United States Office of Research and Office of Water
Environmental Protection Development 4305
Agency Washington DC 20460 Washington DC 20460
EPA/600/R-99/064
March 2000
Methods for Measuring the
Toxicity and
Bioaccumulation of
Sediment-associated
Contaminants with
Freshwater Invertebrates
Second Edition
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EPA 600/R-99/064
MARCH 2000
Methods for Measuring the Toxicity and
Bioaccumulation of Sediment-associated
Contaminants with Freshwater
Invertebrates
Second Edition
Office of Research and Development
Mid-Continent Ecology Division
U.S. Environmental Protection Agency
Duluth, Minnesota 55804
Office of Science and Technology
Office of Water
U.S. Environmental Protection Agency
Washington, D.C. 20460
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Disclaimer
This guidance is designed to describe procedures for testing freshwater
organisms in the laboratory to evaluate the potential toxicity or bioaccumulation
of chemicals in whole sediments. This guidance document has no immediate
or direct regulatory consequence. It does not in itself establish or affect legal
rights or obligations, or represent a determination of any party's liability. The
USEPA may change this guidance in the future.
This guidance document has been reviewed in accordance with USEPA Policy
and approved for publication. Mention of trade names or commercial products
does not constitute endorsement or recommendation for use.
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Foreword
Sediment contamination is a widespread environmental problem that can
potentially pose a threat to a variety of aquatic ecosystems. Sediment functions
as a reservoir for common chemicals such as pesticides, herbicides, polychlo-
rinated biphenyls (PCBs), polycyclic aromatic hydrocarbons (PAHs), and
metals such as lead, mercury, and arsenic. In-place contaminated sediment
can result in depauperate benthic communities, while disposal of contaminated
dredged material can potentially exert adverse effects on both pelagic and
benthic systems. Historically, assessment of sediment quality has been limited
to chemical characterizations. The United States Environmental Protection
Agency (USEPA) is developing methodologies to calculate chemical-specific
sediment quality guidelines (referred to as equilibrium partitioning sediment
guidelines or ESGs) for use in the Agency's regulatory programs. However,
quantifying contaminant concentrations alone cannot always provide enough
information to adequately evaluate potential adverse effects that arise from
interactions among chemicals, or that result from time-dependent availability of
sediment-associated contaminants to aquatic organisms. Because relation-
ships between bioavailability and concentrations of chemicals in sediment are
not fully understood, determination of contaminated sediment effects on aquatic
organisms may require the use of controlled toxicity and bioaccumulation tests.
As part of USEPA's Contaminated Sediment Management Strategy, Agency
programs have agreed to use consistent methods to determine whether
sediments have the potential to affect aquatic ecosystems. More than ten
federal statutes provide authority to many USEPA program offices to address
the problem of contaminated sediment. The sediment test methods in this
manual will be used by USEPA to make decisions under a range of statutory
authorities concerning such issues as: dredged material disposal, registration
of pesticides, assessment of new and existing industrial chemicals, Superfund
site assessment, and assessment and cleanup of hazardous waste treatment,
storage, and disposal facilities. The use of uniform sediment testing proce-
dures by USEPA programs is expected to increase data accuracy and preci-
sion, facilitate test replication, increase the comparative value of test results,
and ultimately increase the efficiency of regulatory processes requiring sedi-
ment tests.
This second edition of the manual is a revision to USEPA (1994a; EPA600/R-
94/024). Primary revisions to the first edition of the manual include:
Section 14: This new section describes methods for evaluating sublethal
effects of sediment-associated contaminants with the amphipod Hyalella azteca.
See also associated revisions to Sections 1.3, 2, 4.3, 7.1.3, and 10.3. Section
11 also outlines methods for measuring growth and survival as primary
endpoints in 10-d tests with Hyalella azteca.
Section 15: This new section describes methods for evaluating sublethal
effects of sediment-associated contaminants with the midge Chironomus
tentans. See also associated revisions to Sections 1.3, 2, 4.3, 7.1.3,10.4, and
Appendix C.
