young organisms are often more sensitive to toxicants than are
adults. For this reason, the use of early life stages, such as first
instars of daphnids and juvenile mysids and fish, is recommended for all
tests. There may be special cases, however, where the limited
availability of organisms will require some deviation from the
recommended life stage. In a given test, all organisms should be
approximately the same age and should be taken from the same source.
Since age may affect the results of the tests, it would enhance the value
and comparability of the data if the same species in the same life stages
were used throughout a monitoring program at a given facility.
6.4 LABORATORY CULTURING
6.4.1 Instructions for culturing and/or holding the recommended test
organisms are included in Appendix A.
6.5 HOLDING AND HANDLING TEST ORGANISMS
6.5.1 Test organisms should not be subjected to changes of more than 3°C
in water temperature or 3 °/oo in salinity in any 12 h period.
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6.5.2 Organisms should be handled as little as possible. When handling
is necessary, it should be done as gently, carefully, and quickly as
possible to minimize stress. Organisms that are dropped or touch dry
surfaces or are injured during handling must be discarded. Dipnets are
best for handling larger organisms. These nets are commercially
available or can be made from small-mesh nylon netting, silk bolting
cloth, plankton netting, or similar material. Wide-bore, smooth glass
tubes (4 to 8 mm inside diameter) with rubber bulbs or pipettors (such as
a PROPIPETTER or other pipetter) should be used for transferring smaller
organisms such as daphnids, mysids, and larval fish.
6.5.3 Holding tanks for fish are supplied with a good quality water (see
Section 5) with a flow-through rate of at least two tank-volumes per day.
Otherwise, use a recirculation system where the water flows through an
activated carbon or undergravel filter to remove dissolved metabolites.
Culture water can also be piped through high intensity ultraviolet light
sources for disinfection, and to photodegrade dissolved organics.
6.5.4 Crowding should be avoided. The DO must be maintained at a minimum
of 4.0 mg/L for marine and warm water, freshwater species, and 6.0 mg/L
for cold-water, freshwater species. The solubility of oxygen depends on
temperature, salinity, and altitude. Aerate if necessary.
6.5.5 Fish should be fed as much as they will eat at least once a day
with live or frozen brine shrimp or dry food (frozen food should be
completely thawed before use). Brine shrimp can be supplemented with
commercially prepared food such as Tetramin or BioRil flake food, or
equivalent. Excess food and fecal material should be removed from the
bottom of the tanks at least twice a week by siphoning.
6.5.6 Fish should be observed carefully each day for signs of disease,
stress, physical damage, and mortality. Dead and abnormal specimens
should be removed as soon as observed. It is not uncommon to have some
fish (5-10%) mortality during the first 48 h in a holding tank because of
individuals that refuse to feed on artificial food and die of starvation.
6.5.7 A daily record of feeding, behavioral observations, and mortality
should be maintained.
6.6 TRANSPORTATION TO THE TEST SITE
6.6.1 Organisms are transported from the base or supply laboratory to a
remote test site in culture water or standard dilution water in plastic
bags or large-mouth screw-cap (500 mL) plastic bottles in styrofoam
coolers. Adequate DO is maintained by replacing the air above the water
in the bags with oxygen from a compressed gas cylinder, and sealing the
bags. Another method commonly used to maintain sufficient DO during
shipment is to aerate with an airstone which is supplied from a portable
pump. The DO concentration must not fall below 4.0 mg/L for marine and
warm-water, freshwater species, and 6.0 mg/L for cold-water, freshwater
species.
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6.6.2 Upon arrival at the test site, organisms are transferred to
receiving water if receiving water is to be used as the test dilution
water. All but a small volume of the holding water (approximately 5%) is
removed by siphoning, and replaced slowly over a 10 to 15 min period with
dilution water. If receiving water is used as dilution water, caution
must be exercised in exposing the test organisms to it, because of the
possibility that it might be toxic. For this reason, it is recommended
that only approximately 10% of the test organisms be exposed initially to
the dilution water. If this group does not show excessive mortality or
obvious signs of stress in a few hours, the remainder of the test
organisms are transferred to the dilution water.
6.6.3 A group of organisms must not be used for a test if they appear to
be unhealthy, discolored, or otherwise stressed, or if mortality appears
to exceed 10% preceding the test. If the organisms fail to meet these
criteria, the entire group must be discarded and a new group obtained.
The mortality may be due to the presence of toxicity, if receiving water
is used as dilution water, rather than a diseased condition of the test
organisms. If the acclimation process is repeated with a new group of
test organisms and excessive mortality occurs, it is recommended that an
alternative source of dilution water be used.
6.6.4 In static tests, marine organisms can be used at all
concentrations of effluent by adjusting the salinity of the effluent to a
standard salinity (such as 25 °/oo) or to the salinity approximating that
of the receiving water, by adding sufficient dry ocean salts, such as
Forty Fathoms", or equivalent, or hypersaline brine.
6.6.5 Saline dilution water can be prepared with deionized water or a
freshwater such as well water or a suitable surface water. If dry ocean
salts are used, care must be taken to ensure that the added salts are
completely dissolved and the solution is aerated 24 h before the test
organisms are placed in the solutions. The test organisms should be
acclimated in synthetic saline water prepared with the dry salts.
Caution: addition of dry ocean salts to dilution water may result in an
increase in pH. (The pH of estuarine and coastal saline waters is
normally 7.5 - 8.3.)
6.6.6 All effluent concentrations and the control(s) used in a test
should have the same salinity. However, if this is impractical because
of the large volumes of water required, such as in flow-through tests,
the highest effluent concentration (lowest salinity) that could be tested
would depend upon the salinity of the receiving water and the tolerance
of the test organisms. The required salinities for toxicity tests with
estuarine and marine species are listed in Tables 15-17. However, the
tolerances of other candidate test species would have to be determined by
the investigator in advance of the test.
6.6.7 Because of the circumstances described above, when performing
flow-through tests of effluents discharged to saline waters, it is
advisable to acclimate groups of test organisms to each of three
different salinities, such as 10, 20, and 30 700, prior to transporting
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them to the test site. It may also be advisable to maintain cultures of
these test organisms at a series of salinity levels, including at least
10, 20, and 30 °/oo, so that the change in salinity upon acclimation at
the desired test dilutions does not exceed 6 °/oo.
6.7 TEST ORGANISM DISPOSAL
6.7.1 Organisms used in a toxicity test are sacrificed when removed from
the test chambers.
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SECTION 7
DILUTION WATER
7.1 TYPES OF DILUTION WATER
7.1.1 The type of dilution water used in effluent toxicity tests will depend
largely on the objectives of the study:
1. If the objective of the test is to estimate the absolute acute
toxicity of the effluent, which is a primary objective of NPDES
permit-related toxicity testing, a synthetic (standard) dilution water
is used. If the test organisms have been cultured in water which is
different from the test dilution water, a second set of controls, using
culture water, should be included in the test.
2. If the objective of the test is to estimate the acute toxicity of
the effluent in uncontaminated receiving water, the test may be
conducted using dilution water consisting of a single grab sample of
receiving water (if non-toxic), collected either upstream and outside
the influence of the outfall, or with other uncontaminated surface
water or standard dilution water having approximately the same
characteristics (hardness or salinity) as the receiving water.
Seasonal variations in the quality of surface waters may affect
effluent toxicity. Therefore, the hardness of fresh receiving water,
and the salinity of saline receiving water samples should be determined
before each use. If the test organisms have been cultured in water
which is different from the test dilution water, a second set of
controls, using culture water, should be included in the test.
3. If the objective of the test is to determine the additive or mitigating
effects of the discharge on already contaminated receiving water, the
test is performed using dilution water consisting of receiving water
collected immediately upstream or outside the influence of the outfall.
A second set of controls, using culture water, should be included in
the test.
7.2 STANDARD, SYNTHETIC DILUTION WATER
7.2.1 Standard, synthetic, dilution water is prepared with deionized water
and reagent grade chemicals or mineral water (Tables 6-8). The source water
for the deionizer can be ground water, surface water, or tap water.
7.2.2 Deionized Water used to Prepare Standard, Synthetic, Dilution Water
7.2.2.1 Deionized water is obtained from a MILLIPORE MILLI-QR or equivalent
system. It is advisable to provide a preconditioned (deionized) feed
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water by using a Culligan, Continental, or equivalent, system in front of the
MILLI-Q* System to extend the life of the MILLI-QR cartridges.
7.2.2.2 The recommended order of the cartridges in a four-cartridge deionizer
(i.e., MILLI-QR System or equivalent) is: (1) ion exchange, (2) ion exchange,
(3) carbon, and (4) organic cleanup (such as ORGANEX-QR, or equivalent),
followed by a final bacteria filter.
7.2.3 Standard, Synthetic Freshwater
7.2.3.1 To prepare 20 L of standard, synthetic, moderately hard,
reconstituted water, use the reagent grade chemicals in Table 6 as follows:
1. Place 19 L of MILLI-QR, or equivalent, deionized water in a properly
cleaned plastic carboy.
2. Add 1.20 g of MgS04, 1.92 g NaHC03, and O.OSOg KC1 to the carboy.
3. Aerate overnight.
4. Add 1.20 g of CaS04 ' 2 H20 to 1 L of MILLI-QR or equivalent
deionized water in a separate flask. Stir on magnetic stirrer until
calcium sulfate is dissolved, add to the 19 L above, and mix well.
5. For Ceriodaphnia culture and testing, add sufficient sodium selenate
(Na2Se04) to provide 2 ug selenium per liter of final dilution water.
6. Aerate the combined solution vigorously for an additional 24 h to
dissolve the added chemicals and stabilize the medium.
7. The measured pH, hardness, etc., should be as listed in Table 6.
7.2.3.2 To prepare 20 L of standard, synthetic, moderately hard,
reconstituted water, using 20% mineral water such as PERRIERR Water, or
equivalent (Table 7), follow the instructions below.
1. Place 16 L of MILLI-QR or equivalent deionized water in a properly
cleaned plastic carboy.
2. Add 4 L of PERRIERR Water, or equivalent.
3. Aerate vigorously for 24 h to stabilize the medium.
4. The measured pH, hardness, and alkalinity of the aerated water will be
as indicated in Table 7.
5. This synthetic water is referred to as diluted mineral water
(DMW) in the toxicity test methods.
7.2.4 Standard, Synthetic Seawater
7.2.4.1 To prepare 20 L of a standard, synthetic, reconstituted seawater
(modified GP2), with a salinity of 31°/oo (Table 8), follow the instructions
below. Other salinities can be prepared by making the appropriate dilutions.
1. Place 20 L of MILLI-QR or equivalent deionized water in a properly
cleaned plastic carboy.
2. Weigh reagent grade salts listed in Table 8 and add, one at a time, to
the deionized water. Stir well after adding each salt.
3. Aerate the final solution at a rate of 1 L/h for 24 h.
4. Check the pH and salinity.
33
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Larger or smaller volumes of modified GP2 can be prepared by using
proportionately larger or smaller amounts of salts and dilution water.
7.2.4.2 Synthetic seawater can also be prepared by adding commercial sea
salts, such as FORTY FATHOMS", or equivalent, to deionized water. For
example, thirty-one parts per thousand (31°/oo) FORTY FATHOMS" can be prepared
by dissolving 31 g of product per liter of deionized water. The salinity of
the resulting solutions should be checked with a refractometer.
7.3 USE OF RECEIVING WATER AS DILUTION WATER
7.3.1 If the objectives of the test require the use of uncontaminated surface
water as dilution water, and the receiving water is uncontaminated, it may be
possible to collect a sample of the receiving water close to the outfall, but
upstream from or beyond the influence of the effluent. However, if the
receiving water is contaminated, it may be necessary to collect the sample in
an area "remote" from the discharge site, matching as closely as possible the
physical and chemical characteristics of the receiving water near the outfall.
7.3.2 [The sample should be collected immediately prior to the testj but never
more than 96 h before the test begins. Except where it is used within 24 h,
or in the case where large volumes are required for flow-through tests, the
sample should be chilled to 4°C during or immediately following collection,
and maintained at that temperature prior to use in the test.
7.3.3 In the case of freshwaters, the regulatory authority may require that
the hardness of the dilution water be comparable to the receiving water at the
discharge site. This requirement can be satisfied by collecting an
uncontaminated surface water with a suitable hardness, or adjusting the
hardness of an otherwise suitable surface water by addition of reagents as
indicated in Table 6.
7.3.4 In an estuarine environment, the investigator should collect
uncontaminated water having a salinity as near as possible to the salinity of
the receiving water at the discharge site. Water should be collected at slack
high tide, or within one hour after high tide. If there is reason to suspect
contamination of the water in the estuary, it is advisable to collect
uncontaminated water from an adjacent estuary. At times it may be
necessary to collect water at a location closer to the open sea, where the
salinity is relatively high. In such cases, deionized water or uncontaminated
freshwater is added to the saline water to dilute it to the required test
salinity. Where necessary, the salinity of a surface water can be increased
by the addition of artificial sea salts, such as FORTY FATHOMS* or equivalent,
a natural seawater of higher salinity, or hypersaline brine. Instructions for
the preparation of hypersaline brine by concentrating natural seawater are
provided below.
7.3.5 Receiving water containing debris or indigenous organisms, that may be
confused with or attack the test organisms, should be filtered through a sieve
having 60 urn mesh openings prior to use.
34
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TABLE 6. PREPARATION OF SYNTHETIC FRESHWATER USING REAGENT GRADE CHEMICALS1
Reaqent Added (mq/L)2
Water
Type
NaHC03
Very soft 12
Soft 48
Moderately Hard 96
Hard 192
Very hard 384
.0
.0
.0
.0
.0
CaS04.2H
7.
30.
60.
120.
240.
^Taken in part from Marking
Add reagent grade chemicals
5
0
0
0
0
and
to
_f 4. _
20 MgS04
7.5
30.0
60.0
120.0
240.0
KC1
0.5
2.0
4.0
8.0
16.0
6
7
7
7
8
PH3
.4-6.
.2-7.
.4-7.
.6-8.
.0-8.
Fi
8
6
8
0
4
nal Water Oualitv
Hardness4
10-13
40-48
80-100
160-180
280-320
Alka-
linity4
10-13
30-35
60-70
110-120
225-245
Dawson (1973).
deionized water.
'Expressed as mg CaC03/L.
TABLE 7. PREPARATION OF SYNTHETIC FRESH WATER USING MINERAL WATER1
Final Water Quality
Water
Type
Volume of
Mineral Water
Added (ml/I)2
Proportion
of Mineral
Water (%)
PH3
Hardness4
Alka-
linity4
Very soft 50
Soft 100
Moderately Hard 200
Hard 400
Very hard5 —
2.5
10.0
20.0
40.0
7.2-8.1
7.9-8.3
7.9-8.3
7.9-8.3
10-13
40-48
80-100
160-180
10-13
30-35
60-70
110-120
jFrom Mount et al., 1987; data provided by Philip Lewis, EMSL-Cincinnati.
Add mineral water to Milli-QR water or equivalent to prepare DMW (Diluted
Mineral Water).
^Approximate equilibrium pH after 24 h of aeration.
^Expressed as mg CaCO,/L.
Dilutions of PERRIER" Water form a precipitate when concentrations equivalent
to "very hard water" are aerated.
35
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TABLE 8. PREPARATION OF SYNTHETIC SEAWATER USING REAGENT GRADE CHEMICALS1'2
Compound
1.
2.
3.
4.
5.
6.
7.
8.
9.
NaCl
Na2S04
KC1
KBr
Na2B407 . 10 H20
MgCl2 . 6 H20
CaCl2 . 2 H20
SrCl2 . 6 H20
NaHC03
Concentration
(9/L)
21.03
3.52
0.61
0.088
0.034
9.50
1.32
0.02
0.17
Amount (g)
Required for
20 L
420.6
70.4
12.2
1.76
0.68
190.0
26.4
0.400
3.40
Modified GP2.
2The constituent salts and concentrations were taken from
USEPA, 1990c. The salinity is 30.89 g/L.
36
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7.3.6 When receiving water is used as dilution water in flow-through tests,
it is preferable to pump the dilution water continuously to the acclimation
chamber and/or dilutor. However, where it is not feasible to pump the
dilution water continuously, grab samples of the dilution water are
transported to the test site in tanks, and continuously pumped from the tanks
to the acclimation chamber and/or dilutor.
7.3.7 Hypersaline Brine
7.3.7.1 Hypersaline brine (HSB) has several advantages that make it desirable
for use in toxicity testing. It can be made from any high quality, filtered
seawater by evaporation, and can be added to deionized water to prepare
dilution water, or to effluents or surface waters to increase their salinity.
7.3.7.2 The ideal container for making HSB from natural seawater is one
that (1) has a high surface to volume ratio, (2) is made of a non-corrosive
material, and (3) is easily cleaned (fiberglass containers are ideal).
Special care should be used to prevent any toxic materials from coming in
contact with the seawater being used to generate the brine. If a heater is
immersed directly into the seawater, ensure that the heater materials do not
corrode or leach any substances that would contaminate the brine. One
successful method used is a thermostatically controlled heat exchanger made
from fiberglass. If aeration is used, use only oil-free air compressors to
prevent contamination.
7.3.7.3 Before adding seawater to the brine generator, thoroughly clean
the generator, aeration supply tube, heater, and any other materials that
will be in direct contact with the brine. A good quality biodegradable
detergent should be used, followed by several thorough deionized water
rinses. High quality (and preferably high salinity) seawater should be
filtered to at least 10 urn before placing into the brine generator. Water
should be collected on an incoming tide to minimize the possibility of
contamination.
7.3.7.4 The temperature of the seawater is increased slowly to 40°C.
The water should be aerated to prevent temperature stratification and to
increase water evaporation. The brine should be checked daily (depending on
the volume being generated) to ensure that the salinity does not exceed
100 °/°o and that the temperature does not exceed 40°C. Additional
seawater may be added to the brine to obtain the volume of brine required.
7.3.7.5 After the required salinity is attained, the HSB should be
filtered a second time through a 1-um filter and poured directly into
portable containers (20-L CUBITAINERS" or polycarbonate water cooler jugs are
suitable). The containers should be capped and labelled with the date the
brine was generated and its salinity. Containers of HSB should be stored in
the dark and maintained under room temperature until used.
7.3.7.6 If a source of HSB is available, test solutions can be made by
following the directions below. Thoroughly mix together the deionized water
and brine before mixing in the effluent.
37
-------
7.3.7.7 Divide the salinity of the HSB by the expected test salinity to
determine the proportion of deionized water to brine. For example, if the
salinity of the brine is 100 °/oo and the test is to be conducted at
25 °/°°> T°° °/°° divided by 25 °/oo = 4.0. The proportion of brine is
1 part in 4 (one part brine to three parts deionized water).
7.3.7.8 To make 1 L of seawater at 25 °/oo salinity from a hypersaline
brine of 100 °/oo, 250 ml of brine and 750 ml of deionized water are
required.
7.4 USE OF TAP WATER AS DILUTION WATER
7.4.1 The use of tap water as dilution water is discouraged unless it is
dechlorinated and fully treated. Tap water can be dechlorinated by
deionization, carbon filtration, or the use of sodium thiosulfate. Use of 3.6
mg/L (anhydrous) sodium thiosulfate will reduce 1.0 mg chlorine/L (APHA, 1989,
pp. 9-32). Following dechlorination, total residual chlorine should not
exceed 0.01 mg/L. Because of the possible toxicity of thiosulfate to test
organisms, a control lacking thiosulfate should be included in toxicity tests
utilizing thiosulfate-dechlorinated water.
7.4.2 To be adequate for general laboratory use following dechlorination, the
tap water is passed through a deionizer and carbon filter to remove toxic
metals and organics, and to control hardness and alkalinity.
7.5 DILUTION WATER HOLDING
7.5.1 A given batch of dilution water should not be used for more than 14
days following preparation because of the possible build-up of bacterial,
fungal, or algal slime growth and the problems associated with it. The
container should be kept covered and the contents should be protected from
light.
38
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SECTION 8
EFFLUENT AND RECEIVING WATER SAMPLING AND SAMPLE HANDLING
8.1 EFFLUENT SAMPLING
8.1.1 The effluent sampling point is ordinarily the same as that specified in
the NPDES discharge permit (USEPA, 1988c). Conditions for exception would be:
(1) better access to a sampling point between the final treatment and the
discharge outfall; (2) if the effluent is chlorinated prior to discharge to
the receiving waters, it may also be desirable to take samples prior to
contact with the chlorine to determine toxicity of the unchlorinated effluent;
or (3) in the event there is a desire to evaluate the toxicity of the influent
to publicly owned treatment works or separate process waters in industrial
facilities prior to their being combined with other process waters or
non-contact cooling water, additional sampling points may be chosen.
8.1.2 The decision on whether to collect grab or composite samples is based
on the requirements of the NPDES permit, the objectives of the test, and an
understanding of the short and long-term operations and schedules of the
discharger. If the effluent quality varies considerably with time, which can
occur where holding times within the treatment facility are short, grab
samples may seem preferable because of the ease of collection and the
potential of observing peaks (spikes) in toxicity. However, the sampling
duration of a grab sample is so short that full characterization of an
effluent over a 24-h period would require a prohibitive number of separate
samples and tests. Collection of a 24-h composite sample, however, may dilute
toxicity spikes, and average the quality of the effluent over the sampling
period. Sampling recommendations are provided below.
8.2 EFFLUENT SAMPLE TYPES
8.2.1 The advantages and disadvantages of effluent grab and composite samples
are listed below:
8.2.1.1 Grab Samples
Advantages:
1. Easy to collect; require a minimum of equipment and on-site time.
2. Provide a measure of instantaneous toxicity. Toxicity spikes are
not masked by dilution.
Disadvantages:
1. Samples are collected over a very short period of time and on a
relatively infrequent basis. The chances of detecting a spike in
toxicity would depend on the frequency of sampling, and the probability
of missing spikes is high.
39
-------
8.2.1.2 Composite Samples:
Advantages:
1. A single effluent sample is collected over a 24-h period.
2. The sample is collected over a much longer period of time than a grab
samples and contains all toxicity spikes.
Disadvantages:
1. Sampling equipment is more sophisticated and expensive, and must be
placed on-site for at least 24 h.
2. Toxicity spikes may not be detected because they are masked by dilution
with less toxic wastes.
8.3 EFFLUENT SAMPLING RECOMMENDATIONS
8.3.1. When tests are conducted on-site, test solutions can be renewed daily
with freshly collected samples.
8.3.2 When tests are conducted off-site, samples are collected once, and used
for test initiation and renewal at 48 h.
8.3.3 Sufficient sample must be collected to perform the required toxicity
and chemical tests. A 4-L (1-gal) CUBITAINERR will provide sufficient sample
volume for most tests (see Tables 11-17).
8.3.4 The following effluent sampling methods are recommended:
8.3.4.1. Continuous Discharges
1. If the facility discharge is continuous, but the calculated retention
time of the continuously discharged effluent is less than 14 days and
the variability of the effluent toxicity is unknown, at a minimum, four
grab samples or four composite samples are collected over a 24-h
period. For example, a grab sample is taken every 6 h (total of four
samples) and each sample is used for a separate toxicity test, or four
successive 6-h composite samples are taken and each is used in a
separate test.
2. If the calculated retention time of a continuously discharged effluent
is greater than 14 days, or if it can be demonstrated that the
wastewater does not vary more than 10% in toxicity over a 24-h period,
regardless of retention time, a single grab sample is collected for a
single toxicity test.
3. The retention time of the effluent in the wastewater treatment facility
may be estimated from calculations based on the volume of the retention
basin and rate of wastewater inflow. However, the calculated retention
time may be much greater than the actual time because of
short-circuiting in the holding basin. Where short-circuiting is
suspected, or sedimentation may have reduced holding basin capacity, a
40
-------
more accurate estimate of the retention time can be obtained by carrying out a
dye study.
8.3.4.2 Intermittent Discharges
8.3.4.2.1 If the facility discharge is intermittent, a grab sample is
collected midway during each discharge period. Examples of intermittent
discharges are:
1. When the effluent is continuously discharged during a single 8-h work
shift (one sample is collected), or two successive 8-h work shifts (two
samples are collected).
2. When the facility retains the wastewater during an 8-h work shift, and
then treats and releases the wastewater as a batch discharge (one
sample is collected).
3. When the facility discharges wastewater to an estuary only during an
outgoing tide, usually during the 4 h following slack high tide (one
sample is collected).
8.3.4.3 At the end of a shift, clean up activities may result in the
discharge of a slug of toxic waste, which may require sampling and testing.
8.4 RECEIVING WATER SAMPLING
8.4.1 Logistical problems and difficulty in securing sampling equipment
generally preclude the collection of composite receiving water samples for
toxicity tests. Therefore, it is common practice to collect a single grab
sample and use it throughout the test.
8.4.2 The sampling point is determined by the objectives of the test. In
rivers, grab samples should be collected at mid-stream and mid-depth, if
accessible. At estuarine and marine sites, samples are collected at mid-
depth.
8.5 EFFLUENT AND RECEIVING WATER SAMPLE HANDLING, PRESERVATION, AND SHIPPING
8.5.1 Unless the samples are used in an on-site toxicity test the day of
collection, it is recommended that they be held at 4°C until used to inhibit
microbial degradation, chemical transformations, and loss of highly volatile
toxic substances.
8.5.2 Composite samples should be chilled as they are collected. Grab
samples should be chilled immediately following collection.
8.5.3 If the effluent has been chlorinated, total residual chlorine must be
measured immediately following sample collection.
8.5.4 Sample "holding time," as defined here, begins when the last grab
sample of a series is collected, or when the composite sampling period is
completed. If the data from the samples are to be acceptable for use in the
41
-------
NPDES Program, the lapsed time from collection to first use of the sample in
test initiation or test solution renewal should not exceed 36 h. The results
of tests using samples held more than 36 h may not reflect the true toxicity
of the effluent at the time of collection. The sampling schedule should be
adjusted so that the lapsed time from sample collection to shipment is held to
a minimum. However, in the event logistical constraints preclude a 36-h
holding time, permission to use samples held longer than 36 h must be obtained
from the NPDES permitting authority (see Paragraph 8.7.1).
8.5.5 To minimize the loss of toxicity due to volatilization of toxic
constituents, all sample containers should be "completely" filled, leaving no
air space between the contents and the lid.
8.5.6 Samples Used in On-Site Tests
8.5.6.1 Samples collected for on-site tests should be used within 24 h.
8.5.7 Samples Shipped to Off-Site Facilities
8.5.7.1 Samples collected for off-site toxicity testing are to be chilled to
4°C during or immediately after collection, and shipped iced to the performing
laboratory. Sufficient ice should be placed with the sample in the shipping
container to ensure that ice will still be present when the sample arrives at
the laboratory and is unpacked.
8.5.7.2 Samples may be shipped in one or more 4-L (1-gal) CUBITAINERSR or new
plastic "milk" jugs. All sample containers should be rinsed with source water
before beinq filled with sample. After use with receiving water or effluents,
CUBITAINERS and plastic jugs are punctured to prevent reuse.
8.5.7.3 Several sample shipping options are available, including Express
Mail, air express, bus, and courier service. Express Mail is delivered seven
days a week. Saturday and Sunday shipping and receiving schedules of private
carriers vary with the carrier.
8.6 SAMPLE RECEIVING
8.6.1 Upon arrival at the laboratory, samples are logged in and the
temperature is measured and recorded. If the samples are not
immediately prepared for testing, they are stored at 4°C until used.
8.6.2 Every effort must be made to initiate the test with an effluent sample
on the day of arrival in the laboratory, and the sample holding time should
not exceed 36 h unless prior arrangements have been made with the NPDES
permitting authority.
8.7 PERSISTENCE OF EFFLUENT TOXICITY DURING SAMPLE SHIPMENT AND HOLDING
8.7.1 The persistence of the toxicity of an effluent prior to its use in a
toxicity test is of interest in assessing the validity of toxicity test data,
and in determining the possible effects of allowing an extension of the
holding time. Where an extension in holding time is requested by a permittee
42
-------
(see Paragraph 8.5.3), information on the effects of the extension on the
toxicity of samples must be obtained by performing a multi-concentration acute
toxicity test on effluent samples held 36 h, and comparing the results with
those obtained in a toxicity test with the same samples after they have been
held for the requested, longer period. The portion of the sample set aside
for the second test must be held under the same conditions as during shipment
and holding.
43
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SECTION 9
ACUTE TOXICITY TEST PROCEDURES
9.1 PREPARATION OF EFFLUENT AND RECEIVING WATER SAMPLES FOR TOXICITY TESTS
I
9.1.1 When aliquots are removed from the sample container, the head space
above the remaining sample should be held to a minimum. Air which enters a
container upon removal of sample should be expelled by compressing the
container before reclosing, if possible (i.e., where a CUBITAINER used), or
by using an appropriate discharge valve (spigot).
9.1.2 It may be necessary to first coarse-filter samples through a sieve
having 2- to 4-mm mesh openings to remove debris and/or break up large
floating or suspended solids. If samples contain indigenous organisms that
may attack or be confused with the test organisms, the samples must be
filtered through a sieve with 60 urn mesh openings. Caution: filtration may
remove some toxicity.
9.1.3 At a minimum, pH, conductivity or salinity, and total residual chlorine
are measured in the undiluted effluent or receiving water, and pH and
conductivity are measured in the dilution water.
9.1.4 It is recommended that total alkalinity and total hardness also be
measured in the undiluted test water (effluent or receiving water) and the
dilution water.
9.1.5 Total ammonia is measured in effluent and receiving water samples where
toxicity may be contributed by unionized ammonia (i.e., where total ammonia
>5 mg/L). The concentration (mg/L) of unionized (free) ammonia in a sample is
a function of temperature and pH, and is calculated using the percentage value
obtained from Table 9, under the appropriate pH and temperature, and
multiplying it by the concentration (mg/L) of total ammonia in the sample.
9.1.6 Effluents and receiving waters can be dechlorinated using 6.7 mg/L
anhydrous sodium thiosulfate to reduce 1 mg/L chlorine (APHA, 1989, pp. 9-32;
note that the amount of thiosulfate required to dechlorinate effluents is
greater than the amount needed to dechlorinate tap water). Since thiosulfate
may contribute to sample toxicity, a thiosulfate control should be used in the
test in addition to the normal dilution water control.
9.1.7 The DO concentration in the samples should be near saturation prior to
use. Aeration will bring the DO and other gases into equilibrium with air,
minimize oxygen demand, and stabilize the pH. However, aeration during
collection, transfer, and preparation of samples should be minimized to reduce
the loss of volatile chemicals.
44
-------
PH
TABLE 9. PERCENT UNIONIZED NH, IN AQUEOUS AMMONIA SOLUTIONS:
TEMPERATURE 15-26°C AND PH 6.0-8.91
TEMPERATURE <°C>
6.0
6.1
6.2
6.3
6.4
6.5
6.6
6.7
6.8
6.9
7.0
7.1
7.2
7.3
7.4
7.5
7.6
7.7
7.8
7.9
8.0
8.1
8.2
8.3
8.4
8.5
8.6
8.7
8.8
8.9
15
0.0274
0.0345
0.0434
0.0546
0.0687
0.0865
0.109
0.137
0.172
0.217
0.273
0.343
0.432
0.543
0.683
0.858
1.08
1.35
1.70
2.13
2.66
3.33
4.16
5.18
6.43
7.97
9.83
12.07
14.7
17.9
16
0.0295
0.0372
0.0468
0.0589
0.0741
0.0933
0.117
0.148
0.186
0.234
0.294
0.370
0.466
0.586
0.736
0.925
1.16
1.46
1.83
2.29
2.87
3.58
4.47
5.56
6.90
8.54
10.5
12.9
15.7
19.0
17
0.0318
0.0400
0.0504
0.0634
0.0799
0.1005
0.127
0.159
0.200
0.252
0.317
0.399
0.502
0.631
0.793
0.996
1.25
1.57
1.97
2.46
3.08
3.85
4.80
5.97
7.40
9.14
11.2
13.8
16.7
20.2
18
0.0343
0.0431
0.0543
0.0683
0.0860
0.1083
0.136
0.171
0.216
0.271
0.342
0.430
0.540
0.679
0.854
1.07
1.35
1.69
2.12
2.65
3.31
4.14
5.15
6.40
7.93
9.78
12.0
14.7
17.8
21.4
19
0.0369
0.0464
0.0584
0.0736
0.0926
0.1166
0.147
0.185
0.232
0.292
0.368
0.462
0.581
0.731
0.918
1.15
1.45
1.82
2.28
2.85
3.56
4.44
5.52
6.86
8.48
10.45
12.8
15.6
18.9
22.7
20
0.0397
0.0500
0.0629
0.0792
0.0996
0.1254
0.158
0.199
0.250
0.314
0.396
0.497
0.625
0.786
0.988
1.24
1.56
1.95
2.44
3.06
3.82
4.76
5.92
7.34
9.07
11.16
13.6
16.6
20.0
24.0
21
0.0427
0.0537
0.0676
0.0851
0.107
0.135
0.170
0.214
0.269
0.338
0.425
0.535
0.672
0.845
1.061
1.33
1.67
2.10
2.62
3.28
4.10
5.10
6.34
7.85
9.69
11.90
14.5
17.6
21.2
25.3
22 23
0.0459 0.0493
0.0578 0.0621
0.0727 0.0781
0.0915 0.0983
0.115 0.124
0.145 0.156
0.182 0.196
0.230 0.247
0.289 0.310
0.363 0.390
0.457 0.491
0.575 0.617
0.722 0.776
0.908 0.975
1.140 1.224
1.43 1.54
1.80 1.93
2.25 2.41
2.82 3.02
3.52 3.77
4.39 4.70
5.46 5.85
6.78 7.25
8.39 8.96
10.3 11.0
12.7 13.5
15.5 16.4
18.7 19.8
22.5 23.7
26.7 28.2
24
0.0530
0.0667
0.0901
0.1134
0.133
0.167
0.210
0.265
0.333
0.419
0.527
0.663
0.833
1.05
1.31
1.65
2.07
2.59
3.24
4.04
5.03
6.25
7.75
9.56
11.7
14.4
17.4
21.0
25.1
29.6
25
0.0568
0.0716
0.0901
0.1134
0.143
0.180
0.226
0.284
0.358
0.450
0.566
0.711
0.893
1.12
1.41
1.77
2.21
2.77
3.46
4.32
5.38
6.68
8.27
10.2
12.5
15.2
18.5
22.2
26.4
31.1
26
0.0610
0.0768
0.0966
0.1216
0.153
0.193
0.242
0.305
0.384
0.482
0.607
0.762
0.958
1.20
1.51
1.89
2.37
2.97
3.71
4.62
5.75
7.14
8.82
10.9
13.3
16.2
19.5
23.4
27.8
32.6
Table provided by Teresa Norberg-King, Environmental Research Laboratory, Duluth,
Minnesota. Also see Emerson, et. al., 1975; Thurston, et. al, 1974; and USEPA, 1985.
45
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9.1.8 If the samples must be warmed to bring them to the prescribed test
temperature, supersaturation of the dissolved oxygen and nitrogen may become a
problem. To avoid this problem, the effluent and dilution water are checked
with a DO probe after reaching test temperature and, if the DO is greater than
100% saturation or lower than 4.0 mg/L, the solutions are aerated moderately
(approximately 500 mL/min) for a few minutes, using an airstone, until the DO
is within the prescribed range (>4.0 mg/L when using warm water species, or
>6.0 mg/L when using cold water species). Caution: avoid excessive aeration.
9.1.9 Mortality due to pH alone may occur if the pH of the sample falls
outside the range of 6.0 - 9.0. Thus, the presence of other forms of toxicity
(metals and organics) in the sample may be masked by the toxic effects of low
or high pH. The question about the presence of other toxicants can be
answered only by performing two parallel tests, one with an adjusted pH, and
one without an adjusted pH. Freshwater samples are adjusted to pH 7.0, and
marine samples are adjusted to pH 8.0, by adding IN NaOH or IN HC1 dropwise,
as required, being careful to avoid overadjustment.
9.2. PRELIMINARY TOXICITY RANGE-FINDING TESTS
9.2.1 EPA Regional and State personnel generally have observed that it is not
necessary to conduct a toxicity range-finding test prior to initiating a
static, acute, definitive toxicity test. However, when preparing to perform a
static test with an sample of completely unknown quality, or before initiating
a flow-through test, it is advisable to conduct a preliminary toxicity range-
finding test.
9.2.2 A toxicity range-finding test ordinarily consists of a down-scaled,
abbreviated static acute test in which groups of five organisms are exposed to
several widely-spaced-sample dilutions in a logarithmic series, such as 100%,
10.0%, 1.00%, and 0.100%, and a control, for 8-24 h. Caution: if the sample
must also be used for the full-scale definitive test, the 36-h limit on
holding time (Section 8) must not be exceeded before the definitive test is
initiated.
9.2.3 It should be noted that the toxicity (LC50) of a sample observed in a
range-finding test may be significantly different from the toxicity observed
in the follow-up definitive test because: (1) the definitive test is usually
longer; and (2) the test may be performed with a sample collected at a
different time, and possibly differing significantly in the level of toxicity.
9.3 MULTI-CONCENTRATION (DEFINITIVE) EFFLUENT TOXICITY TESTS
9.3.1 The tests recommended for use in determining discharge permit
compliance in the NPDES program are multi-concentration, or definitive, tests
which provide (1) a point estimate of effluent toxicity in terms of a LC50, or
(2) a no-observed-adverse-effect concentration (NOAEC) defined in terms of
mortality, and obtained by hypothesis testing. The tests may be static non-
renewal, static renewal, or flow-through.
46
-------
9.3.2 The tests consist of a control and a minimum of five effluent
concentrations commonly selected to approximate a geometric series, such as
100%, 50%, 25%, 12.5%, and 6.25%, by using a dilution factor of 0.5.
9.3.3 These tests are also to be used in determining compliance with permit
limits on the mortality of the "instream" or receiving water
concentration (RWC) of effluents by bracketing the RWC with effluent
concentrations in the following manner: (1) 100% effluent, (2) [RWC + 100]/2,
(3) RWC, (4) RWC/2, and (5) RWC/4. For example, where the RWC = 50%, the
effluent concentrations used in the test would be 100%, 75%, 50%, 25%, and
12.5%.
9.3.4 If acute/chronic ratios are to be determined by simultaneous acute and
short-term chronic tests with a single species, using the same sample, both
types of tests must use the same test conditions, i.e., temperature, water
hardness, salinity, etc.
9.4 RECEIVING WATER TESTS
9.4.1 Receiving water toxicity tests generally consist of 100% receiving
water and a control. The total hardness or salinity of the control should be
comparable to the receiving water.
9.4.2 The data from the two treatments are analyzed by hypothesis testing to
determine if test organism survival in the receiving water differs
significantly from the control. A minimum of four replicates and 10 organisms
per replicate are required for each treatment (see Tables 11-17).
9.4.3 In cases where the objective of the test is to estimate the degree of
toxicity of the receiving water, a definitive, multi-concentration test is
performed by preparing dilutions of the receiving water, using a > 0.5
dilution series, with a suitable control water.
9.5 STATIC TESTS
9.5.1 Static tests may be non-renewal or renewal.
9.5.2 An excess volume of each dilution is prepared to provide sufficient
material for toxicity testing and routine chemical analyses. The solutions
are well mixed with a glass rod, TEFLON" stir bar, or other means. Aliquots
of each sample concentration are delivered to the test chambers, and the
chambers are arranged in random order. The test solutions are brought to the
required temperature, and the test organisms are added. The remaining volumes
of each sample concentration are used, as necessary, for the chemical
analyses.
9.5.3 Saline dilution water can be prepared by adding dry salts (FORTY
FATHOMS" or equivalent, or modified GP2) or hypersaline brine to de-ionized
water, or a suitable surface freshwater, to adjust the salinity of the entire
47
-------
dilution series. If saline receiving water is used as the diluent, a salinity
control must be prepared using deionized water and dried sea salts to
determine if the addition of sea salts alone has an adverse effect on the test
organisms. It may be desirable to conduct static toxicity tests at several
salinities.
9.5.4 If the effluent has low salinity, but the test is to be conducted with
a salt water organism, the test solutions may be prepared by adding dry ocean
salts or hypersaline brine to a sufficient quantity of 100% effluent to raise
the salinity to the required level, which will depend on the objectives of the
test and the policy of the regulatory agency. After the addition of the dried
salts, stir gently for 30 to 60 min, preferably with a magnetic stirrer, to
ensure that the salts are in solution. It is important to check the final
salinity with a refractometer.
9.5.5 Addition of dry salts to effluents and dilution water may change the pH
and affect the toxicity of the waste. If the objective of the test is to
determine the toxicity of the effluent at the original pH, the pH of the
salinity-adjusted solutions can be brought to the required level by dropwise
addition of IN HC1 or IN NaOH. It is recommended that a concurrent test be
conducted with salinity-adjusted effluent in which the pH has not been altered
after adding the salt.
9.5.6 The volume of the effluent used must be sufficient to prepare all
percent concentrations of the effluent needed for the toxicity test and for
routine chemical analysis. For example, to conduct tests with Menidia, the use
of 200 mL of test solution in each of duplicate exposure vessels and five
concentrations of effluent (10 exposure vessels), would require a total of 1 L
of 100% effluent. However, to provide sufficient volumes of test solutions
for routine chemical analysis and for toxicity testing, additional effluent
would be required (1.5-2.0 L).
9.5.7 A standard control lacking thiosulfate should be included in tests
where the dilution water was prepared by dechlorinating tap water with
thiosulfate.
9.5.8 If, within 1 h of the start of the test, 100% mortality has occurred in
the higher effluent concentrations (such as 100% and 50%), additional
concentrations of effluents, such as 3.1%, 1.6%, and 0.8%, are added to the
test at the lower end of the concentration series.
9.6 FLOW-THROUGH TESTS
9.6.1 Flow-through tests are usually performed with the same effluent
concentrations that are used for static tests, except that where the receiving
water is saline and the effluent is not, 100% effluent cannot be tested with a
marine organism. Examples of flow-through test systems are provided in the
Appendix. Small organisms, such as mysids and daphnids, are confined in
screened enclosures placed in the flow-through chambers. More than one
species may be used in the same test chamber in a given test, if segregated.
48
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9.6.2 The dilutor system should be operated long enough prior to adding the
test organisms to calibrate the dilutor and make the necessary adjustments in
the temperature, flow rate through the test chambers, and aeration. The flow
rate through the proportional dilutor must provide for a minimum of five 90%
replacements of water volume in each test chamber every 24 h (see Figure 2).
This replacement rate should provide sufficient flow to maintain an adequate
concentration of dissolved oxygen. The dilutor should also be capable of
maintaining the test concentration at each dilution within 5% of the starting
concentration for the duration of the test. The calibration of the dilutor
should be checked carefully before the test begins to determine the volume
of effluent and dilution water used in each portion of the effluent delivery
system and the flow rate through each test chamber. The general operation of
the dilutor should be checked at least at the beginning and end of each day
during the test.
9.6.3 The control consists of the same dilution water, test conditions,
procedures, and organisms used in testing the effluent. In the event a test
is to be conducted with salt water organisms, where each effluent dilution
has a different salinity, a static control is prepared for the lowest (or
highest, in the case of high salinity, e.g. brine wastes) salinity level
used in the flow-through test to determine if salinity alone has any adverse
effects on the test organisms.
9.7 NUMBER OF TEST ORGANISMS
9.7.1 A minimum of 20 organisms of a given species are exposed to each
effluent concentration (Jensen, 1972). Small fish and invertebrates are
captured with 4- to 8-mm inside diameter pipettes. Organisms larger than 10-
mm can be captured by dip net. In a typical toxicity te"st involving five
effluent concentrations and a control (six concentrations X 20 organisms per
concentration), fish and other large test organisms are captured from a common
pool and distributed sequentially to the test chambers until the required
number of organisms are placed in each. The test chambers are then positioned
randomly. To avoid carryover of excess culture water in transferring small
organisms to the test chambers, it may be advantageous to distribute small
organisms, such as daphnids, mysids, and larval fish, first to small holding
vessels, such as weighing boats, petri dishes, or small beakers. The water in
the intermediary holding vessels is then drawn down to a small volume and the
entire lot is transferred to a test chamber. In the case of daphnids, both
excessive handling and carryover of culture water and can be avoided by
placing the tip of the transfer pipettes below the surface of the water in the
test chambers and allowing the organisms to swim out of the pipettes without
discharging the contents.
9.8 REPLICATE TEST CHAMBERS
9.8.1 Two or more test chambers are provided for each effluent concentration
and the control. Although the data from duplicate chambers are usually
combined to determine the LC50 and confidence interval, the practice of
49
-------
100
0.4 06
4 10
Volume of Water in Tank
Flow of Water Per Hour
20
40 60
100
Figure 2. Approximate times required to replace water in test chambers in
flow-through tests. For example: for a chamber containing 4 L,
with a flow of 2 L/h, the above graph indicates that 90% of the
water would be replaced every 4.8 h. The same time period
(such as hours) must be used on both axes, and the same unit of
volume (such as liters) must be used for both volume and flow.
(From: Sprague, 1969).
50
-------
dividing the test population for each effluent concentration between two or
more replicate chambers has several advantages and is considered good
laboratory practice because it: (1) permits easier viewing and counting of
test organisms; (2) more easily avoids possible violations of loading limits,
which might occur if all of the test organisms are placed in a single test
vessel; and (3) ensures against the invalidation of the test which might
result from accidental loss of a test vessel, where all of the test
organisms for a given treatment are in a single chamber.
9.9 LOADING OF TEST ORGANISMS
9.9.1 A limit is placed on the loading (weight) of organisms per liter of
test solution to minimize the depletion of dissolved oxygen, the accumulation
of injurious concentrations of metabolic waste products, and/or stress
induced by crowding, any of which could significantly affect the test
results. However, the probability of exceeding loading limits is greatly
reduced with the use of very young test organisms.
9.9.2 For both renewal and non-renewal static tests, loading in the test
solutions must not exceed the following live weights: 1.1 g/L at 15°C,
0.65 g/L at 20°C, or 0.40 g/L at 25°C.
9.9.3 For flow-through tests, the live weight of test organisms in the test
chambers must not exceed 7.0 g/L of test solution at 15°C, or 2.5 g/L at 25°C.
9.10 ILLUMINATION
9.10.1 Light of the quality and intensity normally obtained in the laboratory
during working hours is adequate (10-20 uE/nr/s or 50-100 ft-c). A uniform
photoperiod of 16 h light and 8 h darkness can be achieved in the laboratory
or environmental chamber, using automatic timers.
9.11 FEEDING
9.11.1 Where indicated in the test summary tables (Tables 11-17), food is
made available to test organisms while holding before they are placed in the
test chambers. The organisms are fed at test renewal, 48 h after the test is
initiated, if Regional or State policy requires a 96-h test duration.
9.11.2 Where Artemia nauplii are fed, the nauplii are first concentrated on a
NITEXR screen and then are resuspended in fresh or salt water, depending on
the salinity of the test solutions, using just enough water to form a slurry
that can be transferred by pipette. It should be noted that Artemia nauplii
placed in freshwater usually die in 4 h, generally are not eaten after death,
and decay rapidly, whereas those placed in saline water remain viable and can
serve as food for the duration of the test.
9.11.3 Problems caused by feeding, such as the possible alteration of the
toxicant concentration, the build-up of food and metabolic wastes and
resulting oxygen demand, are common in static test systems. Where feeding is
necessary, excess food should be removed daily by aspirating with a pipette.
51
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9.11.4 Feeding does not cause the above problems in flow-through systems.
However, it is advisable to remove excess food, fecal material, and any
particulate matter that settles from the effluent, from the bottom of the test
vessels daily by aspirating with a pipette.
9.12 TEST TEMPERATURE
9.12.1 Test temperature will depend on the test species and objectives of the
test (see Tables 11-17). Where acute and short-term chronic toxicity tests
are performed simultaneously with the same species to determine acute:chronic
ratios, both tests must be performed at the chronic test temperature. The
average daily temperature of the test solutions must be maintained within
+ 1°C of the selected test temperature, for the duration of the test. This
can be accomplished for static tests by use of a water bath or environmental
chamber, and in flow-through tests by passing the effluent and/or dilution
water through separate coils immersed in a heating or cooling water bath prior
to entering the dilutor system. Coils should be made from materials
recommended in Section 5.
9.13 STRESS
9.13.1 Minimize stress on test organisms by avoiding unnecessary
disturbances.
9.14 DISSOLVED OXYGEN CONCENTRATION
9.14.1 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 the
apparent toxicity of the test solutions by stripping them of highly volatile
toxic substances, or increase its toxicity by altering the pH. However, the
DO in the test solution must not be permitted to fall below 4.0 mg/L for warm
water species and 6.0 mg/L for cold water species. Oxygen saturation values
in fresh and saline waters can be determined from Figure 3 and Table 10,
respectively.