Section 2.1.2.1.1: Additional detail has been included on test acceptability
(i.e., control vs. reference sediment).
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Foreword (continued)
Section 6.2.2: The range of acceptable light intensity for culture and testing
has been revised from 500 lux to 1000 lux to 100 to 1000 lux.
Sections 7.2, 8.2, 8.3.2, 8.4.4.7: Additional detail has been added to sections
on formulated sediments, sediment storage, sediment spiking, and interstitial
water sampling.
Sections 9.14,10.3, and 17.4: The requirement to conduct monthly reference-
toxicity tests has been modified to recommend the conduct of reference-
toxicity tests periodically to assess the sensitivity of the test organisms.
Sections 9.14.2 and 17.4.3: These revised sections now state that before
conducting tests with contaminated sediment, it is strongly recommended that
the laboratory conduct the tests with control sediment(s). Results of these
preliminary studies should be used to determine if use of the control sediment
and other test conditions (i.e., water quality) result in acceptable performance
in the tests as outlined in Tables 11.3, 12.3, 13.4, 14.3, and 15.3.
Section 10.3.2: Diatoms are no longer used to culture Hyalella azteca
following procedures of USEPA (1993).
Section 11: In Sectionl 1.2.2 (and associated sections and tables): The
recommended feeding level of 1.5 ml of YCT/day/beaker in the 10-d Hyalella
azteca sediment toxicity test in the first edition of the manual has been revised
to 1.0 ml of YCT/day/beaker. This change was made to make the 10-d test
described in Section 11 consistent with the feeding level recommended in the
42-d test with Hyalella azteca described in Section 14. In Section 11.3:
Additional guidance has been included in the revised manual regarding accli-
mation of test organisms to temperature (see also Section 12.3,13.3,14.3, and
15.3). In Section 11.3.6.1.1: Acceptable concentrations of dissolved oxygen in
overlying water are now expressed in mg/L rather than in a percentage of
saturation. See also Sections 10, 12, 13, 14, and 15.
Sections 12.3.8 and 15.3.8: The recommendation is now made to measure
ash-free dry weight of Chironomus tentans instead of dry weight. See also
Sections 13.3.8 for Lumbriculus variegatus and 14.3.7 for Hyalella azteca.
Section 13.3.7: This section outlines additional guidance on depuration of
Lumbriculus variegatus in bioaccumulation testing.
Section 17.6: This revised section now includes summaries of the results of
round-robin tests using the methods for long-term toxicity tests outlined in
Sections 14 and 15.
Appendix A in the first edition of the manual (USEPA, 1994) was not included in
this edition (summary of a workshop designed to develop consensus for the
10-d toxicity test and bioaccumulation methods). This information has been
cited by reference in this current edition of the manual.
For additional guidance on the technical considerations in the manual, please
contact Teresa Norberg-King, USEPA, Duluth, MN (218/529-5163, fax-5003,
email norberg-king.teresa@epa.gov) or Chris Ingersoll, USGS, Columbia, MO
(573/876-1819, fax-1896, email chris_ingersoll@usgs.gov).
IV
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Abstract
Procedures are described for testing freshwater organisms in the laboratory to
evaluate the potential toxicity or bioaccumulation of chemicals in whole sediments.
Sediments may be collected from the field or spiked with compounds in the
laboratory. Toxicity methods are outlined for two organisms, the amphipod Hyalella
azteca and the midge Chironomus tentans. Toxicity tests with amphipods or midges
are conducted for 10 d in 300-mL chambers containing 100 ml of sediment and
175 ml of overlying water. Overlying water is renewed daily and test organisms are
fed during the toxicity tests. The endpoints in the 10-d toxicity test with H. azteca
and C. tentans are survival and growth. Procedures are primarily described for
testing freshwater sediments; however, estuarine sediments (up to 15%o salinity) can
also be tested in 10-d sediment toxicity tests with H. azteca. Guidance is also
provided for conducting long-term sediment toxicity tests with H. azteca and C. tentans.
The long-term sediment exposures with H. azteca are started with 7- to 8-d-old
amphipods. On Day 28 of the sediment exposure, amphipods are isolated from the
sediment and placed in water-only chambers where reproduction is measured on
Day 35 and 42. Endpoints measured in the amphipod test include survival (Day 28,
35, and 42), growth (on Day 28 and 42), and reproduction (number of young/female
produced from Day 28 to 42). The long-term sediment exposures with C. tentans
start with newly hatched larvae (<24-h old) and continue through emergence,
reproduction, and hatching of the F1 generation (about 60-d sediment exposures).
Survival and growth are determined at 20 d. Starting on Day 23 to the end of the test,
emergence and reproduction of C. tentans are monitored daily. The number of eggs/
female is determined for each egg mass, which is incubated for 6 d to determine
hatching success. The procedures described in Sections 14 and 15 include
measurement of a variety of lethal and sublethal endpoints with Hyalella azteca and
Chironomus tentans; minor modifications of the basic methods can be used in cases
where only a subset of these endpoints is of interest. Guidance for conducting 28-d
bioaccumulation tests with the oligochaete Lumbriculus variegatus is also provided
in the manual. Overlying water is renewed daily and test organisms are not fed
during bioaccumulation tests. Methods are also described for determining
bioaccumulation kinetics of different classes of compounds during 28-d exposures
with L. variegatus.
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VI
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Contents
Foreword iii
Abstract v
Acknowledgments xvii
1 Introduction 1
1.1 Significance of Use 1
1.2 Program Applicability 2
1.3 Scope and Application 4
1.4 Performance-based Criteria 10
2 Summary of Method 11
2.1 Method Description and Experimental Design 11
2.2 Types of Tests 13
2.3 Test Endpoints 13
3 Definitions 14
3.1 Terms 14
4 Interferences 16
4.1 General Introduction 16
4.2 Noncontaminant Factors 17
4.3 Changes in Bioavailability 18
4.4 Presence of Indigenous Organisms 18
5 Health, Safety, and Waste Management 19
5.1 General Precautions 19
5.2 Safety Equipment 19
5.3 General Laboratory and Field Operations 19
5.4 Disease Prevention 20
5.5 Safety Manuals 20
5.6 Pollution Prevention, Waste Management, and Sample Disposal 20
6 Facilities, Equipment, and Supplies 21
6.1 General 21
6.2 Facilities 21
6.3 Equipment and Supplies 21
VII
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Contents (continued)
7 Water, Formulated Sediment, Reagents, and Standards 24
7.1 Water 24
7.2 Formulated Sediment 25
7.3 Reagents 28
7.4 Standards 28
8 Sample Collection, Storage, Manipulation, and Characterization 29
8.1 Collection 29
8.2 Storage 29
8.3 Manipulation 30
8.4 Characterization 31
9 Quality Assurance and Quality Control 33
9.1 Introduction 33
9.2 Performance-based Criteria 33
9.3 Facilities, Equipment, and Test Chambers 33
9.4 Test Organisms 34
9.5 Water 34
9.6 Sample Collection and Storage 34
9.7 Test Conditions 34
9.8 Quality of Test Organisms 34
9.9 Quality of Food 34
9.10 Test Acceptability 34
9.11 Analytical Methods 34
9.12 Calibration and Standardization 34
9.13 Replication and Test Sensitivity 35
9.14 Demonstrating Acceptable Performance 35
9.15 Documenting Ongoing Laboratory Performance 35
9.16 Reference Toxicants 35
9.17 Record Keeping 36
10 Collecting, Culturing, and Maintaining Test Organisms 38
10.1 Life Histories 38
10.2 General Culturing Procedures 40
10.3 Culturing Procedures for Hyalella azteca 41
10.4 Culturing Procedures for Chironomus tentans 42
10.5 Culturing Procedures for Lumbriculus variegatus 46
VIM
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Contents (continued)
11 Test Method 100.1: Hyalella azteca 10-d Survival and Growth Test for Sediments 47