9.14.2 In static tests, low DOs commonly occur in the higher concentrations
of wastewater. Aeration is accomplished by bubbling air through a pipet at
the rate of 100 bubbles/min. If aeration is necessary, all test solutions
must be aerated. It is advisable to monitor the DO closely during the first
few hours of the test. Samples with a potential DO problem generally show a
downward trend in DO within 4 to 8 h after the test is started. Unless
aeration is initiated during the first 8 h of the test, the DO may be
exhausted during an unattended period, thereby invalidating the test.
9.14.3 In most flow-through tests, DO depletion is not a problem in the test
chambers because aeration occurs as the liquids pass through the dilutor
system. If the DO decreases to a level that would be a source of additional
stress, the turnover rate of the solutions in the test chambers must be
increased sufficiently to maintain acceptable DO levels. If the increased
turnover rate does not maintain adequate DO levels, aerate the dilution water
prior to the addition of the effluent, and aerate all test solutions.
52
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CORRECTION FACTORS FOR OXYGEN
SATURATION AT VARIOUS ALTITUDES
ALTITUDE
PRESSURE
FT
MM FACTOR
0
330
655
980
1310
1640
1970
2300
2630
2950
3280
3610
3940
4270
4600
4930
5250
5580
5910
6240
6560
6900
7220
7550
7880
8200
0
100
200
300
400
500
600
700
800
900
1000
1100
1200
1300
1400
1500
1600
1700
1800
1900
2000
2100
2200
2300
2400
2500
760
750
741
732
723
714
705
696
687
679
671
663
655
647
639
631
623
615
608
601
594
587
580
573
566
560
1.00
1.01
1.03
1.04
1.05
1.06
1.08
1.09
1.11
1.12
1.13
1.15
1.16
1.17
1.19
1.20
.22
.24
.25
.26
.28
.30
1.31
1.33
1.34
1.36
5 10 15 20 25 30
Water temperatures °C
10 11 12 13 14 15 16 17
567
Oxygen cc. per liter
Figure 3. Rawson's nomograph for obtaining oxygen saturation values in
freshwater at different temperatures at sea level. When a
straightedge is used to connect the water temperature on the upper
scale and the concentration on the lower scale, the percent
saturation can be read from the point of intersection on the
diagonal scale. To determine the percent saturation at locations
above sea level, factors are provided to convert oxygen
concentrations measured at various altitudes to sea level values in
the table at the upper left. For example, an oxygen concentration
of 6.4 mg/L measured in a body of water at an altitude of 1000 m and
a temperature of 15°C would be equivalent to a concentration of 6.4
X 1.13, or 7.2 mg/L, at sea level. To determine the percent
saturation, a straightedge is used to connect the point at 15°C on
the temperature scale with the point, 7.2 mg/L on the concentration
scale, and the percent saturation is read at the point of
intersection (68%) on the diagonal scale. (From Welch, 1948).
53
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Table 10. OXYGEN SOLUBILITY (MG/L) IN WATER AT EQUILIBRIUM WITH AIR AT
760 MM HG (AFTER RICHARDS AND CORWIN, 1956)
TEMP
0
1
2
3
4
5
6
8
10
12
14
16
18
20
22
24
26
28
30
32
SALINITY (°/oo)
0
14
13
13
13
12
12
12
11
10
10
10
9
9
8
8
8
8
7
7
7
.2
.8
.4
.1
.7
.4
.1
.5
.9
.5
.0
.6
.2
.9
.6
.3
.1
.8
.6
.3
5
13.8
13.4
13.0
12.7
12.3
12.0
11.7
11.2
10.7
10.2
9.7
9.3
9.0
8.6
8.4
8.1
7.8
7.6
7.4
7.1
10
13.4
13.0
12.6
12.3
12.0
11.7
11.4
10.8
10.3
9.9
9.5
9.1
8.7
8.4
8.1
7.8
7.6
7.4
7.1
6.9
15
12.9
12.6
12.2
11.9
11.6
11.3
11.0
10.5
10.0
9.6
9.2
8.8
8.5
8.1
7.9
7.6
7.4
7.2
6.9
6.7
20
12.5
12.2
11.9
11.6
11.3
11.0
10.7
10.2
9.7
9.3
8.9
8.5
8.2
7.9
7.6
7.4
7.2
7.0
6.7
6.5
25
12.1
11.8
11.5
11.2
10.9
10.6
10.3
9.8
9.4
9.0
8.6
8.3
8.0
7.7
7.4
7.2
7.0
6.8
6.5
6.3
30
11.7
11.4
11.1
10.8
10.5
10.2
10.0
9.5
9.1
8.7
8.3
8.0
7.7
7.4
7.2
6.9
6.7
6.5
6.3
6.1
35
11.2
11.0
10.7
10.4
10.1
9.8
9.6
9.2
8.8
8.4
8.1
7.7
7.5
7.2
6.9
6.7
6.5
6.3
6.1
5.9
40
10.8
10.6
10.3
10.0
9.8
9.5
9.3
8.9
8.5
8.1
7.8
7.5
7.2
6.9
6.7
6.5
6.3
6.1
5.9
5.7
43
10.6
10.3
10.0
9.8
9.5
9.3
9.1
8.7
8.3
7.9
7.6
7.3
7.1
6.8
6.6
6.4
6.1
6.0
5.8
5.6
54
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To reduce the potential for driving off volatile compounds in the wastewater,
aeration may be accomplished by bubbling air through a 1-mL pi pet at a rate of
no more than 100 bubbles/min, using an air valve to control the flow.
9.14.4 Caution must be exercised to avoid excessive aeration. Turbulence
caused by aeration should not result in a physical stress to the test
organisms. When aeration is used, the methodology must be detailed in the
report. For safety reasons, pure oxygen should not be used to aerate test
solutions.
9.15 TEST DURATION
9.15.1 Test duration may vary from 24 to 96 h depending on the objectives of
the test and the requirements of the regulatory authority. For specific
information on test duration, see the tables summarizing the test conditions
below.
9.16 ACCEPTABILITY OF TEST RESULTS
9.16.1 For the test results to be acceptable, survival in controls must be at
least 90%. Tests in which the control survival is less than 90% are invalid,
and must be repeated. In tests with specific chemicals, the concentration of
the test material must not vary more than 20% at any treatment level during
the exposure period.
9.16.2 Upon subsequent completion of a valid test, the results of all tests,
valid and invalid, are reported to the regulatory authority with an
explanation of the tests performed and results.
9.17. SUMMARY OF TEST CONDITIONS FOR THE PRINCIPAL TEST ORGANISMS
9.17.1 Summaries of the test conditions for the daphnids, Ceriodaphm'a dubia,
Daphm'a pulex, and D. magna, fathead minnows, Pimephales promelas, rainbow
trout, Oncorhynchus mykiss, brook trout, Salvelinus fontinalis, the mysid,
Mysidopsis bahia, sheepshead minnows, Cypn'nodon variegatus, and silversides,
Menidia beryllina, M. mem'dia, and M. peninsulas, are provided in Tables
11-17.
55
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TABLE 11. SUMMARY OF TEST CONDITIONS AND TEST ACCEPTABILITY CRITERIA FOR
CERIODAPHNIA DUBIA ACUTE TOXICITY TESTS WITH EFFLUENTS AND RECEIVING
WATERS
1. Test type:
2. Test duration:
3. Temperature:1
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:
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 uE/m2/s (50-100 ft-c)
(ambient laboratory levels)
16 h light, 8 h darkness
30 mL (minimum)
15 mL (minimum)
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
1Acute and chronic toxicity tests performed simultaneously to obtain
acute/chronic ratios must use the same temperature and water hardness.
56
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TABLE 11. SUMMARY OF TEST CONDITIONS AND TEST ACCEPTABILITY CRITERIA FOR
CERIODAPHNIA DUBIA ACUTE TOXICITY TESTS WITH EFFLUENTS AND RECEIVING
WATERS (CONTINUED)
14. Feeding regime:
15. Test chamber cleaning:
16. Test chamber 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:
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
SeTenastrum 2 h prior to test solution
renewal at 48 h
Cleaning not required
None
Moderately hard synthetic water prepared
using MILLIPORE MILLI-QR or equivalent
deionized water and reagent grade chemicals
or 20% DMW (see Section 7), receiving water,
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
57
-------
TABLE 12. SUMMARY OF TEST CONDITIONS AND TEST ACCEPTABILITY CRITERIA FOR
DAPHNIA PULEX AND D. MAGNA ACUTE TOXICITY TESTS WITH EFFLUENTS AND
RECEIVING WATERS
1. Test type:
2. Test duration:
3. Temperature:1
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:
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 uE/m2/s (50-100 ft-c)
(ambient laboratory levels)
16 h light, 8 h darkness
30 mL (minimum)
25 mL (minimum)
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 0.2 mL each of YCT and
Selenastrum 2 h prior to test solution
renewal at 48 h
1Acute and chronic toxicity tests performed simultaneously to obtain
acute/chronic ratios must use the same temperature and water hardness.
58
-------
TABLE 12. SUMMARY OF CONDITIONS AND TEST ACCEPTABILITY CRITERIA FOR DAPHNIA
PULEX AND D. HAGNA ACUTE TOXICITY TESTS WITH EFFLUENTS AND RECEIVING
WATERS (CONTINUED)
15. Test chamber cleaning:
16. Test chamber 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:
Cleaning not required
None
Moderately hard synthetic water prepared
using MILLIPORE MILLI-QR or equivalent
deionized water and reagent grade chemicals
or 20% DMW (see Section 7), receiving water,
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
59
-------
TABLE 13. SUMMARY OF TEST CONDITIONS AND TEST ACCEPTABILITY CRITERIA FOR
FATHEAD MINNOW, PINEPHALES PRONELAS, ACUTE TOXICITY TESTS WITH
EFFLUENTS AND RECEIVING WATERS
1. Test type:
2. Test duration:
3. Temperature:1
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:
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 uE/m2/s (50-100 ft-c)
(ambient laboratory levels)
16 h light, 8 h darkness
250 mL (minimum)
200 mL (minimum)
Minimum, after 48 h
1-14 days; 24-h range in age
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
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
1Acute and chronic toxicity tests performed simultaneously to obtain
acute/chronic ratios must use the same temperature and water hardness.
60
-------
TABLE 13. SUMMARY OF TEST CONDITIONS AND TEST ACCEPTABILITY CRITERIA FOR
FATHEAD MINNOW, PIHEPHALES PRONELAS, ACUTE TOXICITY TESTS WITH
EFFLUENTS AND RECEIVING WATERS (CONTINUED)
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:
None, unless DO concentration falls below
4.0 mg/L; rate should not exceed 100
bubbles/min
Moderately hard synthetic water prepared
using MILLIPORE MILLI-QR or equivalent
deionized water and reagent grade chemicals
or 20% DMW (see Section 7), receiving water,
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.
2 L for effluents and receiving waters
90% or greater survival in controls
61
-------
TABLE 14. SUMMARY OF TEST CONDITIONS AND TEST ACCEPTABILITY CRITERIA FOR
RAINBOW TROUT, ONCORHYNCHUS MYKISS, AND BROOK TROUT, SALVELINUS
FONTINALIS, ACUTE TOXICITY TESTS WITH EFFLUENTS AND RECEIVING
WATERS
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:
Static non-renewal, static-renewal, or flow-
through
24, 48, or 96 h
12°C ± 1°C
Ambient laboratory illumination
10-20 uE/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 device.
5 L (minimum) (test chambers should be
covered to prevent fish from jumping out)
4 L (minimum)
Minimum, after 48 h
Rainbow Trout: 15-30 days (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
62
-------
TABLE 14. SUMMARY OF TEST CONDITIONS AND TEST ACCEPTABILITY CRITERIA FOR
RAINBOW TROUT, ONCORHYNCHUS MYKISS, AND BROOK TROUT, SALVELINUS
FONTINALIS, ACUTE TOXICITY TESTS WITH EFFLUENTS AND RECEIVING
WATERS (CONTINUED)
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:
None, unless DO concentration falls below
6.0 mg/L; rate should not exceed 100
bubbles/min
Moderately hard synthetic water prepared
using MILLIPORE MILLI-QR or equivalent
deionized water and reagent grade chemicals
or 20% DMW (see Section 7), receiving water,
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.
20 L for effluents
40 L for receiving waters
90% or greater survival in controls
63
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TABLE 15. SUMMARY OF TEST CONDITIONS AND TEST ACCEPTABILITY CRITERIA FOR
MYSID, HYSIDOPSIS BAHIA, ACUTE TOXICITY TESTS WITH EFFLUENTS AND
RECEIVING WATERS
1. Test type:
2. Test duration:
3. Temperature:1
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:
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 uE/m2/s (50-100 ft-c)
(ambient laboratory levels)
16 h light, 8 h darkness
250 mL (minimum)
200 mL (minimum)
Minimum, after 48 h
1-5 days; 24-h range in age
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
Artemia nauplii are made available while
holding prior to the test; feed 0.2 mL of
concentrated suspension of Artemia nauplii
< 24-h old, daily (approximately 100 nauplii
per mysid)
Cleaning not required
1Acute and chronic toxicity tests performed simultaneously to obtain
acute/chronic ratios must use the same temperature and salinity.
64
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TABLE 15. SUMMARY OF TEST CONDITIONS AND TEST ACCEPTABILITY CRITERIA FOR
MYSID, HYSIDOPSIS BAHIA, ACUTE TOXICITY TESTS WITH EFFLUENTS AND
RECEIVING WATERS (CONTINUED)
16. Test solution aeration:
17. Dilution water:
18. Test concentrations:
19. Dilution series:
None, unless DO concentration falls below
4.0 mg/L; rate should not exceed 100
bubbles/min
5-30 °/oo + 10%; modified GP2, Forty
Fathoms", or equivalent, artificial seawater
prepared with MILLI-QR, or equivalent,
deionized water (see Section 7); or
receiving water
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
20. Endpoint:
21. Sampling and sample
holding requirements:
22. Sample volume required:
23. Test acceptability
criterion:
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 for effluents
2 L for receiving waters
90% or greater survival in controls
65
-------
TABLE 16. SUMMARY OF TEST CONDITIONS AND TEST ACCEPTABILITY CRITERIA FOR
SHEEPSHEAD MINNOW, CYPRINODON VARIEGATUS, ACUTE TOXICITY TESTS WITH
EFFLUENTS AND RECEIVING WATERS
1. Test type:
2. Test duration:
3. Temperature:1
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:
Static non-renewal, static-renewal, or flow-
through
24, 48, 96 h
20°C ± 1°C; or 25°C ± 1°C
Ambient laboratory illumination
10-20 uE/m2/s (50-100 ft-c)
(ambient laboratory levels)
16 h light, 8 h darkness
250 mL (minimum)
200 mL (minimum)
Minimum, after 48 h
1-14 days; 24-h range in age
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
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
1Acute and chronic toxicity tests performed simultaneously to obtain
acute/chronic ratios must use the same temperature and salinity.
66
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TABLE 16. SUMMARY OF TEST CONDITIONS AND TEST ACCEPTABILITY CRITERIA FOR
SHEEPSHEAD MINNOW, CYPRINODON VARIEGATUS, ACUTE TOXICITY TESTS WITH
EFFLUENTS AND RECEIVING WATERS (CONTINUED)
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:
None, unless DO concentration falls below
4.0 mg/L; rate should not exceed 100
bubbles/min
5-32 %>o ± 10%; modified GP2, Forty
Fathoms", or equivalent, artificial seawater
prepared with MILLI-QR or equivalent
deionized water (see Section 7); or
receiving water
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 for effluents
2 L for receiving waters
90% or greater survival in controls
67
-------
TABLE 17. SUMMARY OF TEST CONDITIONS AND TEST ACCEPTABILITY CRITERIA FOR
SILVERSIDE, MENIDIA BERYLLINA, M. HENIDIA, AND H. PENINSULAE, ACUTE
TOXICITY TESTS WITH EFFLUENTS AND RECEIVING WATERS
1. Test type:
2. Test duration:
3. Temperature:1
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:
Static non-revewal, static-renewal, or flow-
through
24, 48, or 96 h
20°C ± 1°C; or 25°C ± 1°C
Ambient laboratory illumination
10-20 uE/m2/s (50-100 ft-c)
(ambient laboratory levels)
16 h light, 8 h darkness
250 mL (minimum)
200 mL (minimum)
Minimum, after 48 h
9-14 days; 24-h range in age
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
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
1Acute and chronic toxicity tests performed simultaneously to obtain
acute/chronic ratios must use the same temperature and salinity.
68
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TABLE 17. SUMMARY OF TEST CONDITIONS AND TEST ACCEPTABILITY CRITERIA FOR
SILVERS IDE, MENIDIA BERYLLINA, M. HENIDIA, AND M. PENINSULAE, ACUTE
TOXICITY TESTS WITH EFFLUENTS AND RECEIVING WATERS (CONTINUED)
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:
None, unless DO concentration falls below 4.0
mg/L; rate should not exceed 100 bubbles/min
Modified GP2, Forty Fathoms", or equivalent,
artificial seawater prepared with MILLI-QR or
equivalent deionized water (see Section 7): or
receiving water:
1-32 °/oo ± 10% for M. beryl!ina;
15-32 °/oo ± 10% for M. menidia
and M. peninsuTae
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 for effluents
2 L for receiving waters
90% or greater survival in controls
69
-------
SECTION 10
TEST DATA
10.1. BIOLOGICAL DATA
10.1.1 Death is the "effect" used for determining toxicity to aquatic
organisms in acute toxicity tests.
10.1.2 Death is not as easily determined for some organisms. The
criteria usually employed in establishing death are: (1) no movement of
gills or appendages; and (2) no reaction to gentle prodding.
10.1.3 The death of some organisms, such as mysids and larval fish, is
easily detected because of a change in appearance from transparent or
translucent to opaque. General observations of appearance and behavior,
such as erratic swimming, loss of reflex, discoloration, excessive mucus
production, hyperventilation, opaque eyes, curved spine, hemorrhaging,
molting, and cannibalism, should also be noted in the daily record.
10.1.4 The test chambers should be checked for early mortality during
the first few hours of the test. The number of surviving organisms in
each test chamber is recorded at the end of each 24-h period (Figure 4).
When recognizable, dead organisms should be removed during each
observation period.
10.1.5 The species, source, and age of the test organisms should be
recorded.
10.2 CHEMICAL AND PHYSICAL DATA
10.2.1 In static tests, at a minimum, pH, salinity or conductivity, and
total residual chlorine are measured in the highest concentration of test
solution and in the dilution water at the beginning of the test, at test
solution renewal, and at test termination. DO, pH, and temperature are
measured in the control and all test concentrations at the beginning of
the test, daily thereafter, and at test termination.
10.2.1.1 It is recommended that total alkalinity and total hardness also
be measured in the control and highest effluent concentration at the
beginning of the test and at test solution renewal.
10.2.1.2 Total ammonia is measured in samples where toxicity may be
contributed by unionized ammonia (where total ammonia might be > 5 mg/L).
70
-------
10.2.1.3 The DO should be monitored closely (every 2 h) for the first
4 to 8 h, to guard against rapid DO depletion, and is measured daily
thereafter in all effluent concentrations in which there are surviving
organisms, and at test termination. It is recommended that test solution
DO be recorded continuously in the test chamber at the highest test
solution concentration or in a surrogate vessel at a comparable test
solution concentration and containing the standard complement of test
organisms.
10.2.1.4 At a minimum, test solution temperature is measured at the
beginning of the test, and daily thereafter. Temperature measurements
are made by placing thermometers or other temperature sensing devices
directly in test solutions or in a comparable volumes of water in
chambers positioned in several locations among the test vessels to
determine test solution temperatures. It is recommended that test
solution temperature be recorded continuously in at least one test
chamber or in a comparable volume of water in a surrogate vessel which is
comparable to the test chambers.
10.2.2 In flow-through tests, at a minimum, pH, salinity or
conductivity, total alkalinity, total hardness, and total residual
chlorine are measured daily in the highest effluent concentration. DO
and temperature are measured at the beginning of the test, daily
thereafter in the control and all test concentrations, and at test
termination.
10.2.3 The measurement of specific conductance is recommended because it
is a very useful parameter in detecting transient fluctuations in the
chemical characteristics of effluents, and will indicate errors in test
dilutions.
10.2.4 Where acute toxicity test methods are utilized to determine
permit limits for toxic chemicals, at a minimum, the concentration of the
test material must be measured in each test concentration at test
initiation, daily thereafter, and at test termination.
10.2.5 Methods used for chemical analysis should be those specified for
Section 304(h) of the CWA (USEPA 1979b, 1982). For salinity
measurements, a refractometer may be used if calibrated with a sample of
known salinity.
71
-------
INDUSTRY/TOXICANT:_
ADDRESS:
CONTACT:
EFFLUENT SERIAL NO:
NPDES PERMIT NO.:
SAMPLE COLLECTOR:
GRAB SAMPLE: COLLECTED (1)
(2)
(3)
(4)
_AM/PM; _/__/_(
AM/PM; _/_/_(DATE)
AM/PM; _/_/_(DATE)
AM/PM; _/__/__
-------
CO
1. EXPOSURE CHAMBER
Total capacity:
Test solution volume:
Test solution surface area:
Water depth (constant):
(cyclic):
3. AERATION
None:
Slow:
ml
mL
cm2
cm
to
(Bubbles or mL/min)
Moderate:
Vigorous:
From:
To:
AM/PM;
"AM/PM;
cm.
(DATE)
(DATE)
2. FEEDING SCHEDULE
Not Fed:
Fed daily:
Fed irregularly:
(describe):
Food used:
4. SCREENED ANIMAL ENCLOSURES
Not used:
Used: (cm) Diameter
5. Condition/appearance of surviving organisms at end of test: (i.e., alive but immobile; loss of
orientation; erratic movement; etc.)
6. Comments:
Figure 5. Check list on back of effluent toxicity data sheet.
-------
SECTION 11
ACUTE TOXICITY DATA ANALYSIS1
11.1 INTRODUCTION
11.1.1 The objective of acute toxicity tests with effluents and receiving
waters is to identify discharges of toxic effluents in acutely toxic amounts.
Data are derived from tests designed to determine the adverse effects of
effluents and receiving waters on the survival of the test organisms. The
recommended effluent toxicity test consists of a control and five or more
concentrations of effluent (i.e., multi-effluent-concentration, or definitive
tests), in which the endpoint is (1) an estimate of the effluent concentration
which is lethal to 50% of the test organisms in the time period prescribed by
the test, expressed as the LC50, or (2) the highest effluent concentration at
which survival is not significantly different from the control (No-Observed-
Adverse-Effect Concentration, or NOAEC). Receiving water tests may be single
concentration or multi-concentration tests. The LC50 is determined by the
Graphical, Spearman-Karber, Trimmed Spearman-Karber, or Probit Method. The
NOAEC is determined by hypothesis testing.
11.1.2 Some states require tests consisting of a control and a single
concentration of effluent with a pass/fail endpoint. Control survival must be
90% or greater for an acceptable test. The test "passes" if survival in the
control and effluent concentration equals or exceeds 90%. The test "fails" if
survival in the effluent is less than 90%, and is significantly different from
control survival (which must be 90% or greater), as determined by hypothesis
testing.
11.1.3 The toxicity of receiving (surface) water can be determined with (1) a
paired test consisting of four replicates each of a suitable control and 100%
surface water, or (2) a multi-concentration test. The results of the first type
of test (100% receiving water and a control) are analyzed by hypothesis testing.
The results of the second type of test may be analyzed by hypothesis testing or
used to determine an LC50.
11.1.4 The data analysis methods recommended in this manual have been chosen
primarily because they are (1) well-tested and well-documented, (2) applicable
to most types of test data sets for which they are recommended, but still
powerful, and (3) most easily understood by non-statisticians. Many other
methods were considered in the selection process, and it is recognized that the
methods selected are not the only possible methods of analysis of acute toxicity
data.
11.1.5 Role of the Statistician
11.1.5.1 The use of the statistical methods described in this manual for
routine data analysis does not require the assistance of a statistician.
Prepared by Florence Fulk with the assistance of Cathy Poore.
74
-------
However, if the data appear unusual in any way, or fail to meet the necessary
assumptions, a statistician should be consulted. The choice of a statistical
method to analyze toxicity test data and the interpretation of the results of
the analysis of the data can become problematic if there are anomalies in the
data. Analysts who are not proficient in statistics are strongly advised
to seek the assistance of a statistician before selecting alternative methods of
analysis and using the results.
11.1.6 Independence, Randomization, and Outliers
11.1.6.1 A critical assumption in the statistical analysis of toxicity data is
statistical independence among observations. Statistical independence means
that given knowledge of the true mean for a given concentration or control,
knowledge of the error in any one actual observation would provide no
information about the error in any other observation. One of the best ways to
insure independence is to properly follow randomization procedures. The purpose
of randomization is to avoid situations where test organisms are placed
serially, by level of concentration, into test chambers, or where all replicates
for a test concentration are located adjacent to one another, which could
introduce bias into the test results.
11.1.6.2 Another area for potential bias of results is the presence of
outliers. An outlier is an inconsistent or questionable data point that appears
unrepresentative of the general trend exhibited by the majority of the data.
Outliers may be detected by tabulation of the data, plotting, and by an analysis
of the residuals. An explanation should be sought for any questionable data
points. Without an explanation, data points should be discarded only with
extreme caution. If there is no explanation, the statistical analysis should be
performed both with and without the outlier, and the results of both analyses
should be reported. For a discussion of techniques for evaluating outliers, see
Draper and John (1981).
11.2 DETERMINATION OF THE LC50 FROM DEFINITIVE, MULTI-EFFLUENT-CONCENTRATION,
ACUTE TOXICITY TESTS
11.2.1 The method used to estimate the LC50 from multi-concentration acute
toxicity tests depends on the shape of the tolerance distribution, and how
well the effluent concentrations chosen characterize the cumulative
distribution function for the tolerance distribution (i.e., the number of
partial mortalities). An review of effluent acute toxicity data from the last
248 tests performed by the Ecological Support Branch, Environmental Services
Division, EPA Region 4, indicated the following pattern in the number of
partial mortalities: (1) no partial mortalities (all or nothing response) -
28%; (2) one partial mortality - 54%; (3) two or more partial mortalities -
16%; (4) LC50 occurring a one of the test concentrations - 2%.
11.2.1.1 Four methods for estimating the LC50 are presented below: the
Graphical Method, the Spearman-Karber Method, the Trimmed Spearman-Karber
Method, and the Probit Method. The analysis scheme is shown in Figure 6.
Included in the presentation of each method is a description of the method,
the requirements for the method, a description of the calculations involved in
the method or a description of the computer program, and an example of the
calculations.
75
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DETERMINATION OF THE LC50
FROM A MULTI-EFFLUENT-CONCENTRATION
ACUTE TOXICITY TEST
MORTALITY DATA
#DEAD
]
r
TWO OR MORE
PARTIAL MORTALITIES?
NO
YES
IS PROBIT MODEL
APPROPRIATE?
(SIGNIFICANTX2TEST)
NO
ONE OR MORE
PARTIAL MORTALITIES?
YES
T
NO
GRAPHICAL METHOD
LC50
YES
PROBIT METHOD
ZERO MORTALITY IN THE
LOWEST EFFLUENT CONG.
AND 100% MORTALITY IN THE
HIGHEST EFFLUENT CONC.?
T
NO
YES
SPEARMAN-KARBER
METHOD
LC50 AND 95%
CONFIDENCE
INTERVAL
^-
TRIMMED SPEARMAN
KARBER METHOD
Figure 6. Flowchart for determination of the LC50 for multi-concentration
acute toxicity tests.
76
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11.2.2 The Graphical Method
11.2.2.1 Description
1. The Graphical Method is a mathematical procedure for estimating
the LC50.
2. The procedure estimates the LC50 by linearly interpolating between
points of a plot of observed percent mortality versus the base 10
logarithm (Iog10) of percent effluent concentration.
3. It does not provide a confidence interval for the LC50 estimate.
4. Use of the Graphical Method is only recommended when there are no
partial mortalities.
11.2.2.2 Requirements
1. The only requirement for the Graphical Method is that the observed
percent mortalities bracket the 50%.
11.2.2.3 General Procedure
1. Let p0, p1? ..., pk denote the observed proportion mortalities for
the control and the k effluent concentrations. The first step is
to smooth the p,. if they do not satisfy p0 < ... < pk. The
smoothing replaces any adjacent p/s that do not conform to p0 <
... < pk, with their average. For example, if p,- is less than
p,-.,, then:
PM = Pi = (P,- + PM)/2
where: p? = the smoothed observed proportion mortality for effluent
concentration i.
2. Adjust the smoothed observed proportion mortality in each effluent
concentration for mortality in the control group using Abbott's formula
(Finney, 1971). The adjustment takes the form:
p° = (p* - p*) / (1 - pS)
where: PQ = the smoothed observed proportion mortality for the control.
3. Plot the smoothed, adjusted data on 2-cycle semi-log graph paper with
the logarithmic axis (the y axis) used for percent effluent
concentration and the linear axis (the x axis) used for observed percent
mortality.
4. Locate the two points on the graph which bracket 50% mortality
and connect them with a straight line.
77
-------
5. On the scale for percent effluent concentration, read the value
for the point where the plotted line and the 50% mortality line
intersect. This value is the estimated LC50 expressed as a
percent effluent concentration.
11.2.2.4 Example Calculation
1. All-or-nothing data (Graphical Method) in Table 18 are used in the
calculations. Note that in this case, the data must be smoothed and
adjusted for mortality in the controls.
2. To smooth the data, the observed proportion mortality for the
control and the lower three effluent concentrations must be
averaged. The smoothed observed proportion mortalities are as
follows: 0.0125, 0.0125, 0.0125, 0.0125, 1.0, and 1.0.
3. The smoothed responses are adjusted for control mortality (see
11.2.2.3), where the smoothed response for the control (p®) = 0.0125.
The smoothed, adjusted response proportions for the effluent
concentrations are as follows: 0.0, 0.0, 0.0, 1.0, and 1.0.
4. A plot of the smoothed, adjusted data is shown in Figure 7.
5. The two points on the graph which bracket the 50% mortality line
(0% mortality at 25% effluent, and 100% mortality at 50% effluent)
are connected with a straight line.
6. The point at which the plotted line intersects the 50% mortality
line is the estimated LC50. The estimated LC50 = 35% effluent
TABLE 18. MORTALITY DATA (NUMBER OF DEAD ORGANISMS) FROM ACUTE TOXICITY
TESTS USED IN EXAMPLES OF LC50 DETERMINATIONS (20 ORGANISMS IN
THE CONTROL AND ALL TEST CONCENTRATIONS)
EFFLUENT
CONC
METHOD OF ANALYSIS
GRAPHICAL
SPEARMAN-
KARBER
TRIMMED-
SPEARMAN
KARBER
PROBIT
CONTROL
6.25%
12.5%
25.0%
50.0%
100.0%
1
0
0
0
20
20
1
1
0
0
13
20
1
0
1
0
0
16
0
3
9
20
20
78
-------
CJ
cc
10 20 30 40 50 60 70 80 90 100
PERCENT MORTALITY
Figure 7. Plotted data and fitted line for Graphical Method,
using all-or-nothing data.
79
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11.2.3 The Spearman-Karber Method
11.2.3.1 Description
1. The Spearman-Karber Method is a nonparametric statistical
procedure for estimating the LC50 and the associated 95%
confidence interval (Finney, 1978).
2. This procedure estimates the mean of the distribution of the
Iog10 of the tolerance. If the log tolerance distribution is
symmetric, this estimate of the mean is equivalent to an
estimate of the median of the log tolerance distribution.
3. If the response proportions are not monotonically non-decreasing with
increasing concentration (constant or steadily increasing with
concentration), the data are smoothed.
4. Abbott's procedure is used to "adjust" the test results for mortality
occurring in the control.
5. Use of the Spearman-Karber Method is recommended when partial
mortalities occur in the test solutions, but the data do not fit
the Probit model.
11.2.3.2 Requirements
1. To calculate the LC50 estimate, the following must be true:
a. The smoothed adjusted proportion mortality for the lowest
effluent concentration (not including the control) must be zero.
b. The smoothed adjusted proportion mortality for the highest
effluent concentration must be one.
2. To calculate the 95% confidence interval for the LC50 estimate,
one or more of the smoothed adjusted proportion mortalities must
be between zero and one.
11.2.3.3 General Procedure
1. The first step in the estimation of the LC50 by the Spearman-
Karber Method is to smooth the observed response proportions,
p,- if they do not satisfy pp < ... < pk (see 11.2.2.3, Step 1).
2. Adjust the smoothed observed proportion mortality in each effluent
concentration for mortality in the control group using Abbott's formula
(see 11.2.2.3, Step 2).
3. Plot the smoothed adjusted data on 2-cycle semi-log graph paper with
the logarithmic axis (the y axis) used for percent effluent
concentration and the linear axis (the x axis) used for observed
percent mortality.
4. Calculate the Iog10 of the estimated LC50, m, as follows:
k-'
•I.
m r-i
80
-------
where p? = the smoothed adjusted proportion mortality at
concentration i
Xj = the log.0 of concentration i
k = the number of effluent concentrations tested, not including
the control.
5. Calculate the estimated variance of m as follows:
K~1 «3 / 1 M3 \ / V V \ 2
r P,-(l - P,-)(X,-+1 - X- ,)
V(m) = I ! U
1=2 4(nf - 1)
where X,- = the Iog10 of concentration i
n, = the number of organisms tested at effluent concentration i
PJ = the smoothed adjusted observed proportion mortality at
effluent concentration i
k = the number of effluent concentrations tested, not including
the control.
6. Calculate the 95% confidence interval for m: m ± 2.0 yv(m)
7. The estimated LC50 and a 95% confidence interval for the
estimated LC50 can be found by taking base10 antilogs of the
above values.
8. With the exclusion of the plot in item 3, the above
calculations can be carried out using the Trimmed Spearman-
Karber computer program mentioned in 11.2.4.3 and 11.2.4.4.
11.2.3.4 Example Calculation
1. Mortality data from a definitive, multi-concentration, acute
toxicity test are given in Table 18. Note that the data must
be smoothed and adjusted for mortality in the controls.
2. To smooth the data, the observed proportion mortality for the
control, and the observed proportion mortality for the 6.25%,
12.5%, and 25% effluent concentrations must be averaged. The
smoothed observed proportion mortalities are as follows:
0.025, 0.025, 0.025, 0.025, 0.65, and 1.00.
3. To adjust the smoothed, observed proportion mortality in each
effluent concentration for mortality in the control group,
Abbott's formula must be used. After smoothing and adjusting,
the proportion mortalities for the effluent concentrations are
as follows: 0.000, 0.000, 0.000; 0.641, and 1.000.
4. The data will not be plotted for this example. For an example
of the plotting procedures, see 11.2.2.4.
5. The Iog10 of the estimated LC50, m, is calculated as follows:
m = [(0.0000 - 0.0000)(0.7959 + 1.0969)]/2 +
[(0.0000 - 0.0000)(1.0969 + 1.3979)]/2 +
[(0.6410 - 0.0000)(1.3979 + 1.6990)]/2 +
[(1.0000 - 0.6410)(1.6990 + 2.0000)]/2
= 1.656527
81
-------
6. The estimated variance of m, V(m), is calculated as follows:
V(m) = (0.0000)(1.0000)(1.3979 - 0.7959)^/4(19) +
(0.0000)(1.0000)(1.6990 - 1.0969)1/4(19) +
(0.6410)(0.3590)(2.0000 - 1.3979)74(19)
= 0.0010977
7. The 95% confidence interval for m is calculated as follows:
1.656527 ± 2 70.0010977 = (1.5902639, 1.7227901)
8. The estimated LC50 is as follows: antilog(l.656527) = 45.3%.
9. The upper limit of the 95% confidence interval for the estimated LC50 is
as follows:
antilog(l.7227901) = 52.8%
10. The lower limit of the 95% confidence interval for the estimated LC50 is
as follows:
antilog(l.5902639) = 38.9%
11.2.4 The Trimmed Spearman-Karber Method
11.2.4.1 Description
1. The Trimmed Spearman-Karber Method is a modification of the Spearman-
Karber, nonparametric statistical procedure for estimating the LC50 and
the associated 95% confidence interval (Hamilton, et al, 1978).
2. This procedure estimates the trimmed mean of the distribution of
the Iog10 of the tolerance. If the log tolerance distribution is
symmetric, this estimate of the trimmed mean is equivalent to an
estimate of the median of the log tolerance distribution.
3. Use of the Trimmed Spearman-Karber Method is only appropriate when
the requirements for the Probit Method and the Spearman-Karber
Method are not met.
11.2.4.2 Requirements
1. To calculate the LC50 estimate with the Trimmed Spearman-Karber
Method, the smoothed, adjusted, observed proportion mortalities must
bracket 0.5.
2. To calculate a confidence interval for the LC50 estimate, one or
more of the smoothed, adjusted, observed proportion mortalities
must be between zero and one.
11.2.4.3 General Procedure
1. Smooth the observed proportion mortalities as described in
11.2.2.3, Step 1.
2. Adjust the smoothed, observed proportion mortalities for mortality
in the control as described in 11.2.2.3, Step 2.
3. Plot the smoothed, adjusted data as described in 11.2.2.3, Step 3.
82
-------
4. Calculate the amount of trim to use in the estimation of the LC50
as follows:
Trim = max(p^, 1 - p£)
where: p* = the smoothed, adjusted proportion mortality for the lowest
effluent concentration, exclusive of the control.
p* = the smoothed, adjusted proportion mortality for the highest
effluent concentration.
k = the number of effluent concentrations, exclusive of the
control.
5. Due to the intensive nature of the calculation for the estimated LC50
and the calculation for the associated 95% confidence interval using
the Trimmed Spearman-Karber Method, it is recommended that the data be
analyzed by computer.
6. A computer program which estimates the LC50 and associated 95%
confidence interval using the Trimmed Spearman-Karber Method,
originally developed by Montana State University, was translated
and modified for EMSL-Cincinnati by Computer Sciences Corporation.
Computer Sciences Corporation, 26 W. Martin Luther King Drive,
Cincinnati, Ohio 45268. The modified program is written in
Borland Turbo C for the IBM compatible PC. A full listing and a
machine-readable, compiled, version of the program can be obtained
from EMSL-Cincinnati by sending a diskette with a written request
to the Quality Research Division, Environmental Monitoring Systems
Laboratory, at the above address.
7. The modified program automatically performs the following
functions:
a. Smoothing.
b. Adjustment for mortality in the control.
c. Calculation of the necessary trim.
d. Calculation of the LC50.
e. Calculation of the associated 95% confidence interval.
8. The modified program does not create a plot of observed proportion
mortality versus the Iog10 of percent effluent concentration.
11.2.4.4 Example Calculation Using the Computer Program
1. Data from Table 18 are used to illustrate the analysis using the
Trimmed Spearman-Karber program.
2. The program requests the following input (see Figure 8):
a. Output destination (disk file or printer).
b. Title for output.
c. Control data.
d. Data for each toxicant concentration.
3. The program output includes the following (see Figure 9):
a. A table of the observed proportion mortality and the smoothed,
adjusted observed proportion mortality for each of the toxicant
concentrations.
b. The amount of trim used in the calculation.
c. The estimated LC50 and the associated 95% confidence interval.
83
-------
UuuuuuuuuOOuuuuuuuuuuuuuuuuOiJuuuuuuuuUUuuiJuuOuuiJOuOtJiJOOOtJOuOuuuu
U EMSL Cincinnati U
U Computer Program for Trimmed Spearman-Karber Method U
U LC50 Estimate and 95% Confidence Interval 0
0 Version 1.5 0
OuuuuuuiJiJuuuOOOuuuuuuuuuuuiJuuOuuOuuuuuuuuuuuuuOOOOOuOuOOuOuuiJuuu
Output to printer or disk file (P/D)? P
Title? Trimmed Spearman-Karber Method Example
Number of animals in the control group = ? 20
Number of deaths in the control group = ? 1
Number of toxicant concentrations = ? 5
Input data starting with the lowest concentration
Toxicant concentration = ? 6.25%
Number of animals exposed = ? 20
Number of deaths = ? 0
Toxicant concentration = ? 12.5%
Number of animals exposed = ? 20
Number of deaths = ? 1
Toxicant concentration = ? 25%
Number of animals exposed = ? 20
Number of deaths = ? 0
Toxicant concentration = ? 50%
Number of animals exposed = ? 20
Number of deaths = ? 0
Toxicant concentration = ? 100%
Number of animals exposed = ? 20
Number of deaths = ? 16
Toxicant
Concentration
Control
6.25
12.50
25.00
50.00
100.00
Do you wish to
Number of
Deaths
1
0
1
0
0
16
modify your data
Number
Exposed
20
20
20
20
20
20
(y/n) ? n
Figure 8. Example of input for computer program for Trimmed
Spearman-Karber Method.
84
-------
uOuuUuuuOuuuuUuuOuuuuOOuuuOuuuuOuuuOuuuuOuuuOuuuuOuuuUUuuOOOuuOO
0 EMSL Cincinnati 0
U Computer Program for Trimmed Spearman-Karber Method U
0 LC50 Estimate and 95% Confidence Interval U
0 Version 1.5 U
uuuuuuuuuuuuuuuuuuuuuuuuuuuuuuuuuuuuuuuuuuuuuuuuuuuuuuuuuuOuuuuu
TRIMMED SPEARMAN-KARBER EXAMPLE
Smoothed,
Observed Adjusted
Cone. Number Number of Proportion Proportion
(%) Exposed Deaths Mortality Mortality
Control
20
1
0.0500
0.0000
6.25
12.50
25.00
50.00
100.00
20
20
20
20
20
0
1
0
0
16
0.0000
0.0500
0.0000
0.0000
0.8000
0.0000
0.0000
0.0000
0.0000
0.7959
Automatic trim used was 20.41%
Estimated LC50 = 77.28
Lower 95% confidence limit = 73.56
Upper 95% confidence limit = 81.19
Figure 9. Example of output from computer program for Trimmed
Spearman-Karber Method.
85
-------
4. The analysis results for this example are as follows:
a. The observed proportion mortalities smoothed and adjusted for
mortality in the control.
b. The amount of trim used to calculate the estimate:
trim = max (0.00, 0.204} = 0.204.
c. The estimate of the LC50 is 77.3%, with a 95% confidence interval
of (73.6%, 81.2%).
11.2.5 The Probit Method
11.2.5.1 Description
1. The Probit Method is a parametric statistical procedure for estimating
the LC50 and the associated 95% confidence interval (Finney, 1978).
2. The analysis consists of transforming the observed proportion
mortalities with a probit transformation, and transforming the effluent
concentrations to Iog10.
3. Given the assumption of normality for the 1og10 of the tolerances, the
relationship between the transformed variables mentioned above is
approximately linear.
4. This relationship allows estimation of linear regression parameters,
using an iterative approach.
5. The estimated LC50 and associated confidence interval are calculated
from the estimated linear regression parameters.
11.2.5.2 Requirements
1. To obtain a reasonably precise estimate of the LC50 with the Probit
Method, the observed proportion mortalities must bracket 0.5.
2. The log.,,, of the tolerance is assumed to be normally distributed.
3. To calculate the LC50 estimate and associated 95% confidence interval,
two or more of the observed proportion mortalities must be between zero
and one.
11.2.5.3 General Procedure
1. Due to the intensive nature of the calculations for the estimated LC50
and associated 95% confidence interval using the Probit Method, it is
recommended that the data be analyzed by a computer program.
2. A computer program to estimate the LC50 and associated 95% confidence
intervals using the Probit Method was developed by EMSL-Cincinnati.
The program was written in IBM PC Basic for the IBM compatible PC by
Computer Sciences Corporation, 26 W. Martin Luther King Drive,
Cincinnati, Ohio 45268. A full listing and a machine-readable,
compiled, version of the program can be obtained from EMSL-Cincinnati
by sending a diskette with a written request to the Quality Assurance
Research Division, Environmental Monitoring Systems Laboratory, at the
above address.
86
-------
11.2.5.4 Example Using the Computer Program
1. Data from Table 18 are used to illustrate the operation of the
Probit program for calculating the LC50 and the associated 95%
confidence interval.
2. The program begins with a request for the following initial input (see
Figure 10):
a. Output designation (P = printer, D = disk file).
b. Title for the output.
c. Control data.
d. Toxicant concentration data.
3. The program output includes the following (see Figure 11):
a. A table of the observed proportion mortality, the adjusted observed
proportion mortality, and the predicted proportion mortality for
each effluent concentration.
b. The calculated chi-squared statistic for heterogeneity and
the tabular value. This test is one indicator of how well
the data fit the model. The program will issue a warning
when the test indicates that the data do not fit the
model.
c. Estimates of the mean (mu) and the standard deviation
(sigma) of the underlying tolerance distribution.
d. Estimates and standard errors of the intercept and slope
of the fitted probit regression line.
e. The estimated LC50 and 95% confidence limits.
f. A plot of the fitted regression line with observed data
overlaid on the plot (see Figure 12).
4. The results of the data analysis for this example are as
follows:
a. The observed proportion mortalities were not adjusted for mortality
in the control.
b. The test for heterogeneity was not significant (the calculated Chi-
square was less than the tabular value), thus the Probit Method
appears to be appropriate for this data.
c. The estimate of the LC50 is 22.9% with a 95% confidence interval of
(18.8%, 27.8%).
87
-------
UUUUUUUUUUUUUUUUUUUUUUUUUUUUUUiJUiJUUUUUUUUUUUUUUUUUUUUUUUUUUUUUUO
U EMSL Cincinnati U
0 Computer Program for the Probit Method U
U LC50 Estimate and 95% Confidence Interval U
U Version 1.5 0
UUUUUUUUUUUUUUUUUUUUUUUUiJUUUUUUUUUOUUOUUUUUUUUUUUUUUUUUUUUUUUUUU
Output to printer or disk file (P/D) ? P
Title? Probit Example
Number of animals in the control group = ? 20
Number of deaths in the control group = ? 0
Number of administered concentrations = ? 5
Input data starting with the lowest concentration
Toxicant concentration = ? 6.25%
Number of animals exposed = ? 20
Number of deaths = ? 0
Toxicant concentration = ? 12.5%
Number of animals exposed = ? 20
Number of deaths = ? 3
Toxicant concentration = ? 25%
Number of animals exposed = ? 20
Number of deaths = ? 9
Toxicant concentration = ? 50%
Number of animals exposed = ? 20
Number of deaths = ? 20
Toxicant concentration = ? 100%
Number of animals exposed = ? 20
Number of deaths = ? 20
Toxicant
Concentration
Control
6.25
12.50
25.00
50.00
100.00
Do you wish to
Number of
Deaths
0
0
3
9
20
20
modify your data
Number
Exposed
20
20
20
20
20
20
(y/n) ? n
Figure 10. Example of input for computer program for Probit Method.
88
-------
uuuuuuuuuuuuOiJOuuuiJuu
u
U Comput
0 LC50 Est
0
uuuuuuuuuuiJUuuuuuuuuu
PROBIT EXAMPLE
Observed Ad
Cone. Number Number of Proportion Pro
(%) Exposed Deaths Mortality Mo
6.25
12.50
25.00
50.00
100.00
20
20
20
20
20
0
3
9
20
20
0.0000
0.1500
0.4500
1.0000
1.0000
uUUuUUuuuiJuUuuuuuuuuuiJuuuuuuuuuUuuiJuuiJuuUiJ
EMSL Cincinnati U
r Program for the Probit Method Li
mate and 95% Confidence Interval U
Version 1.5 U
UUUUUUUUUUUUUUUUUUUUUUUUUUUUUUUUUUUUUUUUUU
usted Predicted
crtion Proportion
tality Mortality
0000
500
4500
0000
0000
0.0022
0.0924
0.5774
0.9570
0.9994
Chi-squared heterogeneity = 3.076
Tabular chi-squared value (df = k - 2 = 3)
Theoretical spontaneous response rate = 0.
Estimated rou (log.,,,) = 1.3593
Estimated LC50 = 22.87%
Estimated sigma (log^g) = 0.1979
Lower 95% confidence limit = 18.79%
Upper 95% confidence limit = 27.85%
= 7.81
000
Figure 11. Example of output for computer program for Probit Method.
89
-------
Probit
10+
9+
8+
7+
6+
5+
4+
3+
2+
1+
O+o
LC01 LC10 LC25 LC50 LC75 LC90 LC99
Figure 12. Plot of adjusted Probits and predicted regression line.