11.1 Introduction 47
11.2 Recommended Test Method for Conducting a 10-d Sediment Toxicity Test
with Hyalella azteca 47
11.3 General Procedures 47
11.4 Interpretation of Results 52
12 Test Method 100.2: Chironomus tentans 10-d Survival and Growth Test for Sediments 55
12.1 Introduction 55
12.2 Recommended Test Method for Conducting a 10-d Sediment Toxicity Test
with Chironomus tentans 55
12.3 General Procedures 55
12.4 Interpretation of Results 60
13 Test Method 100.3: Lumbriculus variegatus Bioaccumulation Test for Sediments 63
13.1 Introduction 63
13.2 Procedure for Conducting Sediment Bioaccumulation Tests with Lumbriculus
variegatus 63
13.3 General Procedures 64
13.4 Interpretation of Results 71
14 Test Method 100.4: Hyalella azteca 42-d Test for Measuring the Effects of Sediment-
associated Contaminants on Survival, Growth, and Reproduction 72
14.1 Introduction 72
14.2 Procedure for Conducting a Hyalella azteca 42-d Test for Measuring the Effects of
Sediment-associated Contaminants on Survival, Growth, and Reproduction 74
14.3 General Procedures 75
14.4 Interpretation of Results 78
15 Test Method 100.5: Life-cycle Test for Measuring the Effects of Sediment-associated
Contaminants on Chironomus tentans 84
15.1 Introduction 84
15.2 Procedure for Conducting a Life-cycle Test for Measuring the Effects of Sediment-
associated Contaminants on Chironomus tentans 84
15.3 General Procedures 87
15.4 Interpretation of Results 92
16 Data Recording, Data Analysis and Calculations, and Reporting 97
16.1 Data Recording 97
16.2 Data Analysis 97
16.3 Data Interpretation 113
16.4 Reporting 114
IX
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Contents (continued)
17 Precision and Accuracy 115
17.1 Determining Precision and Accuracy 115
17.2 Accuracy 115
17.3 Replication and Test Sensitivity 116
17.4 Demonstrating Acceptable Laboratory Performance 116
17.5 Precision of Sediment Toxicity Test Methods: Evaluation of 10-d Sediment
Tests and Reference-toxicity Tests 117
17.6 Precision of Sediment Toxicity Test Methods: Evaluation of Long-term Sediment
Tests 127
18 References 141
Appendices
A. Exposure Systems 157
B. Food Preparation 169
B.1 Yeast, Cerophyl®, and Trout Chow (YCT) for Feeding the Cultures and
Hyalella azteca 169
B.2 Algal Food 170
B.3 Tetrafin® Food (or Other Fish Flake Food) for Culturing and Testing
Chironomus tentans 172
C. Supplies and Equipment for Conducting the Chironomus tentans Long-term
Sediment Toxicity Test 173
C.1 General 173
C.2 Emergence Traps 173
C.3 Reproduction/Oviposit Chambers 173
C.4 Adult Collector Dish 173
C.5 Aspirator 173
C.6 Auxiliary Male Holding Dish 173
C.7 Egg Hatching Chamber 173
C.8 Supplies and Sources 174
C.9 Construction of an Adult Midge Emergence Trap for Use in a "Zumwalt"
Exposure System in Life-cycle Sediment Tests 175
D. Sample Data Sheets 177
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Figures
Figure 10.1 Mean length (+/- 2SD) and relative age of Hyalella azteca collected by sieving
in comparison with length of known-age organisms 43
Figure 10.2 Chironomus tentans larvae 43
Figure 10.3 Aspirator chamber (A) and reproduction and oviposit chamber (B)
for adult midges 45
Figure 11.1 Hyalella azteca 51
Figure 11.2 Lifestage sensitivity of Hyalella azteca in 96-h water-only exposures 53
Figure 11.3 Average recovery of different age Hyalella azteca from sediment
by 7 individuals 54
Figure 12.1 Lifestage sensitivity of chironomids 61
Figure 13.1 Predicted depuration of nonionic organic chemicals from tissue of Lumbriculus
variegatus as a function of Kow and duration of depuration, assuming no
contribution of sediment in the gut 69
Figure 14.1 Relationships between Hyalella azteca length and reproduction 80
Figure 14.2 Relationships between Hyalella azteca dry weight and reproduction 81
Figure 14.3 Relationship between Hyalella azteca length and dry weight 82