90
-------
11.3 DETERMINATION OF NO-OBSERVED-ADVERSE-EFFECT CONCENTRATION (NOAEC) FROM
MULTI-CONCENTRATION TESTS, AND DETERMINATION OF PASS OR FAIL (PASS/FAIL)
FOR SINGLE-CONCENTRATION (PAIRED) TESTS
11.3.1 Determination of the No-Observed-Adverse-Effect Concentration (NOAEC),
for multi-concentration toxicity tests, and pass or fail (Pass/Fail) for
single-concentration toxicity tests is accomplished using hypothesis testing.
The NOAEC is the lowest concentration at which survival is not significantly
different from the control. In Pass/Fail tests, the objective is to determine
if the survival in the single treatment (effluent or receiving water) is
significantly different from the control survival.
11.3.2 The first step in these analyses is to transform the responses,
expressed as the proportion surviving, by the arc-sine-square-root
transformation (Figures 13 and 14). The arc-sine-square-root transformation
is commonly used on proportionality data to stabilize the variance and satisfy
the normality requirement. Shapiro-Milk's test may be used to test the
normality assumption.
11.3.3 If the data do not meet the assumption of normality and there are four
or more replicates per group, then the non-parametric test, Wilcoxon Rank Sum
Test, can be used to analyze the data.
11.3.4 If the data meet the assumption of normality, the F test for equality
of variances is used to test the homogeneity of variance assumption. Failure
of the homogeneity of variance assumption leads to the use of a modified t
test, where the pooled variance estimate is adjusted for unequal variance, and
the degrees of freedom for the test are adjusted.
11.3.5 General Procedure
11.3.5.1 Arc Sine Square Root Transformation
11.3.5.1.1 The arc sine transformation consists of taking the inverse sine of
the square root of the proportion surviving. Whenever the proportion
surviving is 0 or 1, a special modification of the transformation must be used
(Bartlett, 1937). Illustrations of the arc sine square root transformation
and modification are provided below.
1. Calculate the response proportion (RP) for each replicate within a
group, where:
RP = (number of surviving organisms)/(number exposed)
91
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DETERMINATION OF PASS OR FAIL
FROM A SINGLE-EFFLUENT-CONCENTRATION
ACUTE TOXICITY TEST
SURVIVAL DATA
PROPORTION SURVIVING
i
ARC SINE TRANSFORMATION
NORMALITY?
(SHAPIRO-WILK'S TEST)
NO
WILCOXON RANK
SUM TEST
IF YES
HOMOGENEITY OF VARIANCE?
(F-TEST)
YES
SIGNIFICANT DIFF.
IN SURVIVAL?
NO
YES
Figure 13. Flowchart for analysis of single-effluent-concentration
test data.
92
-------
DETERMINATION OF THE NOAEC
FROM A MULTI-EFFLUENT-CONCENTRATION
ACUTE TOXICITY TEST
SURVIVAL DATA
PROPORTION SURVIVING
I
ARC SINE TRANSFORMATION
I
NORMALITY?
(SHAPIRO-WILK'S TEST)
YES
I
NO
YES
HOMOGENEITY OF VARIANCE?
(BARTLETT'S TEST)
NO
NO
EQUAL NUMBER OF
REPLICATES?
EQUAL NUMBER OF
REPLICATES?
YES
NO
STEEL'S MANY-ONE
RANK TEST
WILCOXON RANK SUM
[TEST WITH BONFERRONI
ADJUSTMENTS
ENDPOINT ESTIMATES
NOAEC
Figure 14. Flowchart for analysis of multi-effluent-concentration
test data.
93
-------
2. Transform each RP to arc sine, as follows.
a. For RPs greater than zero or less than one:
Angle (in radians) = arc sine J (RP)
b. Modification of the arc sine when RP = 0.
Angledn radians) = arc sine\\ —
^ 4n
c. Modification of the arc sine when RP = 1.0.
Angle = 1.5708 radians - ( radians for RP = 0)
11.3.5.2 Shapiro Milk's Test
11.3.5.2.1 After the data have been transformed, test the assumption of
normality using Shapiro Milk's test. The test statistic, W, is obtained by
dividing the square of an appropriate linear combination of the sample order
statistics by the usual symmetric estimate of variance (D). The calculated W
must be greater than zero and less than or equal to one. This test is
recommended for a sample size of 50 or less, and there must be more than two
replicates per concentration for the test to be valid.
1. To calculate W, first center the observations by subtracting the mean of
all the observations within a concentration from each observation in
that concentration.
2. Calculate the denominator, D, of the test statistic:
Where: X1 = the ith centered observation
X = the overall mean of the centered observations.
3. Order the centered observations from smallest to largest.
X(D x(2) x(n)
Where: X(l) denotes the ith ordered observation.
4. From Table 19, for the number of observations, n, obtain the
coefficients ax, a2, ..., ak, where k is n/2 if n is even,
and (n - l)/2 if n is odd.
94
-------
TABLE 19. COEFFICIENTS FOR THE SHAPIRO-MILK'S TEST
i
\ n
1
2
3
4
5
2
\
0.7071
—
—
—
—
3
0.7071
0.0000
—
—
—
4
0.6872
0.1667
—
—
—
5
0.6646
0.2413
0.0000
—
—
C
0.6431
0.2806
0.0875
—
—
7
0.6233
0.3031
0.1401
0.0000
—
8
0.6052
0.3164
0.1743
0.0561
—
9
0.5888
0.3244
0.1976
0.0947
0.0000
10
0.5739
0.3291
0.2141
0.1224
0.0399
1
2
3
4
5
6
7
8
9
10
11
\
0.5601
0.3315
0.2260
0.1429
0.0695
0.0000
—
—
—
—
12
0.5475
0.3325
0.2347
0.1586
0.0922
0.0303
—
—
—
—
13
0.5359
0.3325
0.2412
0.1707
0.1099
0.0539
0.0000
—
—
—
14
0.5251
0.3318
0.2460
0.1802
0.1240
0.0727
0.0240
—
—
—
15
0.5150
0.3306
0.2495
0.1878
0.1353
0.0880
0.0433
0.0000
—
—
16
0.5056
0.3290
0.2521
0.1939
0.1447
0.1005
0.0593
0.0196
—
—
17
0.4968
0.3273
0.2540
0.1988
0.1524
0.1109
0.0725
0.0359
0.0000
—
18
0.4886
0.3253
0.2553
0.2027
0.1587
0.1197
0.0837
0.0496
0.0163
—
19
0.4808
0.3232
0.2561
0.2059
0.1641
0.1271
0.0932
0.0612
0.0303
0.0000
20
0.4734
0.3211
0.2565
0.2085
0.1686
0.1334
0.1013
0.0711
0.0422
0.0140
\
i\
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
21
\
0.4643
0.3185
0.2578
0.2119
0.1736
0.1399
0.1092
0.0804
0.0530
0.0263
0.0000
—
—
—
—
22
0.4590
0.3156
0.2571
0.2131
0.1764
0.1443
0.1150
0.0878
0.0618
0.0368
0.0122
—
—
—
—
23
0.4542
0.3126
0.2563
0.2139
0.1787
0.1480
0.1201
0.0941
0.0696
0.0459
0.0228
0.0000
—
—
—
24
0.4493
0.3098
0.2554
0.2145
0.1807
0.1512
0.1245
0.0997
0.0764
0.0539
0.0321
0.0107
—
—
—
25
0.4450
0.3069
0.2543
0.2148
0.1822
0.1539
0.1283
0. 1046
0.0823
0.0610
0.0403
0.0200
0.0000
—
—
26
0.4407
0.3043
0.2533
0.2151
0.1836
0.1563
0.1316
0.1089
0.0876
0.0672
0.0476
0.0284
0.0094
—
—
27
0.4366
0.3018
0.2522
0.2152
0.1848
0.1584
0.1346
0.1128
0.0923
0.0728
0.0540
0.0358
0.0178
0.0000
—
28
0.4328
0.2992
0.2510
0.2151
0.1857
0.1601
0.1372
0.1162
0.0965
0.0778
0.0598
0.0424
0.0253
0.0084
—
29
0.4291
0.2968
0.2499
0.2150
0.1864
0.1616
0.1395
0.1192
0.1002
0.0822
0.0650
0.0483
0.0320
0.0159
0.0000
30
0.4254
0.2944
0.2487
0.2148
0.1870
0.1630
0.1415
0.1219
0.1036
0.0862
0.0697
0.0537
0.0381
0.0227
0.0076
^Taken from: Conover, 1980.
95
-------
TABLE 19. COEFFICIENTS FOR THE SHAPIRO-MILK'S TEST (CONT.)
\. fl
J
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
31
\
0.4220
0.2921
0.2475
0.2145
0.1874
0.1641
0.1433
0.1243
0.1066
0.0899
0.0739
0.0585
0.0435
0.0289
0.0144
0.0000
—
—
—
—
32
0.4188
0.2898
0.2462
0.2141
0.1878
0.1651
0.1449
0.1265
0.1093
0.0931
0.0777
0.0629
0.0485
0.0344
0.0206
0.0068
—
—
—
—
33
0.4156
0.2876
0.2451
0.2137
0.1880
0.1660
0.1463
0.1284
0.1118
0.0961
0.0812
0.0669
0.0530
0.0395
0.0262
0.0131
0.0000
—
—
—
34
0.4127
0.2854
0.2439
0.2132
0.1882
0.1667
0.1475
0.1301
0.1140
0.0988
0.0844
0.0706
0.0572
0.0441
0.0314
0.0187
0.0062
—
—
—
35
0.4096
0.2834
0.2427
0.2127
0.1883
0.1673
0.1487
0.1317
0.1160
0.1013
0.0873
0.0739
0.0610
0.0484
0.0361
0.0239
0.0119
0.0000
—
—
36
0.4068
0.2813
0.2415
0.2121
0.1883
0.1678
0.1496
0.1331
0.1179
0.1036
0.0900
0.0770
0.0645
0.0523
0.0404
0.0287
0.0172
0.0057
—
—
37
0.4040
0.2794
0.2403
0.2116
0.1883
0.1683
0.1505
0.1344
0.1196
0.1056
0.0924
0.0798
0.0677
0.0559
0.0444
0.0331
0.0220
0.0110
0.0000
—
38
0.4015
0.2774
0.2391
0.2110
0.1881
0.1686
0.1513
0.1356
0.1211
0.1075
0.0947
0.0824
0.0706
0.0592
0.0481
0.0372
0.0264
0.0158
0.0053
—
39
0.3989
0.2755
0.2380
0.2104
0.1880
0.1689
0.1520
0.1366
0.1225
0.1092
0.0967
0.0848
0.0733
0.0622
0.0515
0.0409
0.0305
0.0203
0.0101
0.0000
40
0.3964
0.2737
0.2368
0.2098
0.1878
0.1691
0.1526
0.1376
0.1237
0.1108
0.0986
0.0870
0.0759
0.0651
0.0546
0.0444
0.0343
0.0244
0.0146
0.0049
\"
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
43
\
0.3940
0.2719
0.2357
0.2091
0.1876
0.1693
0.1531
0.1384
0.1249
0.1123
0.1004
0.0891
0.0782
0.0677
0.0575
0.0476
0.0379
0.0283
0.0188
0.0094
0.0000
—
42
0.3917
0.2701
0.2345
0.2085
0.1874
0.1694
0.1535
0.1392
0.1259
0.1136
0.1020
0.0909
0.0804
0.0701
0.0602
0.0506
0.0411
0.0318
0.0227
0.0136
0.0045
__
—
43
0.3894'
0.2684
0.2334
0.2078
0.1871
0.1695
0.1539
0.1398
0.1269
0.1149
0.1035
0.0927
0.0824
0.0724
0.0628
0.0534
0.0442
0.0352
0.0263
0.0175
0.0087
0.0000
—
—
—
44
0.3872
0".2667
0.2323
0.2072
0.1868
0.1695
0.1542
0.1405
0.1278
0.1160
0.1049
0.0943
0.0842
0.0745
0.0651
0.0560
0.0471
0.0383
0.0296
0.0211
0.0126
0.0042
—
—
—
45
0.3850
0.2651
0.2313
0.2065
0.1865
0.1695
0.1545
0.1410
0.1286
0.1170
0.1062
0.0959
0.0860
0.0765
0.0673
0.0584
0.0497
0.0412
0.0328
0.0245
0.0163
0.0081
0.0000
—
—
46
0.3830
0.2635
0.2302
0.2058
0.1862
0.1695
0.1548
0.1415
0.1293
0.1180
0.1073
0.0972
0.0876
0.0783
0.0694
0.0607
0.0522
0.0439
0.0357
0.0277
0.0197
0.0118
0.0039
— •
—
47
0.3808
0.2620
0.2291
0.2052
0.1859
0.1695
0.1550
0.1420
0.1300
0.1189
0.1085
0.0986
0.0892
0.0801
0.0713
0.0628
0.0546
0.0465
0.0385
0.0307
0.0229
0.0153
0.0076
0.0000
—
48
0.3789
0.2604
0.22S1
0.2045
0.1855
0.1693
0.1551
0.1423
0.1306
0.1197
0.1095
0.0998
0.0906
0.0817
0.0731
0.0648
0.0568
0.0489
0.0411
0.0335
0.0259
0.0185
0.0111
0.0037
—
49
0.3770
0.2589
0.2271
0.2038
0.1851
0.1692
0.1553
0.1427
0.1312
0.1205
0.1105
0.1010
0.0919
0.0832
0.0748
0.0667
0.0588
0.0511
0.0436
0.0361
0.0288
0.0215
0.0143
0.0071
0.0000
50
0.3751
0.2574
0.2260
0.2032
0.1847
0.1691
9.1554
0.1430
0.1317
0.1212
0.1113
0.1020
0.0932
0.0846
0.0764
0.0685
0.0608
0.0532
0.0459
0.0386
0.0314
0.0244
0.0174
0.0104
0.0035
96
-------
5. Compute the test statistic, W, as follows:
1 k
W [ X a, (XCnH+1) - X(0)]2
D i = l
11.3.5.2.2 The decision rule for the test is to compare the critical value
from Table 20 to the computed W. If the computed value is less than the
critical value, conclude that the data are not normally distributed.
11.3.5.3 F TEST
11.3.5.3.1 The F test for equality of variances is used to test the
homogeneity of variance assumption. When conducting the F test, the
alternative hypothesis of interest is that the variances are not equal.
11.3.5.3.2 To make the two-tailed F test at the 0.05 level of significance,
put the larger of the two sample variances in the numerator of F.
s2
F = _i where S2 > S2
11.3.5.3.3 Compare the calculated F with the 0.05 level of a tabulated F
value with n1 - 1 and n2 - 1 degrees of freedom, where n1 and n2 are the number
of replicates for each of the two groups (Snedecor and Cochran, 1980). If the
calculated F value is less than or equal to the tabulated F, conclude that the
variances of the two groups are equal.
11.3.5.4 T Test
11.3.5.4.1 If the variances for the two groups are found to be statistically
equivalent, then the equal variance t test is the appropriate test.
11.3.5.4.2 Calculate the following test statistic:
t = 1 2
SP
Where: X., = Mean for the control
X2 = Mean for the effluent concentration
?2
•-'2
S,,2 = Estimate of the variance for the control
97
-------
S22 = Estimate of the variance for the effluent concentration
H! = Number of replicates for the control
n2 = Number of replicates for the effluent concentration
11.3.5.4.3 Since we are concerned with a decrease in mortality from the control,
a one-tailed test is appropriate. Thus, compare the calculated t with a critical
t, where the critical t is at the 5% level of significance with n., + n2 - 2
degrees of freedom. If the calculated t exceeds the critical t, the mean
responses are declared different.
11.3.5.5 Modified T Test
11.3.5.5.1 If the F test for equality of variance fails, the t test is still a
valid test. However, the denominator and the degrees of freedom for the test are
modified.
11.3.5.5.2 The t statistic, with the modification for the denominator, is
calculated as follows:
t-
^
Where: %± = Mean for the control
X2 = Mean for the effluent concentration
o
S1 = Estimate of the variance for the control
S22 = Estimate of the variance for the effluent concentration
nj = Number of replicates for the control
n2 = Number of replicates for the effluent concentration
11.3.5.5.3 Additionally, the degrees of freedom for the test are adjusted using
the following formula:
df'= (^-D (fl2-l)
(n2-I) c2 + (l-c)2 (^-l)
Where: si
n,
C=
98
-------
TABLE 20. QUANTILES OF THE SHAPIRO-MILK'S TEST STATISTIC1
n
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
0.01
0.753
0.687
0.686
0.713
0.730
0.749
0.764
0.781
0.792
0.805
0.814
0.825
0.835
0.844
0.851
0.858
0.863
0.868
0.873
0.878
0.881
0.884
0.888
0.891
0.894
0.896
0.898
0.900
0.902
0.904
0.906
0.908
0.910
0.912
0.914
0.916
0.917
0.919
0.920
0.922
0.923
0.924
0.926
0.927
0.928
0.929
0.929
0.930
0.02
0.756
0.707
0.715
0.743
0.760
0.778
0.791
0.806
0.817
0.828
0.837
0.846
0.855
0.863
0.869
0.874
0.879
0.884
0.888
0.892
0.895
0.898
0.901
0.904
0.906
0.908
0.910
0.912
0.914
0.915
0.917
0.919
0.920
0.922
0.924
0.925
0.927
0.928
0.929
0.930
0.932
0.933
0.934
0.935
0.936
0.937
0.937
0.938
0.05
0.767
0.748
0.762
0.788
0.803
0.818
0.829
0.842
0.850
0.859
0.866
0.874
0.881.
0.887
0.892
0.897
0.901
0.905
0.908
0.911
0.914
0.916
0.918
0.920
0.923
0.924
0.926
0.927
0.929
0.930
0.931
0.933
0.934
0.935
0.936
0.938
0.939
0.940
0.941
0.942
0.943
0.944
0.945
0.945
0.946
0.947
0.947
0.947
0.10
0.789
0.792
0.806
0.826
0.838
0.851
0.859
0.869
0.876
0.883
0.889
0.895
0.901
0.906
0.910
0.914
0.917
0.920
0.923
0.926
0.928
0.930
0.931
0.933
0.935
0.936
0.937
0.939
0.940
0.941
0.942
0.943
0.944
0.945
0.946
0.947
0.948
0.949
0.950
0.951
0.951
0.952
0.953
0.953
0.954
0.954
0.955
0.955
0.50
0.959
0.935
0.927
0.927
0.928
0.932
0.935
0.938
0.940
0.943
0.945
0.947
0.9SO
0.952
0.954
0.956
0.957
0.959
0.960
0.961
0.962
0.963
0.964
0.965
0.965
0.966
0.966
0.967
0.967
0.968
0.968
0.969
0.969
0.970
0.970
0.971
0.971
0.972
0.972
0.972
0.973
0.973
0.973
0.974
0.974
0.974
0.974
0.974
0.90
0.998
0.987
0.979
0.974
0.972
0.972
0.972
0.972
0.973
0.973
0.974
0.975
0.975
0.976
0.977
0.978
0.978
0.979
0.980
0.980
0.981
0.981
0.981
0.982
0.982
0.982
0.982
0.983
0.983
0.983
0.983
0.983
0.984
0.984
0.984
0.984
0.984
0.985
0.985
0.985
0.985
0.985
0.985
0.985
0.985
0.985
0.985
0.985
0.95
0.999
0.992
0.986
0.981
0.979
0.978
0.978
0.978
0.979
0.979
0.979
0.980
0.980
0.981
0.981
0.982
0.982
0.983
0.983
0.984
0.984
0.984
0.985
0.985
0.985
0.985
0.985
0.985
0.986
0.986
0.986
0.986
0.986
0.986
0.987
0.987
0.987
0.987
0.987
0.987
0.987
0.987
0.988
0.988
0.988
0.988
0.988
0.988
0.98
1.000
0.996
0.991
0.986
0.985
0.984
0.984
0.983
0.984
0.984
0.984
0.984
0.984
0.985
0.985
0.986
0.986
0.986
0.987
0.987
0.987
0.987
0.988
0.988
0.988
0.988
0.988
0.988
0.988
0.988
0.989
0.989
0.989
0.989
0.989
0.989
0.989
0.989
0.989
0.989
0.990
0.990
0.990
0.990
0.990
0.990
0.990
0.990
0.99
1.000
0.997
0.993
0.989
0.988
0.987
0.986
0.986
0.986
0.986
0.986
0.986
0.987
0.987
0.987
0.988
0.988
0.988
0.989
0.989
0.989
0.989
0.989
0.989
0.990
0.990
0.990
0.990
0.990
0.990
0.990
0.990
0.990
0.990
0.990
0.990
0.991
0.991
0.991
0.991
0.991
0.991
0.991
0.991
0.991
0.991
0.991
0.991
tlaken from Conover, 1980.
99
-------
11.3.5.5.4 The modified degrees of freedom is usually not an integer. Common
practice is to round down to the nearest integer.
11.3.5.5.5 The modified t test is then performed in the same way as the equal
variance t test. The calculated t is compared to the critical t at the 0.05
significance level with modified degrees of freedom. If the calculated t exceeds
the critical t, the mean responses are found to be statistically different.
11.3.5.6 Wilcoxon Rank Sum Test
11.3.5.6.1 If the data fail the test for normality and there are four or more
replicates per group, then the non-parametric test, the Wilcoxon Rank Sum Test
may be used to analyze the data. If less than four replicates were used, a non-
parametric alternative is not available.
11.3.5.6.2 The Wilcoxon Rank Sum Test consists of jointly ranking the data and
calculating the rank sum for the effluent concentration. The rank sum is then
compared to a critical value to determine acceptance or rejection of the null
hypothesis.
11.3.5.6.3 To carry out the test, combine the data for the control and the
effluent concentration and arrange the values in order of size from smallest to
largest. Assign ranks to the ordered observations, a rank of 1 to the smallest,
2 to the next smallest, etc. If ties in rank occur, assign the average rank to
the observation. Sum the ranks for the effluent concentration.
11.3.5.6.4 If the survival in the effluent concentration is significantly less
than that of the control, the rank sum for the effluent concentration would be
lower than the rank sum of the control. Thus, we are only concerned with
comparing the rank sum for the effluent concentration with some "minimum" or
critical rank sum, at or below which the effluent concentration mortality would
be considered to be significantly lower than the mortality in the control. For
a test at the 5% level of significance, the critical rank sum can be found in
Table 21.
11.3.6 Single Concentration Test
11.3.6.1 Data from an acute effluent toxicity test with Ceriodaphnia are
provided in Table 22. The proportion surviving in each replicate is transformed
by the arc sine square root transformation prior to statistical analysis of the
data (Figure 13).
11.3.6.2 After the data have been transformed, test the assumption of normality
via the Shapiro Wilk's test.
11.3.6.2.1 The first step in the test for normality is to center the
observations by subtracting the mean of all observations within a concentration
from each observation in that concentration. The centered observations are
listed in Table 23.
100
-------
11.3.6.2.2 Calculate the denominator, D, of the test statistic:
D = E (xi ~ *>2
i=l
For this set of data, X = 0 and D = 0.060.
11.3.6.2.3 Order the centered observations from smallest to largest. The
ordered observations are listed in Table 24.
11.3.6.2.4 From Table 1, for n = 8 and k = n/2 = 4, obtain the coefficients
a,, a2, ..., ak. The a,- values are listed in Table 25.
11.3.6.2.5 Compute the test statistic, W, as follows:
W= - - - -(0.2200)2 = 0.0807
The differences, x(n"i+1) - X(i), are listed in Table 25.
11.3.6.2.6 From Table 20, the critical W value for n = 8 and a significance
level of 0.01, is 0.749. Since the calculated W, 0.807, is not less than the
critical value the conclusion of the test is that the data are normally
distributed.
11.3.6.3. The F test for equality of variances is used to test the
homogeneity of variance assumption.
11.3.6.3.1 From Table 22, obtain the sample variances for the control and the
100% effluent. Since the variability of the 100% effluent is greater than the
variability of the control, S2 for the 100% effluent concentration is placed
in the numerator of the F statistic and S2 for the control is placed in the
denominator.
11.3.6.3.2 There are four replicates for the control and four replicates for
the 100% effluent concentration. Thus there are three degrees of freedom for
the numerator and the denominator. For a two-tailed test at the 0.01 level of
significance, the critical F value is 47.467. The calculated F, 1.2614, is
less than the critical F, 47.467, thus the conclusion is that the variances of
the control and 100% effluent are equal.
11.3.6.4 The assumptions of normality and homogeneity of variance have been
met for this data set. An equal variance t test will be used to compare the
mean responses of the control and 100% effluent.
101
-------
11.3.6.4.1 To perform the t test, obtain the values for Xr X2, S^, and S22
from Table 22. Calculate the t statistic as follows:
^_ 1.330 - 0.604
0.0997J — + —
4 4
Where:
(4-1) 0.0088 + (4-1) (0.0111)
4+4-2
11.3.6.4.2 For a one-tailed test at the 0.05 level of significance with 6
degrees of freedom, the critical t value is 1.9432. Since the calculated t,
10.298, is greater than the critical t, the conclusion is that the survival in
the 100% effluent concentration is significantly less than the survival in the
control.
11.3.6.5 If the data had failed the normality assumption, the appropriate
analysis would be the Wilcoxon Rank Sum Test. To provide an example of this
test, the mortality data from the t test example will be reanalyzed by the
nonparametric procedure.
11.3.6.5.1 The first step in the Wilcoxon Rank Sum Test is to combine the
data from the control and the 100% effluent concentration and arrange the
values in order of size, from smallest to largest.
11.3.6.5.2 Assign ranks to the ordered observations, a rank of 1 to the
smallest, 2 to the next smallest, etc. The combined data with ranks assigned
is presented in Table 26.
11.3.6.5.3 Sum the ranks for the 100% effluent concentration.
11.3.6.5.4 For this set of data, the test is for a significant reduction in
survival in the 100% effluent concentration as compared to the control. The
critical value, from Table 21, for four replicates in each group and a
significance level of 0.05 is 11. The rank sum for the 100% effluent
concentration is 10 which is less than the critical value of 11. Thus the
conclusion is that survival in the effluent concentration is significantly
less than the control survival.
102
-------
TABLE 21. CRITICAL VALUES FOR WILCOXON'S RANK SUM TEST FIVE PERCENT CRITICAL
LEVEL
NO. REPLICATES
IN CONTROL
NO. OF REPLICATES PER EFFLUENT CONCENTRATION
3456789 10
3
4
5
6
7
8
9
10
6
7
8
8
9
10
10
10
11
12
13
14
15
16
17
16
17
19
20
21
23
24
26
23
24
26
28
29
31
33
35
30
32
34
36
39
41
43
45
39
41
44
46
49
51
54
56
49
51
54
57
60
63
66
69
59
62
66
69
72
72
79
82
TABLE 22. DATA FROM AN ACUTE SINGLE-CONCENTRATION TOXICITY TEST WITH
CERIODAPHNIA
PROPORTION SURVIVING
REPLICATE
CONTROL
100% EFFLUENT
CONCENTRATION
RAW
DATA
A
B
C
D
1.00
1.00
0.90
0.90
0.40
0.30
0.40
0.20
ARC SINE
TRANSFORMED
DATA
A
B
C
D
1.412
1.412
1.249
1.249
0.685
0.580
0.685
0.464
1.330
0.0088
0.604
0.0111
103
-------
TABLE 23. EXAMPLE OF SHAPIRO-WILK'S TEST: CENTERED OBSERVATIONS
TREATMENT
REPLICATE
Control
100% Effluent
0.082
0.081
0.082
-0.024
-0.081
0.081
-0.081
-0.140
TABLE 24. EXAMPLE OF SHAPIRO-WILK'S TEST: ORDERED OBSERVATIONS
1
2
3
4
5
6
7
8
-0.140
-0.081
-0.081
-0.024
0.081
0.081
0.082
0.082
TABLE 25. EXAMPLE OF SHAPIRO-WILK'S TEST: TABLE OF COEFFICIENTS
AND DIFFERENCES
- X
(1)
1
2
3
4
0.6052
0.3164
0.1743
0.0561
0.222
0.163
0.162
0.105
X(8)
X(7)
X(6)
x(5)
- x(1)
- x(2)
- x(3)
- x(4)
104
-------
TABLE 26. EXAMPLE OF WILCOXON'S RANK SUM TEST: ASSIGNING RANKS
TO THE CONTROL AND 100% EFFLUENT CONCENTRATIONS
PROPORTION CONTROL
RANK SURVIVING OR 100%
EFFLUENT
1 0.20 100% EFFLUENT
2 0.30 100% EFFLUENT
3.5 0.40 100% EFFLUENT
3.5 0.40 100% EFFLUENT
5.5 0.90 CONTROL
5.5 0.90 CONTROL
7.5 1.00 CONTROL
7.5 1.00 CONTROL
105
-------
11.3.7 Multi-Concentration Test
11.3.7.1. Formal statistical analysis of the survival data is outlined in
Figure 14. The response used in the analysis is the proportion of animals
surviving in each test or control chamber. Concentrations at which there is no
survival in any of the test chambers are excluded from statistical analysis of
the NOAEC.
11.3.7.2 For the case of equal numbers of replicates across all
concentrations and the control, the determination of the NOAEC endpoint is
made via a parametric test, Dunnett's Procedure, or a nonparametric test,
Steel's Many-one Rank Test, on the arc sine transformed data. Underlying
assumptions of Dunnett's Procedure, normality and homogeneity of variance, are
formally tested. The test for normality is the Shapiro-Wilk's Test, and
Bartlett's Test is used to determine the homogeneity of variance. If either
of these tests fail, the nonparametric test, Steel's Many-one Rank Test, is
used to determine the NOEC and LOEC endpoints. If the assumptions of
Dunnett's Procedure are met, the endpoints are estimated by the parametric
procedure.
11.3.7.3 If unequal numbers of replicates occur among the concentration
levels tested, there are parametric and nonparametric alternative analyses.
The parametric analysis is a t-test with a Bonferroni adjustment. The
Wilcoxon Rank Sum Test with the Bonferroni adjustment is the nonparametric
alternative.
11.3.7.4 Example of Analysis of Survival Data
11.3.7.4.1 This example uses survival data from a fathead minnow test. The
proportion surviving in each replicate must first be transformed by the arc
sine square root transformation procedure. The raw and transformed data,
means and standard deviations of the transformed observations at each toxicant
concentration and control are listed in Table 27. A plot of the survival
proportions is provided in Figure 15.
11.3.7.4.2 Test for Normality
1. The first step of the test for normality is to center the observations
by subtracting the mean of all observations within a concentration from
each observation in that concentration. The centered observations are
summarized in Table 28.
106
-------
TABLE 27. FATHEAD MINNOW SURVIVAL DATA
TOXICANT CONCENTRATION (UG/L)
REPLICATE CONTROL 32 64 128 256 512
RAW
ARC SINE
TRANS-
FORMED
MEANtY,)
S,-
i
A
B
C
D
A
B
C
D
1
1
0
0
1
1
1
1
1.
0.
1
.0
.0
.9
.9
.412
.412
.249
.249
,330
0088
0
0
1
0
1
1
1
1
1,
0.
2
.8
.8
.0
.8
.107
.107
.412
.107
.183
0232
0
1
1
1
1
1
1
1
1.
0.
3
.9
.0
.0
.0
.249
.412
.412
.412
,371
0066
0.9
0.9
0.8
1.0
1.249
1.249
1.107
1.412
1.254
0.0155
4
0
0
1
0
0
1
1
0
1.
0.
5
.7
.9
.0
.5
.991
.249
.412
.785
.109
0768
0.4
0.3
0.4
0.2
0.685
0.580
0.685
0.464
0.604
0.0111
6
TABLE 28. CENTERED OBSERVATIONS FOR SHAPIRO-WILK'S EXAMPLE
TOXICANT CONCENTRATION (UG/L)
REPLICATE CONTROL 32 64 128 256 512
A
B
C
D
0.
0.
-0.
-0.
082
082
081
081
-0.
-0.
0.
-0.
076
076
229
076
-0.122
0.041
0.041
0.041
-0.
-0.
-0.
0.
005
005
147
158
-0
0
0
-0
.118
.140
.303
.324
0.
-0.
0.
-0.
081
024
081
140
107
-------
o
00
o:
o
CL.
O
a:
a.
1.0 »
0.9 »
0.8
0.7-
0.6
0.5-
0.4-
0.3
FPBFwSTIH^^^DYH^,F9? ttCV CONCENTRA
S£?E?IS§R ?ONCi? ^^.^Su^oSTlE"
SIGNIFICANTLY DlPfERENT FROM THE CONTROL)
CONCENTRATION^
0.2-
*
512
32
64
128
256
TOXICANT CONCENTRATION (UG/L)
Figure 15. Plot of mean survival proportion data in Table 27.
-------
2. Calculate the denominator, D, of the statistic:
D = s (X, - X)2
Where: X; = the ith centered observation
X = the overall mean of the centered observations
n = the total number of centered observations
3. For this set of data: n = 24 (number of observations)
X = _1_ (0.000) = 0.000
24
D = 0.4265
4. Order the centered observations from smallest to largest
v(1) V<2) y(n)
A > A , ...» A
Where: Xo> denotes the ith ordered observation.
The ordered observations for this example are listed in Table 29.
5. From Table 17, for the number of observations, n, obtain the
coefficients ap a., ... ak, where k is approximately n/2. For the
data in this example, n = 24 and k = 12. The a, values are listed
in Table 30.
6. Compute the test statistic, W, as follows:
1 k
W [ X a, (X(n'i+1) - X(0)]2
D i=l
The differences X(n'i+1) - X(i) are listed in Table 30. For the data
in this example,
(0.6444)2 = 0.974
0.4265
7. The decision rule for this test is to compare W as calculated in #6
to a critical value found in Table 21. If the computed W is less
than the critical value, conclude that the data are not normally
distributed. For the data in this example, the critical value at a
significance level of 0.01 and n = 24 observations is 0.884. Since
W = 0.974 is greater than the critical value, conclude that the data
are normally distributed.
109
-------
TABLE 29. ORDERED CENTERED OBSERVATIONS FOR THE SHAPIRO-MILK'S EXAMPLE
1
2
3
4
5
6
7
8
9
10
11
12
-0.324
-0.147
-0.140
-0.122
-0.118
-0.081
-0.081
-0.076
-0.076
-0.076
-0.024
-0.005
13
14
15
16
17
18
19
20
21
22
23
24
-0.005
0.041
0.041
0.041
0.081
0.081
0.082
0.082
0.140
0.158
0.229
0.303
TABLE 30. COEFFICIENTS AND DIFFERENCES FOR SHAPIRO-MILK'S EXAMPLE
a..
1
2
3
4
5
6
7
8
9
10
11
12
0.4493
0.3098
0.2554
0.2145
0.1807
0.1512
0.1245
0.0997
0.0764
0.0539
0.0321
0.0107
0.627
0.376
0.298
0.262
0.200
0.163
0.162
0.157
0.117
0.117
0.065
0.0
X(24,
x<23>
x<22)
x<21>
x<20)
X(19)
X(18)
X(17)
X(16)
X(15)
X(H)
X(13)
- x(1)
- x<2>
- x(3>
- x(4)
- x(5)
- x(6)
- x(7)
- x(8)
- x(9)
- x(10)
- x<11>
- x(12)
110
-------
11.3.7.4.3 Test for Homogeneity of Variance
1. The test used to examine whether the variation in mean proportion
surviving is the same across all toxicant concentrations including
the control, is Bartlett's Test (Snedecor and Cochran, 1980). The
test statistic is as follows:
P P
[ ( s V,) In S2 - S V,. In S,-2 ]
B =
Where: V,- = degrees of freedom for each toxicant concentration
and control, V, = (n, - 1)
n,- = the number of replicates for concentration i.
In = loge
i = 1, 2, ..., p where p is the number of concentrations
including the control
c = i + [3(P-i)r1 t s i/v, - (s v,
2. For the data in this example, (See Table 27) all toxicant
concentrations including the control have the same number of
replicates (n,. = 4 for all i). Thus, V,- = 3 for all i.
3. Bartlett's statistic is therefore:
B = [(18)ln(0.0236) - 3 Z ln(S,-2)]/1.1296
= [18(-3.7465) - 3(-24.7516)]/1.1296
= 6.8178/1.1296
= 6.036
111
-------
4. B is approximately distributed as chi square with p - 1 degrees of
freedom, when the variances are in fact the same. Therefore, the
appropriate critical value for this test, at a significance level
of 0.01 with five degrees of freedom, is 15.086. Since B = 6.036
is less than the critical value of 15.086, conclude that the
variances are not different.
11.3.7.4.4 Dunnett's Procedure
1. To obtain an estimate of the pooled variance for the Dunnett's
Procedure, construct an ANOVA table.
TABLE 31. ANOVA TABLE
SOURCE
BETWEEN
WITHIN
Total
DF SUM OF SQUARES
(SS)
p - 1 SSB
N - p SSW
N - 1 SST
MEAN SQUARE (MS)
(SS/DF)
SB2 = SSB/(p-l)
Su2 = SSW/(N-p)
Where: p = number toxicant concentrations including the control
N = total number of observations n1 + n2 ... + np
SSB = S V/X - GZ/N
Between Sum of Squares
ni
SST = S 2 Y,/ - G7N
Total Sum of Squares
SSW = SST - SSB
Within Sum of Squares
G = the grand total of all sample observations, G = s T,-
i=l
n,- = number of observations in concentration i
TJ = the total of the replicate measurements for
concentration i
Yr = the jth observation for concentration i (represents
the proportion surviving for toxicant concentration
i in test chamber j)
112
-------
2. For the data in this example:
'1
n2 = n3 = n4 = n5 = n6 = 4
N = 24
= Y
= Y
= Y
= Y
= Y
= Y
11
21
31
41
51
61
G = T,
12
22
32
42
52
'62
"
*
33
43
r
'63
Y14 = 5.322
Y,,. = 4.733
Y24 -
Y34-
Y-
Y54-
'64 ~
5.485
5.017
4.437
2.414
T6 = 27.408
SSB = X T,2/n, - G2/N
= 1 (131.495) - (27.408)2 = 1.574
4 24
P n;
SST = S 2 '
i=l j=l
- G2/N
33.300 - (27.408)'
24
= 2.000
SSW = SST - SSB = 2.000 - 1.574
0.4260
SB = SSB/(p - 1) = 1.574/(6 - 1) = 0.3150
S2 = SSW/(N - p) = 0.426/(24 - 6) = 0.024
3. Summarize these calculations in the ANOVA table (Table 32)
TABLE 32.
Source
BETWEEN
WITHIN
ANOVA TABLE
DF
5
18
FOR DUNNETT'S
SUM OF SQUARES
(SS)
1.574
0.426
PROCEDURE EXAMPLE
MEAN SQUARE (MS)
(SS/DF)
0.315
0.024
Total
23
2.002
113
-------
4. To perform the individual comparisons, calculate the t statistic for
each concentration, and control combination as follows:
- Yf
Su V (l/n,) + (1/n,)
Where Y,- = mean proportion surviving for concentration i
Y1 = mean proportion surviving for the control
Su = square root of within mean sqaure
n, = number of replicates for control
n,- = number of replicates for concentration i.
5. Table 33 includes the calculated t values for each concentration and
control combination. In this example, comparing the 32 ug/L
concentration with the control the calculation is as follows:
( 1.330 - 1.183 )
t2 = = 1.341
[ 0.155 V (1/4) + (1/4) ]
TABLE 33. CALCULATED T VALUES
TOXICANT CONCENTRATION (UG/L) i t,
32
64
128
256
512
2
3
4
5
6
1.341
-0.374
0.693
2.016
6.624
6. Since the purpose of this test is to detect a significant reduction
in proportion surviving, a one-sided test is appropriate. The
critical value for this one-sided test is found in Table 34. For an
overall alpha level of 0.05, 18 degrees of freedom for error and
five concentrations (excluding the control) the critical value is
2.41. The mean proportion surviving for concentration "i" is
considered significantly less than the mean proportion surviving for
the control if t, is greater than the critical value. Since t is
greater than 2.41, the 512 ug/L concentration has significantly
lower survival than the control. Hence the NOAEC for survival is
256 ug/L.
114
-------
TABLE 34. DUNNETT'S "T" VALUES
(One-tailed) d
X
5
6
7
8
g
10
11
12
13
M
15
16
17
IB
19
20
24
30
40
to
120
•
1
.02
.94
.89
.86
.83
.81
.80
.78
.77
.76
.75
.75
.74
.73
.73
.72
.71
.70
.88
.67
.66
.64
2
.44
.34
.27
.22
.18
.15
.13
.11
.09
.08
.07
.06
.05
.04
.03
.03
.01
.99
.97
.95
1.93
1.92
3
2.68
2.56
2.48,,
2.42
2.37
2.34
2.31
2.29
2.27
2.25
2.24
2.23
2.22
2.21
2.20
2.19
2.17
2.15
2.13
2.10
2.08
2.06
4
2.85
2.71
2.62
2.55
2.50
2.47
2.44
2.41
2.39
2.37
2.36
2.34
2.33
2.32
2.31
2.30
2.28
2.25
2.23
2.21
2.18
2.16
a * .05
5
2.98
2.83
2.73
2.66
2.60
2.56
2.53
2.50
2.48
2.46
2.44
2.43
2.42
2.41.
2.40
2.39
2.36
2.33
2.31
2.28
2.26
2.23
6
3.08
2.92
2.82
2.74
2.68
2.64
2.60
2.58
2.55
2.53
2.51
2.50
2.49
2.48
2.47
2.46
2.43
2.40
2.37
2.35
2.32
2.29
7
3.16
3.00
2.89
2.81
2.75
2.70
2.67
2.64
2.61
2.59
2.57
2.56
2.54
2.53
2.52
2.51
2.48
2.45
2.42
2.39
2.37
2.34
8
3.24
3.07
2.95
2.87
2.81
2.76
2.72
2.69
2.66
2.64
2.62
2.61
2.59
2.58
2.57
2.56
2.53
2.50
2.47
2.44
2.41
2.38
9
3'. 30
3.12
3.01
2.92
2.86
2.81
2.77
2.74
2.71
2.69
2.67
2.65
2.64
2.62
2.61
2.60
2.57
2.54
2.51
2.48
2.45
2.42
1
.37
.14
.00
.90
.82
.76
.72
.68
.65
.62
.60
.58
.57
.55
.54
.53
.49
.46
.42
.39
.36
.33
2
.90
.61
.42
.29
.11
.11
.06
.01
.97
.94
.91
.88
.86
.84
.83
.81
.77
.72
.68
.64
.60
.56
3
4.21
.88
.66
.51
.40
.31
.25
.19
.15
.11
.08
.05
.03
.01
.99
.97
.92
.87
.82
.78
.73
.68
4
.43
.07
.83
.87
.55
.45
.38
.32
.27
.23
.20
.17
.14
.12
.10
.08
.03
.97
.92
.87
2.82
2.77
a -.01
I
4.60
4.21
3.96
3.79
3.66
3.56
3.48
3.42
S.37
3.32
3.29
3.26
3.23
3.21
3.18
3.17
3.11
3.05
2.99
2.94
2.89
2.84
6
4.73
4.33
4.07
3.88
3.75
3.64
3.56
3.50
1.44
1.40
3.36
3.33
1.30
3.27
3.25
3.23
3.17
3.11
3.05
3.00
2.94
2.89
7
4.85
4.43
4.15
3.96
1.82
3.71
3.63
1.56
3.51
3.46
3.42
3.39
3.36
3.33
3.31
3.29
3.12
3.16
1.10
3.04
2.99
2.93
1
.94
.51
.23
.03
.89
.78
.69
.62
.56
.51
.47
.44
.41
.38
.36
.34
.27
.21
.14
.08
3.03
2.97
9
5.03
4.59
4.30
4.09
1.94
3.83
1.74
1.67
3.61
1.56
3.52
1.48
1.45
1.42
1.40
1.18
1.31
3.24
1.18
3.12
3.06
3.00
1From Miller, 1981.
115
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7. To quantify the sensitivity of the test, the minimum significant
difference (MSD) that can be detected statistically may be
calculated.
MSD = d Sw y (1/n,) + (1/n)
Where: d = the critical value for the Dunnett's procedure
Su = the square root of the within mean square
n = the common number of replicates at each concentration
(this assumes equal replication at each concentration)
n( = the number of replicates in the control.
8. In this example:
MSD = 2.41 (0.155) V (1/4) + (1/4)
= 2.41 (0.155)(0.707)
= 0.264
9. The MSD (0.264) is in transformed units. To determine the MSD in
terms of percent survival, carry out the following conversion.
1. Subtract the MSD from the transformed control mean.
1.330 - 0.264 = 1.066
2. Obtain the untransformed values for the control mean and the
difference calculated in 1.
[Sine ( 1.330) ]2 = 0.943
[Sine ( 1.066) ]2 = 0.766
3. The untransformed MSD (MSD ) is determined by subtracting the
untransformed values from 2.
MSDU = 0.943 - 0.766 = 0.177
10. Therefore, for this set of data, the minimum difference in mean
proportion surviving between the control and any toxicant
concentration that can be detected as statistically significant is
0.177.
11. This represents a decrease in survival of 19% from the control.
116
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SECTION 12
REPORT PREPARATION1
The following general format and content are recommended for the report:
12.1 INTRODUCTION
1. Permit number
2. Toxicity testing requirements of permit
3. Plant location
4. Name of receiving water body
5. Contractor (if contracted)
a. Name of firm
b. Phone number
c. Address
12.2 PLANT OPERATIONS
1. Product(s)
2. Raw materials
3. Operating schedule
4. Description of waste treatment
5. Schematic of waste treatment
6. Retention time (if applicable)
7. Volume of discharge (MGD, CFS, GPM)
8. Design flow of treatment facility at time of sampling
12.3 SOURCE OF EFFLUENT, RECEIVING WATER, AND DILUTION WATER
1. Effluent Samples
a. Sampling point
b. Sample collection method
c. Collection dates and times
d. Mean daily discharge on sample collection date
e. Lapsed time from sample collection to delivery
f. Sample temperature when received at the laboratory
g. Physical and chemical data
2. Receiving Water Samples
a. Sampling point
b. Sample collection method
c. Collection dates and times
d. Streamflow at time of sampling and 7Q10
e. Lapsed time from sample collection to delivery
f. Sample temperature when received at the laboratory
g. Physical and chemical data
1Adapted from USEPA, 1989c.
117
-------
3. Dilution Water Samples
a. Source
b. Collection date(s) and time(s) (where applicable)
c. Pretreatment
d. Physical and chemical characteristics (pH, hardness, salinity, etc.)
12.4 TEST CONDITIONS
1. Toxicity test method used (title, number, source)
2. Endpoint(s) of test
3. Deviations from reference method, if any, and reason(s)
4. Date and time test started
5. Date and time test terminated
6. Type and volume of test chambers
7. Volume of solution used per chamber
8. Number of organisms per test chamber
9. Number of replicate test chambers per treatment
10. Feeding frequency, and amount and type of food
11. Acclimation temperature of test organisms (mean and range)
12. Test temperature (mean and range)
12.5 TEST ORGANISMS
1. Scientific name
2. Age
3. Life stage
4. Mean length and weight (where applicable)
5. Source
6. Diseases and treatment (where applicable)
12.6 QUALITY ASSURANCE
1. Reference toxicant used routinely; source
2. Date and time of most recent reference toxicant test;
test results and current cusum chart
3. Dilution water used in reference toxicant test
4. Physical and chemical methods used
12.7 RESULTS
1. Provide raw toxicity data in tabular form, including daily
records of affected organisms in each concentration (including
controls)
2. Provide table of endpoints: LC50, NOAEC, Pass/Fail.
3. Indicate statistical methods used to calculate endpoints
4. Provide summary table of physical and chemical data
5. Tabulate QA data
12.8 CONCLUSIONS AND RECOMMENDATIONS
1. Relationship between test endpoints and permit limits.
2. Action to be taken.
118
-------
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Skarheim, H.P. 1973. Tables of the fraction of ammonia in the undissociated
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fishes. Spec. Publ. No. 5, Amer. Fish. Soc., Washington, D.C. 526 pp.
Sprague, J.B., and A. Fogels. 1977. Watch the Y in bioassay. Proceedings of
the 3rd Aquatic Toxicity Workshop, Halifax, N.S., Nov. 2-3, 1976.
Environm. Prot. Serv. Tech. Rpt. No. EPS-5-AR-77-1, Halifax, Canada.
pp.107-118.
Stephan, C.E. 1977. Methods for calculating an LC50. In: F.L. Mayer and
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27, 1982. Spec. Publ. Soc. Environ. Tox. Chem. Pergamon Press, New York,
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USEPA. 1972. Recommended bioassay procedure for fathead minnow Pimephales
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127
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APPENDICES
A. Distribution, Life Cycle, Taxonomy, and Culture Methods . . 131
1. Ceriodaphm'a dubia 131
2. Daphnia (Daphnia magna and D. pulex) 148
3. Mysids (Mysidopsis bahia) 169
4. Brine Shrimp (Artemia salina) 189
5. Fathead Minnows (Pimephales promelas) 198
6. Rainbow Trout (Oncorhynchus mykiss) and Brook
Trout (Salvelinus fontinalis) 217
7. Sheepshead Minnows (Cyprinodon van'egatus) 227
8. Silversides: Inland Silverside (Menidia
beryllina), Atlantic Silverside, (M. mem'dia,
and Tidewater Silverside (M. peninsulae) 246
B. Supplemental List of Acute Toxicity Test Species 263
C. DiTutor Systems 266
1. Solenoid and Vacuum Siphon Dilutor Systems 266
2. Solenoid System Equipment List 270
3. Vacuum System Equipment List 273
4. Dilutor Control Panel Equipment List 278
D. Plans for Mobile Toxicity Test Laboratory 279
1. Tandem-axle Trailer 279
2. Fifth-wheel Trailer 282
E. Check Lists and Information Sheets 283
1. Toxicity Test Field Equipment List 283
2. Information Check List for On-site Industrial
and Municipal Waste Toxicity Tests 285
3. Daily Events Log 291
4. Dilutor Calibration Form 292
5. Daily Dilutor Calibration Check 293
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APPENDIX A
SYSTEMATICS, ECOLOGY, LIFE HISTORY, AND CULTURE METHODS
A.I. CERIODAPHNIA DUBIA*
1. SYSTEMATICS
1.1. Morphology and Taxonomy
1.1.1 Cen'odaphnia are closely related and morphologically similar to
Daphm'a, but are smaller and have a shorter generation time (Berner, 1986).
They are generally more rotund, lack the prominent rostral projection typical
of Daphm'a, and do not develop the dorsal helmets and long posterior spines
often observed in Daphm'a.
1.1.2 With Cen'odaphnia dubia, the female has a heavy, setulated pecten on
the postabdominal claw (Figure 1A), and the male was long antennules (Figure
1C), in contrast to the closely related C. reticulata, where the female has
heavy, triangular denticles in the pecten of the postabdominal claw (Figure
2D), and the male has very short antennules (Figure 2C). Some clones having
intermediate characters may be hybrids or phenotypic variants of C. dubia
(Berner, 1986). Detailed descriptions of the males and females of both
species and the variant were given by Berner (1986).
1.1.3 Although males are very similar to females, they can be recognized by
their rapid, erratic swimming habit, smaller size, denser coloration,
extended antennules and claspers, and rostrum morphology.
2. ECOLOGY AND LIFE HISTORY
2.1 Distribution
2.1.1 C. dubia, has been reported from littoral areas of lakes, ponds, and
marshes throughout most of the world, but it is difficult to ascertain its
true distribution because it has been reported in the literature under several
other names (C. affim's, C. quadrangula, and C. reticulata. It has also been
suggested that reports of C. dubia in New Zealand and parts of Asia may be yet
another unnamed species (Berner, personal communication).
2.2 Ecology
2.2.1 Cen'odaphnia ecology and life history are very similar to those of
other daphnids. Specific information on the ecology and life history of
Cen'odaphnia dubia is either not available or is widely scattered throughout
Prepared by Philip A. Lewis and James M. Lazorchak.
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Figure 1. Cen'odaphnia dubia. A. (1) parthenogenetic female; (2) postabdomen,
and (3) claw; B. ephippial female; C. male. (From Berner, 1986)
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Figure 2. Cen'odaphm'a reticulata. A. (1) parthenogenetic female,
(2) postabdomen, (3) and claw; B. ephippial female;
C. male. (From Berner, 1986).
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the literature. However, it is known to be a pond and lake dwelling species
that is usually common among the vegetation in littoral areas (Fairchild,
1981). In the Lake of Velence, Hungary, C. dubia was most common in regions
where "grey" and "dark brown" waters merged (Pal, 1980). In Par Pond
(Savannah River Plant, Aiken, SC) the Ceriodaphm'a were much more abundant in
the heated water (effluent from the nuclear reactor) than in the ambient area
(Vigerstad and Tilly, 1977), and in a reservoir in Russia, animals from the
heated water were larger and heavier than those living under normal water
temperatures (Kititsyna and Sergeeva, 1976). In Iran they are common in
warmer, montane, oligotrophic lakes (Smagowicz, 1976).
2.2.2 In Lake Kinneret, Israel, Cen'odaphnia reticulata are abundant only
between March and June, with a peak in May when the temperature ranges between
20 and 22°C. When summer temperatures reached 27-28°C, the Ceriodaphm'a were
reduced in size and egg production became significantly less, leading to a
progressive decline of the population (Gophen, 1976). In Lake Parvin, France,
the period of development was from June to September (Devaux, 1980).
2.2.3 Ceriodaphm'a typically swim with an erratic, jerking motion for a
period of time, and hang motionless in the water between swimming bouts. This
swimming behavior results in a mean speed of 1.5-2.5 mm/s. When approached by
a predator, however, it flees by swimming away quickly along a straight path
(Wong, 1981).
2.2.4 During most of the year, populations of Ceriodaphm'a consist almost
entirely of females; the males appearing principally in autumn. Production of
males appears to be induced primarily by low water temperatures, high
population densities, and/or a decrease in available food. As far as is
presently known, C. dubia reproduce only by cyclic parthenogenesis in which
the males contribute to the genetic makeup of the young during the sexual
stage of reproduction.
2.2.5 The females tend to aggregate during sexual reproductive activity, when
ephippia are produced (Brandl and Fernando, 1971). Ephippia are embryos
encased in a tough covering, and are resistent to drying. They can be stored
for long periods and shipped through the mail in envelopes, like seeds. When
placed in water at the proper temperature, ephippia hatch in a few days
producing a new parthenogenetic population.
2.2.6 Ceriodaphm'a have many predators, including fish, the mysid Mysis
relicta, Chaoborus larvae, and copepods. As with Daphnia, it also reacts to
intense predation with defensive strategies. Ceriodaphm'a reticulata
(possibly C.dubia) in a Minnesota lake, reacted to the copepod, Cyclops
vernal is, by producing large offspring and growing to a large size at the
expense of early reproduction (Lynch, 1979). They reacted to fish predators
by producing smaller offspring in larger numbers.
2.3 Food and Feeding
2.3.1 Cladocera are polyphagous feeders and find their food in the seston.
Daphnids, including the Ceriodaphm'a, are classified as fine mesh filter
feeders by Geller and Mueller (1981). These fine mesh filter feeders are most
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abundant in eutrophic lakes during summer phytoplankton blooms when suspended
bacteria are available as food only for filter-feeding species with fine mesh.
2.3.2 Lynch (1978) examined the gut contents of Cen'odaphnia reticulata
(possibly C. dubia) from a Minnesota pond and found bacteria, detritus and
partially digested algae. In this pond, Cen'odaphnia and Daphm'a pulex shared
the same resource base and had very similar diets, but the Cen'odaphnia fed
more intensively on diatoms. The Cen'odaphnia were considered to be less
sensitive to low food levels than Daphm'a, because of their high rate of
population growth during periods of low food levels in late summer.
2.4 Life Cycle
2.4.1 Four distinct periods may be recognized in the life cycle of
Cen'odaphnia: (1) egg, (2) juvenile, (3) adolescent, and (4) adult. The life
span of Cen'odaphnia, from the release of the egg into the brood chamber until
the death of the adult, is highly variable depending on the temperature and
other environmental conditions. Generally the life span increases as
temperature decreases, due to lowered metabolic activity. For example, the
average life span of Cen'odaphnia dubia is about 30 days at 25°C, and 50 days
at 20 C. One female was reported to have lived 125 days and produced 29
broods at 20°C (Cowgill et al., 1985).
2.4.2 Typically, a clutch of 4 to 10 eggs is released into the brood chamber,
but clutches with as many as 20 eggs are common. The eggs hatch in the brood
chamber and the juveniles, which are already similar in form to the adults,
are released in approximately 38 h, when the female molts (casts off her
exoskeleton or carapace). The total number of young produced per female
varies with temperature and other environmental conditions. The most young
are produced in the range of 18-25°C (124 young per female in a 28-day life
span at 24°C) (113 young per female in a 77-day life span at 18°C) but
production falls off sharply below 18°C (13 young per female in a 24-day life
span at 12°C) (McNaught and Mount, 1985).
2.4.3 The time required to reach maturity (produce their first offspring) in
C.dubia varies from three to five days and appears to be dependent on body
size and environmental conditions. A study of the growth and development of
parthenogenetic eggs by Shuba and Costa (1972) revealed that at 24°C the
embryos matured to free-swimming juveniles in approximately 38 h. The eggs
that did not develop fully usually were aborted after 12 hours.
2.4.4 The growth rate of the organism is greatest during its juvenile stages
(early instars), and the body size may double during each of these stages.
Each instar stage is terminated by a molt. Growth occurs immediately after
each molt while the new carapace is still elastic.
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2.4.5 Following the juvenile stages, the adolescent period is very short, and
consists of a single instar. It is during the adolescent instar that the
first clutch of eggs reaches full development in the ovary. Generally, eggs
are deposited in the brood chamber within minutes after molting, and the young
which develop are released just before the next molt.
2.4.6 In general, the duration of instars increases with age, but also
depends on environmental conditions. A given instar usually lasts
approximately 24 h under favorable conditions. However, when conditions are
unfavorable, it may last as long as a week. Four events take place in a
matter of a few minutes at the end of each adult instar: (1) release of young
from the brood chamber to the outside, (2) molting, (3) increase in size, and
(4) release of a new clutch of eggs into the brood chamber. The number of
young per brood is highly variable, depending primarily on food availability
and environmental conditions. C.dubia may produce as many as 25 young in a
single brood, but more commonly the number is six to ten. The number of young
released during the adult instars reaches a maximum at about the fourth
instar, after which there is a gradual decrease.
3. CULTURING METHODS
3.1 Cen'odaphm'a are available from commercial biological supply houses.
Guidance on the source of culture animals to be used by a permittee for self-
monitoring effluent toxicity tests should be obtained from the permitting
authority. Only a small number of organisms (20-30) are needed to start a
culture. Before test organisms are taken from a culture, the culture should
be maintained for at least two generations using the same food, water, and
temperature as will be used in the toxicity tests.
3.2 Cultures of test organisms should be started at least three weeks before
the brood animals are needed, to ensure an adequate supply of neonates for the
test. Only a few individuals are needed to start a culture because of their
prolific reproduction.
3.3 Starter animals may be obtained from an outside source by shipping in
polyethylene bottles. Approximately 20-30 animals and 3 mL of food (see
below) are placed in a 1-L bottle filled full with culture water. Animals
received from an outside source should be transferred to new culture media
gradually over a period of 1-2 days to avoid mass mortality.
3.4 It is best to start the cultures with one animal, which is sacrificed
after producing young, embedded, and retained as a permanent microscope slide
mount to facilitate identification and permit future reference. The species
identification of the stock culture should be verified by a taxonomic
authority. The following procedure is recommended for making slide mounts of
Cen'odaphm'a (Beckett and Lewis, 1982):
1. Pipet the animal onto a watch glass.
2. Reduce the water volume by withdrawing excess water with the
pi pet.
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3. Add a few drops of carbonated water (club soda or seltzer
water) or 70% ethanol to relax the specimen so that the
post-abdomen is extended. (Optional: with practice,
extension of the postabdomen may be accomplished by putting
pressure on the cover slip).
4. Place a small amount (one to three drops) of mounting medium
on a glass microscope slide. The recommended mounting
medium is CMCP-9/9AF Medium, prepared by mixing two parts
of CMCP-9 with one part of CMCP-9AF. For more viscosity and
faster drying, CMC-10 stained with acid fuchsin may be used.
CMCP-9 and 9AF are available from Polysciences, Inc., Paul Valley
Industrial Park, Warrington, Pennsylvania, 18976 (215-343-6484).
5. Using a forceps or a pipet, transfer the animal to the drop
of mounting medium on the microscope slide.
6. Cover with a cover slip and exert minimum pressure to remove
any air bubbles trapped under the cover slip. Slightly more
pressure will extend the postabdomen.
7. Allow mounting medium to dry.
8. Make slide permanent by placing CMC-10 around the edges of
the coverslip.
9. Identify to species (see Pennak, 1989; Berner, 1985).
10. Label with waterproof ink or diamond pencil.
11. Store for permanent record.
3.5 Culture Media
3.5.1 Although Cen'odaphm'a stock cultures can be successfully maintained in
some tap waters, well waters, and surface waters, use of synthetic water as
the culture medium is recommended because (1) it is easily prepared, (2) it is
of known quality, (3) it yields reproducible results, and (4) allows adequate
growth and reproduction. Culturing may be successfully done in hard,
moderately hard or soft reconstituted water, depending on the hardness of the
water in which the test will be conducted. The quality of the dilution water
is extremely important in Cen'odaphm'a culture. The use of Millipore Milli-QR
or Super-QR, or equivalent, to prepare reconstituted water is highly
recommended. The use of diluted mineral water (DMW) for culturing and testing
is widespread due to the ease of preparation.
3.5.2 The chemicals used and instructions for preparation of reconstituted
water are given in Section 7. The compounds are dissolved in distilled or
deionized water and the media are vigorously aerated for several hours before
using. The initial pH of the media is between 7.0 and 8.0, but it will rise
as much as 0.5 unit after the test is underway.
3.6 MASS CULTURE
3.6.1 Mass cultures are used only as a "backup" reservoir of organisms.
Neonates from mass cultures are not to be used directly in toxicity tests.
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3.6.2 One-liter or 21 glass beakers, crystallization dishes, "battery jars,"
or aquaria may be used as culture vessels. Vessels are commonly filled to
three-fourths capacity. Cultures are fed daily. Four or more cultures are
maintained in separate vessels and with overlapping ages to serve as back-up
in case one culture is lost due to accident or other unanticipated problems,
such as low DO concentrations or poor quality of food or laboratory water.
3.6.3 Mass cultures which will serve as a source of brood organisms for
individual culture should be maintained in good condition by frequent renewal
of the medium and brood organisms. Cultures are started by adding 40-50
neonates per liter of medium. The stocked organisms should be transferred to
new culture medium at least twice a week for two weeks. After two weeks, the
culture is discarded and re-started with neonates in fresh medium. Using this
schedule, 1-L cultures will produce 500 to 1000 neonate Ceriodaphnia each
week.
3.6.4 Reserve cultures also may be maintained in large (80-L) aquaria or
other large tanks.
3.7 INDIVIDUAL CULTURE
3.7.1 Individual cultures are used as the immediate source of neonates for
toxicity tests.
3.7.2 Individual organisms are cultured in 15 mL of culture medium in 30-mL
(1 oz) plastic cups or 30-mL glass beakers. One neonate is placed in each
cup. It is convenient to place the cups in the same type of board used for
toxicity tests (see Figure 1 in this Section).
3.7.3 Organisms are fed daily and are transferred to fresh medium a minimum
of three times a week, typically on Monday, Wednesday, and Friday. On the
transfer days, food is added to the new medium immediately before or after the
organisms are transferred.
3.7.4 To provide cultures of overlapping ages, new boards are started weekly,
using neonates from adults which produce at least eight young in their third
or fourth brood. These adults can be used as sources of neonates until 14
days of age. A minimum of two boards are maintained concurrently to provide
backup supplies of organisms in case of problems.
3.7.5 Cultures which are properly maintained should produce at least 20 young
per adult in three broods (seven days or less at 25°C). Typically, 60 adult
females (one board) will produce more than the minimum number of neonates
(120) required for two tests.
3.7.6 Records should be maintained on the survival of brood organisms and
number of offspring at each renewal. Greater than 20% mortality of adults or
less than an average of 20 young per adult on a board at 25°C during a
one-week period would indicate problems, such as poor quality of culture media
or food. Organisms on that board should not be used as a source of test
organisms.
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3.8 CULTURE MEDIUM
3.8.1 Moderately hard synthetic water prepared using MILLIPORE MILLI-QR or
equivalent deionized water and reagent grade chemicals or 20% DMW is recommended
as a standard culture medium (see Section 7, Dilution Water).
3.9 CULTURE CONDITIONS
3.9.1 Ceriodaphm'a should be cultured at the temperature at which they will be
used in the toxicity tests (20°C or 25°C ± 2°C).
3.9.2 Day/night cycles prevailing in most laboratories will provide adequate
illumination for normal growth and reproduction. A 16-h/8-h day/night cycle is
recommended.
3.9.3 Clear, double-strength safety glass or 6 mm plastic panels are placed on
the culture vessels to exclude dust and dirt, and reduce evaporation.
3.9.4 The organisms are delicate and should be handled as carefully and as
little as possible so that they are not unnecessarily stressed. They are
transferred with a pipet of approximately 2-mm bore, taking care to release the
animals under the surface of the water. Any organism that is injured during
handling should be discarded.
3.10 FOOD PREPARATION AND FEEDING
3.10.1 Feeding the proper amount of the right food is extremely important in
Ceriodaphm'a culturing. The key is to provide sufficient nutrition to support
normal reproduction without adding excess food which may reduce the toxicity of
the test solutions, clog the animal's filtering apparatus, or greatly decrease
the DO concentration and increase mortality. A combination of Yeast,
CEROPHYLLR, and Trout chow (YCT) or flake food, along with the unicellular green
alga, Selenastrum capn'cornutum, will provide suitable nutrition if fed daily.
3.10.2 The YCT and algae are prepared as follows:
3.10.2.1 Digested trout chow (or flake food):
1. Preparation of trout chow requires one week. Use starter or No. 1 pellets
prepared according to current U.S. Fish and Wildlife Service
specifications, or flake food. Suppliers of trout chow include Zeigler
Bros., Inc., P.O. Box 95, Gardners, Pennsylvania, 17324 (717-780-9009);
Glencoe Mills, 1011 Elliott, Glencoe, Minnesota, 55336 (612-864-3181); and
Murray Elevators, 118 West 4800 South, Murray, Utah 84107 (800-521-9092).
2. Add 5.0 g of trout chow pellets or flake food to 1 L of MILLI-QR water. Mix
well in a blender and pour into a 2-L separatory funnel. Digest prior to
use by aerating continuously from the bottom of the vessel for one week at
ambient laboratory temperature. Water lost due to evaporation is replaced
during digestion. Because of the offensive odor usually produced during
digestion, the vessel should be placed in a fume hood or other isolated,
ventilated area.
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3. At the end of digestion period, place in a refrigerator and allow to
settle for a minimum of 1 h. Filter the supernatant through a fine mesh
screen (i.e., NITEXR 110 mesh). Combine with equal volumes of
supernatant from CEROPHYLLR and yeast preparations (below). The
supernatant can be used fresh, or frozen until use. Discard the
sediment.
3.10.2.2 Yeast:
1. Add 5.0 g of dry yeast, such as FLEISCHMANN'SR to 1 L of MILLI-QR
water.
2. Stir with a magnetic stirrer, shake vigorously by hand, or mix with a
blender at low speed, until the yeast is well dispersed.
3. Combine the yeast suspension immediately (do not allow to settle) with
equal volumes of supernatant from the trout chow (above) and CEROPHYLLR
preparations (below). Discard excess material.
3.10.2.3 CEROPHYLL" (Dried, Powdered, Cereal Leaves):
1. Place 5.0 g of dried, powdered, cereal leaves in a blender. (Available as
"CEREAL LEAVES," from Sigma Chemical Company, P.O. Box 14508, St. Louis,
Missouri, 63178, (800-325-3010); or as CEROPHYLL", from Ward's Natural
Science Establishment, Inc., P.O. Box 92912, Rochester, New York,
14692-9012, (716-359-2502). Dried, powdered, alfalfa leaves obtained from
health food stores have been found to be a satisfactory substitute for
cereal leaves.
Add 1 L of MILLI-Q" water.
Mix in a blender at high speed for 5 min, or stir overnight at medium
speed on a magnetic stir plate.
4. If a blender is used to suspend the material, place in a refrigerator
overnight to settle. If a magnetic stirrer is used, allow to settle for
1 h. Decant the supernatant and combine with equal volumes of
supernatant from trout chow and yeast preparations (above). Discard
excess material.
3.10.2.4 Combined YCT Food:
1. Mix equal (approximately 300 mL) volumes of the three foods as described
above.
2. Place aliquots of the mixture in small (50-mL to 100-mL) screw-cap
plastic bottles and freeze until needed.
3. Freshly prepared food can be used immediately, or it can be frozen until
needed. Thawed food is stored in the refrigerator between feedings, and
is used for a maximum of two weeks.
4. It is advisable to measure the dry weight of solids (dry 24 h at 105°C)
in each batch of YCT before use. The food should contain 1.7 - 1.9 g
solids/L. Cultures or test solutions should contain 12-13 mg solids/L.
2.
3.
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3.10.3 Algal (Selenastrum) Food
3.10.3.1 Algal Culture Medium
1. Prepare (five) stock nutrient solutions using reagent grade chemicals as
de-serHs^d in Table 1.
2/Add 1 mp of each stock solution, in the order listed in Table 1, to
StpfH=ex1mately 900 ml of MILLI-Q water. Mix well after the addition of
each solution. Dilute to 1 L and mix well. The final concentration of
macronutrients and micronutrients in the culture medium is given in
Table 2.
filter the medium through a 0.45um pore diameter
a vacuum of not more than 380 mm (15 in.) mercury, or at a
Immediately
membrane at
pressure of
with 500 ml
If the
can be
not more than one-half atmosphere (8 psi). Wash the filter
deionized water prior to use .
filtration is carried out with sterile apparatus, filtered medium
used immediately, and no further sterilization steps are required
before the inoculation of the medium. The medium can also be sterilized
by autoclaving after it is placed in the culture vessels.
5. Unused sterile medium should not be stored more than one week prior to
use, because there may be substantial loss of water by evaporation.
3.10.3.2 Algal Cultures
3.10.3.2.1 Two types of algal cultures are maintained: (1) stock cultures,
and,(2) "food" cultures.
3.10.3.2.2 Establishing and Maintaining Stock Cultures of Algae
1. Upon receipt of the "starter" culture (usually about 10 ml), a stock
culture is initiated by aseptically transferring one milliliter to each
of several 250-mL culture flasks containing 100 ml algal culture medium
(prepared as described above). The remainder of the starter culture can
be held in reserve for up to six months in a refrigerator (in the dark)
at 4°C.
2. The stock cultures are used as a source of algae to initiate "food"
cultures for Ceriodaphnia toxicity tests. The volume of stock culture
maintained at any one time will depend on the amount of algal food
required for the Ceri'odaphm'a cultures and tests. Stock culture volume
may be rapidly "scaled up" to several liters, if necessary, using 4-L
serum bottles or similar vessels, each containing 3 L of growth medium.
3. Culture temperature is not critical. Stock cultures may be maintained in
environmental chambers with cultures of other organisms if the
illumination is adequate (continuous "cool-white" fluorescent lighting of
approximately 86 ± 8.6 uE/m/s, or 400 ft-c).
4. Cultures are mixed twice daily by hand or stirred continuously.
5. Stock cultures can be held in the refrigerator until used to start "food"
cultures, or can be transferred to new medium weekly. One-to-three
milliliters of 7-day old algal stock culture, containing approximately
1.5 X 10 cells/ml, are transferred to each 100 mL of fresh culture
medium. The inoculum should provide an initial cell density of
approximately 10,000-30,000 cells/ml in the new stock cultures. Aseptic
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techniques should be used in maintaining the stock algal cultures, and
care should be exercised to avoid contamination by other microorganisms.
6. Stock cultures should be examined microscopically weekly, at transfer,
for microbial contamination. Reserve quantities of culture organisms can
be maintained for 6-12 months if stored in the dark at 4°C. It is
advisable to prepare new stock cultures from "starter" cultures obtained
from established outside sources of organisms every four
to six months.
3.10.3.2.3 Establishing and Maintaining "Food" Cultures of Algae
1. "Food" cultures are started seven days prior to use for Cen'odaphm'a
cultures and tests. Approximately 20 ml of 7-day-old algal stock culture
(described in the previous paragraph), containing 1.5 X 106 cells/ml,
are added to each liter of fresh algal culture medium (i.e., 3 L of
medium in a 4-L bottle, or 18 L in a 20-L bottle). The inoculum should
provide an initial cell density of approximately 30,000 cells/ml. Aseptic
techniques should be used in preparing and maintaining the cultures, and
care should be exercised to avoid contamination by other microorganisms.
However, sterility of food cultures is not as critical as in stock
cultures because the food cultures are terminated in 7-10 days. A
one-month supply of algal food can be grown at one time, and the excess
stored in the refrigerator.
2. Food cultures may be maintained at 25°C in environmental chambers with
the algal stock cultures or cultures of other organisms if the
illumination is adequate (continuous "cool-white" fluorescent lighting of
approximately 86 + 8.6 uE/m2/s, or 400 ft-c).
3. Cultures are mixed continuously on a magnetic stir plate (with a medium
size stir bar) or in a moderately aerated separatory funnel, or are mixed
twice daily by hand. If the cultures are placed on a magnetic stir
plate, heat generated by the stirrer might elevate the culture
temperature several degrees. Caution should be exercised to prevent the
culture temperature from rising more than 2-3°C.
3.10.3.3 Preparing Algal Concentrate for Use as Cen'odaphm'a Food
1. An algal concentrate containing 3.0 to 3.5 X 107 cells/ml is prepared
from food cultures by centrifuging the algae with a plankton or
bucket-type centrifuge, or by allowing the cultures to settle in a
refrigerator for approximately two-to-three weeks and siphoning off the
supernatant.
2. The cell density (cells/ml) in the concentrate is measured with an
electronic particle counter, microscope and hemocytometer, fluorometer,
or spectrophotometer, and used to determine the concentration required to
achieve a final cell count of 3.0 to 3.5 X 107/mL.
3. Assuming a cell density of approximately 1.5 X 106 cells/ml in the
algal food cultures at 7 days, and 100% recovery in the concentration
process, a 3-L, 7-10 day culture will provide 4.5 X 109 algal cells.
This number of cells would provide approximately 150 mL of algal cell
concentrate (1500 feedings at 0.1 ml/feeding) for use as food. This
would be enough algal food for four Cen'odaphm'a tests.
4. Algal concentrate may be stored in the refrigerator for one month.
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TABLE 1. NUTRIENT STOCK SOLUTIONS FOR MAINTAINING ALGAL STOCK CULTURES
AND TEST CONTROL CULTURES.
STOCK
SOLUTION
1. MACRONUTRIENTS
A.
B.
C.
D.
2. MICRONUTRIENTS
COMPOUND
MgCl2.6H20
CaCl2.2H20
NaN03
MgS04.7H20
K2HP04
NaHC03
•
H,B03
MnCl2.4H20
ZnCl2
FeCl,.6H20
CoCl2.6H20
Na2MoO,.2H20
CuCl2.2H20
Na2EDTA.2H20
Na2Se04
AMOUNT DISSOLVED IN
500 ML MILLI-Q* WATER
6.08 g
2.20 g
12.75 g
7.35 g
0.522 g
7.50 g
92.8 mg
208.0 mg
1.64 mga
79.9 mg
0.714 mgb
3.63 mgc
0.006 mgd
150.0 mg
1.196 mge
aZnd2 - Weigh out 164 mg and dilute to 100 mL. Add 1 mL of this
solution to Stock #1.
bCod2.6H20 - Weigh out 71.4 mg and dilute to 100 mL. Add 1 mL of
this solution to stock #1.
cNa2Mo04.2H20 - Weigh out 36.6 mg and dilute to 10 mL. Add 1 mL
of this solution to stock #1.
dCud2.2H20 - Weigh out 60.0 mg and dilute to 1000 mL. Take 1 mL of
this solution and dilute to 10 mL. Take 1 mL of the second dilution and
add to Stock #1.
6Na2Se04 - Weigh out 119.6 mg and dilute to 100 mL. Add 1 mL of this solution
to Stock #1.
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TABLE 2. FINAL CONCENTRATION OF MACRONUTRIENTS AND MICRONUTRIENTS
IN THE CULTURE MEDIUM
MACRONUTRIENT
NaN03
MgCl2.6H20
CaCl2.2H20
MgS04.7H20
K2HP04
NaHC03
MICRONUTRIENT
H3B03
MnCl2.4H20
ZnCl2
CoCl2.6H20
CuCl2.2H20
Na2Mo04.2H20
FeCl3.6H20
Na2EDTA.2H20
Na,SeO,
CONCENTRATION
(MG/L)
25.5
12.2
4.41
14.7
1.04
15.0
CONCENTRATION
(UG/L)
185
416
3.27
1.43
0.012
7.26
160
300
2.39
ELEMENT
N
Mg
Ca
S
P
Na
K
C
ELEMENT
B
Mn
Zn
Co
Cu
Mo
Fe
--
Se
CONCENTRATION
(MG/L)
4.20
2.90
1.20
1.91
0.186
11.0
0.469
2.14
CONCENTRATION
(UG/L)
32.5
115
1.57
0.354
0.004
2.88
33.1
1.00
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3.4 FEEDING
3.4.1 Cultures should be fed daily to maintain the organisms in optimum
condition so as to provide maximum reproduction. Stock cultures which are
stressed because they are not adequately fed may produce low numbers of young,
large number of males, and ephippial females. Also, their offspring may
produce few young when used in toxicity tests.
1. If YCT is frozen, remove a bottle of food from the freezer 1 h before
feeding time, and allow to thaw.
2. Mass cultures are fed daily at the rate of 7 ml YCT and 7 mL algae
concentrate/L culture.
3. Individual cultures are fed at the rate of 0.1 ml YCT and 0.1 mL algae
concentrate per 15 mL culture.
4. YCT and algal concentrate should be thoroughly mixed by shaking before
dispensing.
5. Return unused YCT food mixture and algae concentrate to the refrigerator.
Do not re-freeze YCT. Discard unused portion after one week.
3.5 FOOD QUALITY
3.5.1 The quality of food prepared with newly acquired supplies of yeast, trout
chow, dried cereal leaves, or algae, should be determined in side-by-side
comparisons of Cen'odaphm'a survival and reproduction, using the new food and
food of known, acceptable quality, over a seven-day period in control medium.
3.6 VIDEO TRAINING TAPE AVAILABLE FOR CULTURING METHODS
3.6.1 A video training tape and supplemental report (USEPA, 1989) on culturing
Cen'odaphm'a dubia are available from the National Audiovisual Center, Customer
Services Section, 8700 Edgeworth Drive, Capitol Heights, Maryland, 20743-3701,
(Phone 301-763-1891), as part of a video package on short-term chronic toxicity
tests for freshwater organisms (Order No. EPA18036), which costs $45.00.
4. TEST ORGANISMS
4.1 Neonates, or first instar Cen'odaphm'a less than 24 hours old, taken from
the 3rd or 4th brood, are used in toxicity tests. To obtain the necessary
number of young for an acute toxicity test, it is recommended that the animals
be cultured in individual 30 mL beakers or plastic cups for seven days prior to
the beginning of the test. Neonates are used from broods of at least eight
young. Fifty adults in individual cultures will usually supply enough neonates
for one toxicity test.
4.2 Use a disposable, widemouth pipette to transfer Cen'odaphm'a . The
diameter of the opening should be approximately 4 mm. The tip of the pipette
should be kept under the surface of the water when the Cen'odaphm'a are released
to prevent air from being trapped under the carapace. Liquid containing adult
Cen'odaphm'a can be poured from one container to another without risk of
injuring the animals.
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SELECTED REFERENCES
Brand!, Z. and C.H. Fernando. 1971. Microaggregation of the cladoceran
Cen'odaphnia affinis Lilleborg with a possible reason for
microaggregation of zooplankton. Can. J. Zool. 49:775.
Cowgill, U.M., K.I. Keating and I.T. Takahashi. 1985. Fecundity and
longevity of Cen'odaphnia dubia/affinis in relation to diet at two
different temperatures. J. Crust. Biol. 5(3):420-429.
Devaux, J. 1980. Contribution to limnological study of Lake Pavin, France.
2: Relationship between abiotic parameters, phytoplankton and
zooplankton in the 0-20 meter zone. Hydrobiologia 68(l):17-34.
Fairchild, G.W. 1981. Movement and micro distribution of Sidia crystalline
and other littoral micro Crustacea. Ecology 62(5)-.1341-1352.
Geller, W. and H. Mueller. 1981. The filtration apparatus of Cladocera
filter mesh sizes and their implications on food selectivity. Oecologia
(Berl.) 49(3):316-321.
Gophen, M. 1976. Temperature dependence of food intake, ammonia excretion,
and respiration in Cen'odaphnia reticulata, Lake Kinneret, Israel.
Freshwat. Biol. 6(5):451-455.
Kititsyna, L.A. and O.A. Sergeeva. 1976. Effect of temperature increase on
the size and weight of some invertebrate populations in the cooling
reservoir of the Kinakhovsk State Regional Electric Power Plant.
Ekologyia 5:99-102.
Lynch, M. 1978. Complex interactions between natural coexploiters -
Daphm'a and Cen'odaphnia. Ecology 59(3) :552-564.
Lynch, M. 1979. Predation, competition, and zooplankton community
structure: An experimental study. Limnol. Oceanogr. 24(2):253-272.
McNaught, D.C. and D.I. Mount. 1985. Appropriate durations and measures for
Cen'odaphnia toxicity tests. In: R.C. Bahner and D.J. Hansen (eds.),
Aquatic Toxicology and Hazard Assessment: Eighth Symposium. ASTM STP
891, American Society for Testing and Materials, Philadelphia,
Pennsylvania, pp. 375-381.
Norberg, T.J. and D.I. Mount. 1985. Diets for Cen'odaphnia reticulata
life-cycle tests. In: R.D. Cardwell, R. Purdy and R.C. Bahner (eds.),
Aquatic Toxicology and Hazard Assessment: Seventh Symposium. ASTM STP
854. American Society for Testing and Materials, Philadelphia,
Pennsylvania pp. 42-52.
146
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Pal, G. 1980. Characterization of water quality regions of the Lake of
Velence, Hungary with planktonic crustaceans. Allattani Kozl.
67(l-4):49-58.
Pennak, R.W. 1989. Fresh-water invertebrates of the United States. 3rd ed.
Protozoa to Mollusca. John Wiley & Sons, New York, NY.
Shuba, T. and R.R. Costa. 1972. Development and growth of Cen'odaphnia
reticulata embryos. Trans. Am. Microsc. Soc. 91(3):429-435.
Smagowicz, K. 1976. On the zooplankton of Lake Zeribar, Western Iran. Acta
Hydrobiol. 18(1):89-100.
USEPA. 1986. Taxonomy of Cen'odaphnia (Crustacea: Cladocera) in U.S.
Environmental Protection Agency cultures. D.B. Berner. U. S.
Environmental Protection Agency, Environmental Monitoring and Support
Laboratory, Cincinnati, OH 45268. EPA/600/4-86/032
USEPA. 1989. Culturing of Cen'odaphnia dubia. Supplemental report for video
training tape. U. S. Environmental Protection Agency, Washington, D.C.
EPA/505/8-89/002a.
Vigerstad, T.J. and L.J. Tilly. 1977. Hyper-thermal effluent effects on
heleo planktonic Cladocera and the influence of submerged macrophytes.
Hydrobiologia 55(l):81-86.
Wong, C.K. 1981. Predatory feeding behavior of Epischura lacustris
(Copepoda: Calanoida) and prey defense. Can. J. Fish. Aquat. Sci.
38:275-279.
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APPENDIX A
SYSTEMATICS, ECOLOGY, LIFE HISTORY, AND CULTURE METHODS
A.2. DAPHNIA (D. MAGNA AND D. PULEX)^
1. SYSTEMATICS
1.1 Morphology and Taxonomy
1.1.1 The generalized anatomy of a parthenogenetic female is shown in
Figure 1. Daphm'a pulex is an extremely variable species consisting of
several reproductively isolated clonal groups and is often not distinguishable
from other species (such as D. obtusa) that have large teeth on the middle
pecten of the postabdomenal claw (Figure 2C) (Lynch, 1985; Dodson, 1981).
Probably the most distinctive feature of the parthenogenetic female D. pulex
is the long second abdominal process of the abreptor (postabdomen) that
extends beyond the base of the anal setae (Figure 2A).
1.1.2 D. pulex is a wide ranging species that shows little variation
throughout its range. Two of its most distinctive characteristics are the
deeply sinuate posterior margin of the abreptor (Figures 3A and 3D) and the
ridges on the head which run parallel to the mid-dorsal line (Figure 3B).
1.1.3 D. pulex is much smaller than D. magna, attaining a length of up to 3.5
mm compared to 5.0 or 6.0 mm for D. magna. Although the two species can often
be separated by size, they can be differentiated with certainty only by
examining the postabdominal claws for size and number of spines using a
compound microscope. D. pulex has 5-7 stout teeth on the middle pecten
(Figure 2C) while D. magna has a uniform row of 20 or more small teeth (Figure
3E). Another characteristic for separating the neonates of the two species is
the location of the nuchal organ which is higher up on the posterior margin of
the head in D. magna than in D. pulex (Schwartz and Hebert, 1984). For a more
complete taxonomic discussion of the two species see Brooks (1957).
2. DISTRIBUTION
2.1 D. magna has a worldwide distribution in the northern hemisphere. In
North America it appears to be absent from the eastern United States (except
for Northern New England) and Alaska (Figure 4). D. pulex occurs over most of
North America except the tropics and high arctic (Figure 5), and probably
occurs in Europe and South America as well. Both species often occur in the
same pools but D. pulex usually out-competes D. magna in mixed populations and
takes over as the sole inhabitant by summer's end (Modi in, 1982; Lynch, 1983).
Prepared by Philip A. Lewis and James M. Lazorchak.
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Figure 1. Generalized anatomy of a female Daphm'a, X70; A, antenna; AS, anal
setae; BC, brood chamber; H, heart; INT, intestine; L, legs; 0V,
ovary; P, postabdomen; PC, postabdominal claw. (From Pennak, 1989)
--777777
Figure 2. Female Daphm'a pulex. A, lateral aspect (note smoothly rounded
posterior margin of postabdomen); B, ephippial female; C,
postabdomen showing large spines on the claw. (From Brooks, 1957)
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Figure 3. Female D. magna. A. Lateral aspect; B. dorsal aspect; C. ephippial
female; D. postabdomen showing sinuate posterior margin; E.
postabdominal claw. (From Brooks, 1957).
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i\'t^^^^^r\Y N *t(W£>
>7
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3. ECOLOGY AND LIFE HISTORY
3.1 General Ecology
3.1.1 D. magna is principally a lake dweller and is restricted to waters in
northern and western North America exceeding a hardness of 150 mg/L (as CaCO,)
(Pennak, 1989). In the Netherlands, D. magna are found in shallow ponds witn
muddy bottoms rich in organic matter and with low oxygen demand (3 to 4 mg/L).
D. pulex is principally a pond dweller where the oxygen content is higher, but
is also found in lakes. It is generally considered a clean water species
being dominant in nature during periods of low turbidity. However, Scholtz,
Seaman and Pieterse (1988) found that high turbidity had little effect on
survival and reproduction in laboratory studies.
3.1.2 Daphm'a populations are generally sparse in winter and early spring,
but as water temperatures reach 6°C to 12°C, they increase in abundance and
subsequently may reach population densities as high as 200 to 500
individuals/! (Pennak, 1989). Populations in ponds decline to very low
numbers during the summer months. In autumn there may be a second population
pulse, followed by a decline to winter lows.
3.1.3 During most of the year, populations of Daphm'a consist almost entirely
of females, the males being abundant only in spring or autumn when up to 56%
of the offspring of D. magna may be males (Barker and Hebert, 1986). Males
are distinguished from females by their smaller size, larger antennules,
modified postabdomen, and first legs, which are armed with a stout hook used
in clasping. Production of males appears to be induced principally by low
temperatures or high densities and subsequent accumulation of excretory
products, and/or a decrease in available food. These conditions may induce
the appearance of sexual (resting) eggs (embryos) in cases called ephippia
(Figures 2B and 3C), which are cast off during the next molt. It appears that
the shift toward male and sexual egg production is related to the metabolic
rate of the parent. Any factor which tends to lower metabolism may be
responsible. Ruvinsky et al. (1978) suggested that the genome of the animal
has two developmental programs based on identical sets of chromosomes. The
female program consistently functions under a wide range of conditions,
whereas the male program is turned on by specific ecological stimuli. The
eggs from which the males and females develop have identical chromosome sets.
Sex determination is based on changes in chromatin structure when the mother
receives a specific signal that sexual reproduction is needed for adaptation
to extreme conditions.
3.1.4 D. magna reproduce only by cyclic parthenogenesis in which males
contribute to the genetic makeup of the young during the sexual stage of
reproduction, whereas D. pulex may reproduce either by cyclic or obligate
parthenogenesis in which the zygotes develop within the ephippium by ameoitic
parthenogenesis with no genetic contribution from the males. Thus, the
ephippial and live-born offspring are genetically identical to their mothers.
Both forms may be present in the same population resulting in cyclic
populations exhibiting considerable genetic variation early in the year and an
obligate population with a low range of genotypic values. After 25
generations of asexual reproduction the variation in the cyclic
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parthenogenesis group becomes about the same as that in the obligate group
(Lynch, 1984). These populations exhibiting a low range of genotypic values
are much more vulnerable to perturbations such as nutrient introduction or
toxic discharges. The clonal makeup of a Daphm'a population is effected by
food, oxygen, temperature and predation (Weider, 1985; Brookfield, 1984).
3.1.5 Ephippia are small and lightweight and can be dried and stored for long
periods making them easy to transport. They may be shipped in envelopes like
seeds. Upon arrival at the new location the ephippia can be hatched in a few
days when placed in water at the proper temperature (Schwartz and Hebert,
1987).
3.1.6 Daphm'a are preyed upon by many predators and have developed behavioral
and morphological antipredator defenses to make themselves more difficult to
catch and consume. Dodson (1988) showed that D. pulex responded to a possible
chemical stimuli released by the predator which resulted in the daphnids
retreating from the vicinity of the predators. Certain clones of D. pulex may
develop morphological changes when predators are present but not when they are
absent from the pond. Some of these changes are of such magnitude that they
have been described as separate species. D. minnehaha is a morphological
variation of D. pulex which develops spines in response to the stimuli of
predators (Krueger and Dodson, 1981). Different genotypes of D. pulex react
in different ways to the predator (Chaoborus) factor and to temperature
(Havel, 1985).
3.2 Food and Feeding
3.2.1 Both D. pulex and D. maqna are well adapted to live in algal blooms,
which are high in proteins and carbohydrates, where they feed on algae and
bacteria. D. magna prefers bacteria to algae as food (Ganf, 1983; Hadas et
al., 1983) while D. pulex uses bacteria as food only when algal biomass
declines (Borsheim and 01 sen, 1984). Food type and abundance affect the
sensitivity of Daphm'a to pollutants and their reproduction rate. Keating and
Dagbusan (1986) showed that both D. pulex and D. magna fed diatoms were more
tolerant of pollutants than those fed only green algae. Lipid reserves are a
good indication of the nutritional condition of the animals (Holm and Shapiro,
1984; Tessier and Goulden, 1982).
3.3 Life History
3.3.1 Four distinct periods may be recognized in the life history of Daphm'a:
(1) egg, (2) juvenile, (3) adolescence, and (4) adult (Pennak, 1989). The
life span of Daphm'a, from the release of the egg into the brood chamber until
the death of the adult, is highly variable depending on the species and
environmental conditions (Pennak, 1989). Generally the life span increases as
temperature decreases, due to lowered metabolic activity. The average life
span of D. magna is about 40 days at 25°C, and about 56 days at 20°C. The
average life span of D. pulex at 20°C is approximately 50 days. Typically, a
clutch of 6 to 10 eggs is released into the brood chamber, but as many as 57
have been reported. The eggs hatch in the brood chamber and the juveniles,
which are already similar in form to the adults, are released in approximately
two days when the female molts (casts off her exoskeleton or carapace). The
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time required to reach maturity (produce their first offspring) in D. pulex
varies from six to 10 days (mean = 7.78 days) and also appears to be dependent
on body size. The growth rate of the organism is greatest during its juvenile
stages (early instars), and the body size may double during each of these
stages. D. pulex has three to four juvenile instars, whereas D. magna has
three to five instars. Each instar stage is terminated by a molt. Growth
occurs immediately after each molt while the new carapace is still elastic.
3.3.2 Following the juvenile stages, the adolescent period is very short, and
consists of a single instar. It is during the adolescent instar that the
first clutch of eggs reaches full development in the ovary. Generally, eggs
are deposited in the brood chamber within minutes after molting, and the young
which develop are released just before the next molt.
3.3.3 D. magna usually has 6-22 adult instars, and D. pulex has 18-25. In
general, the duration of instars increases with age, but also depends on
environmental conditions. A given instar generally lasts approximately two
days under favorable conditions, but when conditions are unfavorable, may last
as long as a week.
3.3.4 Four events take place in a matter of a few minutes at the end of each
adult instar: (1) release of young from the brood chamber to the outside, (2)
molting, (3) increase in size, and (4) release of a new clutch of eggs into
the brood chamber. The number of young per brood is highly variable for
Daphnia, depending primarily on food availability and environmental
conditions. D. magna and D. pulex may both produce as many as 30 young during
each adult instar, but more commonly the number is six to 10. The number of
young released during the adult instars of D. pulex reaches a maximum at the
tenth instar, after which there is a gradual decrease (Anderson and Zupancic,
1937). Scholtz et al. (1988) reported that nearly all of the eggs that are
oviposited by D. pulex became neonates, indicating a highly successful
hatching rate. The maximum number of young produced by D. magna occurs at the
fifth adult instar, after which it decreases (Anderson and Jenkins, 1942).
4. CULTURING METHODS
4.1 Sources of Organisms
4.1.1 Daphnia are available from commercial biological supply houses. Only a
small number of organisms (20-30) are needed to start a culture. D. pulex is
preferred over D. magna by some biologists because it is more widely
distributed, is tolerant of a wider range of environmental conditions, and is
easier to culture. However, the neonates are smaller, swim faster and are
more difficult to count, and produce more "floaters" than D. magna and,
therefore, are somewhat more difficult to use in toxicity tests. Guidance on
the source and species of Daphnia to be used by a permittee for effluent
toxicity tests should be obtained from the permitting authority.
4.1.2 Cultures of test organisms should be started at least three weeks
before the brood animals are needed, to ensure an adequate supply of neonates
for the test.
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4.1.3 Starter animals may be obtained from an outside source by shipping in
polyethylene bottles. Approximately 20-30 animals and 3 ml of food (see
below) are placed in a 1-L bottle filled full with culture water. Animals
received from an outside source should be transferred to new culture media
gradually over a period of 1-2 days to avoid mass mortality.
4.1.4 It is best to start the cultures with one animal, which is sacrificed
after producing young, embedded, and retained as a permanent microscope slide
mount to facilitate identification and permit future reference. The species
identification of the stock culture should be verified by a taxonomic
authority. The following procedure is recommended for making slide mounts of
Daphnia (Beckett and Lewis, 1982):
1. Pipet the animal onto a watch glass.
2. Reduce the water volume by withdrawing excess water with the
pipet.
3. Add a few drops of carbonated water (club soda or seltzer
water) or 70% ethanol to relax the specimen so that the
post-abdomen is extended. (Optional: with practice,
extension of the postabdomen may be accomplished by putting
pressure on the cover slip).
4. Place a small amount (one to three drops) of mounting medium
on a glass microscope slide. The recommended mounting
medium is CMCP-9/9AF Medium, prepared by mixing two parts
of CMCP-9 with one part of CMCP-9AF. For more viscosity and
faster drying, CMC-10 stained with acid fuchsin may be used.
CMCP-9 and 9AF are available from Polysciences, Inc., Paul Valley
Industrial Park, Harrington, Pennsylvania, 18976 (215-343-6484).
5. Using a forceps or a pipet, transfer the animal to the drop
of mounting medium on the microscope slide.
6. Cover with a cover slip and exert minimum pressure to remove
any air bubbles trapped under the cover slip. Slightly more
pressure will extend the postabdomen.
7. Allow mounting medium to dry.
8. Make slide permanent by placing CMC-10 around the edges of
the coverslip.