Figure 15.1 Relationship between weight and emergence of Chironomus tentans 94
Figure 15.2 Relationship between weight and reproduction of Chironomus tentans 94
Figure 15.3 Relationship between ash-free dry weight (AFDW) and length of Chironomus
tentans 95
Figure 15.4 Relationship between ash-free dry weight (AFDW) and intrinsic rate of natural
increase of Chironomus tentans 95
Figure 16.1 Treatment response fora Type I and Type II error 100
Figure 16.2 Power of the test vs. percent reduction in treatment response relative
to the control mean at various CVs (8 replicates, alpha = 0.05 [one-tailed]) 101
Figure 16.3 Power of the test vs. percent reduction in treatment response relative to the
control mean at various CVs (5 replicates, alpha = 0.05 [one-tailed]) 101
Figure 16.4 Power of the test vs. percent reduction in treatment response relative to the
control mean at various CVs (8 replicates, alpha = 0.10 [one-tailed]) 102
Figure 16.5 Effect of CV and number of replicates on the power to detect a
20% decrease in treatment response relative to the control mean
(alpha = 0.05 [one-tailed]) 102
Figure 16.6 Effect of alpha and beta on the number of replicates at various CVs
(assuming combined alpha + beta = 0.25) 103
Figure 16.7 Decision tree for analysis of survival, growth, and reproduction data subjected
to hypothesis testing 104
Figure 16.8 Decision tree for analysis of point estimate data 108
XI
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Figures (continued)
Figure 17.1 Control (cusum) charts: (A) hypothesis testing results; and (B) point estimates
(LC, EC, or 1C) 116
Figure A.1 Portable table top STIR system described in Benoit etal. (1993) 158
Figure A.2 Portable table top STIR system with several additional options as described in
Benoit etal. (1993) 159
Figure A.3 Tanks forthe STIR system in Benoit et al. (1993) 160
Figure A.4 Water splitting chamber described in Zumwalt et al. (1994) 165
Figure A.5 Diagram of in-line flow splitter used to deliver overlying water in the sediment
exposures of Lumbriculus variegatus (Brunson et al., 1998) 167
Figure C.1 Emergence trap used in the life-cycle Chironomus tentans sediment test 174
Figure C.2 The reproduction/oviposit chamber with the double stack support stand 174
Figure C.3 Adult collection/transfer equipment 175
Figure C.4 Emergence traps that can be used with the Zumwalt water-delivery system
described in Section A.4 176
Figure D.1 Data sheet forthe evaluation of a Chironomus tentans culture 178
Figure D.2 QA/QC data sheet for Chironomus tentans culture 179
Figure D.3 QA/QC data sheets for Chironomus tentans culture 180
Figure D.4 Data sheet for performing reference-toxicity tests 181
Figure D.5 Data sheet for temperature and overlying water chemistry measurements 182
Figure D.6 Data sheet for daily checklist for sediment tests 183
Figure D.7 Data sheet for water quality parameters 184
Figure D.8 Chemistry data sheet 185
Figure D.9 Daily comment data sheet 186
Figure D.10 Weight data sheet 187
Figure D.11 Data sheets for Chironomus tentans tests 188
Figure D.12 Instructions for terminating a Chironomus tentans test 189
Figure D.13 Data sheet forthe Chironomus tentans life-cycle test 190
Figure D.14 Example entries for a Chironomus tentans life-cycle test data sheet 191
Figure D.15 Instructions for completing the Chironomus tentans life-cycle test data sheet.... 192
XII
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Tables
Table 1.1 Sediment Quality Assessment Procedures 3
Table 1.2 Statutory Needs for Sediment Quality Assessment 4
Table 1.3 Rating of Selection Criteria for Freshwater Sediment Toxicity Testing Organisms . 7
Table 1.4 Water-only, 10-d LC50 (u,g/L) Values for Hyalella azteca, Chironomus tentans,
and Lumbriculus variegatus 7
Table 4.1 Advantages and Disadvantages for Use of Sediment Tests 16