9. Identify to species (see Pennak, 1989).
10. Label with waterproof ink or diamond pencil.
11. Store for permanent record.
4.2 Culture Media
4.2.1 Although Daphnia stock cultures can be successfully maintained in some
tap waters, well waters, and surface waters, use of synthetic water as the
culture medium is recommended because (1) it is easily prepared, (2) it is of
known quality, (3) it yields reproducible results, and (4) allows adequate
growth and reproduction. Reconstituted hard water (total hardness of 160 -180
mg/L as CaC03j is recommended for D. magna culturing, and reconstituted
moderately hard water (total hardness of 80-90 mg/L CaC03) is recommended for
D. pulex culturing. The quality of the dilution water is important in Daphnia
culture. The use of Millipore Milli-QR or Super-QR, or equivalent, to prepare
155
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reconstituted water is highly recommended. The use of diluted mineral water
(DMW) for culturing and testing is widespread due to the ease of preparation.
4.2.2 The chemicals used and instructions for preparation of reconstituted
water are given in Section 7. The compounds are dissolved in distilled or
deionized water and the media are vigorously aerated for several hours before
using. The initial pH of the media is between 7.0 and 8.0, but it will rise
as much as 0.5 unit after the test is underway.
4.3 Culture Conditions
4.3.1 Daphnia can be cultured successfully over a wide range of temperatures,
but should be protected from sudden changes in temperature, which may cause
death. The optimum temperature is approximately 20°C, and if ambient
laboratory temperatures remain in the range of 18-26°C, normal growth and
reproduction of Daphnia can be maintained without special temperature control
equipment. D. magna can survive when the DO concentration is as low as 3 mg/L
but D. pulex does best when the DO concentration is above 5 mg/L. Therefore
it is recommended that the DO concentration in the culture be maintained at 5
mg/L or above. Unless the cultures are too crowded or overfed, aeration is
usually not necessary.
4.3.2 Illumination
4.3.2.1 The variations in ambient light intensities (10-20 uE/m2/s, or 50-100
ft-c) and prevailing day/night cycles in most laboratories do not seem to
affect Daphnia growth and reproduction significantly. However, a minimum of
16 h of illumination should be provided each day.
4.3.3 Culture Vessels
4.3.3.1 Culture vessels of clear glass are recommended since they allow easy
observation of the Daphnia. A practical culture vessel is an ordinary 4-L
glass beaker, which can be filled with approximately 3 L medium (reconstituted
water). Maintain several (at least five) culture vessels, rather than only
one. This will ensure back-up cultures so that in the event of a population
"crash" in one or several chambers, the entire Daphnia population will not be
lost. If a vessel is stocked with 30 adult Daphnia, it will provide
approximately 300 young each week.
4.3.3.2 Initially, all culture vessels should be washed well (see Section 5).
After the culture is established, clean each chamber weekly with distilled or
deionized water and wipe with a clean sponge to rid the vessel of accumulated
food and dead Daphnia (see section on culture maintenance below). Once per
month, wash each vessel with detergent during medium replacement. Rinse three
times with tap water and then with culture medium to remove all traces of
detergent.
4.3.4 Weekly Culture Media Replacement
4.3.4.1 Careful culture maintenance is essential. The medium in each stock
culture vessel should be replaced three times each week with fresh medium.
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This is best accomplished by changing solutions Monday, Wednesday, and Friday,
as follows:
1. Place about 300 ml of the old media in a temporary holding vessel.
2. Transfer about 25 or 30 adults from the old culture vessel to the
holding vessel using a wide bore pipette.
3. Discard the remaining Daphm'a along with the media.
4. Clean the culture vessel as described above.
5. Fill the newly-cleaned vessel with fresh medium.
6. Gently transfer (by pouring) the contents of the temporary holding
vessel (old medium with the Daphm'a) into the vessel containing the
new medium making sure that none of the animals stick to the sides
of the vessel.
7. Feed the animals
4.3.4.2 If the medium is not replaced three times weekly, waste products will
accumulate, which could cause a population crash or the production of males
and/or sexual eggs.
4.3.4.3 Daphm'a cultures should be thinned whenever the population exceeds
200 individuals per stock vessel to prevent over-crowding, which may cause a
population crash, or the production of males and/or ephippia. A good time to
thin the populations is on Monday, Wednesday, and Friday, before feeding. To
transfer Daphm'a, use a 15-cm disposable, jumbo bulb pipette, or 10-mL "serum"
pipette which has had the delivery tip cut off and fire polished. The
diameter of the opening should be approximately 5 mm. A serum pipette, a
pipette bulb, such as a PROPIPETTER, or (MOPET ) portable, motorized pipetter,
will provide the controlled suction needed when selectively collecting
Daphm'a.
4.3.4.4 Liquid containing adult D, pulex and D. magna can be poured from one
container to another without risk of air becoming trapped under their
carapaces. However, the very young Daphm'a are much more susceptible to air
entrapment and for this reason should be transferred from one container to
another using a pipette. The tip of the pipette should be kept under the
surface of the liquid when the Daphm'a are released.
4.3.4.5 Each culture vessel should be covered with a clear plastic sheet or
glass plate to exclude dust and dirt, and minimize evaporation.
4.4 FOOD PREPARATION AND FEEDING
4.4.1 Feeding the proper amount of the right food is extremely important in
Daphm'a culturing. The key is to provide sufficient nutrition to support
normal reproduction without adding excess food which may reduce the toxicity
of the test solutions, clog the animal's filtering apparatus, or greatly
decrease the DO concentration and increase mortality. YCT, a combination of
Yeast, CEROPHYLL*, and Trout chow (or flake food), along with the unicellular
green alga, Selenastrum capn'cornutum, will provide suitable nutrition if fed
daily.
4.4.2 The YCT and algae are prepared as follows:
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4.4.2.1 Digested trout chow (or flake food):
1. The preparation requires one week. Use starter or No. 1 pellets prepared
according to current U.S. Fish and Wildlife Service specifications, or
flake food. Suppliers of trout chow include Zeigler Bros., Inc., P.O.
Box 95, Gardners, Pennsylvania, 17324 (717-780-9009); Glencoe Mills, 1011
Elliott, Glencoe, Minnesota, 55336 (612-864-3181); and Murray Elevators,
118 West 4800 South, Murray, Utah 84107 (800-521-9092).
2. Add 5.0 g of trout chow pellets or flake food to 1 L of MILLI-Q" water.
Mix well in a blender and pour into a 2-L separatory funnel. Digest
prior to use by aerating continuously from the bottom of the vessel for
one week at ambient laboratory temperature. Water lost due to
evaporation is replaced during digestion. Because of the offensive odor
usually produced during digestion, the vessel should be placed in a fume
hood or other isolated, ventilated area.
3. At the end of digestion period, place in a refrigerator and allow to
settle for a minimum of 1 h. Filter the supernatant through a fine mesh
screen (i.e., NITEXR 110 mesh). Combine with equal volumes of
supernatant from CEROPHYLLR and yeast preparations (below). The
supernatant can be used fresh, or frozen until use. Discard the
sediment.
4.4.2.2 Yeast:
1. Add 5.0 g of dry yeast, such as FLEISCHMANN'SR to 1 L of MILLI-QR
water.
2. Stir with a magnetic stirrer, shake vigorously by hand, or mix with a
blender at low speed, until the yeast is well dispersed.
3. Combine the yeast suspension immediately (do not allow to settle) with
equal volumes of supernatant from the trout chow (above) and CEROPHYLLR
preparations (below). Discard excess material.
4.4.2.3 CEROPHYLL" (Dried, Powdered, Cereal Leaves):
1. Place 5.0 g of dried, powdered, cereal leaves in a blender. (Available
as "CEREAL LEAVES," from Sigma Chemical Company, P.O. Box 14508, St.
Louis, Missouri, 63178, (800-325-3010); or as CEROPHYLLR, from Ward's
Natural Science Establishment, Inc., P.O. Box 92912, Rochester, New York,
14692-9012, (716-359-2502). Dried, powdered, alfalfa leaves obtained
from health food stores have been found to be a satisfactory substitute
for cereal leaves.
2. Add 1 L of MILLI-QR water.
3. Mix in a blender at high speed for 5 min, or stir overnight at medium
speed on a magnetic stir plate.
4. If a blender is used to suspend the material, place in a refrigerator
overnight to settle. If a magnetic stirrer is used, allow to settle for
1 h. Decant the supernatant and combine with equal volumes of
supernatant from trout chow and yeast preparations (above). Discard
excess material.
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4.4.2.4 Combined YCT Food:
1. Mix equal (approximately 300 ml) volumes of the three foods as described
above.
2. Place aliquots of the mixture in small (50-mL to 100-mL) screw-cap
plastic bottles and freeze until needed.
3. Freshly prepared food can be used immediately, or it can be frozen until
needed. Thawed food is stored in the refrigerator between feedings, and
is used for a maximum of one week.
4. It is advisable to measure the dry weight of solids in each batch of YCT
before use. The food should contain 1.7 - 1.9 g solids/L. Cultures or
test solutions should contain 12-13 mg solids/L.
4.4.3 Algal (Selenastrum) Food
4.4.3.1 Algal Culture Medium
1. Prepare (five) stock nutrient solutions using reagent grade chemicals as
described in Table 1.
2. Add 1 ml of each stock solution, in the order listed in Table 1, to
approximately 900 ml of MILLI-Q water. Mix well after the addition of
each solution. Dilute to 1 L and mix well. The final concentration of
macronutrients and micronutrients in the culture medium is given in
Table 2.
3. Immediately filter the medium through a 0.45um pore diameter
membrane at a vacuum of not more than 380 mm (15 in.) mercury, or at a
pressure of not more than one-half atmosphere (8 psi). Wash the filter
with 500 ml deionized water prior to use.
4. If the filtration is carried out with sterile apparatus, filtered medium
can be used immediately, and no further sterilization steps are required
before the inoculation of the medium. The medium can also be sterilized
by autoclaving after it is placed in the culture vessels.
5. Unused sterile medium should not be stored more than one week prior to
use, because there may be substantial loss of water by evaporation.
4.4.3.2 Algal Cultures
4.4.3.2.1 Two types of algal cultures are maintained: (1) stock cultures,
and,(2) "food" cultures.
4.4.3.2.2 Establishing and Maintaining Stock Cultures of Algae
1. Upon receipt of the "starter" culture (usually about 10 ml), a stock
culture is initiated by aseptically transferring one milliliter to each
of several 250-mL culture flasks containing 100 ml algal culture medium
(prepared as described above). The remainder of the starter culture can
be held in reserve for up to six months in a refrigerator (in the dark)
at 4°C.
2. The stock cultures are used as a source of algae to initiate "food"
cultures for Daphm'a toxicity tests. The volume of stock culture
maintained at any one time will depend on the amount of algal food
159
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required for the Daphm'a cultures and tests. Stock culture volume
may be rapidly "scaled up" to several liters, if necessary, using 4-L
serum bottles or similar vessels, each containing 3 L of growth medium.
3. Culture temperature is not critical. Stock cultures may be maintained in
environmental chambers with cultures of other organisms if the
illumination is adequate (continuous "cool-white" fluorescent lighting of
approximately 86 + 8.6 uE/m2/s, or 400 ft-c).
4. Cultures are mixed twice daily by hand or stirred continuously.
5. Stock cultures can be held in the refrigerator until used to start "food"
cultures, or can be transferred to new medium weekly. One-to-three
milliliters of 7-day old algal stock culture, containing approximately
1.5 X 106 cells/mL, are transferred to each 100 ml of fresh culture
medium. The inoculum should provide an initial cell density of
approximately 10,000-30,000 cells/mL in the new stock cultures. Aseptic
techniques should be used in maintaining the stock algal cultures, and
care should be exercised to avoid contamination by other microorganisms.
6. Stock cultures should be examined microscopically weekly, at transfer,
for microbial contamination. Reserve quantities of culture organisms can
be maintained for 6-12 months if stored in the dark at 4°C. It is
advisable to prepare new stock cultures from "starter" cultures obtained
from established outside sources of organisms every four
to six months.
4.4.3.2.3 Establishing and Maintaining "Food" Cultures of Algae
1. "Food" cultures are started seven days prior to use for Daphm'a
cultures and tests. Approximately 20 ml of 7-day-old algal stock culture
(described in the previous paragraph), containing 1.5 X 106 cells/ml,
are added to each liter of fresh algal culture medium (i.e., 3 L of
medium in a 4-L bottle, or 18 L in a 20-L bottle). The inoculum should
provide an initial cell density of approximately 30,000 cells/mL. Aseptic
techniques should be used in preparing and maintaining the cultures, and
care should be exercised to avoid contamination by other microorganisms.
However, sterility of food cultures is not as critical as in stock
cultures because the food cultures are terminated in 7-10 days. A
one-month supply of algal food can be grown at one time, and the excess
stored in the refrigerator.
2. Food cultures may be maintained at 25°C in environmental chambers with
the algal stock cultures or cultures of other organisms if the
illumination is adequate (continuous "cool-white" fluorescent lighting of
approximately 86 + 8.6 uE/m2/s, or 400 ft-c).
3. Cultures are mixed continuously on a magnetic stir plate (with a medium
size stir bar) or in a moderately aerated separatory funnel, or are mixed
twice daily by hand. If the cultures are placed on a magnetic stir
plate, heat generated by the stirrer might elevate the culture
temperature several degrees. Caution should be exercised to prevent the
culture temperature from rising more than 2-3°C.
4.4.3.3 Preparing Algal Concentrate for Use as Daphm'a Food
1. An algal concentrate containing 3.0 to 3.5 X 107 cells/mL is prepared
from food cultures by centrifuging the algae with a plankton or
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TABLE 1. NUTRIENT STOCK SOLUTIONS FOR MAINTAINING ALGAL STOCK CULTURES
AND TEST CONTROL CULTURES.
STOCK
SOLUTION
1. MACRONUTRIENTS
A.
B.
C.
D.
2. MICRONUTRIENTS
COMPOUND
MgCl,.6H20
CaCl2.2H20
NaN03
MgS04.7H20
K2HP04
NaHC03
•
H,BO,
MnCl2.4H20
ZnCl2
FeCl,.6H20
CoCl2.6H20
Na2Mo04.2H20
CuCl2.2H20
Na2EDTA.2H20
Na2Se04
AMOUNT DISSOLVED IN
500 ML MILLI-Q* WATER
6.08 g
2.20 g
12.75 g
7.35 g
0.522 g
7.50 g
92.8 mg
208.0 mg
1.64 mga
79.9 mg
0.714 mgb
3.63 mgc
0.006 mgd
150.0 mg
1.196 mge
aZnC!2 - Weigh out 164 mg and dilute to 100 mL. Add 1 mL of this
solution to Stock #1.
bCod2.6H20 - Weigh out 71.4 mg and dilute to 100 mL. Add 1 mL of
this solution to stock #1.
cNa2Mo04.2H20 - Weigh out 36.6 mg and dilute to 10 mL. Add 1 mL
of this solution to stock #1.
dCud2.2H20 - Weigh out 60.0 mg and dilute to 1000 mL. Take 1 mL of
this solution and dilute to 10 mL. Take 1 mL of the second dilution and
add to Stock #1.
eNa2Se04 - Weigh out 119.6 mg and dilute to 100 mL. Add 1 mL of this solution
to Stock #1.
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TABLE 2. FINAL CONCENTRATION OF MACRONUTRIENTS AND MICRONUTRIENTS
IN THE CULTURE MEDIUM
MACRONUTRIENT
NaN03
MgCl2.6H20
CaCl2.2H20
MgS04.7H20
K2HP04
NaHC03
MICRONUTRIENT
H3B03
MnCl2.4H20
ZnCl2
CoCl2.6H20
CuCl2.2H20
Na2Mo04.2H20
FeCl3.6H20
Na2EDTA.2H20
Na,SeO,
CONCENTRATION
(MG/L)
25.5
12.2
4.41
14.7
1.04
15.0
CONCENTRATION
(UG/L)
185
416
3.27
1.43
0.012
7.26
160
300
2.39
ELEMENT
N
Mg
Ca
S
P
Na
K
C
ELEMENT
B
Mn
Zn
Co
Cu
Mo
Fe
--
Se
CONCENTRATION
(MG/L)
4.20
2.90
1.20
1.91
0.186
11.0
0.469
2.14
CONCENTRATION
(UG/L)
32.5
115
1.57
0.354
0.004
2.88
33.1
1.00
162
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bucket-type centrifuge, or by allowing the cultures to settle in a
refrigerator for approximately two-to-three weeks and siphoning off the
supernatant.
2. The cell density (cells/ml) in the concentrate is measured with an
electronic particle counter, microscope and hemocytometer, fluorometer,
or spectrophotometer, and used to determine the concentration required to
achieve a final cell count of 3.0 to 3.5 X 107/mL.
3. Assuming a cell density of approximately 1.5 X 106 cells/ml in the
algal food cultures at 7 days, and 100% recovery in the concentration
process, a 3-L, 7-10 day culture will provide 4.5 X 109 algal cells.
This number of cells would provide approximately 150 ml of algal cell
concentrate.
4. Algal concentrate may be stored in the refrigerator for one month.
4.5 Feeding
4.5.1 Feeding rate and frequency are important in maintaining the organisms
in optimal condition so that they achieve maximum reproduction. Stock cultures
which are stressed because they are not adequately fed may produce low numbers
of young, large numbers of males, and ephippial females. When the young taken
from these inadequately fed Daphnia cultures are used in toxicity tests, they
may show higher than acceptable mortality in controls and greater than normal
sensitivity to toxicants. Steps to follow when feeding the YCT and algal diet
are as follows:
1. If YCT is frozen, remove a bottle of the food from the freezer at
least 1 h before feeding time, and allow to thaw.
2. Mass cultures are fed Monday, Wednesday, and Friday at the rate of 4.5
ml YCT and 2 ml of algae concentrate per 3-L culture.
3. On Tuesday and Thursday the culture water is stirred to re-suspend
the settled algae and another 2 mL of algal concentrate is added.
4. The YCT and algal concentrate is thoroughly mixed by shaking before
dispensing.
5. Return unused YCT food mixture and algal concentrate to the
refrigerator. Do not re-freeze the YCT. Discard unused portion
of YCT after one week.
4.5.2 The quality of food prepared with newly acquired supplies of yeast,
trout chow, and dried cereal leaves, or algae, should be determined in
side-by-side comparisons of Daphnia survival and reproduction tests, using the
new food and food of known, acceptable quality, over a seven-day period in
control medium.
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ultrastructure. Arch. Environ. Contam. Toxicol. 9:23-40.
Schwartz, S.S. and P.D.N. Hebert. 1984. Subgeneric distinction in the genus
Daphnj'a: A new diagnostic trait. Trans. Am. Microsc. Soc.
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Schwartz, S.S. and P.D.N. Hebert. 1987. Methods for the activation of the
resting eggs of Daphm'a. Freshwat. Biol. 17:373-379.
Tessier, A.J. and C.E. Goulden. 1982. Estimating food limitation in
cladoceran populations. Limnol. Oceanogr. 27(4):707-717.
USEPA. 1977. 1977. Chemical/biological implications of using chlorine and
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55804. EPA-600/3-77-066. 87 pp.
USEPA. 1978a. The Selenastrum capn'cornutum Printz, algal assay
W.E. Miller, J.C. Greene and T. Shiroyama.Environmental
Research Laboratory, U. S. Environmental Protection Agency,
Corvallis, Oregon. EPA-600/9-78-018. 126 pp.
bottle test
USEPA. 1978b. Methods for measuring the acute toxicity of effluents
to aquatic organisms. 2nd ed. W.H. Peltier. Environmental
Monitoring and Support Laboratory, U. S. Environmental Protection
Agency, Cincinnati, Ohio. EPA-600/4-78-012.
USEPA. 1981. Effluent toxicity screening test using Daphm'a and mysid shrimp.
Weber, C.I., and W.H. Peltier. Environmental Monitoring and Support
Laboratory, U. S. Environmental Protection Agency, Cincinnati, Ohio.
167
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Vijverberg, 0. 1989. Culture techniques for studies on the growth, development
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conditons: a review. Freshw. Biol. 21:317-373.
Walton, VI.E., S.M. Compton, J.D. Allan and R.E. Daniels. 1982. The
effect of acid stress on survivorship and reproduction of Daphnia pulex
(Crustacea-.Cladocera). Can. J. Zool. 60:573-579.
Weider, L.J. 1985. Spatial and temporal genetic heterogeneity in a natural
Daphnia population. J. Plankton Res. 7(1):101-123.
Winner, R.W. 1984. Selenium effects on antennal integrity and chronic
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APPENDIX A
DISTRIBUTION, LIFE CYCLE, TAXONOMY, AND CULTURE METHODS
A.3. MYSIDS (MYSIDOPSIS BAHIA)1
1. DISTRIBUTION
1.1 Mysids (Figure 1) are small shrimp-like crustaceans found in both
the marine and freshwater environments. The mysid that currently is of
primary interest in the NPDES program is the estuarine species.
Mysidopsis bahia. It occurs primarily at salinities above 15 /00; Stuck
et al. (1979a) and Price (1982) found greatest abundances at salinities
near 30 °/oo- Three sympatric species of Mysidopsis, M. almyra, M. bahia,
and M. bigelowi, have been cultured and used in toxicity testing. The
distribution of Mysidopsis species has been reported by Stuck et al.
(1979b), Price (1982), and Heard et al. (1987).
1.2 Other marine mysids that have been used in toxicity testing and held
or cultured in the lab include Metamysidopsis elongata, Neomysis
americana, Neomysis awatschensis, Neomysis intermedia, and recently for
the Pacific coast, Holmesimysis sculpta and Neomysis mercidis. A
freshwater species, Mysis relicta, presently not used in toxicity
testing, but found in the same habitat as Daphm'a pulex, might be
considered in the future for toxicity testing.
2. LIFE CYCLE
2.1 In laboratory culture, Mysidopsis bahia reach sexual maturity in 12
to 20 days, depending on water temperature and diet (Nimmo et al., 1977).
Normally, the female will have eggs in the ovary at approximately 12 days
of age. The lamellae of the marsupium pouch have formed or are in the
process of forming when the female is approximately 4 mm in length (Ward,
1991b). Unlike Daphm'a, the eggs will not develop unless fertilized.
Mating takes place at night and lasts only a few minutes (Mauchline,
1980).
2.2 Brood pouches are normally fully formed at approximately 15 days
(approximately 5 mm in body length), and young are released in 17 to 20
days (Ward, 1991b). The number of eggs deposited in the brood and the
number of young produced per brood are a direct function of body length
as well as environmental conditions. Mature females have produced as
many as 25 Stage I larvae (egg-shaped embryo) per brood (8-9 mm in body
length) in natural and artificial seawater (FORTY FATHOMS") but average
11+6 Stage III larvae (final stage before larvae are released), with
increasing numbers correlated with increasing body length (Ward, 1991b).
A new brood is produced every 4 to 7 days.
Prepared by Stephen H. Ward.
169
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anlennuJe
antenna
antenna! scale
X
dorsal process
statocvsl
8 thoracic limb pleopods
abdominal segments
\
thoracic segments dorsal process
uropod
Figure 1. Lateral and dorsal view of a mysid with morphological
features identified (from Stuck et al., 1979).
170
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2.3 At time of emergence, juveniles are immobile, making them
susceptible to predation by adult mysids. The juveniles are planktonic
for the first 24-48 hours and then settle to the bottom, orient to the
current, and actively pursue food organisms such as Artemia. Carr et al.
(1980) reported that the stage in the life cycle of M. almyra most
sensitive to drilling mud was the juvenile molt, which occurs between 24
and 48 hours after release from the brood pouch. Ward (1989) found a
relationship between CaC03 level and growth and reproduction and that M.
bahia were more sensitive to cadmium during molting (24-72 h post
release) in high or low levels of CaC03. Work done by Lee and Buikema
(1979) for Daphnia pulex also showed increased sensitivity during
molting.
3. MORPHOLOGY AND TAXONOMY
3.1 Since Mysidopsis bahia occur with two other species of Mysidopsis,
an understanding of the taxonomy of M. almyra, M. bahia, and M. bigelowi
is important for culturing and testing practices. The taxonomic key of
Heard, Price, and Stuck (1987) is suggested (see Table 1 for
morphological guide to Mysidopsis).
3.2 Adults of M. bahia range in length from 4.4 mm to 9.4 mm (Molenock,
1969), measured from the anterior margin of the carapace to the end of
uropods. The mature females are normally larger than the males and the
pleopods of the female are smaller than those of the male (Ward, 1991b)
(Figure 2). Mysidopsis bahia can be positively identified as male or
female when they are 4 mm in body length (Ward, 1991b). Living organisms
are usually transparent, but may be tinted yellow, brown or black.
Mysidopsis bigelowi can be readily distinguished from M. almyra and M.
bahia by the morphology of the second thoracic leg. Mysidopsis bigelowi
has a greatly enlarged endopod of the thoracic limb 2 ("first leg") and
the limb has a distinctive row of 6 to 12 spiniform setae on the inner
margin of the sixth segment (Heard et al., 1987). Mysidopsis bahia can
also be distinguished from other species of Mysidopsis by the number of
apical spines on the telson (4-5 pairs) and the number of spines on the
inner uropods distal to the statocyst (normally 2-3) (Figure 2).
3.3 Heard et al. (1987) state that the most reliable character for
separating adult M. almyra and M. bahia is the number of spines on the
inner uropods (M. almyra will always have a single spine). Further,
Price (1982) found that for all stages of development for both species,
the shape of the anterior margin of the carapace (rostral plate) could be
used to distinguish M. almyra (broadly rounded) from M. bahia (more
produced). Figure 2 illustrates the morphological features most useful
in identifying M. bahia (redrawn Molenock, 1969; Heard et al., 1987).
4. CULTURE METHODS
4.1 Source of Organisms
4.1.1 Starter cultures of mysids can be obtained from commercial
sources, particularly in the Gulf of Mexico region for M. almyra and M,
bahia.
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TABLE 1. GENUS MYSIDOPSIS: COMMON AND SPECIFIC CHARACTERS OF SPECIES KNOWN FROM
U.S. ATLANTIC AND GULF OF MEXICO WATERS1'2
M. eclipes M. brattstroemi
Anterior dorsal margin Triangular Triangular
of carapace
Presence of distal seg- yes yes
ment on antenna I
Length/breadth ratio of 2-2.5 3-3.5
antenna I scale
# segments in carpo- 3 3
propodus of thoracic
endopods 3-8
# segments in exopod ? 6
of male pleopod 4
t^ # spines on uropodal 8-9 10-20
ro endopod
Length of terminal pair(s) Almost Slightly
of telson spines relative twice longer
to lateral margin spines as long
# of pairs of apical 1 1
telson spines
# setae on inner margin ca 3 ca 4
of segment 6 of second
thoracic endopod
# setae on inner margin ca 5 ca 4
of segment 5 of second
thoracic endopod
M. mortenseni M. furca
Triangular Rounded
yes yes
3.5-4 4
3 3
(2 in juv.)
6-7 7
18-31 20-40
Slightly > twice
longer as long
1 2
ca 3 ca 3
ca 2 ca 3
M. almvra
M. bahia M
Rounded Triangular
no
6
2
7
1
Gradually
increasing
4-8
2-3
7-18
no
6
2
7
1-4
Gradually
increasing
3-6
2-3
7-18
. bigelowi
Triangular
no
5.4
2
7-8
5
(occ. 3-4)
Abruptly
increasing
3
6-12
2
M. sp.
(Inshore)
Triangular
no
5.4
2
7
3-4
(occ. 2 or 5)
Abruptly
increasing
3-4
6-12
2
M. sp.
(Offshore)
Triangular
no
5.2
2
7
5
(occ. 4)
Abruptly
increasing
3
6-12
5-7
'From Heard et al., 1987
Modified in part from Brattegard, 1969
-------
Figure 2. Morphological features most useful in identifying Mysidopsis bahia.
a. male; b. female; c. thoracic leg 2; d. telson; e. right uropod,
dorsal; f. male, dorsal (redrawn from Molenock, 1969; Heard et al.,
1987). Note gonad in area where marsupium is located on female and
length of male pleopods as compared to female. Also note the 3
spines on the endopod of the uropod (e).
173
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4.1.2 Mysids of different species can also be collected by plankton tows or
dip nets (approximately 1.0 mm mesh size) in estuarine systems. Heard et al.
(1987) have identified specimens of N. bahia along the eastern coast, however,
it has been principally identified as a subtropical species found in the Gulf
of Mexico and along the east coast of Florida. Since many species of mysids
may be present at a given collection site, the identification of the organisms
selected for culture should be verified by an experienced taxonomist. The
permittee should consult the permitting authority for guidance on the source
of test organisms (indigenous or laboratory reared) before use.
4.2 Culturing System
4.2.1 Stock cultures can be maintained in continuous-flow or closed
recirculating systems. In laboratory culture of M. bahiat recirculating
systems are probably the most common practice. During the past ten years, a
number of closed recirculating systems have been described (Leger and
Sorgeloos, 1982; Nimmo et al., 1991; Ward 1984, 1991a). Since no single
recirculating technique is the best in all respects, the system adopted will
depend on the facilities and equipment available and the objectives of the
culturing activities. Two other species of mysid, M. almyra and M. bigelowi,
have also been successfully reared in the system described in this section
(Ward, 1991a). Further, there now exist a number of review papers (Venables,
1987 and Lussier et al., 1988) that describe in detail techniques developed
by others that will be very helpful in culturing Mysidopsis.
4.2.2 Closed recirculating systems are unique because the re-used seawater
they contain develops an unusual set of characteristics caused primarily by
metabolic waste produced by the mysids. The accumulation of waste products
and suspended particles in the water column is prevented by passing the
seawater through a biological filtration system, in which ammonia and nitrite
are oxidized by nitrifying bacteria.
4.3 Culture Tanks
4.3.1 Stock cultures of mysids are maintained in a closed recirculating
system. The system should consist of four 200-L glass aquaria. However,
smaller tanks, such as 80-L glass aquaria, can be used. When setting up a
system, it is important to consider surface to volume ratio since this will
determine how many mysids can be held in each aquarium. If smaller tanks must
be used, the 20-gallon "high" form is recommended. Figure 3 (Ward, 1984;
1991a) illustrates the main components of the biological filtration system.
The flow rate through the filter is controlled by the water valve and is
maintained between 4-5 L/min. This flow will be sufficient to establish a
moderate current (from the filter return line) in the aquarium to allow the
mysids (which are positively rheotactic) to align themselves with the current
formed.
4.3.2 The filtration system consists of commercially-available under-gravel
filter plates and external power filter. Each aquarium has two filter plates,
forming a false bottom on each side of the tank, on which 2 cm of crushed
174
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coral are placed. The external power filter (Eheim, model 2017) canister is
layered as shown in Figure 3 with a thin layer of filter fiber between each
layer of carbon and crushed oyster shells. There has been some modification
of the original filtration system (Ward, 1984), with crushed coral instead of
oyster shells used on the filter bed, because crushed coral does not dissolve
in seawater as readily as crushed oyster shells. If the system described
above cannot be used, an acceptable alternative is an airlift pumping
arrangement (Spotte, 1979). Crushed coral and oyster shells are commercially
available and should be washed with deionized water and autoclaved before use.
4.4 Culture Media
4.4.1 A clean source of filtered natural seawater (0.45 urn pore diameter)
should be used to culture Mysidopsis bahia, however, artificial seasalts
(FORTY FATHOMS") have also been successfully used (Ward, 1991b). A salinity
range between 20 and 30 °/oo can De useo< (25 °/0o is suggested) to culture M.
bahia. Leger and Sorgeloos (1982) reported success in culturing M. bahia in a
formula following Dietrich and Kalle (Kalle, 1971), and still report continued
use of this formula (Leger et al., 1987b). Other commercial brands have also
been used (Reitsema and Neff, 1980; Nimmo and Iley, 1982; Nimmo et al., 1991)
with varying degrees of success. The culture methods presented in Ward (1984;
1991a) have been tried with a number of commercial brands of artificial
seawater listed in Bidwell and Spotte (1985). Commercial brands of seasalts
can be extremely variable in the amount of NaHCO, they provide, which, if not
controlled, can affect growth and reproduction (Ward, 1989; 1991a). In a
comparative study, Ward (1991b) found normal larval development within the
marsupium using both natural seawater and FORTY FATHOMS" (i.e., Stage I -
embryo; Stage II - eyeless larva; Stage III - eyed larva which is the final
stage before release) and stressed the importance of proper preparation of the
seasalts and monitoring of conditions in the tank.
4.4.2 The culture media should be aged to allow the build-up of nitrifying
bacteria in the filter substrate. To expedite the aging process, 15 ml of a
concentrated suspension of Artemia should be added daily. If using natural or
artificial seawater, the carbonate alkalinity level should be maintained
between 90 and 120 mg/L. It is also important to establish an algal
community, Spirulina subsalsa, in the filter bed (Ward, 1984) and a healthy
surface dwelling diatom community, Nitzchia sp., on the walls (Ward, 1991a) in
conjunction with the transfer of part of the biological filter from a healthy
tank, when possible. After seven days, the suitability of the medium is
checked by adding 20 adult mysids. If the organisms survive for 96 hours, the
culture should be suitable for stocking.
4.4.3 If brine solutions are used, 100°/0o salinity must not be exceeded.
This corresponds to a carbonate alkalinity value of approximately 50 mg/L,
which will allow relatively normal physiological mechanisms associated with
CaCO, to occur during certain phases of the life cycle for M. bahia (Ward,
1989).
175
-------
Filter return line
Power filter
Water valve
Power filter feed line
Filter bed
^ ../... f"N
l\NN. N V X\ V \ VV* v x v x A x ^
Charcoal
Filter plates
Oyster shells
Figure 3. Closed recirculating system showing the two phases of
the biological filtration system which consists of the
filter bed and external power filter (from Ward, 1984, 1991a)
176
-------
4.5 Environmental Factors
4.5.1 Temperature must be maintained within a range of 24° C to 26° C.
Twelve to sixteen hours illumination should be provided daily at 50 to 100 ft-
c. The daily light cycle can be provided by combining overhead room lights,
cool-white fluorescent bulbs (approx. 50 ft-c, 12L:12D), with individual Grow-
lux fluorescent bulbs placed horizontally over each tank (approx. 65 ft-c,
10L:14D). This procedure will avoid acute illumination changes by allowing
the room lights to turn on one hour before and one hour after the aquaria
lights. A timing device, such as an electronic microprocessor-based timer
(ChronTrol", model CD, or equivalent) can be used to control the light cycle.
These procedures are fully outlined in Ward (1984; 1991a).
4.5.2 Good aeration (> 60% saturation by vigorous aeration with an air
stone), a 10-20 percent exchange of seawater per week, and carbonate in the
filtration system are essential in helping to control pH drops caused by
oxidation of NH4-N and N02-N by bacteria.
4.5.3 The single most important environmental factor when culturing
Mysidopsis bahia or other organisms in recirculators is the conversion of
ammonia to nitrite, and nitrite to nitrate by nitrifying bacteria. Spotte
(1979) has suggested upper limits of 0.1 mg total NH4-N/L, 0.1 mg N02-N/L and
20 mg N03-N/L for good laboratory operation of recirculating systems. For the
recirculating system and techniques described here for mysids, the levels of
ammonia, nitrite and nitrate never exceeded 0.05 mg of total ammonia-N/L
(NH3(aq)and NH4+), 0.08 mg NO?-N/L and 18 mg N03-N/L (Ward, 1988). The toxicity
of ammonia is based primarily on unionized ammonia (NH3) and the proportion of
NH3 species to NH4+ species is dependent on pH, ionic strength and
temperature. It is strongly recommended that the concentrations of total
ammonia, nitrite and nitrate do not exceed those reported here. The ammonia,
nitrite, and nitrate levels can be checked by using color comparison test kits
such as those made by LaMotte Chemical or equivalent methods.
4.5.4 Bacterial oxidation of excreted ammonia by two groups of autotrophic
nitrifying bacteria (Nitrosomonas and Nitrobacter), results in an increase of
hydrogen ions, which causes a drop in pH and subsequent loss of buffering
capacity. Typically, the culturist responds to the change in pH by adding
Na2C03 or NaHC03. However, such efforts to buffer against a drop in pH will
result in an increase in alkalinity and the uncontrolled use of carbonates can
affect reproduction, especially at higher alkalinity values (Ward 1989;
1991a). Therefore, when using carbonates to buffer against pH changes,
alkalinity values should not exceed 120 mg/L, which is easily measured by
using a titrator kit such as that available from LaMotte Chemical or
equivalent methods.
4.5.5 Figure 4 (from Ward, 1991a) depicts juvenile production per aquarium,
no buffer added, over a period of 24 weeks. A regression line was calculated
for these data and the slope and correlation coefficient were analyzed by
Student's t test. The data showed that even when the pH dropped as low of
7.5, there was a significant increase (P < 0.001) in
177
-------
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juvenile production. However, the pH should be maintained above 7.8 by the
controlled use of NaHC03 and frequent water exchanges.
4.6 Feeding
4.6.1 Frequent feeding with live food is necessary to prevent cannibalism of
the young by the adults. McKenny (1987) suggests feeding densities of 2-3
Artemia per ml of seawater and Lussier et al. (1988) suggest a feeding rate of
150 Artemia nauplii per mysid daily.
4.6.2 In the M. bahia-Artemia predator-prey relationship, it is also
important to provide sufficient quantities of nutritionally viable free-
swimming stage-I nauplii (Ward, 1987); final hatching from the membranous-sac
(pre-nauplii) into stage-I nauplii does not always occur. Artemia cysts that
have been incubated for 24 h should be periodically examined with a stereozoom
microscope to enumerate free-swimming stage-I nauplii and prenauplii
(membranous-sac stage).
4.6.3 It has also been found that heavy metals can affect the hatchability of
Artemia (Rafiee et al., 1986; Liu and Chen, 1987), therefore, when using
natural seawater the level of metals should always be checked.
4.6.4 Ward (1987; 1991a) has tried different brands of Artemia from different
geographic origins and lot numbers; many achieved stage I nauplii and still
caused variability in production of mysids which suggests that they were
nutritionally lacking,. Leger et al. (1985; 1987) have drawn attention to poor
larval survival of M. bahia and low levels of certain polyunsaturated fatty
acids found in the Artemia fed. The enhancement of Artemia has also been
studied and there are numerous techniques that have been successful (Leger et
al., 1986).
4.6.5 Ward (1987; 1988) has found that it is important to control the flow of
seawater in recirculating systems (keep below 5 L/min) so that Artemia does
not become limiting to the mysid. Newly hatched Artemia should be fed to
mysids at least twice a day. To supply Artemia to the mysid population on the
weekend and prevent cannibalism of newly released mysids, an automatic feeder
such as described by Schimmel and Hansen (1975) or Ward (1984; 1991a) could be
used. Ward (1991a) designed a system to hatch Artemia when personnel were
not available to set up Artemia for the following morning and afternoon
feeding, such as Monday. Cysts were placed in two 4-L Erlenmeyer flasks
(dry), an airstone was placed in each flask, and two vessels overhead were
filled with 3500 mL of 30°/0o seawater each. The previously described timer
(ChronTrolR, Model CD) was used to open the normally closed solenoids,
allowing the seawater to gravity feed and hydrate the cysts.
4.6.6 It is possible that a surface dwelling diatom community acts as a
secondary food that supplements deficient brands of Artemia, especially for
newly released juveniles. Ward (1991a) has observed that a strong fertilizing
action is caused by the excretory products of the mysid population. As the
concentration of nitrate increases (nitrification) to about 5 mg/L (in
approximately 7-10 weeks in an aquarium), a bloom of surface dwelling
179
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diatoms, principally Nitzschia, but including Amphora and Cocconeis, occurs in
natural or artificial seawater (Ward, 1991b). It is interesting to note that,
at the same time, there is a dramatic increase in the number of juveniles
observed in the aquaria (Figure 4). The diatoms form layers on the walls of
the aquarium and swarms of newly released juveniles have been found among
them, possibly feeding upon them.
4.6.7 Nitzschia has been identified as a food source for the marine mud
snail, Ilyanassa obsoleta (Collier, 1981), and the sea urchin, Lytechinus
pictus (Hinegardner and Tuzzi, 1981). The diatom, Skeletonema, has also been
used as a supplemental food for M. bahia (Venables, 1987). Del isle and
Roberts (1986) reported on the use of rotifers, Branchionus plicatilis, as a
superior food for juvenile mysids. Rotifers are active swimmers, ranging in
size from 100-175 urn as compared to 420-520 urn for Artemia, and would provide
a good alternative food source if their fatty acid profile is adequate.
4.7 Culture Maintenance
4.7.1 To avoid an excessive accumulation of algal growth on the internal
surfaces of the aquaria, the walls and internal components should be scraped
periodically and the shell substrate (coral or oyster) turned over weekly.
Also, the filter plates must be completely covered so that the biological
filter functions properly. After a culture tank has been in operation for
approximately 2-3 months, detritus builds up on the bottom, which is removed
with a fish net after first removing the mysids. The rate of water flow
through the tanks should be maintained between 4-5 L/min, and 10-20% of the
seawater in each aquarium should be exchanged weekly.
4.7.2 Some culturists have noted problems with hydrozoan pests in their
cultures and there are procedures for their eradication, if necessary (Lawler
and Shepard, 1978; Mutton et al., 1986).
4.8 Production Level
4.8.1 At least four aquaria should be maintained to insure a sufficient
number of organisms on a continuing basis. If each 200-L aquarium is
initially stocked with between 200 and 500 adults (do not exceed 500 adults),
they will provide sufficient numbers of test organisms (Figure 4) each month.
If the cultures are correctly maintained, at least 20 percent of the adult
population should consist of gravid females (have a visible oostegite brood
pouch with young). It is also advantageous to cull older mysids in the
population every 4-6 weeks and to move mysids among the four aquaria to
diversify the gene pool.
5. VIDEO TRAINING TAPE AVAILABLE FOR CULTURING METHODS
5.1 A video training tape and supplemental report (USEPA, 1990) on culturing
Mysidopsis bahia are available from the National Audiovisual Center, Customer
Services Section, 8700 Edgeworth Drive, Capitol Heights, Maryland, 20743-3701,
(Phone 301-763-1891), as part of a video package on culturing and short-term
chronic toxicity test methods (Order No. A18657; cost $75.00).
180
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6. TEST ORGANISMS
6.1 Juvenile Mysidopsis bahia, one to five days old, are used in the tests.
To obtain the necessary number for a test, several techniques are available.
A mysid generator such as the one described by Reistsema and Neff (1980) has
been successfully used. Another method to obtain juveniles is to take
approximately 200 adult females (bearing embryos in their brood pouches) from
the stock culture and place them in a large (10 cm X 15 cm) standard fish
transfer net (2.0 to 3.0 mm openings) that is partially submerged in an 8-L
aquarium containing 4 L of clean culture medium. As the juveniles are
released from the brood pouches, they drop through the fish net into the
aquarium. The adults and juveniles in the aquarium are fed twice daily 24-
hour post hydrated Artemia. The adults are allowed to remain in the net for
48 h, and are then returned to the stock tanks. The juveniles that are
produced in the small tank may be used in the toxicity tests over a five-day
period. Another method for obtaining juveniles (Ward 1987; 1989) is simply to
remove juveniles from the stock culture with a fine mesh net, place them in 2-
L Pyrex crystalline dishes with media, positioned on a light table that has
an attached viewing plate (2 mm squares), and remove juveniles less than 2 mm
in length (approximately 24 h old).
181
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SELECTED REFERENCES
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APPENDIX A
SYSTEMATICS, ECOLOGY, LIFE HISTORY, AND CULTURE METHODS
A.4. BRINE SHRIMP (ARTEMIA SALINA^
1. SYSTEMATICS
1.1 Morphology and Taxonomy
1.1.1 The taxonomic status of Artemia has long been controversial because
there is considerable morphological variability over parts of its range. The
present consensus is that there is a single cosmopolitan species, Artemia
salina, which has numerous intergrading physiological and morphological
varieties (Pennak, 1978). Brine shrimp belong to the subclass Branchiopoda
which is characterized by many pairs of flattened appendages on the thorax
(Figure 1), in contrast to other members of the Crustacea that have no more
than six pairs. Probably the most distinctive feature of Artemia salina is
the compressed, triangular, and blade-shaped distal segment of the second
antenna of the male (Figure 2). The mature adult is 8 to 10 mm long with a
stalked lateral eye, sensorial antennulae,a linear digestive tract and 11
pairs of thoracopods. In the male the antennae are transformed into muscular
claspers used to secure the female during copulation.
2. DISTRIBUTION
2.1 Artemia are found nearly worldwide in saline lakes and pools. In North
America, they have been reported throughout the western United States, in
Nebraska and Connecticut and in Saskatchewan, Canada. They are probably more
widely distributed than indicated because of limited effort in collecting from
many areas of the country. They are absent from many suitable habitats,
probably because of their limited dispersal methods.
3. ECOLOGY AND LIFE HISTORY
3.1 General Ecology
3.1.1 The ecological conditions under which brine shrimp live are highly
variable. The salinity can exceed 300 °/oo, where most other life cannot
survive. Favored by the absence of predators and food competitors in such
places, Artemia develop very dense populations. Although not a marine
species, they sometimes occur in bays and lagoons where brines are formed by
evaporation of seawater (salt pans). They are more commonly found in highly
saline lakes, such as the Great Salt Lake, where the shoreline may become
ringed with brown layers of accumulated brine shrimp cysts. Brine shrimp are
also common in evaporation basins used for the commercial production of salt.
Prepared by Philip A. Lewis and David A. Bengtson.
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Figure 1. Drawing of male (A) and
female (B) brine shrimp
(From Kuenen and Bass-
Becking, 1938) .
Figure 2. Head of adult male showing
triangular distal segment
of antennae modified as
claspers (From Persoone,
et al., 1980).
--- -V
Figure 3. Pre-nauplius and freshly
hatched first instar
(From Persoone, et al.,
1980).
Figure 4. Male and female brine
shrimp nauplius preparing
to copulate (From Persoone,
et al., 1980).
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3.1.2 The reproductive habits of different populations vary considerably. In
parts of Europe parthenogenesis is the rule, males being rare or absent, but
in North America most Artemia populations seem to be diploid with males
common.
3.1.3 The principal mechanism of Artemia dispersion is transportation of the
cysts by wind or waterfowl and by deliberate or accidental human inoculation.
3.1.4 Growth of brine shrimp is influenced by many factors and the tolerance
of these factors is strain dependent. Optimum temperature for most strains
ranges between 25 and 35°C but strains have been reported thriving at 40°C.
Most geographical strains do not survive temperatures below 6°C except as
cysts. These cysts are tolerant of temperatures from far below 0°C to near
the boiling point of water. Although Artemia can survive and reproduce under a
wide range of salinity, they are seldom found in nature in salinities below
45 °/°o or above 200 /oo. The pH tolerance of Artemia varies from neutral to
highly alkaline but the cysts will hatch best at a pH of 8 or higher.
3.1.5 Many predators including many zooplankton that populate natural salt
waters, many salt water fish, several insect groups (odonates, hemipterans and
beetles), and birds feed on brine shrimp in situations where they can tolerate
the conditions.
3.2 Food and Feeding
3.2.1 Brine shrimp are typically filter-feeders that consume organic
detritus, microscopic algae and bacteria. Blooms of microscopic algae are
favorite habitats of Artemia, and large populations develop in such areas
where they feed on the algae and heterotrophic bacteria that are produced by
these blooms. Brine shrimp populations have done well in cultures when fed
algae, rice bran (Sorgeloos et al., 1979), soybean meal or whey powder
(Bossuyt and Sorgeloos, 1979). The nauplii do not need food for four days
after hatching.
3.3 Life History
3.3.1 Most strains of Artemia produce cysts that float (cysts from the Mono
Lake, California strain sink). These cysts remain in diapause as long as they
are kept dry or under anaerobic conditions. Upon hydration, the embryo in the
cyst becomes activated. After several hours the outer membrane bursts and the
embryo emerges still encased in the hatching membrane. Soon the hatching
membrane is ruptured and the free-swimming nauplius is born. The first instar
is brownish-orange colored and has three pairs of appendages (Figure 3). The
larva grows through about 15 molts and becomes differentiated into male or
female after the tenth molt. Copulation is initiated when the male grasps the
female with its modified antennae (Figure 4). The fertilized eggs develop
either into free-swimming nauplii, or they are surrounded by a thick shell and
deposited as cysts which are in diapause.
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4. METHODS FOR HATCHING ARTEMIA CYSTS
4.1 Sources of Cysts
4.1.1 Brine shrimp cysts are available from many commercial sources,
representing several geographical strains. The cysts from any source can vary
from batch to batch in terms of nutritional quality for the test organisms.
Therefore, it is recommended that each new batch purchased should be analyzed
chemically, and that a side-by-side feeding test be performed on their
nutritional suitability by comparing the response of the test organisms with
the new cysts and cysts of known quality (ASTM, 1988). A list of sources of
cysts is provided at the end of this chapter.