Table 6.1 Equipment and Supplies for Culturing and Testing Specific Test Organisms 23
Table 7.1 Characteristics of Three Sources of Clays and Silts
Used in Formulated Sediments 26
Table 7.2 Carbon, Nitrogen, Phosphorus Levels for Various Sources of Organic Carbon.... 26
Table 7.3 Sources of Components Used in Formulated Sediments 27
Table 9.1 Recommended Test Conditions for Conducting Reference-toxicity Tests
with One Organism/Chamber 36
Table 9.2 Recommended Test Conditions for Conducting Reference-toxicity Tests
with More Than One Organism/Chamber 37
Table 10.1 Sources of Starter Cultures of Test Organisms 40
Table 10.2 Chironomus tentans Instar and Head Capsule Widths 43
Table 11.1 Test Conditions for Conducting a 10-d Sediment Toxicity Test
with Hyalella azteca 48
Table 11.2 General Activity Schedule for Conducting a 10-d Sediment Toxicity Test
with Hyalella azteca 48
Table 11.3 Test Acceptability Requirements for a 10-d Sediment Toxicity Test
with Hyalella azteca 49
Table 12.1 Recommended Test Conditions for Conducting a 10-d Sediment Toxicity Test
with Chironomus tentans 56
Table 12.2 General Activity Schedule for Conducting a 10-d Sediment Toxicity Test
with Chironomus tentans 57
Table 12.3 Test Acceptability Requirements for a 10-d Sediment Toxicity Test
with Chironomus tentans 58
Table 13.1 Recommended Test Conditions for Conducting a 28-d Sediment
Bioaccumulation Test with Lumbriculus variegatus 64
Table 13.2 Recommended Test Conditions for Conducting a Preliminary 4-d Sediment
Toxicity Screening Test with Lumbriculus variegatus 65
XIII
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Tables (continued)
Table 13.3 General Activity Schedule for Conducting a 28-d Sediment Bioaccumulation
Test with Lumbriculus variegatus 66
Table 13.4 Test Acceptability Requirements for a 28-d Sediment Bioaccumulation Test
with Lumbriculus variegatus 67
Table 13.5 Grams of Lumbriculus variegatus Tissue (Wet Weight) Required for
Various Analytes at Selected Lower Limits of Detection 70
Table 13.6 Detection Limits (ng) of Individual PAHs by HPLC-FD 70
Table 14.1 Test Conditions for Conducting a 42-d Sediment Toxicity Test
with Hyalella azteca 73
Table 14.2 General Activity Schedule for Conducting a 42-d Sediment Toxicity Test
with Hyalella azteca 74
Table 14.3 Test Acceptability Requirements for a 42-d Sediment Toxicity Test
with Hyalella azteca 75
Table 14.4 Percentage of Paired Tests or Paired Endpoints Identifying Samples as Toxic
in Hyalella azteca 14-d or 28-d Tests 83
Table 15.1 Test Conditions for Conducting a Long-term Sediment Toxicity Test with
Chironomus tentans 85
Table 15.2 General Activity Schedule for Conducting a Long-term Sediment Toxicity Test
with Chironomus tentans 86
Table 15.3 Test Acceptability Requirements for a Long-term Sediment Toxicity Test
with Chironomus tentans 87
Table 15.4 Endpoints for a Long-term Sediment Toxicity Test with Chironomus tentans 88
Table 16.1 Suggested a Levels to Use for Tests of Assumptions 105
Table 16.2 Estimated Time to Obtain 95 Percent of Steady-state Tissue Residue 112
Table 17.1 Intralaboratory Precision for Survival of Hyalella azteca and
Chironomus tentans in 10-d Whole-sediment Toxicity Tests, June 1993 117
Table 17.2 Participants in 1993 Round-robin Studies 118
Table 17.3 Interlaboratory Precision for Hyalella azteca 96-h LC50s from Water-only
Static Acute Toxicity Tests Using a Reference Toxicant (KCI) (October 1992) ..118
Table 17.4 Interlaboratory Precision for Survival of Hyalella azteca in 10-d
Whole-sediment Toxicity Tests Using Four Sediments (March 1993) 119
Table 17.5 Interlaboratory Precision for Chironomus tentans 96-h LC50s from
Water-only Static Acute Toxicity Tests Using a Reference Toxicant
(KCI) (December 1992) 120
Table 17.6 Interlaboratory Precision for Chironomus tentans 96-h LC50s from
Water-only Static Acute Toxicity Tests Using a Reference Toxicant
(KCI) (May 1993)) 120
Table 17.7 Interlaboratory Precision for Survival of Chironomus tentans in 10-d
Whole-sediment Toxicity Tests Using Three Sediments (May 1993) 121
XIV
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Tables (continued)
Table 17.8 Interlaboratory Precision for Growth of Chironomus tentans in 10-d
Whole-sediment Toxicity Tests Using Three Sediments (May 1993) 121
Table 17.9 Interlaboratory Precision for Survival (%) of Hyalella azteca in 10-d
Whole-sediment Toxicity Tests (1996/1997) 123
Table 17.10 Interlaboratory Precision for Survival (%) of Chironomus tentans in 10-d
Whole-sediment Toxicity Tests (1996/1997) 124
Table 17.11 Interlaboratory Precision for Growth (mg/lndividual dry weight) of
Chironomus tentans in 10-d Whole-sediment Toxicity Tests (1996/1997) 125
Table 17.12 Interlaboratory Precision for Growth (mg/lndividual as ash-free dry weight)
of Chironomus tentans in 10-d Whole-sediment Toxicity Tests (1996/1997) 126
Table 17.13 Physical Characteristics of the Sediments Used in the Preliminary
and Definitive Round-robin Evaluations of Long-term Methods for
Sediment Toxicity Testing (Section 17.6) 128
Table 17.14 Percentage of Laboratories Meeting Performance Levels for the Following
Endpoints in the WB Control Sediment Evaluated in the Long-term
Round-robin Tests 128
Table 17.15 Interlaboratory Comparison of Day 28 Percent Survival (Mean ±SD) of
H. azteca in a Long-term Sediment Exposure Using Five Sediments 129
Table 17.16 Interlaboratory Comparison of Day 35 Percent Survival (Mean ±SD) of
H. azteca in a Long-term Sediment Exposure Using Five Sediments 130
Table 17.17 Interlaboratory Comparison of Day 42 Percent Survival (Mean ±SD) of
H. azteca in a Long-term Sediment Exposure Using Five Sediments 131
Table 17.18 Interlaboratory Comparison of Day 28 Length (Mean mm/Individual ±SD)
of/-/, azteca in a Long-term Sediment Exposure Using Five Sediments 132
Table 17.19 Interlaboratory Comparison of Day 28 Dry Weight (Mean mg/lndividual ±SD)
of/-/, azteca in a Long-term Sediment Exposure Using Five Sediments 133
Table 17.20 Interlaboratory Comparison of Reproduction (Mean Number of Young/Female
±SD) of/-/, azteca in a Long-term Sediment Exposure Using Five Sediments ... 134
Table 17.21 Interlaboratory Comparison of Day 20 Percent Survival (Mean ±SD) of
C. tentans in a Long-term Sediment Exposure Using Five Sediments 135
Table 17.22 Interlaboratory Comparison of Dry Weight (Mean mg/lndividual ±SD) of
C. tentans in a Long-term Sediment Exposure Using Five Sediments 136
Table 17.23 Interlaboratory Comparison of Ash-free Dry Weight (Mean mg/lndividual ±SD)
of C. tentans in a Long-term Sediment Exposure Using Five Sediments 137
xv
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Tables (continued)
Table 17.24 Interlaboratory Comparison of Percent Emergence (Mean ±SD) of C. tentans
in a Long-term Sediment Exposure Using Five Sediments 138
Table 17.25 Interlaboratory Comparison of the Number of Eggs/Female (Mean ±SD)
in a Long-term Sediment Exposure Using Five Sediments 139
Table 17.26 Interlaboratory Comparison of Percent Hatch (Mean ±SD) of C. tentans in a
Long-term Sediment Exposure Using Five Sediments 140
Table A.1 Sediment Copper Concentrations and Organism Survival and Growth
at the End of a 10-d Test 163
Table A.2 Sediment Dieldrin Concentrations and Organism Survival and Growth
at the End of a 10-d Test 163
Table A.3 Materials Needed for Constructing a Zumwalt et al. (1994) Delivery System 166
Table B.1 Nutrient Stock Solutions for Maintaining Algal Stock Cultures 170
Table B.2 Final Concentration of Macronutrients and Micronutrients in the
Algal Culture Medium 170
XVI
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Acknowledgments
This document is a general purpose testing manual for freshwater sediments.