4.2 Storage of Cysts
4.2.1 Sealed cans of Artemia cysts can be stored for years at room
temperature, but once opened, should be used up within two months. After each
use, the can should be tightly covered with a plastic lid and stored in the
refrigerator. If the entire contents of a can will not be used up in two
months, it is recommended that the portion that is expected to be unused be
placed in a tightly closed container and frozen until needed.
4.3 Hatching of Cysts
4.3.1 A 2-1 separatory funnel makes a convenient brine shrimp hatching
vessel, but nearly any transparent or translucent (preferably colorless)
conical shaped container that will hold water may be used. A satisfactory
apparatus can be prepared by removing the bottom of a 2-L plastic soft drink
bottle and inserting a rubber stopper with a flexible tube and pinch cock.
The hatching chambers must be clean and free from toxic material. All
detergents should be completely removed by rinsing well with deionized water.
4.3.2 Salinity of the water used for hatching brine shrimp cysts should be
between 25 and 35 /oo. Natural sea water or water made up from artificial
sea salts may be used. The hatching medium can be prepared by placing 1800 ml
of deionized water in the hatching chamber and adding 50-70 g non-iodized
salt. After the salt is added, lower a 1 ml pipette or glass tube fitted to
an air supply into the vessel, so that the tip rests on the bottom, and bubble
air vigorously through it to dissolve the salt.
4.3.3 Add the desired quantity of cysts to the vessel. Approximately 15 ml
of cysts in a 2-L hatching vessel will provide enough brine shrimp nauplii to
feed three large stock cultures of mysids in 76-L aquaria, or 1000 to 1500
newly hatched fish in four to six 8-L tanks.
4.3.4 Continue the aeration to keep the cysts and newly hatched nauplii from
settling to the bottom where the DO would quickly be depleted and the newly
hatched animals would die.
4.3.5 The area in which the cysts are hatched should be provided with
approximately 20 uE/m /s (100 ft-c) of illumination.
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4.3.6 The cysts will hatch in about 24 h at a temperature of 25°C. Hatching
time varies with incubation temperature and the geographic strain of Artemia
used.
4.4 Harvesting the Nauplii
4.4.1 When the brine shrimp nauplii first emerge from the cyst, they are
enclosed in a membranous sac (Figure 3). To be taken as food by the test
organisms, the pre-nauplii must emerge from the sac and swim about (Stage-I or
first instar nauplius).
4.4.2 The first instar (Stage-I) nauplii do not feed. Their value as food
for the test organisms decreases from birth until they begin feeding. Because
they do not feed in the hatching vessels, it is important to harvest and use
the nauplii soon after hatching. The nauplii can be easily harvested in the
following manner:
1. After approximately 24 h at 25°C, remove the pipet supplying air and
allow the nauplii to settle to the bottom of the hatching chamber. The
empty egg shells will float to the top and the newly hatched nauplii and
unhatched eggs will settle to the bottom. A light trained on the bottom
of the separatory funnel will hasten the settling process.
2. After approximately 5 min, using the stopcock, drain off the nauplii into
a 250-mL beaker.
3. After another 5 min, again drain the nauplii into the beaker.
4. The nauplii are further concentrated by pouring the suspension into a
small cylinder which has one end closed with #20 plankton netting or they
may be washed through a 150-um net or screen.
5. The concentrate is resuspended in 50 mL of appropriate culture water,
mixed well, and dispensed with a pipette. (Mysids require approximately
100 to 150 nauplii/mysid/day).
6. Discard the remaining contents of the hatching vessel and wash the
vessel with hot soap and water.
7. Prepare fresh salt water for each new hatch.
4.4.3 To have a fresh supply of Artemia nauplii daily, at least two
hatching vessels should be used, so that the newly-hatched can be harvested
daily.
4.5 Feeding Assay
4.5.1 Before using brine shrimp nauplii from a new batch of cysts for
routine feeding of cultures and test organisms, they should be tested for
their ability to support life, growth, and reproduction of the test animals.
Two treatments with four replicates each are required for this test. In
Treatment (A), the test organisms are fed the nauplii from the new batch of
Artemia cysts, and in Treatment (B), the test organisms are fed nauplii of
known, good quality, such as from the reference Artemia cysts or from a batch
of Artemia cysts that have been successfully used in culturing and testing.
4.5.2 If there is no significant difference in the survival, growth, and/or
reproduction of the organisms in the two treatments at the end of a 7-day
193
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period, it is assumed that the new batch of Artemia cysts is satisfactory. If
the survival, growth, and/or reproduction in treatment A is significantly less
than the response in treatment B over a 7-day test period it is assumed that
the new batch of brine shrimp cysts are unsuitable for use as a food source
for the organisms tested.
4.5.3 Test chambers and all test conditions during the feeding assay should
be similar to those planned for use in the subsequent toxicity tests.
4.6 List of commercial sources of Artemia cysts.
Aquafauna Biomarine
P.O. Box 5
Haawthorne, CA 90250
Tel. (213) 973-5275
Fax (212) 676-9387
(Great Salt Lake, North Arm
San Francisco Bay)
Argent Chemical
8702 152nd Ave. NE
Redmond, WA 98052
Tel. (800) 4266258
Tel. (206) 855-3777
Fax (206)885-2112
(Platinum Label - San francisco Bay;
Gold Lable - San Francisco Bay,
Brazil, other; Silver Label - Great
Salt Lake, Australia, other; Bronze
label - China, Canada, other]
Bonneville Artemia International, Inc.
P.O. Box 511113
Salt Lake City, UT 84151-1113
Tel. (801) 972-4704
Fax (801) 972-4795
Ocean Star International
P.O. Box 643
Snowville, UT
Tel. (801) 872-8217
Fax (801) 872-8272
(Great Salt Lake)
Sanders Brine Shrimp Co.
3850 South 540 West
Ogden, UT 84405
Tel. (801) 393-5027
(Great Salk Lake)
Aquarium Products
180L Penrod Court
Glen Burnie, MD 21061
Tel. (800) 368-2507
Tel. (301) 761-2100
(Colombia)
Artemia Systems
Wiedauwkaai 79
B-9000 Ghent, Belgium
Tel 011-32-91-534142
Fax 011-32-91-536893
(For marine species - AF grade
[small nauplii], UL grade [large
nauplii], for freshwater species -
IH grade [small nauplii], EG grade
[large nauplii]
Golden West Artemia
411 East 100 South
Salk Lake City, UT 84111
Tel. (801) 532-1400
Fax (801) 531-8160
Pennsylvania Pet Products
Box 191
Spring City, PA
(Great Salt Lake)
San Francisco Bay Brand
8239 Enterprise Drive
Newark, CA 94560
Tel. (415) 792-7200
(Great Salt Lake, San Francisco
Bay)
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Sea Critters Inc. Western Brine Shrimp
P.O. Box 1508 957 West South Temple
Tavernier, FL 33070 Salt Lake City, UT 84104
305-367-2672 Tel. (801) 364-3k642
Fax (801) 534-0211
(Great Salt Lake)
195
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SELECTED REFERENCES
ASTM. 1988. Standard practice for using brine shrimp nauplii as food for
test animals in aquatic toxicology. Standard E1203-87, American
Society for Testing and Materials, Annual Book of Standards,
Philadelphia, Pennsylvania.
Beck, A.D., and D.A. Bengtson. 1982. International study on Artemia
XXII: Nutrition in aquatic toxicology - Diet quality of geographical
strains of the brine shrimp, Artemia. In: J.G. Pearson, R.B. Foster,
and W.E. Bishop (eds.), Aquatic Toxicology and Hazard Assessment: Fifth
Conference. ASTM STP 766, American Society for Testing and Materials,
Philadelphia, Pennsylvania, pp. 161-169.
Beck, A.D., D.A. Bengtson, and W.H. Howell. 1980. International study on
Artemia. V. Nutritional value of five geographical strains of Artemia:
Effects of survival and growth of larval Atlantic silversides,
Menidia menidia. In: G. Persoone, P. Sorgeloos, D.A. Roels, and E.
Jaspers, eds. The brine shrimp, Artemia. Vol. 3. Ecology, culturing,
use in aquaculture. Universa Press, Wetteren, Belgium, pp. 249-259.
Bengtson, D.A.S., A.D. Beck, S.M. Lussier, D. Migneault, and C.E. Olney.
1984. International study on Artemia. XXXI. Nutritional effects in
toxicity tests: Use of different Artemia geographical strains. In: G.
Persoone, E. Jaspers, and C. Claus, (eds.). Ecotoxicological testing
for the marine environment, Vol. 2. State Univ. Ghent and Inst. Mar.
Sci. Res., Bredene, Belgium, pp. 399-416.
Bossuyt, E. and P. Sorgeloos. 1979. Technological aspects of the batch
hatching of Artemia in high densities. In: G. Persoone, P. Sorgeloos,
0. Roels and E. Jaspers (eds.), The brine shrimp Artemia. Vol. 3.
Ecology, culturing, use in aquaculture. Universa Press, Wetteren,
Belgium, pp. 133-152.
Browne, R.A. 1982. The cost of reproduction in brine shrimp. Ecology
63(l):43-47.
Johns, D.M., W.J. Berry, and W. Walton. 1981. International study on
Artemia. XVI. Survival, growth and reproductive potential of the
mysid, Mysidopsis bahia Molenock fed various geographical strains of
the brine shrimp, Artemia. J. Exp. Mar. Biol. Ecol. 53:209-219.
Kuenen, D.J. and L.G.M. Baas-Becking. 1938. Historical notes on Artemia
salina (I.). Zool. Med. 20:222-230.
Leger, P., D.A. Bengtson, K.L. Simson and P. Sorgeloos. 1986. The use and
nutritional value of Artemia as a food source. Oceanogr. Mar. Biol.
Ann. Rev. 24:521-623.
196
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Lenz, P.M. 1980. Ecology of an alkali-adapted variety of Artemia from
Mono Lake, California, U.S.A. In: G. Persoone, P. Sorgeloos, 0. Roels
and E. Jaspers (eds.), The brine shrimp Artemia. Vol. 3. Ecology,
culturing, use in aquaculture. Universa Press, Wetteren, Belgium, pp.
79-96.
Nikonenko, Ye. M. 1986. Adaptation of Artemia salina to toxicants.
Hydrobiol. J. 22(5):94-98.
Pennak, R.W. 1989. Fresh-water invertebrates of the United States.
Protozoa to mollusca. John Wiley and Sons, New York, New York.
pp. 358-359.
Persoone, G., P. Sorgeloos, 0. Roels, and E. Jaspers, (eds.). 1980. The
brine shrimp Artemia. Vol. 1. Morphology, genetics, radiobiology,
toxicology. Universa Press, Wetteren, Belgium. 318 pp.
Persoone, G., P. Sorgeloos, 0. Roels, and E. Jaspers, (eds.). 1980. The
brine shrimp Artemia. Vol. 2. Physiology, biochemistry, molecular
biology. Universa Press, Wetteren, Belgium. 636 pp.
Persoone, G., P. Sorgeloos, 0. Roels, and E. Jaspers, (eds.). 1980. The
brine shrimp Artemia. Vol. 3. Ecology, culturing, use in aquaculture.
Universa Press, Wetteren, Belgium. 428 pp.
Sorgeloos, P., M. Baesa-Mesa, E. Bossuyt, E. Bruggeman, J. Dobbeler, D.
Versichele, E. Lavina and A. Bernardine. 1979. Culture of Artemia on
rice bran: The conversion of waste-products into highly nutritive
animal protein. Aquaculture 21:393-396.
Sorgeloos, P. 1980. Life history of the brine shrimp Artemia. In: G.
Persoone, P. Sorgeloos, D.A. Roels, and E. Jaspers (eds.), The brine
shrimp, Artemia. Vol. 1. Morphology, genetics, radiobiology,
toxicology. Universa Press, Wetteren, Belgium, pp. ixx-xxii.
Usher, R.R., and D.A. Bengtson. 1981. Survival and growth of sheepshead
minnow larvae and juveniles on diet of Artemia nauplii. Prog.
Fish-Cult. 43:102-105.
197
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APPENDIX A
MORPHOLOGY, TAXONOMY, DISTRIBUTION, GENERAL LIFE HISTORY,
AND CULTURE METHODS
A.5 FATHEAD MINNOW (PIMEPHALES PROMELAS^
1. MORPHOLOGICAL AND ANATOMICAL CHARACTERISTICS
1.1 Fathead minnows vary greatly in many characteristics throughout their
wide geographic range. The morphology and characters for identification are
taken from Clay (1962), Hubbs and Lagler (1964), Eddy and Hodson (1961), Scott
and Grossman (1973), and Trautman (1981). Adults (Figure 1) are small fish,
typically 43 mm to 102 mm, and averaging about 50 mm, in total length. The
standard lengths are usually less than four and one-half times the body depth.
The first rudimentary ray of the dorsal fin is more or less thickened and
distinctly separated from the first well-developed ray by a membrane. The
lateral line is usually incomplete, but may be complete in specimens from some
geographic areas. The scales are cycloid and moderate in size. Andrews
(1970), reporting on fish collected in Colorado, noted that no scales were
found on fish smaller than 14 mm, and the average length for first scale
formation was 16.3 mm. The scales in the lateral series number 41 to 54.
1.2 The mouth is terminal. The snout does not extend beyond the upper lip
and is decidedly oblique. Nuptial tubercles occur on mature males only, are
large and well-developed on the snout, and rarely extend beyond the nostrils.
Figure 1. Fathead minnow: adult female (left) and breeding male
(right). (From Eddy and Hodson, 1961).
Prepared by Donald J. Klemm, Quentin H. Pickering, and Mark E. Smith.
198
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They occur in three main rows, with a few on the lower jaw. In addition to
nuptial tubercles, the males have an elongate, fleshy, or spongy pad extending
in a narrow band from the nape to the dorsal fin. The pad is wide anteriorly,
and narrows to engulf the first dorsal ray. In addition, the sides of the
body become almost black except for two wide vertical bars which are light in
color. In contrast to the males, the mature females remain quite drab.
1.3 The peritoneum is brownish-black, and the intestine is long and coiled
one or more times.
1.4 Some external markings occur infrequently. Young occasionally have a
dusky band on the snout and opercules. Other young and adults, from clear and
weedy waters, have a distinct, lateral band across the body. The band may be
absent in breeding males or, if present, becomes very diffuse anteriorly.
This band is usually most apparent on preserved specimens. Dymond (1926),
Trautman (1981), and others described the saddle-like pattern often associated
with breeding males in which a light area develops just behind the head and
another beneath the dorsal fin, the areas between producing a saddle affect.
A dark spot is usually present in front of the dorsal fin in mature males, and
a narrow, dark, vertical bar or spot is present at the base of the caudal fin,
but often is not very distinct.
2. TAXONOMY
2.1 The specific name (Pimephales promelas) appears to be incorrectly applied
to this fish because the fathead minnow does not fit the description
originally given by Rafinesque (1820) (Lee et al_- > 1980). Common names
include "northern fathead minnow", and "blackhead minnow," in addition to
fathead minnow. The holotype was collected near Lexington, Kentucky.
2.2 Some geographic variations have been noted in the morphology of the
fathead minnow. Vandermeer (1966) indicated that the introduction of this
species outside its native range may have resulted in some local deviations
from the broad patterns of geographic variation in taxonomic characters. Some
populations have been designated as subspecifically distinct: Pimephales
promelas promelas. the northern form; P. j). harveyensis, the Harvey Lake form,
from Isle Royal in Lake Superior and P. £. confertus, the southern form (Hubbs
and Lagler, 1949, 1964). However, Taylor (1954), Vandermeer (1966), and
others expressed doubt concerning the validity of assigning subspecific status
to the variants and recommended against their recognition. Vandermeer (1966),
in a statistical analysis of the geographic variations in taxonomic
characters, stated that two of the three described subspecies intergrade
clinally.
2.3 Of the eight characters measured, two showed a north-south trend; (1) eye
diameter, with the northern fish having smaller eyes, and (2) completeness of
the lateral line, with the northern fish having the least complete lateral
line. However, Scott and Grossman (1973), indicated that some Canadian
populations exhibit a nearly complete lateral line. The American Fisheries
Society (1980) does not recognize any of the fathead minnow subspecies.
199
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3. DISTRIBUTION
3.1 The fathead minnow is widely distributed in North America (Figure 2). It
is a popular bait fish, and the ease with which it is propagated has led to
its widespread introduction both within and outside the native range of the
species. It has been so widely distributed in the eastern and southwestern
United States by bait transportation that it is difficult to determine its
original range. The presumed native distribution (Vandermeer, 1966; Scott and
Grossman, 1973; Lee, et al., 1980) extended from the Great Slave Lake in the
northwest to New Brunswick, in eastern Canada, southward throughout the
Mississippi valley in the United States, to southern Chihuahua in Mexico.
Distribution records for this species also now include Oregon (Andreasen,
1975), and the Central Valley (Kimsey and Fisk, 1964) and other locations in
California (Andreasen, 1975), but there are no records for British Columbia.
3.2 This species is found in a wide range of habitats. It is most abundant
in muddy brooks, streams, creeks, ponds, and small lakes, is uncommon or
absent in streams of moderate and high gradients and in most of the larger and
deeper impoundments, and is tolerant of high temperature and turbidity, and
low oxygen concentrations.
3.3 Species associated with the fathead minnow seem to vary greatly
throughout its range (Scott and Crossman, 1973; Trautman 1981). Trautman
(1981) reported that fathead minnows and bluntnose minnows, Pimephales notatus
(Rafinesque), were competitors, and that fathead minnows occurred in greatest
numbers only where bluntnose minnows were absent or comparatively few in
number. He also stated that the fathead minnow may hybridize with the
bluntnose minnow
3.4 The fathead minnow is primarily omnivorous, although Coyle (1930)
reported algae to be one of its main foods in Ohio. Elsewhere in the United
States, young fish have been reported to feed on organic detritus from bottom
deposits, and unicellular and filamentous algae and planktonic organisms.
Adults feed on aquatic insects, worms, small crustaceans, and other animals.
Scott and Crossman (1973) and others regard the fathead minnow as a highly
desirable forage fish, providing food for other fishes and birds.
4. GENERAL LIFE HISTORY
4.1 The natural history and spawning behavior (Markus, 1934; Flickinger,
1973; Andrews and Flickinger, 1974; and others) of the fathead minnow are well
known because of the early interest in raising the fish for bait and for
feeding other pond fish, such as bass. Sexual dimorphism occurs at maturity.
Breeding males develop a conspicuous, narrow, elongated, gray, fleshy pad of
spongy tubercles on the back, anterior to the dorsal fin, and two or three
rows of strong nuptial tubercles across the snout. The sides of the body
become almost black except for two wide vertical bars which are light in
color. In contrast, the females remain quite drab.
200
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^(vv ^ .-'^
^LV^T?-- •
'^IV^-^^f^f *---• - <*-,
^ dan -<;<^:\ A L'-/-' i--6. -TX4
-, ^-SJV) ' v. ' • <
y 'j v-
v:?W^
•T*^
t~ IN r ' /r>\~ -«••'- .*r*- --% ••»•> K ^K»» "Vi. '- - ff- i «T-.-««?=»*"*••
; :----
b/:vW?i^P?S^
Figure 2. Map showing the distribution of the fathead
minnow in North America. Open circles represent
transplanted populations. Most Atlantic slope
records are probably transplanted populations.
(From Lee et al.., 1980).
4.2 The initiation of spawning varies with temperature throughout its
geographic range. Isaak (1961), Carlander (1969), and others reported that,
in the wild, fathead minnows begin spawning in the spring, when the water
temperature reaches 16-18°C, and continue to spawn throughout most of the
summer. The minimum spawning temperature, however, may vary with population
and latitude.
4.3 Markus (1934) reported that spawning always occurred at night, whereas
Isaak (1961) observed spawning during the day, as well as at night. Gale and
Buynak (1982) and others reported that spawning often began before dawn and
usually was completed before noon. Observations of the fathead minnow
cultures at EPA's Newtown Facility also indicate the majority of fathead
minnows spawn in early morning.
201
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4.4 Breeding males are very territorial and select sites for spawning, such
as the underside of a log or branch, rock, board, tin can, or almost any other
solid inanimate object, usually in water from 7 cm to 1 m in depth. A
receptive female is sought out and brought into position below the nest site.
After circling below the nesting site, the female is nudged and lifted on the
male's back until she lies on her side immediately below the undersurface of
the spawning substrate, where she releases a small number of eggs (usually 100
to 150) at a time. The eggs are adhesive and attach to the underside of the
spawning substrate. The females have a urogenital structure (ovipositor) to
help deposit the eggs on the underside of objects. Flickinger (1966)
indicated that the ovipositor is noticeable at least a month prior to
spawning. The reported size of the eggs varies from 1.15 mm (Markus, 1934) to
1.3 mm in diameter (Wynne-Edwards, 1932).
4.5 Immediately after the eggs are laid, they are fertilized by the male, and
the female is driven off. Once eggs are deposited in the nest, the male
becomes very aggressive and will use the large tubercles on his snout to help
drive off all intruding small fishes. In addition to fertilizing and guarding
the eggs, the male agitates the water around the eggs, which ventilates them
and keeps them free of detritus. Some males will spawn with several females
on the same substrate, so that the nest may contain eggs in various stages of
development. The number of eggs per nest may vary from as few as nine or 10
to as many as 12,000.
4.6 The ovaries of the females contain eggs in all stages of development, and
they spawn repeatedly as the eggs mature. A female may deposit eggs in more
than one nest. Although the average number of eggs per spawn is generally 100
to 150, large females may lay 400 to 500 eggs per spawn.
4.7 Gale and Buynak (1982), in a study using five captive pairs of fathead
minnows in separate outdoor pools, observed that each pair produced 16 to 26
clutches of eggs between May and August. The time between spawns, which
ranged from two to 16 days, was affected by water temperature. As the
temperature increased, the intervals between spawning sessions become shorter
and more uniform. In their study, from nine to 1,136 (mean of 414) eggs were
deposited per spawn. The average number of eggs deposited per spawn ranged
from 371 to 480, and the total number of eggs spawned per female ranged from
6,803 to 10,164 (mean of 8,604). The length of the spawning period during a
given season also varied greatly between females. The authors suggested that
the fecundity of fathead minnows is much higher than has generally been
recognized, but they noted that fecundity of fish in the natural environment,
where conditions might be more or less favorable, might differ from that of
captive fish.
4.8 The incubation time depends on temperature, and is 4.5 to 6 days at 25°C.
The newly-hatched young (larvae) are about 5 mm long, white in color, and have
large black eyes. The general appearance and typical pigmentation of the
various larval stages are illustrated in Figures 3A-3M). In a warm, food-rich
environment, growth is rapid. Markus (1934) stated that fish hatched in
202
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Figure 3. Fathead minnow (Pimephales promelas) larvae: A. protolarva,
lateral view, 4.3 mm TL; B. protolarva, dorsal view, 5.6 mm
TL; C. protolarva, lateral view, 5.6 mm TL; D. protolarva,
ventral view, 5.6 mm TL; E. mesolarva, lateral view, 6.9 mm TL;
F. mesolarva, dorsal view, 7.9 mm TL; G. mesolarva, lateral
view, 7.9 mm TL; (from Snyder et al., 1977).
203
-------
rr~ *:•;•• -•''.•'-' :..'.".••'—-• ^C?. -n^ ^^ -riftK
;..r ... y,^.V^«^,..^^^^^__~rjj.-;^._.-;_tg?S^
* .••:5..'J-*T>l.,vVsir*^>'>-<~ -iS^-, ^^ • ^'S'W-
^KS\
K
M
Figure 3. (Cont.)
Fathead minnow (Pimephales promelas) larvae. H. mesolarva,
ventral view, 7.9 mm TL; I. metal arva, lateral view, 9.3 mm
TL; J. metalarva, dorsal view, 14.3 mm TL; K. metalarva,
lateral view, 14.3 mm TL; L. metalarva, ventral view, 14.3
mm TL; M. late metalarva, lateral view, 19.6 mm TL. (from
Snyder, et al . , 1977).
204
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May in Iowa reached adult size and were spawning by late July. Hubbs and
Cooper (1935) and others noted that such rapid growth is unlikely in more
northerly waters, and that the young do not spawn the first year. In cooler
water the adult size is probably not reached until the second year. The males
generally grow faster than the females, a characteristic of minnow species.
4.9 The fathead minnow is short lived, and rarely survives to the third year.
However, Scott and Crossman (1973) stated that longevity varies throughout
the geographic range of the species. Post-spawning mortality was reported to
be great by several authors, but was not observed by Gale and Buynak (1982).
However, in defending their territory, male fish, may become weakened by a
lack of food over a prolonged period and their resistance to disease may be
lowered. Also, at spawning time, many waters are warm and somewhat stagnate,
favoring the spread of fish parasites and disease.
5. CULTURE METHODS
5.1 Outside Sources of Fathead Minnows
5.1.1 Fathead minnows are available from commercial biological supply houses.
Fish obtained from outside sources for use as brood stock or in toxicity tests
may not always be of suitable age and quality. Fish provided by supply houses
should be guaranteed to be of (1) the correct species, (2) disease free, (3)
in the requested age range, (4) and in good condition. The latter can be done
by providing the record of the date on which the eggs were laid and hatched,
and information on LC50 of contemporary fish using reference toxicants.
5.2 Inhouse Sources of Fathead Minnows
5.2.1 Problems in obtaining suitable fish from outside laboratories can be
avoided by developing an inhouse laboratory culture facility. Fathead minnows
can be easily cultured in static, recirculating, or flow-through systems.
5.2.2 Flow-through systems require large volumes of water and may not be
feasible in some laboratories. The culture tanks should be shielded from
extraneous disturbances using opaque curtains, and should be isolated from
toxicity testing activities to prevent contamination.
5.2.3 To avoid the possibility of inbreeding of the inhouse brood stock, fish
from an outside source should be introduced yearly into the culture unit.
5.2.4 The inhouse culture facility consists of the following components:
5.2.4.1 Water Supply
5.2.4.1.1 Water Quality
5.2.4.1.1.1 Reconstituted (synthetic) water or dechlorinated tap water can
be used, but natural water may be preferred. To determine water quality, it
is desirable to analyze the water for toxic metals and organics quarterly (see
Section 4).
205
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Temperature, dissolved oxygen, pH, hardness, and alkalinity should also be
measured periodically.
5.2.4.1.1.2 If a static or recirculating system is used, it is necessary to
equip each tank with an outside activated carbon filter system, similar to
those sold for tropical fish hobbyists (or one large activated carbon filter
system for a series of tanks) to prevent the accumulation of toxic metabolic
wastes (principally nitrite and ammonia) in the water.
5.2.4.1.2 Dissolved oxygen
5.2.4.1.2.1 The DO concentration in the culture tanks should be maintained
near saturation, using gentle aeration with 15 cm air stones if necessary.
Brungs (1971), in a carefully controlled long-term study, found that the
growth of fathead minnows was reduced significantly at all DO concentrations
below 7.9 mg/L. Soderberg (1982) presented an analytical approach to the re-
aeration of flowing water for culture systems.
5.2.4.2 Maintenance
5.2.4.2.1 Adequate procedures for culture maintenance must be followed to
avoid poor water quality in the culture system. The spawning and brood stock
culture tanks should be kept free of debris (excess food, detritus, waste,
etc.) by siphoning the accumulated materials (such as dead brine shrimp
nauplii or cysts) from the bottom of the tanks daily with a glass siphon tube
attached to a plastic hose leading to the floor drain. The tanks are more
thoroughly cleaned as required, Algae, mostly diatoms and green algae,
growing on the glass of the spawning tanks are left in place, except for the
front of the tank, which is kept clean for observation. To avoid excessive
build-up of algal growth, the walls of the tanks are periodically scraped.
The larval culture tanks are cleaned once or twice a week to reduce the mass
of fungus growing on the bottom of the tank.
5.2.4.2.2 Activated charcoal and floss in the tank filtration systems should
be changed weekly, or more often if needed. Culture water may be maintained
by preparation of reconstituted water or use of dechlorinated tap water.
Distilled or deionized water is added as needed to compensate for evaporation.
5.2.4.2.3 Before new fish are placed in tanks, salt deposits are removed by
scraping or with 5% acid solution, the tanks are washed with detergent,
sterilized with a hypochlorite solution, and rinsed well with hot tap water
and then with laboratory water.
5.2.5 Spawning Tanks and Culture Conditions
5.2.5.1 For breeding tanks, it is convenient to use 60 L (15 gal) or 76 L (20
gal) aquaria. The spawning unit is designed to simulate conditions in nature
conductive to spawning, such as water temperature and photoperiod. Spawning
tanks must be held at a temperature of 25 + 2°C. Each aquarium is equipped
with a heater, if necessary, a continuous filtering unit, and spawning
substrates.
206
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The photoperiod for the culture system should be maintained at 16 h light and
8 h darkness. For the spawning tanks, this photoperiod must be rigidly
controlled. A convenient photoperiod is 5:00 AM to 9:00 PM. Fluorescent
lights should be suspended about 60 cm above the surface of the water in the
brood and larval tanks. Both DURATESTR and cool-white fluorescent lamps have
been used, and product similar results. An illumination level of 10-20
uE/m2/s (50 to 100 ft-c) is adequate.
5.2.6 Spawning Behavior and Conditions
5.2.6.1 To simulate the natural spawning environment, it is necessary to
provide substrates (nesting territories) upon which the eggs can be deposited
and fertilized, and which are defended and cared for by the males. The
recommended spawning substrates consist of inverted half-cylinders, such as
7.6 cm X 7.6 cm (3 in. X 3 in.) sections of schedule 40, PVC pipe. The
substrates should be placed equi-distant from each other on the bottom of the
tanks.
5.2.6.2 To establish a breeding unit, 15-20 pre-spawning adults six to eight
months old are taken from a "holding" or culture tank and placed in a 76-L
spawning tank. At this point, it is not possible to distinguish the sexes.
However, after less than a week in the spawning tank, the breeding males will
develop their distinct coloration and territorial behavior, and spawning will
begin. As the breeding males are identified, all but two are removed,
providing a final ratio of 5-6 females per male. The excess spawning
substrates are used as shelter by the females.
5.2.6.3 Sexing of the fish to ensure a correct female/male ratio in each tank
can be a problem. However, the task usually becomes easier as experience is
gained (Flickinger, 1966). Sexually mature females usually have large bellies
and a tapered snout. The sexually mature males are usually distinguished by
their larger overall size, dark vertical color bands, and the spongy nuptial
tubercles on the snout. Unless the males exhibit these secondary breeding
characteristics, no reliable method has been found to distinguish them from
females. However, using the coloration of the males and the presence of an
enlarged urogenital structures and other characteristics of the females, the
correct selection of the sexes can usually be achieved by trial and error.
5.2.6.4 Sexually immature males are usually recognized by their aggressive
behavior and partial banding. These undeveloped males must be removed from
the spawning tanks because they will eat the eggs and constantly harass the
mature males, tiring them and reducing the fecundity of the breeding unit.
Therefore, the fish in the spawning tanks must be carefully checked
periodically for extra males.
5.2.6.5 A breeding unit will remain in their spawning tank about four months.
Thus, each brood tank or unit is stocked with new spawners about three times a
year. However, the restocking process is rotated so that at any one time the
spawning tanks contain different age groups of brood fish.
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5.2.7 Embryo Collection
5.2.7.1 Fathead minnows spawn mostly in the early morning hours. They should
not be disturbed except for a morning feeding (approximately 8:00 AM) and
daily examination of substrates for eggs in late morning or early afternoon.
In nature, the male protects, cleans, and aerates the eggs until they hatch.
In the laboratory, however, it is necessary to remove the eggs from the tanks
to prevent them from being eaten by the adults, and for ease of handling for
purposes of recording embryo count and hatchability, and for the use of the
newly hatched for young fish for toxicity tests.
5.2.7.2 Daily, beginning six to eight hours after the lights are turned on
(i.e., 11:00 AM - 1:00 PM), the substrates in the spawning tanks are each
lifted carefully and inspected for embryos. Substrates without embryos are
immediately returned to the spawning tank. Those with embryos are immersed in
clean water in a collecting tray, and replaced with a clean substrate. A
daily record is maintained of each spawning site and estimated number of
embryos on the substrate.
5.2.8 Embryo Incubation
5.2.8.1 Three different methods are described for embryo incubation.
5.2.8.1.1 Incubation of Embryos on the Substrates: Several (2-4) substrates
are placed on end in a circular pattern (with the embryos on the inner side)
in 10 cm of water in a tray. The tray is then placed in a constant
temperature water bath, and the embryos are aerated with a 2.5 cm airstone
placed in the center of the circle. The embryos are examined daily, and the
dead and fungused embryos are counted, recorded, and removed with forceps. At
an incubation temperature of 25°C, 75-100% hatch occurs in five days. At
22°C, embryos incubated on aerated tiles require seven days for 50% hatch.
5.2.8.1.2 Incubation of Embryos in a Separatory Funnel: The embryos are
removed from the substrates with a rolling action of the index finger ("rolled
off")(Gast and Brungs, 1973), their total volume is measured, and the number
of embryos is calculated using a conversion factor of approximately 430
embryos/mL. The embryos are incubated in about 1.5 L of water in a 2 L
separatory funnel maintained in a water bath. The embryos are stirred in the
separatory funnel by bubbling air from the tip of a plastic micro-pipette
placed at the bottom, inside the separatory funnel. During the first two
days, the embryos are taken from the funnel daily, those that are dead and
fungused are removed, and those that are alive are returned to the separatory
funnel in clean water. The embryos hatch in four days at a temperature of
25°C. However, usually on day three the eyed embryos are removed from the
separatory funnel and placed in water in a plastic tray and gently aerated
with an air stone. Using this method, the embryos hatch in five days.
5.2.8.1.2.1 Hatching time is greatly influenced by the amount of agitation of
the embryos and the incubation temperature. If on day three the embryos are
transferred from the separatory funnel to a static, unaerated container, a 50%
hatch will occur in six days (instead of five) and a 100% hatch will occur in
seven days.
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5.2.8.1.3 Incubation in Embryo Incubation Cups: The embryos are "rolled off"
the substrates, and the total number is estimated by determining the volume.
The embryos are then placed in incubation cups attached to a rocker arm
assembly (Mount, 1968). Both flow-through and static renewal incubation have
been used. On day one, the embryos are removed from the cups and those that
are dead and fungused are removed. After day one, only dead embryos are
removed from the cups. Most of the embryos will hatch in five days if
incubated at 25°C.
5.2.8.1.4 During the incubation period, the eggs are examined daily for
viability and fungal growth, until they hatch. Unfertilized eggs, and eggs
that have become infected by fungus, should be removed with forceps using a
table top magnifier-illuminator. Non-viable eggs become milky and opaque, and
are easily recognized. The non-viable eggs are very susceptible to fungal
infection, which may then spread throughout the egg mass. Removal of fungused
eggs should be done quickly, and the spawning substrates should be returned to
the incubation tanks as quickly as possible so that the good eggs are not
damaged by desiccation.
5.2.9 Larvae Rearing Tanks
5.2.9.1 Newly-hatched larvae are transferred daily from the egg incubation
apparatus to small rearing tanks, using a large bore pipette, until the hatch
is complete. New rearing tanks are set up on a daily basis to separate fish
by age group. Approximately 1500 newly hatched larvae are placed in a 60-L
(15-gal) or 76-L (20-gal) all-glass aquarium for 30 days. A density of 150
fry per liter is suitable for the first four weeks. The water temperature in
the rearing tanks is allowed to follow ambient laboratory temperatures of 20-
25°C, but sudden, extreme, variations in temperature must be avoided.
5.2.10 Holding or Culture Tanks for Replacement Spawners
5.2.10.1 Replacement spawners (brood stock) are cultured from larvae produced
in the spawning tanks. After 30 days in a larval rearing tank, a number of
juveniles, equivalent to 2-4 days hatch are transferred to brood stock tanks
for a 30- to 60-day growth period. The sub-adults then are transferred to
500-L brood stock tanks to provide about 500 sub-adult fish per month for the
brood tank rotation. The surplus fish are transferred to 2000-L fiber glass,
or equivalent, holding tanks.
5.2.10.2 Surplus young males removed from spawning tanks, and other surplus
mature males, are placed in all-male holding tanks for future use as spawners.
Similarly, young and surplus mature females are held in all-female holding
tanks until needed as spawners. Tanks holding replacement spawners need not
be temperature-controlled, but for ease of transfer to the spawning tanks, it
is preferable to hold the water temperature close to that of the spawning
tanks (25 ± 2°C).
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5.2.11 Food and Feeding
5.2.11.1 Newly hatched brine shrimp nauplii or frozen adult brine shrimp and
commercial fish starter are fed to the fish cultures in volumes based on age,
size, and number of fish in the tanks. The amount of food and feeding schedule
affects both growth and egg production.
5.2.11.2 Fish from hatch to 30 days old are fed starter food at the beginning
and end of the work day, and newly hatched brine shrimp nauplii (from the
brine shrimp culture unit) twice a day, usually mid-morning and mid-afternoon.
Utilization of older (larger) brine shrimp nauplii may result in starvation of
the young fish because they are unable to ingest the larger food organisms
(see Appendix A.4 for instructions on the preparation of brine shrimp
nauplii). Avoid introducing Artemia cysts and empty shells when the brine
shrimp nauplii are fed to the fish larvae. Some of the mortality of the
larval fish observed in cultures could be caused from the ingestion of these
materials.
5.2.11.3 Fish older than four weeks are fed frozen brine shrimp and
commercial fish starter (#1 and #2), which is ground fish meal enriched with
vitamins. As the fish grow, larger pellet sizes are used, as appropriate.
(Starter, No. 1 and N. 2 granules, U.S. Fish and Wildlife Service
Formulation Specification Diet SD9-30, can be obtained from Zeigler Bros.,
Inc., P.O. Box 90, Gardners, PA 17324.
5.2.11.4 The spawning fish and pre-spawners in holding tanks usually are fed
all the adult frozen brine shrimp and tropical fish flake food or dry
commercial fish food (No. 1 or No. 2 granules) that they can eat (ad libitum)
at the beginning of the work day and in the late afternoon (i.e., 8:00 AM and
4:00 PM). The fish are feed twice a day, twice with dry food and once with
adult shrimp, during the week, and once a day on weekends.
5.2.12 Disease Control
5.2.12.1 Fish are observed daily for abnormal appearance or behavior.
Bacterial or fungal infections are the most common diseases encountered.
However, if normal precautions are taken, disease outbreaks will rarely, if
ever, occur. Hoffman and Mitchell (1980) have put together a list of some
chemicals that have been used commonly for fish diseases and pests.
5.2.12.2 Treatment of individual lots of infected fish should be carried out
separate from the main culture. Use of treated fish should be avoided, if
possible, and diseased cultures should be replaced.
5.2.12.3 In aquatic culture systems where filtration is utilized, the
application of certain antibacterial agents should be used with caution. A
treatment with a single dose of antibacterial drugs can interrupt nitrate
reduction and stop nitrification for various periods of time, resulting in
changes in pH, and in ammonia, nitrite and nitrate concentrations (Collins
et al., 1976). These changes could cause the death of the culture
organisms.
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with sodium hypochlorite. Also to avoid the contamination of cultures or
spread of disease, each time nets used to remove live or dead fish from tanks,
they are first sterilized with sodium hypochlorite or formalin, and rinsed in
hot tap water. Before a new lot of fish is transferred to culture tanks, the
tanks are cleaned and sterilized as described above.
5.2.13 Recording Keeping
5.2.13.1 Records are kept in a bound notebook, include: (1) type of food and
time of feeding for all fish tanks; (2) time of examination of the tiles
for embryos, the estimated number of embryos on the tile, and the tile
position number; (3) estimated number of dead embryos and embryos with
fungus observed during the embryonic development stages; (4) source of all
fish; and (5) daily observation of the condition and behavior of the fish.
6. VIDEO TRAINING TAPE AVAILABLE FOR CULTURIN6 METHODS
6.1 A video training tape and supplemental report (USEPA, 1989) on culturing
fathead minnows are available from the National Audiovisual Center, Customer
Services Section, 8700 Edgeworth Drive, Capitol Heights, Maryland, 20743-3701,
(Phone 301-763-1891), as part of a video package on short-term chronic
toxicity tests for freshwater organisms (Order No. EPA18036), which costs
$45.00.
7. REFERENCE TOXICANTS
7.1 It is recommended that static acute toxicity tests be performed monthly
with a reference toxicant. Fathead minnow larvae one to 14 days old are used
to monitor the acute toxicity of the reference toxicant to the test fish
produced by the culture unit.
8. TEST ORGANISMS
8.1 Fish one to 14 days old are used in acute toxicity tests.
8.2 If the fish are kept in a holding tank or container, most of the water
should be siphoned off to concentrate the fish. The fish are then transferred
one at a time randomly to the test chambers until each chamber contain 10
fish. Alternately, fish may be placed one to two at a time into small beakers
or plastic containers until they each contain five fish. Two of these
beakers/plastic containers (total of 10 fish) are then assigned to each
randomly-arranged control and exposure chamber.
8.3 The fish are transferred directly to the test vessels or intermediate
chambers using a large-bore, fire-polished glass tube (6 mm to 9 mm I.D. X 30
cm long) equipped with a rubber bulb, or a large volumetric pipet with tip
removed and fitted with a safety type bulb filler. The glass or plastic
containers should only contain a small volume of dilution water.
8.4 It is important to note that larvae should not be handled with a
dip net. Dipping small fish with a net may result in damage to the fish
and cause mortality.
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ASTM, Philadelphia, Pennsylvania.
Andreasen, J.K. 1975. Occurrence of the fathead minnow, Pimephales
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Andrews, A.K. 1970. Squamation chronology of the fathead minnow,
Pimephales promelas. Trans. Amer. Fish. Soc. 99(2):429-432.
Andrews, A.K. 1971. Altitudinal range extension for the fathead minnow
(Pimephales promelas). Copeia 1:169.
Andrews, A., and S. Flickinger. 1974. Spawning requirements and
characteristics of the fathead minnow. Proc. Ann. Conf. Southeastern
Assoc. Game Fish Comm. 27:759-766.
Benoit, D.A. and R.W. Carlson. 1977. Spawning success of fathead
minnows on selected artificial substrates. Prog. Fish. Cult.
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Brown, B.E. 1970. Exponential decrease in a population of fathead
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Brungs, W.A. 1971 a. Chronic effects of elevated temperature on the
fathead minnow (Pimephales promelas Rafinesque). U.S. Environmental
Protection Agency. EPA/600/8-81/011.
Brungs, W.A. 1971b. Chronic effects of low dissolved oxygen
concentrations on fathead minnows (Pimephales promelas). J. Fish.
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Buttner, J.K. and S.W. Duda. 1988. Maintenance and reproduction of
fathead minnows in the laboratory. Aquatic Ecology Section,
Department of Biological Sciences, SUNY College at Brockport,
Brockport, NY 14420.
Carlander, K. 1969. Handbook of freshwater fishery biology, Vol. 1.
Iowa State Univ. Press, Ames, Iowa.
Chiasson, A.G. and J.H. Gee. 1983. Swim bladder gas composition and
control of buoyancy by fathead minnows (Pimephales promelas) during
exposure to hypoxia. Can. J. Zool. 61(10):2213-2218.
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Clay, W. 1962. The Fishes of Kentucky. Kentucky Dept. Fish and
Wildlife Res., Frankfort, Kentucky.
Coble, D.W. 1970. Vulnerability of fathead minnows infected with yellow
grub to largemouth bass predation. J. Parasitol. 56(2):395-396.
Collins, M.T., J.B. Gratzer, D.L. Dawe, and T.G. Nemetz. 1976.
Effects of antibacterial agents on nitrification in aquatic
recirculating systems. J. Fish. Res. Bd. Can. 33:215-218.
Coyle, E.E. 1930. The algal food of Pimephales promelas (fathead
minnow). Ohio J. Sci. 30(l):23-35.
Cross, F.B. 1967. Handbook of fishes of Kansas. Univ. Kansas Mus.
Natur. Hist. Misc. Publ. 45:1-357.
Denny, J.S. 1987. Guidelines for the culture of fathead minnows
Pimephales promelas for use in toxicity tests. Environmental Research
Laboratory, U.S. Environmental Protection Agency, Duluth, Minnesota.
EPA/600/3-87-001.
Dixon, R.D. 1971. Predation of mosquito larvae by the fathead minnow,
Pimepha1es prome1 as Rafinesque. Manit. Entomol. 5:68-70.
Drummond, R.A. and W.F. Dawson. 1970. An inexpensive method for
simulating diel patterns of lighting in the laboratory. Trans. Am.
Fish Soc. 99:434-435.
Dymond, 1926. The Fishes of Lake Nipigon. Univ. Toronto Stud. Biol.
Ser. 27 Publ. Ont. Fish. Res. Lab 27:1-108.
Eddy, S., and A.C. Hodson. 1961. Taxonomic keys to the common animals of
the north central states. Burgess Publ. Co., Minneapolis, Minnesota.
Flickinger, S.A. 1966. Determination of sexes in the fathead minnow.
Trans. Amer. Fish. Soc. 98(3):526-527.
Flickinger, S.A. 1973. Investigation of pond spawning methods for
fathead minnows. Proc. Ann. Conf. Southeast. Assoc. Game and Fish
Commiss. 26: 376-391.
Gale, W.F., and G.L. Buynak. 1982. Fecundity and spawning frequency of
the fathead minnow--A fractional spawner. Trans. Amer. Fish. Soc.
111:35-40.
Gast, M.H. and W.A. Brungs. 1973. A procedure for separating eggs of
the fathead minnow. Prog. Fish. Cult. 35:54.
Guest, W.C. 1977. Technique for collecting and incubating eggs of the
fathead minnow. Prog. Fish. Cult. 39(4):188.
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Hedges, S., and R. Ball. 1953. Production and harvest of bait fishes in
ponds. In.: Michigan Dept. Conservation. Misc. Publ. 6, Lansing,
Michigan, pp. 1-30.
Held, J.W. and J.J. Peterka. 1974. Age, Growth, and food habits of the
fathead minnow, Pimephales promelas, in North Dakota saline lakes.
Trans. Am. Fish. Soc. 103(4):743-756.
Hendrickson, G.L. 1979. Ornithodiplostomum ptychocheilus: migration to
the brain of the fish intermediate host, Pimephales promelas. Exp.
Parasit. 48:245-258.
Herwig, N. 1979. Handbook of drugs and chemicals used in the treatment
of fish diseases. Charles C. Thomas, Publ., Springfield, Illinois.
272 pp.
Hoffman, G.L. 1958. Studies on the life cycle of Ornithodiplostomum
ptychocheilus (Faust) (Trematoda: Strigeoidea) and the "self cure" in
infected fish. J. Parasitol. 44(4):416-421.
Hoffman, G.L., and A.J. Mitchell. 1980. Some chemicals that have been
used for fish diseases and pests. Fish Farming Exp. Sta., Stuttgart,
Arkansas 72160. 8 pp.
Hubbs, C.L., and G.P. Cooper. 1935. Age and growth of the long eared and
the green sunfishes in Michigan. Pap. Mich. Acad. Sci. Arts. Letts.
20:669-696.
Hubbs, C.L, and K.F. Lagler. 1949. Fishes of Isle Royale, Lake Superior,
Michigan. Pap. Mich. Acad. Sci. Arts. Letts. 33:73-133.
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Univ. Mich. Press, Ann Arbor, Michigan.
Ingram. R. and W.D. Wares, II. 1979. Oxygen consumption in the fathead
minnow (Pimephales promelas Rafinesque) II: Effects of pH, osmotic
pressure, and light level. Comp. Biochem. Physio!. 62A: 895-8971
Isaak, D. 1961. The ecological life history of the fathead minnow,
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Arbor, Michigan. 150 pp.
Kimsey, J.B., and L.O. Fisk. 1964. Freshwater nongame fishes of
California. Calif. Dept. Fish and Game., Sacramento, California.
Klak, G.E. 1940. Neascus infestation of blackhead, blunt nosed, and
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Klinger, S.A., J.J. Magnuson, and G.W. Gallepp. 1982. Survival
mechanisms of the central mudminnow (Umbra limi), fathead minnow
(Pimephales promelas), and brook stickleback (Culea inconstans) for low
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Konefes, J.L. and R.W. Bachmann. 1972. Growth of the fathead minnow
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Carolina. 27611.
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Pimephales promelas, as a forage fish. Trans. Am. Fish. Soc. 57:92-99.
Manner, H.W. and C.M. Casimira. 1974. Early embryology of the fathead
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Manner, H.W., M. VanCura, and C. Muehlman. 1977. The ultrastructure of
the chorion of the fathead minnow, Pimephales promelas. Trans. Amer.
Fish. Soc. 106(1):110-114.
Markus, H. 1934. Life history of the blackhead minnow (Pimephales
promelas). Copeia 1934:116-122.
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APPENDIX A
SYSTEMATICS AND TAXONOMY, DISTRIBUTION, LIFE HISTORY, GENERAL DESCRIPTION,
AND HOLDING AND ACCLIMATION PROCEDURES
A.6 RAINBOW TROUT, ONCORHYNCHUS MYKISS AND
BROOK TROUT, SALVELINIUS FONTINALIS 1
1. RAINBOW TROUT
1.1 SYSTEMATICS AND TAXONOMY
1.1.1 Rainbow trout are native to the streams of the Pacific coast where
several varieties or strains have developed. The seagoing form is known as
the steel head trout and is thought to be identical to the strictly freshwater
rainbow form. Many other strains, for example, the inland lake form (Kamloops
trout) are found in other watersheds. Because of the ease with which the eggs
can be transported, different strains have been distributed all over the
world.
1.1.2 Rainbow trout are a variable species that differ considerably over the
whole of their range. Populations in different regions and watersheds of
North America have been referred to over the years by different scientific
names (e.g. species, distinct subspecies, or variants of a single species and
different regional common names). In recent years the validity of the generic
name, Salmo, for some western North American trout species has been
questioned. Fish taxonomists agree that native "Salmo" trouts of the northern
Pacific Ocean drainage are closely related with Pacific salmon Oncorhynchus
spp. The American Society of Ichthyologists and Herpetologists and the
American Fisheries Society have accepted Oncorhynchus as the appropriate
generic name for all native Pacific drainage trouts that are presently called
Salmo, based on new data and evidence by Smith and Stearly (1989).
Furthermore, the Names of Fishes Committee of the American Fisheries Society
has adopted the specific name, Oncorhynchus mykiss, for the rainbow trout and
its anadromous form, steelhead trout. The new names for the other North
American species affected are the following: Apache trout (0. apache),
cutthroat trout (0. clarki), Gila trout (0. gilae), golden trout (0.
aguabonita), and Mexican golden trout (0. chrysogaster).
1.2. DISTRIBUTION
1.2.1 The native range of the rainbow trout group (all varieties) in North
America is west of the Rocky Mountains and along the eastern Pacific Ocean,
but the species (Oncorhynchus mykiss) has now been introduced into many parts
of the continent (Figure 1). Except for the northern and southern extremes of
1 Prepared by Donald J. Klemm.
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the rainbow trout range, anadromous populations occur in all coastal rivers.
This species, under all its common names (rainbow trout, Kamloops trout,
steelhead trout, steelhead, coast rainbow trout, and silver trout), has been
so widely introduced in North America outside its natural range as to suggest
it may occur throughout the United States in all suitable habitats. Rainbow
trout are widely introduced and established in appropriate cold water habitats
all over the world.
Figure 1. Map showing the distribution of the rainbow trout in North
America. (Modified from Lee et al., 1980).
1.3. GENERAL LIFE HISTORY
1.3.1 In its natural environment of flowing streams of the western mountains,
the rainbow trout (Figure 2) thrives best at temperatures ranging from 3°C in
the winter to 21°C in the summer, but the optimum temperature is between 10-
16°C. The rainbow trout can withstand higher and lower temperature if it is
acclimated gradually. However, the rainbow trout's growth is impeded by
extremes of temperature, for example, above 27°C which it can tolerate only
for short periods of time.
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Figure 2. Rainbow trout (Modified from Eddy and Underbill, 1974)
1.3.2 Rainbow trout are basically spring spawners, but they can spawn at the
beginning of summer or early winter, depending on climate, elevation, and
genetic strain. If the spawning occurs in late fall or in winter, the eggs do
not hatch until spring. Prior to the spawning season adult males develop a
kype (elongated hooked snout) on the lower jaw and their colors intensify.
Males and females usually migrate upstream and select spawning sites in beds
of fine, clean gravel in riffles or runs above pools in streams. Long
journeys may be made by lake-dwelling rainbow (or Kamloops) and steelhead
trouts or anadromous, ocean-run rainbow steelheads. If the rainbow trout are
confined to land-locked lakes, they move into shallow shoals or reefs of
gravel and sand for spawning. Females dig out pits or sweep out depressions
(redds) in the gravel or sand and later spawn with males. Males are capable
of displaying aggressive behavior on the spawning grounds and can drive other
males away from a redd occupied by a female. In general, one or more males
court the digging female by sliding along side and crossing over her body and
rubbing their snout against her caudal peduncle with body pressing and body
vibrations. The female deposits her eggs, which are 3-5 mm in diameter,
demersal, and pink to orange in color. The eggs are immediately fertilized by
one or more males, fall into spaces between the gravel, and are covered with
loose gravel or sand to depths of 20 cm or more by the female. Females are
capable of digging and spawning in several redds with the same male or
different males. The number of eggs released can range from 400-3000,
depending on the size of the female.
1.3.3 Eggs usually hatch in approximately four to seven weeks. The time of
hatching, however, varies greatly with region and habitat. If the stream
temperature averages 7°C, eggs will hatch in about 48 days. The newly hatched
fish, called alevins, have a yolk sac, which is absorbed in three to seven
days. After the yolk sac is absorbed, the young are called fry, and begin
feeding in 10-15 days. In general, rainbow trout feed on a variety of
invertebrates. Also, depending on their size and the habitat in which they
live, other fishes and fish eggs, especially salmon, can be important food.
The fry of lake-resident spawners move up or down the spawning river to the
lake, or they may spend as much as one to three years in the streams. The
stream-resident spawners remain in the streams, whereas the steelhead trout,
which are stream-spawners, migrate to the sea, usually after 1-4 years in
freshwater.
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1.3.4 The growth of rainbow trout is highly variable with the area, habitat,
type of life history, and quantity and type of food. Some males may be good
breeders at two years of age, but few females produce eggs until their third
year of life. Rainbow trout young attain finger!ing size of about 76 mm by
the end of their first summer. The length may range between 178-204 mm at the
end of the second year, 279-382 mm after the third year, 356-406 mm after the
fourth year, and 406 mm or more after the fifth year. Lake- and ocean-run
rainbows may grow over twice as fast as this. However, the average length of
rainbow trout (or Kamloops trout) is 305-458 mm and that of steelhead trout is
508-762 mm. Under favorable conditions of artificial propagation, yearlings
average about 28 g, 2-year-olds about 255 g, 3-year-olds between .45-9 kg, and
4-year-olds between 1.4-1.8 kg. Returning sea-run individuals weigh up to 18
kg, or even more, but usually between 1.4-9 kg with the majority weighing less
than 5.4 kg. Some western varieties weight up to 23 kg, but the midwest
rainbows are much smaller. Those in streams are rarely over 1.4 kg, but in
some large lakes (e.g., Lake Superior) and in some western lakes they may
reach 7 kg or much larger. The life expectancy of rainbow trout can be as low
as three or four years in many streams and Lake populations, but that of
seagoing steelhead rainbow trout and Great lakes populations would appear to
be 6 to 8 years (Scott and Crossman, 1973).
1.4. GENERAL DESCRIPTION
1.4.1 Adult rainbow trout are bluish or olive green above and silvery on the
sides, with a broad pink lateral stripe that is enhanced during the spawning
season. The back, the sides, and the dorsal and caudal fins are profusely
dotted with small dark spots. Their color is variable with habitat, size, and
sexual condition. Stream forms and spawners are generally darker with color
more intense, lake forms lighter, brighter, and more silvery. Different color
types are often called by different names, e.g., darker stream fish often
called rainbows; larger, brighter, silvery fish in western lakes often called
Kamloops trout, and large silvery specimens returning from the sea and in the
Great Lakes or tributaries called steelhead trout. The scales are large,
numbering 120 to 150 in the lateral line. The caudal fin is very slightly
forked. The dorsal fin has 11 rays, and the anal fin has from 10 to 12 rays.
1.4.2 Young rainbow trout are typically blue to green on the dorsal surface,
silver to white on the sides and white ventrally. There are 5-10 dark marks
on the back between the head and dorsal fin. Also, there are 5-10 short,
dark, oval parr marks widely spaced on the sides, straddling the lateral line
with some small dark spots above but not below the lateral line. The dorsal
fin has a white to orange tip and a dark leading edge, or a series of bars or
spots. The adipose fin is edged with black, and the anal fin has an orange to
white tip.
2. BROOK TROUT
2.1 SYSTEMATICS AND TAXONOMY
2.1.1 Brook trout can be found exhibiting some variation in growth rate and
color throughout its range, but is considered a stable and well-defined
species (American Fishery Society, 1980). Male brook trout may be crossed
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with female lake trout (Salvelinus namaycush) to produce fertile hybrids that
are known as splake. Troutman (1981) and other papers cited in this section
indicate that brook trout can naturally and artificially hybridize with brown
trout (Salmo trutta) and rainbow trout (Oncorhynchus mykiss). For additional
information and discussion on freshwater and anadromous brook trout stocks and
systematic notes of brook trout, see Scott and Crossman (1973) and other
papers cited in this section.
2.2 DISTRIBUTION
2.2.1 The native range of the brook trout (Figure 3) is eastern North America,
extending throughout much of eastern Canada from Hudson Bay and Ungava Bay
drainages and Labrador; southward through the New England States and in the
Appalachian Mountains to the headwaters of the Savannah, Chattahoochee, and
Tennessee Rivers in the Carolinas and Georgia. In the Great Lakes, brook
trout are native to Lake Superior and tributaries to the northern tip of the
Lower Peninsula, the interior of the Great Lakes basin. They are also native
to a few far-northern headwaters of the upper Mississippi river system, and
western Minnesota and northeastern Iowa.
2.2.2 The brook trout has been widely introduced to higher elevations in
western North America. This species is also found in temperate regions of
other continents. Inland forms are found in colder lakes and streams, and
sea-run (anadromous) forms are found in the northeastern North American
coastal water areas.
Figure 3. Map showing the distribution of the brook trout in North
America. (Modified from Lee et al., 1980).
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2.3 GENERAL LIFE HISTORY
2.3.1 Brook trout (Figure 4) are generally found in clear brooks, streams,
and rivers in which the mean temperature rarely exceeds 10°C. The optimum
temperature is reported as ranging from 7 to 13°C, but they may be found
living in waters with temperatures ranging from 1 to 22°C (Piper et al. 1982)
The brook trout usually inhabits waters which flow less swiftly than those
inhabited by the rainbow. Brook trout also thrive in the small cold-water
lakes of the Great Lakes region, provided that suitable spawning conditions
exist.
Figure 4. Brook trout (Modified from Eddy and Underbill, 1974)
2.3.2 Brook trout spawn in late summer or autumn, the date varying with
latitude and temperature, usually from late October to December when the water
temperature is suitable although some may start spawning in September in
certain streams flowing into large lakes. Some females are capable of
spawning when they are a year old, while others do not mature until the second
year. When the spawning season occurs, brook trout move upstream into small
head waters or brooks where they select gravel and sand substrates usually in
shallow riffle areas or the tail-ends of pools for the spawning beds.
Spawning usually occurs during the day.
2.3.3 The female prepares a nest (redd), similar to those of the rainbow
trout, by sweeping out a depression in the gravel and sand substrate. During
preparation of the redd, the male starts courtship by quivering around the
female and driving off all intruders. When the female is ready to spawn, she
takes a position above and close to the redd. The male gets close to her side
and arches his body over hers, discharging milt as the female deposits her
eggs. Occasionally second male may join them in the spawning. After
spawning, the male leaves.
2.3.4 The eggs are 3.5 to 5.0 mm in diameter, are adhesive, and adhere to the
gravel at the bottom of the redd. The female pushes loose gravel and sand to
the center, covering the entire redd, and then desert the nest. A female may
spawn several times, and the number of eggs can vary from 100 to 5000,
depending on the size of the female.
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2.3.5 The eggs remain in the redd until the water temperature rises during
the following spring. If the level of DO is adequate, the eggs will hatch in
approximately 75 days at an average water temperature of 6.1 C, and in
approximately 50 days at an average temperature of 10°C. The upper lethal
temperature limit for developing eggs is about 11.7°C (Scott and Grossman,
1973).
2.3.6 After the eggs hatch, the larvae (sac fry) remain in the gravel of the
redd until the yolk is absorbed. Depending on the water temperature, it may
take from one to three months for the yolk sac to be absorbed (Lagler, 1956).
While the yolk sac is absorbed, the fry work themselves free from the gravel
and start feeding. They become free swimming at about 38 mm long. Under
natural conditions, newly hatched brook trout establish small feeding
territories in the stream and feed on small aquatic insects, insect larvae,
and other organisms.
2.3.7 Growth of brook trout is extremely variable, depending on the
suitability of the environment. The average length attained at various ages
may approximate 8.9 cm the first year; 15.2 cm the second year, 22.9 cm the
third year, 30.5 cm the fourth year, and 33 cm the fifth year. Brook trout
generally do not exceed a length of 54 cm and a weight of 1.5 kg (Troutman,
1981). However, Scott and Crossman (1973) reported a brook trout as large as
6.6 kg. Rumors of larger brook trout have been circulated, but none have been
verified. Brook trout may overpopulate small streams, resulting in large
numbers of small trout less than 25.4 cm long. Wild brook trout seldom live
longer than five years, and rarely live more than eight years.
2.4 GENERAL DESCRIPTION
2.4.1 The sides of large young and adult brook trout are dark olive,
sprinkled with light spots and red spots outlined with purplish or blue hue.
Some forms have red spots with light brown margins. The scales are cycloid,
small, in about 215 to 250 rows at the lateral line. The top of the head and
back is dark olive and heavily vermiculated. There are no black or brown
spots on the head, back, adipose, or caudal fin. The anterior rays of the
pectoral, pelvic, and anal fins are milk-white, bordered posteriorly with a
dusky hue and the remainder of the fins yellowish or reddish.
2.4.2 The back of young or immature brook trout is olive, the sides are
lighter and more silvery, and the belly is whitish. There are between 8-12
rectangular parr marks on the sides, also a few yellow and blue spots, but no
black spots.
2.4.3 The dorsal fin has 10 rays, and the anal fin has 9 rays. The belly of
breeding males is red, and some males may develop a hook (or kype) at the
front of the lower jaw. The tail or caudal fin is slightly notched in the
young but is generally square in older brook trout.
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3. HOLDING AND ACCLIMATION PROCEDURES FOR TROUT STOCKS
3.1 SOURCES OF ORGANISMS
3.1.1 Trout fry are obtained from commercial hatcheries during March through
July. However, if trout are needed for toxicity testing, it is advisable to
contact the hatchery for its trout hatching and rearing schedule. If trout
must be ordered from out-of-state, the State Fish and Game Agency should be
contacted concerning regulations on fish importation. The recommended age for
test organisms is approximately 15-30 days (after yolk sac absorption to 30
days) for rainbow trout and 30-60 days for brook trout. Trout are purchased
36 to 48 h prior to their use as testing organisms, but they must have time to
stabilize over the acclimation period. Trout should appear disease-free and
unstressed, with fewer than 5% of the animals dying during the 24-48 hours
preceding use in a toxicity test.
3.1.2 Trout fry are usually transported in plastic bags of at least 4-mil
plastic or thicker in shipping containers. The bags are partially filled with
water saturated with oxygen. During warm weather the shipping containers are
cooled with ice or cold packs to prevent temperature increases which will
result in the loss of fish. Trout should be acclimated gradually from the
temperature of the transportation unit to that of holding environment. Upon
arrival at the destination the plastic bags should be allowed to float
unopened in the holding tank for about 30 minutes to acclimate the fish.
3.2 HOLDING CONDITIONS
3.2.1 Trout are held in 200-L (50-gal) or larger tanks supplied with a flow-
through water system, or with recirculated water and a biological filtration
system. The holding water should be moderately hard and free of chlorine,
have low concentrations of metals, and should have a pH between 6 and 9.
Provide a daily photoperiod of 16 hours light, 8 hours darkness with an
illumination at 10-20 uE/m2/s (50-100 ft-c, or ambient laboratory levels). A
15-min dimmer timer should be used to gradually increase or decrease the
illumination when lights are turned on or off. The gradual increase and
decrease of illumination at the beginning and ending of the photoperiod is
important because trout tend to jump when startled by a sudden change in light
intensity. Holding water temperature is maintained at 12°C ± 2°C and is
aerated as close as possible to saturation. Measurements of temperature, DO,
pH, conductivity, and ammonia are made on holding water daily.
3.3 FEEDING
3.3.1 Trout are fed fine texture trout chow which can be obtained from
Zeigler Bros., Inc., P.O. Box 95, Gardners, PA, (717) 677-6181 or Rangen Inc.,
Buhl, ID, (208) 543-6421. The fry in the holding tank are fed (ad libitum) up
to 24 hours before the start of the acute toxicity test. Dead or moribund
fish should be removed from the holding tanks every day. Excess food and
feces are vacuum-siphoned off the bottom of the tank daily.
3.3.2 Daily records should be maintained for organism survival, health, and
acclimation conditions.
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4. TEST ORGANISMS
4.1 Rainbow trout fry 15 to 30 days old, and Brook trout 30-60 days old, are
used in acute tests (see summary tables of test conditions in Section 9). The
fry in the holding tank are not fed for 24 hours prior to the start of the
test. The fry are caught carefully with a fine mesh net and placed gently in
the 5 L (4 L test solution volume) test chambers, until 10 fish are reached
per test chamber. Larger test chambers or 5 fish/chambers may be necessary if
DO or pH problems are encountered. Placement of the test chambers is random.
4.2 After the fish are introduced, the behavior should be noted and recorded
throughout the test period. At the beginning and ending of the photoperiod,
during the test, the light intensity should be raised and lowered gradually
over a 15-min period using a dimmer switch or suitable device. Between
observations the test vessels are covered to act as a dust barrier and to
prevent fish from jumping out.
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SELECTED REFERENCES
American Fisheries Society. 1980. A list of common and scientific names of
fishes from the United States and Canada. Special Publication No. 12.
Amer. Fish. Soc., Bethesda, Maryland.
Bailey, R.M and C.R. Robins. 1989. Changes in North American fish names,
especially as related to the International Code of Zoological
Nomenclature, 1985. Bull. Zool. Nomencl. 45(2):92-103.
Eddy, S. and A.C. Hodson. 1970. Taxonomic keys to the common animals of the
north central states. Burgess Publ. Co., Minneapolis, Minnesota.
Eddy, S. and J.C. Underfill!. 1974. Northern fishes. Univ. Minnesota Press,
Minneapolis, Minnesota.
Hubbs, C.L. and K.F. Lagler. 1967. Fishes of the Great Lakes Region. Univ.
Michigan Press, Ann Arbor, Michigan.
Lagler, K.F. 1956. Freshwater Fishery Biology. Wm. C. Brown Co., Publ.,
Dubuque, Iowa.
Lee, D.S., C.R. Gilbert, C.H. Hocutt, R.E. Jenkins, D.E. McAllistger, and
R. Stauffer, Jr. 1980. Atlas of North American freshwater fishes. Publ.
1980-12, North Carolina State Museum Nat. Hist., Raleigh, North Carolina.
Leitritz, E. and R.C. Lewis. 1976. Trout and salmon culture (Hatchery
methods). California Dept. Fish and Game, Fish Bulletin 164. Sacramento,
California.
National Academy of Sciences. 1974. Fishes - Guidelines for the breeding,
care, and management of laboratory animals. Printing and Publishing
Office, National Academy of Sciences, Washington, D.C.
Piper, R.G., I.B. McElwain, L.E. Orme, J.P.McCraren, L.G. Fowler, and
J.R. Leonard. 1982. Fish hatchery management. U.S. Dept. Interior, Fish
and Wildlife Service, Washington, D.C.
Scott, W.B. and E.J. Crossman. 1973. Freshwater fishes of Canada. Fisheries
Research Board of Canada, Ottawa, Canada.
Smith, G.R. and R.F. Stearly. 1989. The classification and scientific names
of rainbow and cutthroat trouts. Fisheries 14(1):4-10.
Trautman, M.B. 1981. The fishes of Ohio. Ohio State Univ. Press and Ohio
Sea Grant Program, Center Lakes Erie Area Research, Columbus, Ohio.
Millers, B. 1991. Trout Biology: A natural history of trout and salmon.
Lyons and Burfoud, 31 West 21 Street, New York, New York 10010.
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APPENDIX A
MORPHOLOGY AND TAXONOMY, LIFE HISTORY, AND CULTURE METHODS
A.7 SHEEPSHEAD MINNOW (CYPRINODON VARIEGATUS)^
1. MORPHOLOGY AND TAXONOMY
1.1 The sheepshead minnow (Cypn'nodon van'egatus) belongs to the family
Cyprinodonitidae (killifishes), which includes 45 genera and 300 species
worldwide, occurring on all continents except Australia. Most species
are freshwater, but some occur in brackish and coastal marine waters.
There are thirteen species in the Genus Cypn'nodon in the United States
(American Fisheries Society, 1980). The sheepshead minnow is the only
marine species, and is widely distributed in the coastal waters of the
Atlantic and Gulf of Mexico.
1.2 Adult sheepshead minnows (see Hardy, 1978, for a complete
description.) can attain a total length of 93 mm, but the average
standard length report for adults is 35-50 mm. The males are usually
somewhat longer than females. The fish have the following morphological
characteristics: lack a lateral line; have 24-29 lateral scale rows; have
a large elongate humeral scale just above the pectoral base; the dorsal
fin has nine to 13 rays; the anal fin has nine to 12 rays; the caudal fin
has 14-16 principal rays and a total of 28-29 rays; the pectoral fin has
14-17 rays, and the ventral fin has five to seven rays.
1.3 The body of males is short, compressed, and deep. The depth
increases with age. The upper profile is evenly elevated. The males are
olivaceous above with a lustrous steel blue or bluish green area on the
back from nape to dorsal or beyond, and have a series of poorly defined
dark bars on the sides and a belly that is yellowish white to deep
orange. The dorsal fin ocellus on posterior rays is lacking or developed
as faint dusky spot.
1.4 The females are light olive, brown, brassy, or light orange above
with 14 dark crossbars on the lower sides alternating with seven to eight
crossbars on the back. The lower sides and belly are yellowish or white.
The dorsal fin is olive or dusky and has one or two prominent
ocelli on the posterior rays.
Prepared by Donald J. Klemm.
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2. LIFE HISTORY
2.1. Distribution and General Ecology
2.1.1 Sheepshead minnows occur in estuaries along the Atlantic and Gulf
coasts (Figure 1). They are a schooling, euryhaline species that inhabit
a variety of shallow water habitats, such as coves, bays, ponds, inlets,
harbors, bayous, salt marshes, and along open beaches. In some cases,
they may be very abundant where the bottom is partially sandy, emergent
vegetation lacking, and little current or wave action are present. This
species may establish populations in inland lakes containing relatively
high concentrations of dissolved salts. They are tolerant of extreme
Figure 1. Map showing the distribution of the sheepshead minnow
(Cyprinodon variegatus) in North America. Open circles
represent transplanted populations. (From Lee et al., 1980)
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changes in water temperatures, ranging from 0-40°C, and in salinities,
ranging from 0.1 to 149 °/oo (Simpson and Grunter, 1956; Nordlie, 1987).
2.1.2 This omnivorous fish is an important component of the estuarine
ecosystem serving as a link in transferring energy from lower trophic
levels, detritus and benthic plants and animals, to carnivores in higher
trophic levels (Hansen and Parrish, 1977). Sheepshead minnows serve as
forage fish for commercially and recreationally valued fish species, such
as the black drum (Pogonias croon's), red drum (Sciaenops ocellata),
bluefish (Pomatomus saltatrix), spotted seatrout (Cynoscion nebulosus),
striped bass (Morone saxatilis), and snook (Centropomus undecimalis)
(Gunter, 1945; Darnell, 1958; Grant, 1962; Sekavec, 1974, and Carter et
al., 1973).
2.2 General Spawning Behavior
2.2.1 Sheepshead minnows (Figures 2, 3, 4) spawn at depths of 2.5 to 60
cm in shallow bays, tide pools, mangrove lagoons, and pools in shallow,
gently flowing streams, and other similar habitats over bottoms of sand,
black silt, or mud. Males occupy territories up to 0.3-0.6 no in diameter
and may or may not construct nest pits. Spawning may take place out of
both pit and territory. Besides temperature, Martin (1972) reported that
sudden changes in salinity can initiates spawning activities. Eggs
(Figure 2) are demersal, adhesive or semi-adhesive with very minute
attachment filaments (threads) more or less evenly distributed over the
chorion. They stick to a variety of substrates, such as plants, sand,
rocks, logs, and to each other. Sometimes they stick to plants near the
surface, and at other times become partially buried in the bottom. The
yolk contains one very large and many minute oil globules. Adults spawn
possibly throughout the year on the Gulf coast of the United States.
Hansen and Parrish (1977) reported that in an estuary near Pensacola,
Florida, spawning may occur during any month of the year. Ripe females
are found April to October in North Carolina, throughout the summer in
the Chesapeake Bay, May to August in Delaware Bay, May to September in
New Jersey and New York, and June to mid-July in Massachusetts.
3. CULTURE METHODS AND FACILITIES
3.1. Sources of Organisms
3.1.1 Juvenile and adult Sheepshead minnows (Figure 4) for use as brood
stock spawners may be obtained from commercial biological supply houses
or taken by seine in coastal estuaries of the Atlantic coast and Gulf of
Mexico. They may also be obtained from young fish raised to maturity in
the laboratory. Feral brood stock and first generation laboratory fish
may be preferred, to minimize inbreeding. A continuous supply of wild
stock, however, may be more cost effective. Neither fish nor eggs of
feral stock should contain excessive contaminants nor exhibit excessive
mortality, and the fish should demonstrate normal behavior. Before being
used as a source of gametes, field-caught adults should be maintained and
observed in the laboratory for at least one week to permit detection of
disease and to allow time for acute mortality resulting from stress
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Figure 2. Sheepshead minnow (Cvprinodon varieqatus). A. unfertilized egg, B,
9 blastodisc stage; C-D. 8-cell stage; E. 16-cell stage; F late
cleavage; G. germ ring formed; H. Blastoderm over 1/4 of yolk, I.
early embryo; J. embryo 48-hours old; K. tail-free embryo. (From
Kuntz, 1916).
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B
3.4 mm TL
4 .0 mm
4.0 mm
D
Foster, 1974)
E. yolk-sac larvae;
D' from
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T
-&.
CD
Co
0)
•^ •
00
cu
Q.
-------
of capture. Injured or diseased fish should be discarded.
3.2. Laboratory Culture Facilities
3.2.1 Sheepshead minnows can be cultured in a static, recirculated, or
flow-through systems. Flow-through systems require large volumes of water and
may not be feasible in some laboratories.
3.3. Laboratory Year-round Spawning
3.3.1 In the laboratory, adults may be kept in breeding condition year round.
Females may spawn a number of times at intervals of one to seven days, and
will generally produce an average of 10 to 30 eggs per spawning (Hansen,
1978). To obtain large number of eggs at one particular time, adult fish of
27 mm standard length or greater should be used. If fish are taken in the
field, they should be acclimated for at least one to two weeks in 20-30 °/oo
salinity, a water temperature of 25-28°C, and a photoperiod of 14 h light and
10 h dark.
3.3.2 Sheepshead minnows can be continuously cultured in the laboratory from
eggs to adults. The eggs (embryos), larvae, juveniles, and adults (Figures 2,
3, 4) should be kept in rearing and holding tanks of appropriate size and
maintained at ambient laboratory temperature. The larvae should be fed
sufficient newly-hatched Artemia nauplii daily to assure that live nauplii are
always present. At the juvenile stage, they are fed frozen adult brine shrimp
and a commercial flake food, such as TETRA SM-80R (available from Tetra Sales,
(U.S.A.), 201 Tabor Rd, Morris Plains, New Jersey 07950, phone: 800-526-0650),
MARDEL AQUARIAM" Tropical Fish Flakes (available from Mardell Laboratories,
Inc., 1958 Brandon Court, Glendale Heights, Illinois 60139, phone:
312-351-0606), or equivalent. Adult fish are fed flake food two or three
times daily, supplemented with frozen adult brine shrimp.
3.3.3 Sheepshead minnows normally reach sexual maturity three to five months
after hatching, and have an average standard length of approximately 27 mm for
females and 34 mm for males, if held at a temperature of 25-30°C in rearing
tanks of adequate size, and fed adequately. At this time, the males begin to
exhibit sexual dimorphism and initiate territorial behavior. When the fish
reach sexual maturity, and are to be used to obtain large number of embryos by
natural spawning, the brood stock should be kept in a temperature controlled
system at 18-20oC. To initiate spawning, the spawners are moved to spawning
tanks with a temperature of 25°C. Adults can be maintained in natural or
artificial seawater in a flow-through, static, or recirculating, aerated
system consisting of an all-glass aquarium, or a LIVING STREAM fiberglass,
circular or rectangular tank (Figid Unit, Inc., 3214 Sylvania Ave., Toledo,
Ohio 43613, phone 419-474-6971), or equivalent (see Middaugh, 1985,
EPA/600/4-85-013, and Middaugh, Hemmer, and Goodman, 1987, EPA/600/87/004, for
a recirculating system).
3.3.4 Static systems are equipped with an undergravel filter. Recirculating
systems are equipped with an outside biological filter constructed in the
laboratory using a reservoir system of crushed coral, crushed oyster shells,
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or dolomite and gravel, charcoal, floss, (see Spotte, 1973; 1979, Bower, 1983
for information on filters and conditioning the biological filter), or a
commercially available cartridge filter, such as a MAGNUM" Filter, available
from Carolina Biological Supply Co., Burlington, North Carolina 27215, phone
800-334-5551, an EHEIMR Filter, available from Hawaiian Marine Imports Inc.,
P.O. Box 218687, Houston, Texas 77218, phone 713-492-7864, or an equivalent
system. The culture conditions should include seawater at 20-30 °/°o, and a
photoperiod of 14 h light and 10 h dark. Water temperature may be controlled
or maintained at ambient laboratory levels.
3.4 Obtaining Eggs (Embryos) for Toxicity Tests
3.4.1 Embryos can be shipped to the laboratory from an outside source or
obtained from adults held in the laboratory. Ripe eggs can be obtained either
by natural spawning or by intraperitoneal injection of the females
with human chorionic gonadotrophin (HCG) hormone, available from United States
Biochemical Corp., Cleveland, Ohio 44128, phone, 216-765-5000. If the
culturing system for adults is temperature controlled, natural spawning can be
induced to obtain large number of embryos by raising the temperature to 25°C.
Natural spawning is preferred because repeated spawning can be obtained from
the same brood stock, whereas with hormone injection, the brood stock is
sacrificed in obtaining gametes. It should be emphasized that the injection
and hatching schedules given below are to be used only as guidelines.
Response to the hormone varies with brood stock and temperature. Time-to-
hatch and percent hatch also vary among stocks and among batches of embryos
obtained from the same stock, and are dependent on temperature, DO, and
salinity.
3.5. Natural Spawning
3.5.1 Adult fish should be maintained at 18-20°C in a temperature controlled
system. The number of spawning chambers and fish to be spawned should be
based on the requirements for providing sufficient numbers of viable embryos.
As indicated above, an adult female in spawning condition will generally
produce an average 10 to 30 eggs per spawn. To obtain embryos for a test,
adult fish (generally, at least eight-to-ten females and three males) are
transferred to a spawning chamber in a 57 L (15 gal) aquarium with the correct
photoperiod and temperature (14 h light/10 h dark, and a temperature of 25°C),
seven to eight days before the larval fish are needed. The spawning tank is
fitted with a spawning chamber and an embryo collection tray. The spawning
chamber consists of a basket of 3-5 mm NITEXR mesh, approximately 20 X 35 X
22 cm high (Hansen, et al., 1978), designed to fit into the aquarium.
Spawning generally will begin within 24 h or less. The embryos will fall
through the bottom of the spawning chamber and lightly adhere onto a
collecting screen or tray placed on the bottom of the tank. The collecting
tray should be checked for embryos the next morning. The number of eggs
produced is highly variable. The number of spawning units required to provide
the fish needed to perform a toxicity test (generally two to four) as
determined by experience. If the collecting trays do not contain sufficient
embryos after the first 24 h, discard the embryos, replace the tray, and
collect the embryos for another 24 h. To help keep the embryos clean, the
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adults are fed while the screens are removed. Spawning fish should be
shielded from excessive outside disturbance, e.g. an opaque curtain should
surround the entire culture system. Care should also be taken so that outside
light sources do not interfere with the photoperiod.
3.5.2 The embryos are collected in a tray placed on the bottom of the tank.
The collecting trays are fabricated from plastic fluorescent light fixture
diffusors (grids), with cells approximately 14 mm deep X 14 mm square. A
screen consisting of 250-500 urn mesh is attached to one side (bottom) of the
grid with silicone adhesive. The depth and small size of the grid protects
the embryos from predation by the adult fish. The collecting trays with
newly-spawned embryos are removed from the spawning tank, and the embryos are
collected from the screens by washing them with a wash bottle or removing them
gently with a fine brush. The embryos from several spawning units are
generally pooled in a single container to provide a sufficient number to
conduct the test(s). The embryos are transferred to a petri dish, or
equivalent, filled with fresh culture water, and are examined using a
dissecting microscope or other suitable magnifying device. Damaged and
infertile eggs are discarded (see Figure 2). The embryos are then placed in
incubation dishes (e.g. KIMAX or PYREXR crystallizing dishes, Carolina
culture dishes, or equivalent; see 3.8, Embryo Incubation and Hatching
Facility). It is recommended that the embryos be obtained from fish cultured
inhouse, rather than from outside sources, to eliminate the uncertainty of
damage caused by shipping and handling that may not be observable, but which
might affect the results of the test. After sufficient number embryos are
collected for the test, the adult fish are returned to the (18-20°C) culture
holding tanks.
3.6 Sustained Natural Embryo Production
3.6.1 Sustained (long-term), daily, embryo production can be achieved by
maintaining mature fish (ratio of approximately 12-15 males to 50-60 females)
in tanks, such as a 285-L LIVING STREAM" tank, or equivalent, at a temperature
of 23-25°C. Embryos are collected seven or eight days prior to starting the
acute or chronic toxicity tests for less than 24 hour or older larvae.
Embryos are produced daily, and when needed, collecting trays are placed on
the bottom of the tank. The next morning, the embryo collectors are removed,
and the embryos are washed into a shallow glass culture dish using artificial
seawater. Four collecting trays, each approximately 20 cm X 45 cm, will cover
the bottom of a 285-L tank.
3.7 Forced Spawning
3.7.1 Human chorionic gonadotrophin (HCG) is reconstituted with sterile
saline or Ringer's solution immediately before use. The standard HCG vial
contains 1,000 IU, which is reconstituted in 10 Ml of saline. Freeze-dried
HCG, which comes with premeasured and sterilized saline, is the easiest to
use. The reconstituted HCG may be used for several weeks if kept in the
refrigerator.
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3.7.2 Each female is injected with HCG on two consecutive days. The HCG is
injected into the peritoneal cavity, just below the skin, using the smallest
needle possible. A 50 IU dose (0.5 Ml of reconstituted hormone solution) is
recommended for females approximately 27 mm in standard length. A larger or
smaller dose may be used for fish which are significantly larger or smaller
than 27 mm. It may be helpful if fish that are to be injected are maintained
at 20°C before injection, and the temperature raised to 25°C on the day of the
first injection. Injected females should be isolated from males.
3.7.3 With injections made on days one and two, females which are held at
25°C should be ready for stripping on Days 4, 5, or 6. Ripe females should
show pronounced abdominal swelling, and release at least a few eggs in
response to a gentle squeeze. Eggs are stripped from the ripe females and
mixed with sperm derived from excised, macerated testes. At least ten females
and five males are used per test to ensure that there is a sufficient number
of viable embryos.
3.7.4 Prepare the testes immediately before stripping the eggs from the
females. Remove the testes from three-to-five males. The testes are paired,
dark-grey organs along the dorsal midline of the abdominal cavity. If the
head of the male is cut off and pulled away from the rest of the fish, most of
the internal organs can be pulled out of the body cavity, leaving the testes
behind. The testes are placed in a few Ml of seawater until the eggs are
ready.
3.7.5 Strip the eggs from the females into a dish containing 50-100 Ml of
seawater, by firmly squeezing the abdomen. Sacrifice the females and remove
the ovaries if all the ripe eggs do not flow out freely. Break up any clumps
of ripe eggs and remove clumps of ovarian tissue and under-ripe eggs. Ripe
eggs are spherical, approximately 1.0-1.7 mm in diameter, and almost clear.
Place the testes in a fold of NITEXR screen (250-500 urn mesh), dampen with
seawater, and macerate while holding over the dish containing the eggs. Rinse
the testes with seawater to remove the sperm from the tissue, and wash the
remaining sperm and testes into the dish with the eggs. Let the eggs and
sperm stand together for 10-15 minutes, swirling occasionally.
3.7.6 Pour the contents of the dish into a crystallizing dish or equivalent
and insert an airstone. Aerate gently, so that the water moves slowly over
the eggs, and incubate at 25°C for 60-90 min. After this period of time, wash
the fertilized eggs on a NITEXR screen, place them in clean seawater in an
incubation chamber.
3.8. Embryo Incubation and Hatching Facility
3.8.1 Embryos are incubated in KIMAXR or PYREXR crystallizing dishes,
Carolina culture dishes, or equivalent, at a temperature of 25°C and 14-h
light/10-h dark photoperiod. An air stone is placed in each dish, and the
contents are gently aerated for the duration of the incubation. The water in
the incubation chambers is replaced daily. Approximately 24 h prior to
hatching, the salinity of the seawater in the incubation chambers is changed
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to that of the test salinity, if different. The salinity must remain within
the 20 to 30 °/oo range. The embryos should hatch in 6-7 days at 25°C, and in
4 to 5 days at 30°C.
3.9 Feeding and Stocking Density
3.9.1 The sheepshead minnow cultures should be provided a sufficient amount
of high quality nutrition without over-feeding. The adult and juvenile
sheepshead minnows are fed, frozen adult brine shrimp and flake food, ad
libitum, daily. The larvae are fed newly hatched Artemia nauplii and crushed
flake food, ad libitum, daily. Methods for culturing brine shrimp are
discussed in Appendix A.4. The stocking of adult fish in the holding tanks
depends on the biological filter system (see Biological Filters and Substrate
Conditioning). A circular, 1.3 m (48 in.) diameter, 880 L (235 gal),
fiberglass tank will hold approximately 30-50 adult fish with a varied sex
ratio. A stocking density of about 300 larvae is suitable in a 76 L aquarium.
Brood stock should be replaced with feral fish annually, or whenever the
fecundity of the females diminishes, and they appear spent with age and from
frequent breeding.
3.10 Culture Tanks
3.10.1 Larvae, juvenile, and adult fish should be kept in holding and rearing
tanks of appropriate size. The tanks can be all-glass aquaria, fiberglass
tanks, or equivalent. All tanks should have appropriate biological filtration
systems, and the culture filtration system should be conditioned properly
before adding the fish (see Spotte, 973, 1979; Bower, 1983).
3.11. Biological Filters and Substrate Conditioning
3.11.1 Holding and rearing aquaria and tanks can accommodate as many fish as
its biological filter will permit. The substrate conditioning for the
undergravel or outside filters is also important to the life and health of the
fish. Substrate conditioning is the process to develop nitrifying
bacteria (Nitrosomonas and Nitrobacter) that can convert ammonia and nitrite
to nitrate. A conditioned filter bed is defined as one in which the capacity
for ammonia and nitrite oxidation is sufficient to keep pace with the
production of ammonia by the fish. Consult Spotte (1970; 1979) or Bower
(1983) for a thorough understanding of the biological filter and conditioning
process.
3.12 Culture Water
3.12.1 Artificial seawater is prepared by dissolving FORTY-FATHOMS" or
equivalent artificial sea salts in deionized water to a salinity of
20-30 °/°o- Synthetic sea salts are packaged in plastic bags and mixed with
deionized (MILLI-QR or equivalent) water. The instructions on the package of
sea salts should be followed carefully, and the salts should be
mixed in a separate container, and not in the culture tank. The deionized
water used in hydration should be in the temperature range of 21-26°C.
Seawater
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made from artificial sea salts is conditioned (see Spotte, 1973, 1979; Bower,
1983) before it is used for culturing by aerating mildly for at least 24 h.
3.12.2 Adequate aeration will bring the Ph and concentration of dissolved
oxygen and other gases into equilibrium. The concentration of dissolved
oxygen in the water supply should be 90-100% saturation before it is used. If
a residue or precipitate is present, the solution should be filtered before
use. The seawater should be monitored periodically to insure a constant
salinity.
3.13 Culture Conditions
3.13.1 Holding and rearing tanks and any area used for manipulating live
sheepshead minnows should be located in a room or space separated from that in
which toxicity test(s) are to be conducted. The salinity of the culture
systems should be between 20 and 30 °/oo. Water temperature for the brood
stock should be maintained at 18-20°C. A photoperiod of 14 h illumination
(10-20 uE/m2/s, or 50-100 ft-c) and 10 h dark, should be provided. The
holding and rearing tanks should be aerated so that the DO is not less than
1.0 ppm below saturation at any given temperature, with 5.0 ppm (60%
saturation) being the absolute lowest limit.
3.14 Culture Maintenance
3.14.1 Replace approximately 10% of the culture water every two weeks, or 25%
monthly. The culture water should be clear. If the water appears cloudy or
discolored, replace at least 50% of it. Replacement water should be well
oxygenated and at the same temperature and salinity as the culture water.
Salinity is maintained at the proper level by adding deionized water to
compensate for evaporation. A replenisher, made of the trace elements, iodine
(Kl) and bromine (KBr), is added (1 mL/400 L) to the culture water each week,
or commercial trace elements replenisher should be used as directed by the
artificial sea salt manufacturer.
3.14.2 To avoid excessive build up of algal growth, periodically scrape the
walls of culture system. Some of the algae will serve as a supplement to the
diet of the fish. A partial activated carbon "charcoal" change in the
filtration systems should be done monthly or as needed. The detritus (dead
brine shrimp nauplii and cysts, adult brine shrimp, other organic material
accumulation) should be siphoned from the bottom of rearing and holding
aquaria or tanks each week or as needed.
3.15 Water Quality Monitoring
3.15.1 Checking the chemistry of the sea water is critical to the success of
the marine culture system. The water quality will determine whether the life
support processes in the filter bed work at reasonable and steady rates. The
culture water is checked routinely for temperature, alkalinity, pH, DO, total
ammonia, nitrite, and nitrate. More frequent monitoring of these parameters
is recommended during periods of organism procurement and
starting new culture systems with inside underground filters and outside-of-
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tank biological filtration. The DO should be maintained at greater than 60%
saturation. The pH should not go below 7.5 with an acceptable range between
7.5 to 8.3. Low pH levels can result from overcrowding, overfeeding, or waste
accumulation, especially in static or recirculating culture systems.
3.15.2 Acceptable pH levels can be re-established by siphoning off 50-75% of
the water and replacing it with conditioned artificial seawater of the same
temperature. Also, sodium bicarbonate or commercially available liquid
buffers can be added to the tanks whenever the pH falls below 7.5. Un-ionized
ammonia, total (NH3 + NH4), and nitrite ion (N02) levels should not exceed 0.1
ppm in the holding tanks. It is recommended that the ammonia and nitrite
concentrations be determined prior to starting new culture systems. It is
recommended that nitrate (N03) concentrations be determined prior to starting
new culture systems, and the nitrate ion concentrations should not exceed
20 mg/L.
3.15.3 A specific schedule for water quality monitoring should be established
for each culture system. All water quality measurements and data are recorded
in the culture and environmental conditions log books.
3.16 Disease Control and Treatment
3.16.1 Discussions of identification and treatment of common parasites of
marine fish culturing can be found in Spotte (1973), Sindermann (1970), and
Bower (1983). Several commercial companies, e.g. Aquatronics, P.O. Box 12107,
La Costa Station, Malibu, California 09265; Marine Enterprises, Inc.,
Baltimore, Maryland (301) 321-1189; and Hawaiian Marine Imports, Inc.,
Houston, Texas (713) 492-7864, sell various kinds of medication to treat
common parasites of marine fish.
3.16.2 A colorless medication, FORMALITE IIR, available from Aquatronics,
has been used successfully for the treatment of the protozoan parasites,
ChilodoneTla, Cost/a, Tn'choina, Scyphidia, Trichophrya, and Ichyophirius.
4. VIDEO TRAINING TAPE AVAILABLE FOR CULTURING METHODS
4.1 A video training tape and supplemental report (USEPA, 1990) on culturing
sheepshead minnows are available from the National Audiovisual Center,
Customer Services Section, 8700 Edgeworth Drive, Capitol Heights, Maryland,
20743-3701, (Phone 301-763-1891), as part of a video package on short-term
chronic toxicity test methods for marine organisms (Order No. A18545). The
package includes methods for sheepshead minnows, inland silversides, sea
urchins, and Champia, and costs $85.00.
5. TEST ORGANISMS
5.1 Sheepshead minnows 1-14 days old are used in the acute toxicity test. If
the larvae are used one or two days after hatching, they can be held in the
crystallizing or culture dishes. If they are to be used later, they should be
placed in larger holding aquarium or tanks. Prior to beginning the test, the
larvae can be transferred to small beakers or plastic cups, using a
large-bore, fire-polished glass tube (6 mm to 9 mm I.D. X 30 cm long) equipped
with a rubber bulb.
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5.2 If the larvae are to be moved to holding aquaria, a large-bore,
fire-polished glass tube should also be used to move them. It is important to
note that larvae and fry should not be handled with a dip net. Dipping larvae
and fry with a net can result in very high mortality. Some of the water in
the holding aquarium or tank containing the larvae should be siphoned off
before they are transferred using the large-bore tube. This should make them
easier to catch. The same large-bore, fire-polished glass tube discussed
above should be used to gently transfer the fish from the holding vessels to
the test vessels. As the fish are counted, they can be transferred to small
plastic cups before they are added to the test vessels. It is more convenient
to first transfer five fish to each of several small beakers or plastic
containers with a few ml of 20-30 °/°° saline dilution water. The appropriate
number of fish (multiples of five) can then be added to the test vessels.
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APPENDIX A
MORPHOLOGY AND TAXONOMY, LIFE HISTORY, AND CULTURE METHODS
A.8 SILVERSIDES (MENIDIA SPP)1
1. MORPHOLOGY AND TAXONOMY
1.1 Adult Atlantic silversides attain a total length of up to 117 mm (Figure
1A and IB). Females in general are slightly larger than males. The first
dorsal fin has three to seven, usually four or five spines. The second dorsal
fin has one spine and eight or nine rays; the anal fin has one spine and 19 to
29, usually 21 to 26, rays; and the pectoral fin has 12 to 16, usually 14 or
15, rays (Robbins, 1969). Atlantic silverside embryos are easily
distinguished from those of the closely related inland silverside, Mem'dia
beryllina. The former have a bundle of elastic filaments attached to the
chorion at one small area of insertion (Figure 1C and ID). These filaments,
typically longer than the diameter of the egg, are all the same diameter. In
contrast, inland silverside eggs posses one or two thick, elongated filaments,
up to 50 mm long and four to nine shorter, thinner filaments (Figure IE and
IF).
2. GENERAL LIFE HISTORY
2.1 Distribution
2.1.1 Silversides occur in estuaries along the Atlantic, Gulf, and Pacific
coasts (Figures 2-4). The Atlantic silverside, Mem'dia menidia, is a resident
of estuaries from Maine to northern Florida. It occurs at intermediate to
high salinities, typically of 12 to 30 parts per thousand (ppt), and remains
in Atlantic estuaries throughout most of the year (De Sylva et al., 1962;
Dahlberg, 1972). Recent evidence indicates an offshore migration at northern
latitudes in the fall and reappearance of adults in estuaries in late spring
(Conover and Kynard, 1981). This species is an important component in
estuarine ecosystems, serving as forage fish for commercially and
recreationally valued species such as striped bass, bluefish and spotted
seatrout (Merriman, 1941; Bayliff, 1950; Middaugh, 1981).
2.1.2 Although the culturing methods described in this section were written
primarily for Menidia menidia, they are also suitable for the inland
silverside, M. beryllina, and the tidewater silverside, M. peninsulae
(Middaugh et al., 1987). The staff of the Environmental Research Laboratory,
Gulf Breeze, Florida, have developed procedures for spawning, culturing, and
testing of other fishes, including the California grunion, Leuresthes tenuis.
Prepared by Douglas P. Middaugh and Donald J. Klemm.
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Figure 1. Silverside (Menidia): A-D, M. menidia, (Atlantic silverside); A,
adult; ca. 95 mm SL (Massachusetts); B, adult, ca. 102 mm SL
(Florida); C, unfertilized egg (diagrammatic); D, developing embryo
(note that filaments are all equal in diameter); E-F, M. beryl!ina
(inland silverside); E, unfertilized egg (diagrammatic); F,
developing embryo (note one thick filament and several thin
filaments). (A,B from Kendall, 1902; C from Wang, 1974; D from
Ryder, 1883; E from Wang, 1974; F from Hildebrand, 1922.)
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B
Figure 2. Biographical Distribution: A, inland silverside, Menidia beryl!ina;
B, Atlantic silverside, M. menidia. (From Middaugh et al., 1987).
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Figure 3. Biogeographical distribution: A, tidewater silverside, Menidia
peninsulae; B, California grunion, Leuresthes tenuis. (From
Middaugh et al., 1987).
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Figure 4. Biogeographical distribution: Topsmelt, Athen'nops affinis.
and the topsmelt, Athen'nops affim's. The availability of these fishes as
test organisms will permit the use of indigenous fish in toxicity tests of
wastes discharged along the entire coast line of the contiguous United States
and Alaska.
2.2 Spawning Behavior
2.2.1 The Atlantic silverside spawns during spring and summer. Spawning runs
generally occur during April - June or July at northern latitudes, and March
through July or August at southern latitudes (Bayliff, 1950; Hildebrand and
Schroeder, 1928; Middaugh and Lempesis, 1976). Spawning occurs in the upper
intertidal zone during daytime high tides (Middaugh, 1981). Eggs are
deposited on a variety of substrates which provide protection from thermal
stress and desiccation (Middaugh et al., 1981; Conover and Kynard, 1981).
Females typically release 200 to 800 eggs, 1.0-1.2 mm diameter, as they spawn.
Individuals may spawn up to five or six times, at two week intervals, during
the reproductive season. The life span is generally 12-15 months, although
year class-2 fish are occasionally found (Beck, 1979),
3. CULTURING METHODS
3.1 Sources of Organisms
3.1.1 Mem'dia may be obtained from commercial biological supply houses or
collected in the field.
3.1.1.1 The optimal time for collecting ripe M. mem'dia in the field is just
prior to daytime high tides between 8:00 AM and noon (usually one to four days
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after the occurrence of a new or full moon), when prespawning schools move
into the upper intertidal zone (Middaugh, 1981; Middaugh et al., 1981). Since
the Atlantic silverside prefers relatively high salinities, it is recommended
that collections be made in areas with salinities of 20 °/oo or greater.
Sandy beaches, bordering open but protected estuarine bays, are suitable for
collecting adults. A 1 X 10-m bag seine with knotless 5-mm mesh is ideal for
collecting. Since Atlantic silversides typically reside in shallow water, 1.5
m deep, they are easily captured by seining close to shore. It is important
to avoid total beaching of the bag seine when collecting M. mem'dia. These
fragile fish will quickly die if removed from water and, more importantly,
ripe females often abort their eggs if stranded. Ideally, the bag portion of
the seine, containing captured adults, should remain in water 5-15 cm deep
(Middaugh and Lempesis, 1976).
3.1.1.2 It is possible to transport the spawn (fertilized eggs) or adults to
the laboratory. The following procedure is recommended for stripping,
fertilizing and transporting eggs from the field to the laboratory:
1. Immediately after seining (while still on the beach) three to five ripe
females should be dipped into a bucket of seawater to remove sand and
detritus.
2. Eggs are stripped into a glass culture dish containing seawater or onto a
nylon screen (0.45 to 1.0 mm mesh) (Figure 5), which is then gently
lowered into a culture dish of seawater with the eggs on the upper surface
of the screen (Barkman and Beck, 1976). If excessive pressure is required
to strip the eggs, the female should be discarded. Mature eggs, 1.0-1.2
mm in diameter, are clear, and have an amber hue.
3. Milt from several males can then be stripped into the culture dish and
mixed with the eggs by gently tilting the dish from side to side. Upon
contact with seawater, adhesive threads on mature eggs uncoil, making
enumeration and separation difficult. If eggs are stripped directly into
the culture dish, one end of a nylon string may be dipped into the dish
and gently rolled so the embryos adhere (Middaugh and Lempesis, 1976).
The Barkman and Beck (1976) technique for attaching the eggs to nylon
screening minimizes the natural clumping tendency due to entanglement of
the filaments on M. menidia eggs.
4. Strings of embryos or embryos on screens may be transported to the
laboratory by placing them in an insulated glass container filled with
seawater at the approximate temperature and salinity of fertilization.
If gravid fish are transported to the laboratory for subsequent spawning,
care must be taken to avoid overcrowding of fish in transport containers.
Continuous, vigorous aeration is required and any increase in container
water temperature should be minimized (Beck, 1979).
A mass culture system for incubating the screen-adhered eggs and
collecting the hatched larvae in a flowing seawater system (Figure 5)
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HAND STRIPPING EGGS
HAND STRIPPING MALES
ONTO 500/1 NYLON SCREEN
IMMERSED IN SEA WATER
SPERM SUSPENSION
IN SEA WATER
FERTILIZED EGGS
ADHERED TO SCREEN
ADD SPERM SUSPENSION TO
TRAY CONTAINING EGGS
AERATION
.1
15 MINUTES
FILTERED
SEA WATER (FSW)
HATCHED LARVAE
= 11 DAYS AT 20° C
^SCREEN WITH
EGGS
400/i
SCREENED
DISCHARGE
EGG INCUBATION JAR
CONSTANT
LEVEL SIPHON
LARVAE HARVESTED
WITH NET WITH
PLASTIC FILM
BOTTOM
USE
FOOD: BRINE
SHRIMP NAUPLII
TRANSFER TO
LARVAE
STANDP1PE- SCREENED
LARVAL REARING
TANK- FILLED WITH SEA WATER
CONTINUOUS FLOW
SCREENED
OUTFLOW
Figure 5. Techniques for collection of silverside eggs in the field, and
production of larvae in the laboratory (From Beck, 1979).
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was described in detail by Beck (1979). A similar procedure utilizing a
recirculating system was described by Middaugh and Lempesis (1976).
3.2 Laboratory Year-round Spawning
3.2.1 Atlantic, inland, and tidewater silversides may be spawned in the
laboratory on a year-round basis. Procedures described by Middaugh and Takita
(1983), and Middaugh and Hemmer (1984), provide for maintenance of a brood
stock of 30 to 50 fish, sex ratio 1:1, in 1.3 m diameter, circular holding
tanks which are part of a recirculating seawater system (Figure 6). The
photoperiod should be adjusted to 14 L:10 D (lights on at 5:00 AM and off at
7:00 PM, intensity 10-20 uE/m2/s, or 50-100 ft c), with the water temperature
maintained at 18-20°C for fish from northern latitudes, and 20-25°C for
southern latitudes. Suitable salinities for the culture units would be 25-30
ppt for the Atlantic and tidewater silversides, and 7 °/oo for the inland
silverside. Fish are fed 8 g TetraminR each morning and afternoon, and
concentrated Artemia nauplii (hatch obtained from approximately 15 ml of eggs
after 48 h of incubation at 25°C) in mid-afternoon (see section on Artemia
culture). Excess food should be siphoned from the holding tanks weekly.
Filter media (activated charcoal) located in a reservoir tray should be
changed weekly, immediately after cleaning the holding tanks. To induce
spawning by the Atlantic silverside, the circulation current velocity in the
holding tanks should be reduced to zero (from 8 to 0 cm/sec) twice daily by
turning off the seawater circulation pump from midnight to 1:00 AM, and from
noon to 1:00 PM. Atlantic silversides will spawn in response to interrupted
current velocities during daytime (noon to 1:00 PM). Spawning of the tidewater
silverside also is enhanced by reducing the current velocity twice daily, but
spawns primarily during nighttime. No interruption in current is necessary to
enhance spawning by the inland silverside.
3.2.2 A suitable spawning substrate can be made by cutting enough 25 cm
lengths of No. 18 nylon string to form a small bundle, and tying a string
around the middle of the bundle to form a "mop." The mop is suspended just
below the surface of the water, in contact with the side of the holding tanks.
Spawning fish will deposit eggs on this substrate. The mops are removed from
the holding tanks daily and suspended in incubation vessels. Typical egg
production ranges from 300 to 1200 per spawn. Fish generally can be expected
to spawn three to four days each week.
3.2.3 It is essential that light-tight curtains surround the holding tanks.
These curtains should remain closed except during periodic feedings, tanks
cleaning, and during removal and replacement of spawning substrates.
3.2.4 Embryos attached to nylon screening or nylon string may be suspended in
a culture system such as shown in Figure 6. The culture chambers for embryos
should be constructed of glass. Upon hatching, larvae may be transferred from
the collection container to a 90-cm diameter glass or fiberglass tank with a
volume of 350 L. Tanks receive a continuous flow of seawater at 2 L/min.
Water is introduced at the tank periphery causing a gentle current sufficient
to induce orientation to water movement and normal schooling behavior. Water
is discharged from the tank by two automatic siphons. Siphon openings are
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Figure 6. Holding and spawning system utilized in the culture of silversides
(Menidia). A, 1.3 m diameter tanks; B, circulation pump; C,
reservoir; D, seawater distribution system; E, by-pass line; F,
seawater return line; and G, reservoir filter system. (From
Middaugh and Hemmer, 1984).
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protected by a 400 urn nylon screen to prevent escape of larvae. An inverted
funnel is used at the siphon to decrease the velocity of discharge water, thus
preventing impingement of larvae.
3.2.5 Embryos can also be incubated in small (4-10 L) glass aquaria, by
placing the nylon screening or strings just below the surface of the water.
Gentle aeration should be provided by an airstone positioned near the bottom
of the holding aquaria.
3.3 Culture Media
3.3.1 Use natural seawater if it is available and unpolluted. Otherwise use
synthetic seawater prepared by adding artificial marine salts, such as FORTY
FATHOMS", to deionized water. If synthetic seawater is used, it should be
aged for a least one week before being utilized in culture aquaria.
3.4 Culture Conditions
3.4.1 The salinity maintained during incubation should be similar to that of
the water from which the adults were taken, if collected in the field, or at
which the adults are being maintained in the laboratory, if the embryos
originate from laboratory brood stock. Water temperature should be maintained
at 20 to 25°C depending upon the latitude where fish are collected. Provide a
photoperiod of 12-14 h of illumination daily at 10-20 uE/m2/s, or 50-100 ft-c
(12 h minimum light/24 h). Embryos will hatch in seven to 14 days, depending
upon the incubation temperature and salinity (Middaugh and Lempesis, 1976).
3.5 Feeding and Stocking Density
3.5.1 Upon hatching, Mem'dia larvae should be fed immediately. Newly hatched
brine shrimp (Artemia) nauplii (less than eight hours old) are fed to the
larvae twice daily. It is essential to feed M. mem'dia and M. peninsulae
larvae newly-hatched brine shrimp nauplii (Middaugh et al., 1987).
Utilization of older, larger, brine shrimp nauplii will result in starvation
of the larvae since they are unable to ingest the larger food organisms.
Three to four days after hatching, the fish are able to consume older (larger)
brine shrimp nauplii. Because of their small size M. beryllina larvae must be
fed a mixohaline rotifer, Branchionus plicatilus from day of hatch through day
five. Thereafter, they are able to consume newly-hatched and older Artemia
nauplii (see Middaugh et al., 1987; Weber et al., 1987). Methods for
culturing brine shrimp are discussed in the Appendix. A stocking density of
about 300 larvae is suitable in an 76-L aquarium.
3.6 Culture Maintenance
3.6.1 To avoid excessive build up of algal growths, periodically scrape the
walls of aquaria. Activated charcoal in the aquarium filtration systems
should be changed weekly and detritus (dead brine shrimp nauplii or cysts)
siphoned from the bottom of holding aquaria each week. Salinity may be
maintained at the proper level by addition of distilled or deionized water to
compensate for evaporation.
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4. VIDEO TRAINING TAPE AVAILABLE FOR CULTURING METHODS
4.1 A video training tape and supplemental report (USEPA, 1990) on culturing
inland silversides are available from the National Audiovisual Center,
Customer Services Section, 8700 Edgeworth Drive, Capitol Heights, Maryland,
20743-3701, (Phone 301-763-1891), as part of a video package on short-term
chronic toxicity test methods for marine organisms (Order No. A18545). The
package includes methods for inland silversides, sheepshead minnows, sea
urchins, and Champia, and costs $85.00.
5. TEST ORGANISMS
5.1 Fish 9-14 days old are used in acute toxicity tests (see Section 9).
Most of the water in the holding aquarium should be siphoned off before
removal of larvae. Larvae can then be siphoned from the holding tanks into a
holding vessel. It is essential that larvae not be handled with a dip net,
because it will result in very high mortality. A large-bore, fire-polished
glass tube, 6 mm I.D. x 500 mm long (1/4 in. ID X 18 in. long), equipped with
a rubber squeeze bulb should be used to transfer the larvae from the holding
vessel to the test vessels. It is more convenient to first transfer five fish
to each of several small beakers containing 20 mL of saline dilution water.
The appropriate number of fish (multiples of five) can then be added to test
vessels.
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Stoeckel, J.N. and R.C. Heidinger. 1988. Overwintering of the inland
silverside in southern Illinois. North Amer. J. Fish. Manage.
8:127-131.
Thomson, D.A., and K.A. Muench. 1976. Influence of tides and waves on
the spawning behavior of the Gulf of California grunion, Leuresthes
sardina (Jenkins and Evermann). Southern Calif. Acad. Sci. Bull.
75:198-203.
Thompson, W.F., and J.B. Thompson. 1919. The spawning of the grunion.
Calif. Fish and Game Comm. Bull. 3:1-29.
USEPA. 1981. Nutritional requirements of marine larval and juvenile fish.
Environmental Research Laboratory, K.L. Simpson, P.S. Schauer, C.R.
Seidel, and L.M. Richardson. U. S. Environmental Protection Agency,
Narragansett, Rhode Island. EPA-600/S3-81/049.
USEPA. 1987. Methods for spawning, culturing and conducting toxicity-tests
with early life stages of four atherinid fishes: the inland silverside,
Mem'dia beryllina, Atlantic silverside, M. mem'dia, tidewater silverside,
M. pem'nsulae, and California grunion, Leuresthes tenuis. D.P. Middaugh,
M.J. Hemmer, and L.R. Goodman. Office of Research and Development, U. S.
Environmental Protection Agency, Washington, D.C. 20460.
EPA/600/8-87-004.
USEPA. 1988. Short-term methods for estimating the chronic toxicity of
effluents and receiving waters to marine and estuarine organisms. C.I.
Weber, W.B. Horning, II, D.J. Klemm, T.W. Neiheisel, P.A. Lewis, E.L.
Robinson, J. Menkedick, and F. Kessler. Environmental Monitoring and
Support Laboratory, U. S. Environmental Protection Agency, Cincinnati,
Ohio 45268. EPA-600/4-87/028.
261
-------
USEPA. 1990. Sheepshead minnow and inland silverside larval survival and
growth toxicity tests. Supplemental report for training videotape.
Office of Research and Development, U. S. Environmental Protection Agency,
Washington, D.C. EPA/600/3-90/075.
Walker, B.W. 1949. Periodicity of spawning by the grunion, Leuresthes
tenuis, an atherine fish. Ph.D. Dissertation, Univ. California, Los
Angeles. 166 pp.
Walker, B.W. 1952. A guide to the grunion. Calif. Fish and Game
38:409-420.
Wang, J.C. 1974. Antherinidae - silversides. In: Lippson, A.O., and
R.L. Moran, Manual for identification of early developmental stages of
fishes of the Potomac River estuary, pp. 143-151. Power Plant Siting
Program, Maryland Dept. Nat. Resour., Baltimore, Maryland. PPSP-MP-13. 282
pp.
Wexler, M. 1983. The fish that spawns on land. Nat. Wildl. 21(3):33-36.
Wurtzbaugh, W., and H. Li. 1985. Die! migration of a zooplanktivorous fish
(Menidia beryllina) in relation to the distribution of its prey in a large
eutrophic lake. Limnol. Oceanogr. 30(3):565-576.
Van, H-Y. 1984. Occurrence of spermatozoa and eggs in the gonad of a
tidewater silverside, Menidia beryllina. Copeia 2:544-545.
262
-------
APPENDIX B
SUPPLEMENTAL LIST OF ACUTE TOXICITY TEST SPECIES
TEST ORGANISM
TEST
TEMP
LIFE
STAGE
FRESHWATER SPECIES: VERTEBRATES - WARMWATER
Notropis Teedsi
Lepomis macrochirus
Ictalurus punctatus
Bannerfin shiner 25
Bluegill sunfish 20,25
Channel catfish "
FRESHWATER SPECIES: INVERTEBRATES - COLDWATER
Pteronarcys spp.
Pacifastacus
lem'usculus
Baetis spp.
Ephemerella spp.
Stoneflies*
Crayfish*
Mayflies*
12
1-14 days
larvae
juveniles
nymphs
FRESHWATER SPECIES: INVERTEBRATES - WARMWATER
20,25
HyaTella spp.
Gaimarus lacustris
G. fasciatus
G. pseudolimnaeus
Hexagenia limbata
H. bilineat a
Chironomus spp.
Amphipods
Mayflies
Midges
juveniles
nymphs
larvae
*Stoneflies, crayfish, and mayflies may have to be field
collected and acclimated for a period of time to ensure the
health of the organisms and that stress from collection is
past. Species identification must be verified.
263
-------
SUPPLEMENTAL LIST OF ACUTE TOXICITY TEST SPECIES (CONTINUED)
TEST ORGANISM
TEST
TEMP
SALIN-
LIFE
STAGE
MARINE AND ESTUARINE SPECIES: VERTEBRATES - COLDWATER
English sole 12 32-34
Parophrys vetulus
Citharichys
sitigmaeus Sanddab
Pseudopleuronectes
americanus Winter flounder
MARINE AND ESTUARINE SPECIES: VERTEBRATES - WARMWATER
20,25
Paralichthys
dentatus
P. lethostigma
Fundulus simillis
Fundulus
heteroclitus
Lagodon rhomboides
Orthipristis
chrysoptera
Leostomus xanthurus
Gasterosteus
aculeatus
Flounder
Killifish
Mummichog
Pinfish
Pigfish
Spot
Threespine
stickleback
32-34
20-32
25-32
20-32
15-30
10-30
1-90 days
post meta-
morphosis
1-90 days
1-30 days
1-90 days
20-32 1-30 days
264
-------
SUPPLEMENTAL LIST OF ACUTE TOXICITY TEST SPECIES (CONTINUED)
TEST ORGANISM
TEST
TEMP
SALIN-
LIFE
STAGE
MARINE AND ESTUARINE SPECIES: INVERTEBRATES - COLDWATER
Pandalus jordani
Strongylocentrotus
droebachiensis
Strongylocentrotus
purpuratus
Dendraster
excentricus
Cancer magister
Holmesimysis
costata
Oceanic shrimp
Green sea urchin
Purple sea urchin
Sand dollar
Dungeness crab
Mysid
12
25-32
32-34
MARINE AND ESTUARINE SPECIES: INVERTEBRATES - WARMWATER
Callinectes sapidus
Palaemonetes pugio,
P. vulgaris,
P. intermedius
Penaeus setiferus
Penaeus duorarum
Penaeus aztecus
Crangon
septemspinosa
Nysidopsis almyra
Neomysis americana
Hetamysidopsis
elongata
Crassostrea
virginica
Crassostrea gigas
Arbacia punctulata
Blue crab
Grass shrimp
White shrimp
Pink shrimp
Brown shrimp
Sand shrimp
Mysid
American oyster
Pacific oyster
Purple sea urchin
20,25 10-30
10-32
20-32
25-32
10-32
20-32
25-32
32-34
juvenile
gametes/embryo
ii n
juvenile
1-5 days
juvenile
1-10 days
post-larval
1-5 days
embryo
gametes/embryo
265
-------
APPENDIX C
DILUTOR SYSTEMS
Two proportional dilutor systems are illustrated: the solenoid valve
system, and the vacuum siphon system.
1. Solenoid and Vacuum Siphon Dilutor Systems
The designs of the solenoid and vacuum siphon dilutor systems incorporate
features from devices developed by many other Federal and state programs, and
have been shown to be very versatile for on-site bioassays in mobile
laboratories, as well as in fixed (central) laboratories. The Solenoid Valve
system is fully controlled by solenoids (Figures 1, 2, and 3), and is
preferred over the vacuum siphon system. The Vacuum Siphon system (Figures 1,
4, and 5), however, is acceptable. The dilution water, effluent, and
pre-mixing chambers for both systems are illustrated in Figures 6, 7, and 8.
Both systems employ the same control panel (Figure 9).
If in the range-finding test, the LC50 of the effluent falls in the
concentration range, 6.25% to 100%, pre-mixing is not required. The
pre-mixing chamber is bypassed by running a TYGON" tube directly from the
effluent in-flow pipe to chamber E-2 (see Figures 3 and 5), and Chambers E-l
and D-l and the pre-mixing chamber are deactivated.
The dilutor systems described here can also be used to conduct tests of
the toxicity of pure compounds by equipping the control panel with an
auxiliary power receptacle to operate a metering pump to deliver an aliquot of
the stock solution of the pure compound directly to the mixing chamber during
each cycle. In this case, chamber E-l is de-activated and chamber D-l is
calibrated to deliver a volume of 2000 ml, which is used to dilute the aliquot
to the highest concentration used in the toxicity test.
266
-------
ro
Figure 1. Photographs of the solenoid valve system (left), and the vacuum siphon system (right),
-------
FLOW CONTROL
VALVES
NORMALLY OPEN
SOLENOID VALVES
7 mm (9/32 in.)
DILUTION WATER
INFLOW-*.
EFFLUENT
INFLOW
CYCLE-
COUNTER
DILUTION WATER
CHAMBERS
LAPSE
TIME
CLOCK
MAGNETIC
STIRRER
TEST CHAMBERS 1-20 LITERS CAPACITY
LIDUID LEVEL
SWITCH
DILUTION WATER OVERFLOW
19 mm (3/4 in.) HOLE
12 mm (1/2 In. ) WASTE LINE
EFFLUENT
CHAMBER
PRE-MIXING
CHAMBER
NORMALLY OPEN SOLENOID VALVE
7 mm (9/32 in.) ID
EFFLUENT OVERFLOW
19 mm (3/4 in.) HOLE
12 mm (1/2 in.j LINE
MIXING CHAMBERS
Figure 2. Solenoid valve dilutor system, general diagram (not to scale)
268
-------
EFFLUENT
CHAMBER'
i i
D-l
D-2
n
i i
10 mm OD
ADJUSTABLE
STANDPIPE DRAIN
PRE-MIXING
CHAMBER
MAGNETIC
STIRRER
DILUTION WATER
CHAMBERS
NORMALLY CLOSED
SOLENOID VALVES
El , D2 - D6 = 7 mm (9/32 in. ) ID
Dl = 9.5 mm (3/8 in.) ID
6 mm OD DELIVERY TUBE
NORMALLY OPEN SOLENOID VALVE
7 mm (9/32 in.) ID
6 mm 00 DELIVERY TUBE
EFFLUENT CHAMBERS
NORMALLY CLOSED
SOLENOID VALVE
7 mm (9/32 in.) ID
6 mm 00 DELIVERY TUBE
MIXIN G CHAMBER
1 200 ml CAPACITY
10 mm OD DELIVERY TUBE
TEST CHAMBER
1-20 LITER CAPACITY
NOTE: WHEN 100* EFFLUENT IS USED AS THE HIGHEST EFFLUENT CONCENTRATION,
E-l, D-l, AND THE PRE-MIXING CHAMBER ARE BYPASSED BY CONNECTING A
TYGON TUBE TO THE EFFLUENT INFLOW, AND RUNNING IT DIRECTLY TO E-2.
IN THIS CASE, SOLENOIDS FOR E-l AND D-l, AND THE PRE-MIXING CHAMBER
ARE DISCONNECTED. D-2 + E-3 = 50% EFFLUENT; D-3 + E-4 = 25% EFFFLUENT,
ETC.
Figure 3. Solenoid valve dilutor system, detailed diagram (not to scale),
269
-------
SOLENOID SYSTEM EQUIPMENT LIST
for dilution water and
wall thickness, for dilution water and
1. Diluter Glass.
2. Stainless Steel Solenoid Valves
a. 3, normally open, two-way, 55 psi, water, 1/4" pipe size, 9/32"
orifice size, ASCO 8262152, for incoming effluent and dilution
water pipes and mixing chamber pipe.
b. 1, normally closed, two-way, 15 psi, water, 3/8" pipe size, 3/8"
orifice size, ASCO 8030B65, for D-l chamber evacuation pipe.
c. 12, normally closed, two-way, 36 psi, water, 1/4" pipe size,
9/32" orifice size. ASCO 8262C38, for remaining dilution
chambers (D2 - D6) and effluent chamber (E1-E6) evacuation pipes.
3. Stainless steel tubing, seamless, austenetic, 304 grade for freshwater
and 316 grade for saline water.
a. 10 ft of 3/8" OD, 0.035" wall thickness,
effluent pipes.
b. 60 ft of 1/4" OD, 0.035'
effluent pipes.
c. 1 ft of 3/4" OD, 0.035" wall thickness, for standpipe in Dl
chamber.
4. Swagelok tube connectors, stainless steel.
a. 4, male tube connectors, male pipe size 1/4", tube OD 3/8."
b. 2, male tube connectors, male pipe size 1/2", tube OD 3/8."
c. 26, male tube connectors, male pipe size 1/4, tube OD 1/4."
d. 2, male tube connectors, male pipe size 3/8", tube OD 3/8."
e. 2, male adapter, tube to pipe, male size 1/2", tube OD 3/8."
5. 7, 1200 mL stainless steel beakers.
6. Several Ibs each of Neoprene stoppers, sizes 00, 0, and 1; 1 Ib of
size 5.
7. 14 - aquarium (1-20 liters).
8. Magnetic stirrer.
9. 2 - PVC ball valves, 1/2" pipe size.
10. Dilutor control panel - see Fig. 32 and equipment list.
11. Plywood sheeting, exterior grade: one - 4' x 8' x 3/4", one - 4' >
8' x 1/2".
12. Pine or redwood board, 1" x 8", 20 ft.
13. Epoxy paint, 1 gal.
14. Assorted wood screws, nails, etc.
15. 25 ft - 1/4" ID, TEFLONR tubing, to connect the mixing chambers to
the test chambers.
270
-------
NORMALLY OPEN
SOLENOID VALVES
mm (9/32 in.) ID
V _
L
DILUTION WATER CHAMBERS
^- EFFIUENT CHAM8EB
•*• PRE-MIXING CHAMBER
<
N
/!
)
u
LIOUID LEVEL SWITCH
NORMALLY CLOSED
SOLENOID VALVE
5 mm (3/8 in.) ID
nl
EFFLUENT CHAMBERS
MIXING CHAMBERS
TEST CHAMBERS 1-20 LITERS CAPACITY
Figure 4. Vacuum siphon dilutor system, general diagram (not to scale),
271
-------
0-1
D-2
NORMALLY CLOSED
SOLENOID VALVE
9.5 mm (3/8 in.) ID
DILUTION WATER CHAMBERS
VACUUM LINE
tmm 00 CONNECTING TUBE T FORM
10 Him OD U SHAPE SYPHON TUBE
6mm 0 0 VACUUM LINE TUBE
STAINLESS STEEL HOSE CLAMP
10 Mm ID CONNECTING TUBES Y FORM
10 mm 0 D DELIVERY TUBE
120ml BOTTLE VACUUM BLOCK
IO mm 0 D DELIVERY TUBE
10 mm O.D. DELIVERY TUBE
IO mm O.D AUTOMATIC SYPHON TUBE
EFFLUENT CHAMBERS
O.D U SHAPE SYPHON TUBE
IO mm I D CONNECTING TUBE Y FORM
IO mm 00 DELIVERY TUBE
MIXING CHAMBER 1200 ml CAPACITY
10 mm 0 D DELIVERY TUBE
TEST CHAMBERS CAPACITY
1- 20 LITERS
Figure 5. Vacuum siphon dilutor system, detailed diagram (not to scale),
272
-------
VACUUM SIPHON SYSTEM EQUIPMENT LIST
1. Diluter Glass.
2. Stainless steel solenoid valves.
a. 2, normally open, two-way, 55 psi, water, 1/4" pipe size, 9/32"
orifice size, ASCO 8262152, for incoming effluent and dilution
water pipes.
b. 2, normally closed, two-way, 15 psi, water, 3/8" pipe size, 3/8"
orifice size, ASCO 8030B65, for dilution water chamber D-6 and
effluent chamber E-2.
3. Stainless steel tubing, seamless, austenetic, 304 grade for freshwater
and 316 grade for saline water.
a. 60 ft of 3/8" OD, 0.035" wall thickness, for dilution water and
effluent pipes.
b. 20 ft of 5/16" OD, 0.035" wall thickness, for standpipes in
mixing chambers.
c. 1 ft of 3/4" OD, 0.035" wall thickness, for standpipe in Dl
chamber.
4. Swagelok tube connectors, stainless steel
a. 4, male tube connectors, male pipe size 1/4", tube OD 3/8."
b. 2, male tube connectors, male pipe size 3/8", tube OD 3/8."
c. 2, male adapter, tube to pipe, male pipe size 1/2", tube OD 3/8."
d. 2, male tube connectors, male pipe size 1/2", tube OD 3/8."
5. 7, 1,200 mL stainless steel beakers.
6. Several Ibs each of NEOPRENER stoppers, sizes 00, 0 and 1; 1 Ib of
size 5.
7. 14, aquarium (1-20 liters)
8. Magnetic stirrer.
9. 2, PVC Ball valves, 1/2" pipe size.
10. Dilutor control panel equipment - see Fig. 32 and equipment list.
11. 7, 120 ml NALGENER bottles.
12. 3 ft, l-in-2 aluminum bar, for siphon support brackets.
13. Stainless steel set screws, box of 50, for securing SS tubing in
siphon support brackets.
14. Stainless steel hose clamps, box of 10, size #4 or 5,(need 3 boxes).
15. 6, NALGENER T's, 5/16" OD.
16. 12, TYGONR Y connectors, 3/8" I.D.
17. TYGONR tubing, 3/8" OD, 10 ft.
18. Plywood sheeting, exterior grade: one - 4' x 8x x 3/4", one - 4' x
8' x 1/2".
19. Pine or redwood board, 1" x 8", 20 ft.
20. Epoxy paint, 1 gal.
21. Assorted wood screws, nails, etc.
22. 25 ft of 5/16" ID, TEFLONR tubing, to connect the mixing chambers
to the test chambers.
273
-------
EFFLUENT CHAMBER
DILUTION WATER CHAMBERS
STANDPIPE
DRAIN
OVERFLOH
19 mm (3/4 in.) DIAMETER HOLE
12 mm (1/2 in.) WASTE LINE
C1 C2
19 mm (3/4 fn.) HOLE
12 mm (1/2 In.) LIKE
(FOR VACUUM SIPHON SYSTEM ONLY)
BOTTOM PLATES
C2
00 0000
T
107
i
L..33_H-»J U-38 157 208—268 335-4
DRAIN HOLES IN BOTTOM PLATE (Cl AND C2) SHOWN FOR SOLENOID
VALVE DILUTOR SYSTEM. FOR VACUUM SIPHON DILUTOR SYSTEM,
DRAIN HOLE IS REQUIRED ONLY FOR CHAMBER El.
INDIVIDUAL PART SIZE AND NUMBER OF PIECES USING 6 mm
(1/4 in.) PLATE GLASS. NOTE: 1.6 mm (1/16 in.) No. 304
GRADE (FOR FRESH WATER) OR No. 316 GRADE (FOR SALINE HATER)
STAINLESS STEEL MAY BE SUBSTITUTED FOR GLASS
Cl
C2
Fl
F2:
231 nn X 95 mm
200 mm X 95 mm
107 mm X 107 mm
447 mm X 107 mm
107 mm X 231 mm
447 mm X 231 mm
(15)
4 (END PLATES)
5 (PARTITIONS)
1 (BOTTOM PLATE FOR El)
1 (BOTTOM PLATE FOR D1-D6)
2 (FRONT AND BACK PANELS FOR El)
2 (FRONT AND BACK PANELS FOR Dl-06)
INSIDE CELL MEASUREMENTS AND APPROXIMATE VOLUMES
WIDTH LENGTH HEIGHT VOLUME
El: 95 mm X 95 mm X 231 mm • 2085 mL
2375 mL
760 mL
950 mL
1140 mL
1140 mL
1330 mL
Dl:
02:
D3:
D4:
05:
D6:
125 m
40 m
50 m
60 m
60 m
X 95
X 95
X 95
X 95
X 95
70 mm X 95
m
m
m
m
m
m
X 200
X 200
X 200
X 200
X 200
X 200
Figure 6. Effluent and dilution water chambers (not to scale)
274
-------
EFFLUENT OVERFLOW
19 mm (3/4 1r>.) HOLE
12 ITW (1/2 In.) LINE
BOTTOM PLATE (C)
T
92
r
-» • JB/ *n >•
1* '> 61 &-1S2
o o
0
— »237H
O
O
DRAIN HOLES IN BOTTOM PLATE (C) SHOWN FOR SOLENOID VALVE
DILUTOR SYSTEM ONLY. FOR VACUUM SIPHON DILUTOR SYSTEM,
A DRAIN HOLE IS REQUIRED ONLY FOR CHAMBER E2.
INDIVIDUAL PART SIZE AND NUMBER OF PIECES USING 6 mm (1/4 in.)
PLATE GLASS ARE SHOWN BELOW. NOTE: 1/16 in. No. 304 (FOR FRESH WATER)
OR No. 316 STAINLESS STEEL (FOR SALINE WATER) MAY BE SUBSTITUTED
FOR GLASS.
LENGTH WIDTH
180 mm
155 mm
296 mm
80 mm
80 mm
92 mm
296 mm X 180 mm
NO. PIECES (9)
2 (END PLATES)
4 (PARTITIONS)
t (BOTTOM PLATE)
2 (FRONT AND BACK PLATES)
INSIDE CHAMBER MEASUREMENTS AND APPROXIMATE VOLUMES:
WIDTH LENGTH HEIGHT VOLUME
E2: 110 mm X 80 mm X 155 mm = 1364 mL
E3: 60 mm X 80 mm X 155 mm = 744 mL
E4: 30 mm X 80 mm X 155 mm = 372 mL
E5: 30 mm X 80 mm X 155 mm = 372 mL
E6: 30 mm X 80 mm X 155 mm = 372 mL
Figure 7. Effluent chambers (not to scale),
275
-------
PRE-MIXING CHAMBER
M-3
M-1
240 mm
SIDE VIEW
END VIEW
1 25 mm
M-1
19 mm (3/4 in.) HOLE
12 mm (1/2/in.) LINE
15 mm
1 65 mm
INDIVIDUAL PART SIZE AND NUMBER OF PIECES
USING 6 mm (1/4 in.) PLATE GLASS. APPROXIMATE
CAPACITY 4360 mL.
M-1 125 mm X 153 mm
M-2 125 mm X 153 mm
M-3 240 mm X 165 mm
M-4 240 mm X 125 mm
1 (END PLATE, WITH HOLE)
1 (END PLATE)
1 (BOTTOM PLATE)
2 (SIDE PLATES)
Figure 8. Pre-mixing chamber (not to scale),
276
-------
TDR-I
NOTE: I. 7 DENOTES CHASSIS GROUND
ALL SOLENOIDS
Figure 9. Dilutor control panel wiring diagram.
-------
DILUTOR CONTROL PANEL EQUIPMENT LIST*
Designation
A]
CTR-1
ET
Fl
Jl
J2
J3
Ll
L2
L.S.
Pi
Si
SJ2
SJ3
SJ4 -
TDR-1
TDR-2
CKT Description
Encapsulated amplifier
Cycle counter
Elapsed time indicator
Input power fuse
Recepticle
Aux A.C. output jack
Main input power cord
Fill indicator light
Emptying indicator light
Liquid level sensor
(Dual Sensing Probe)
Plug
On-off main power switch (spst)
On-off aux power switch (spst)
Solenoid
Additional Solenoids for
Solenoid Valve System
Time delay relay
Aux time delay relay
Manufacturer
Cutler Hammer 13535H98C
Redington #P2-1006
Conrac #636W-AA H&T
Little fuse 342038
Amphenol 91PC4F
Stand. 3-prong AC Rcpt.
Stand. 3-prong AC male plug
Dialco 95-0408-09-141
Dialco 95-0408-09-141
Cutler Hammer 13653H2
Amphenol 91MC4M
Cutler Hammer 7580 K7
Cutler Hammer 7580 K7
(See Solenoid and Vacuum
System equipment lists)
Dayton 5x829
Dayton 5x829
*Consult local electric supply house.
278
-------
APPENDIX D
PLANS FOR MOBILE TOXICITY TEST LABORATORY
1. TANDEM-AXLE TRAILER
SPOTLIGHT
f
REGION IV
ATHENS GEORGIA
AIR CONDITIONER
WINDOW
Q
75'
EXTERNAL SIDE VIEW
18 5'
CABINETS
DOOR
1 SINK 1 3£
WALL
DILUTERS
WINDOW
" STANDING CABINETS - _,...._
CABINETS OVER DRAWERS
TOP VIEW
Figure 1. Mobile bioassay laboratory, tandem-axle trailer. Above
external side view; below - internal view from above.
279
-------
L|GHTS
SWITCH'S
OUTSIDE SPOTLIGHT
/^CEILING LIGHTS
WEATHERPROOF SPOTLIGHT
REAR INSIDE
o
LICENSE PLATE REAR OUTSIDE
o
n
.AIR CONDITIONER,
. DRAWERS •
D
FRONT INSIDE
FRONT OUTSIDE
Figure 2. Mobile bioassay laboratory, tandem-axle trailer, external
and internal end views.
280
-------
LEFT SIDE
. ELECTRIC OUTLETS,
1
1
1
1
1
1
m^
i — i — i
g
I Y
-^
'— ' DILUTER BOARD
1 Ir
;<[
AIR CONDITIONER^
STAINLESS STEEL TROUGH STAINLESS
WHEEL WELL
FRONT £>
STEEL TROUGH
-
4 1 24" 1
CABINETS WITH 6" OPENING DUAL WHEEL WELL
SLIDING DOORS WITH
SPRING COVER
ON OUTSIDE
DRAWERS
24"X 16"
RIGHT SIDE
CABINETS 18" X 12"
SLIDING DOORS
t
FRONT
ELECTRIC OUTLET:
2 DRAWERS
18"X 18"
SWITCH'S P.UMP UNDER SINK
CABINET LIGHTS
3 DRAWERS
24"X 16"
DUAL WHEEL WELL
36" HIGH CABINETS
SLIDING DOORS
• 18'-
Figure 3. Mobile bioassay laboratory, tandem-axle trailer, internal
views of side walls.
281
-------
APPENDIX 0
PLANS FOR MOBILE TOXICITY TEST LABORATORY
2. FIFTH WHEEL TRAILER
DILUTOR SYSTEM
UPPER CABINETS
COUNTER TOP
DRAWERS
AIR CONDITIONER
DILUTOR SYSTEM
DOUBLE SHELVES
FOR STATIC TESTS
PLAN A
PLAN B
•125 FEET-
SINK
WATER TANKX
32" DOOR
CABINET
UPPER CABINETS
COUNTER TOP
SINK
*
DRAWERS'^ WATER TANK^
DOOR
r
DOUBLE SHELVES
FOR STATIC TEST
UPPER CABINETS
COUNTER TOP
AIR CONDITIONER^
X
X
DILUTER SYSTEMS
-
5 5 FEET
Figure 4. Mobile bioassay trailer, fifth-wheel trailer, internal
view from above.
282
-------
APPENDIX E
CHECK LISTS AND INFORMATION SHEETS1
1. TOXICITY TEST FIELD EQUIPMENT LIST
Truck
Boards
Cinder blocks
Drums: 500 gal nalgene
55 gal metal - diesel fuel
22 gal
5 gal
Gas can
Jacks
Jumper cables
Oil
Pumps:
_(2) Homelite
_Hoses & couplings
_Shovels
_Spare tires (trailer, generator)
Trailer
Acetone
"Aerators (battery operated)
Air line: Clamps
Aerators (battery operated)
~Air line: Clamps
Stones
Tubing
Valves
_Alcohol
_Aluminum foil
_Alkalinity analysis (0.02 N
Boots: safety
wading
Batteries
"Beakers:
Bottles:
0 cell
^T50 mL nalgene
200 ml glass (3 boxes)
_D.O.
_Uash
_S ample
_VOA vials
500 ml plastic
"Glass organic
Qt. w/teflon liner
_Brine shrimp eggs
"Broom
"Brushes (wash)
_Buckets
"Camera
"Chlorine kit (w/chem)
"Cleanser
_Clip board (Ig, sm)
_Cork borer set
_Culture dishes (200 ml, Daphnia)
Daphnia food
_Data sheets: Bioassay (static)
Bioassay (flow-thru)
Diluter volume delivery
Calibarator delivery sheet
Daily events log
_Dish pan
_Dish rack
_Dissolved oxygen:
~ KCL membrane solution
Membranes
Meter (YSI)
_
_Probes
Reagent:
_
_Alkaline azide
H2S04
"0.0375 Na thiousulfate
Starch
_nied H20
Emergency road kit
"Enamel pans (Ig.sm)
"Erlenmeyer flasks:
500 ml (2)
TOGO mL
"2000 ml
_Extension cords
"Fire extinguishers (2)
_First aid kit
_Fish nets, (Ig.sm)
_Flash light
Generator: Oil
Jilter - fuel
_Funnel
_Grease gun(wheels)
^Credit card
_Lock/key
Siphon hose
^Prepared by the staff of the Environmental Biology Section,
Ecological Support Branch, Environmental Services Division,
U.S. Environmental Protection Agency, Athens, Georgia 30613
283
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TOXICITY TEST FIELD EQUIPMENT LIST (CONT.)
_Glass cutter
_Gloves (plastic)
_Graduated cylinders:
25 mi, 50mL, lOOmL
250mL, 500mL, lOOOmL, 2000ml
_Ground wire & rod
_Hand soap
_Hard hats
_Hardness analysis: Buffer
HC1 (20%
Heaters:
EDTA
indicator
Aquarium
Space
_Hose: Clamps
Connectors
Ice chests
jars: 750 ml (4 boxes)
3 gal (glass) (1)
5 gal (glass) (1)
Sample jugs (2)
_Kimwipes (Ig.sm)
_Lab coats (2)
_Level
_Lignt 110 V
_Log book
_Magnetic stirrers:
Mop
_l_ighted
Other
_Paper towels
_Parachute cord
_Parafilm
_Pencils, pens
_pH: Meters, Orion
_ Meters, corning
Buffers,4,7,10
Probes (extras)
_Pipets: Bulbs
Eyedroppers
Volumetric |
_Plastic bags
_Quality assurance - SPCP
_Rain gear
_Reconstituted hard water
_Refactometer
_Respirators (cartridges)
1 ml, 5 ml, 10 ml)
_Rubber bands
_Ruler
_Safety glasses
_Safety manual
_Sample labels
_Scissors
_Screen bioassay cups
_Sea salts (Instant Ocean)
_Separatory funnels & racks
_Silent giants
_Silicon sealant
_Solenoids (spare)
_stainless steel tubing pieces
_Stanoard Methods Hand Book
_Stirring bars
_Stoppers (assorted)
_Submersible pumps: _ Ig, sm.
screens
_Super ice
_Tablets (paper)
_Tape: _ Cellophane
_ Color coded
_ Electrician
_ Masking
_ Nylon
Therometers: Dial
Glass
_Tools (lock/key)
Jygon tubing, 1/8"
_Volumetric flasks
_WD40
_Weigh boats
_Wire tags
1/4", 3/8"
1000 ml, 2000 ml)
284
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APPENDIX E
CHECK LISTS AND INFORMATION SHEETS
2. INFORMATION CHECK LIST FOR ON-SITE INDUSTRIAL
OR MUNICIPAL WASTE TOXICITY TEST
1. PRE-TRIP INFORMATION
Facility Name:
Address:
Phone number:
Plant Representative(s):
Names, Titles, Addresses of Company Personnel:
A. To Receive Correspondence:
B. To Receive Carbons:
Date of Notification Letter:
State Making Notification and Arrangements:
Special Plant Safety/Security Requirements for EPA Personnel to Observe:
Local Accomodation Recommendations:
Directions to Plant:
Availability of Power Hookups (three 20-amp, 110-V Circuits)
285
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Distance from Power Source to Trailer:
Trailer Location:
(Feet)
Possible Source of Dilution Water:
Major Products:
Raw Materials:
Name of Receiving Water:
Schedule of Plant Operation (continuous, weekdays only, etc.)
Treatment Steps:
Treatment Level (BPT, BAT, etc.):
Wastewater Retention Time by Lagoon or Treatment Step:
Lagoon
Designation
Retention Time
Hours Days
Total Wastewater Retention Time: Hours;_
Retention Time Determination: Calculated;
Calculation method:
_Days
Actual
286
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Description of Wastewater Tap Point:
Description of Outfall (surface, submerged diffuser, etc.):
Description of Other Waste Disposal Alternatives in Use (spray irrigation,
deepwell, municipal discharge, etc.):
2. ON-SITE INFORMATION
Wastewater General Characteristics:
Color:
Odor:
Solids:_
Other:
Serial Number(s) of Discharge(s) to be Tested:
Description of Receiving Water: Uniflow; Tidal;
Approximate amplitude, feet
Color:
Odor:
Solids:
Salinity: High tide ; Low tide
Other:
7Q10: ; Ave. flow
287
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Description of Receiving Water Zone of Dilution:
Location and Description of Water Sampling Point(s):
Fresh:
Salt:
Dilution Waste General Characteristics:
Color:
Odor:
Solids:_
Other:
Description of Toxicity Test Anomalies (plant production changes, power
failure, rain events, etc.):
Duration
Time & Date Time & Date Anomaly
Description of Plant maintenance:
288
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DIAGRAM OF WASTEWATER TREATMENT FACILITIES:
289
-------
3. FOLLOW-UP INFORMATION
Date of follow-up letter:
Wastewater Flow (data supplied by discharger):
Week Prior to Testing Week of Testing
Date Discharge (MGD) Date Discharge (MGD)
Average Discharge (MGD):
Organisms Tested On-site or In-Lab:
Species
Flow-thu
test
duration
(h)
Static
test
duration Test
(h) Location Dates
Results
Possible Recommended Action as a Result of These Findings:
290
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APPENDIX E
CHECK LISTS AND INFORMATION SHEETS
3. DAILY EVENTS LOG
Date: Page of Pages
Site: Day # of Study
Initials:_ Day # of Flow-through Test
Time: Notes:
291
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APPENDIX E
CHECK LISTS AND INFORMATION SHEETS
4. DILUTOR CALIBRATION FORM
Calibration Site:.
Diluter Number:
Calibrator:
Date:"
Effluent
Concentration (%)
Dilution Water (mL)
Trial 1
2
3
Average
Effluent (ml)
Trial 1
2
3
Average
100.0
0
1000
50.0
500
500
25.0
750
250
12.5
875
125
6.25
938
62
3.12
969
31
1.56
984
16
Mixing Chamber (%):.
Waste water (ml):
Dilution Water (ml)
Vol (ml)
Trial 1
2
*
Average
Dilution
Water
Effluent
Remarks:
292
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DILUTOR VOLUME DELIVERY INFORMATION
Industry:
Date:
Test Start Date:
Hie:
CO
Observation
Number
Final
Date
Tine
Volume Delivery to
Metering Boxes
Each 10 Seconds (mL)
Dilution Hater
Hastewater
Measured
Cycle Tine
(min. )
Control Box
Hour
Cycles
Comments
Ave 4-day turnover =
Min/turnover
USEPA
Athens, GA
STANDARD DILUTOR FLOW RATES
Flovi rate (raL/10 seconds)
Cycle Time (min) Dilution Water Wastewater Turnover Tine (rain)
2.5
5.0
7.5
10.0
12.5
15.0 a
532
199
123
89
69
67
246
92
57
41
32
22
43
85
128
170
213
300
o o
> 7^
co
o o -^
70 m
N-4 Z
O Z O
> Tl l-l
r- o x
i— SO
1-1 o
O Z
O X
=n m
m m
o *H
This turnover rate provides approximately five 90% turnovers per day.
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