This manual is a revision to a previously published edition of this manual
(USEPA, 1994a). The approaches described in this manual were developed
from ASTM (1999a), ASTM (1999b), ASTM (1999c), ASTM (1999d), Ankley et
al. (1993), Phipps et al. (1993), USEPA (1994b), USEPA (1994c), Ingersoll et
al. (1995), Ingersoll et al. (1998), Sibley et al. (1996), Sibley et al. (1997a),
Sibley et al. (1997b), and Benoit et al. (1997).
This second edition of the manual reflects the consensus of the Freshwater
Sediment Toxicity Assessment Committee and the U.S. Environmental Protec-
tion Agency (USEPA) Program Offices. Members of the Freshwater Sediment
Toxicity Assessment Committee for the second edition of this manual included
G.A. Burton, Wright State University, Dayton, OH; T.D. Dawson, Integrated
Laboratory Systems (ILS), Duluth, MN; F.J. Dwyer, U.S. Fish & Wildlife Service
(USFWS), Columbia, MO; C.G. Ingersoll, U.S. Geological Survey (USGS),
Columbia, MO; D.S. Ireland, USEPA, Washington, D.C.; N.E. Kemble, USGS,
Columbia, MO; D.R. Mount, USEPA, Duluth, MN; T.J. Norberg-King, USEPA,
Duluth, MN; P.K. Sibley, University of Guelph, Guelph, Ontario, and Leanne
Stahl, USEPA, Washington, D.C.
The principal authors of the first edition of this manual (USEPA, 1994a)
included C.G. Ingersoll, G.T. Ankley, G.A. Burton, F.J. Dwyer, R.A. Hoke, T.J.
Norberg-King, and P.V. Winger. Principal authors to the second edition of the
manual included C.G. Ingersoll, G.A. Burton, T.D. Dawson, F.J. Dwyer, D.S.
Ireland, N.E. Kemble, D.R. Mount, T.J. Norberg-King, P.K. Sibley, and L. Stahl.
Contributors to specific sections of the manual are:
1. Sections 1-9; General Guidelines
G.T. Ankley, USEPA, Duluth, MN
G.A. Burton, Wright State University, Dayton, OH
T.D. Dawson, ILS, Duluth, MN
F.J. Dwyer, USGS, Columbia, MO
R.A. Hoke, SAIC, Hackensack, NJ
C.G. Ingersoll, USGS, Columbia, MO
D.S. Ireland, USEPA, Washington, DC
N.E. Kemble, USGS, Columbia, MO
D.R. Mount, USEPA, Duluth, MN
T.J. Norberg-King, USEPA, Duluth, MN
C.E. Schlekat, SAIC, Narragansett, Rl
K.J. Scott, SAIC, Narragansett, Rl
L. Stahl, USEPA, Washington, DC
2. Sections 10-15; Culture and Test Methods
G.T. Ankley, USEPA, Duluth, MN
D.A. Benoit, USEPA, Duluth, MN
T.D. Dawson, ILS, Duluth, MN
E.L. Brunson, USGS, Columbia, MO
XVII
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Acknowledgements (continued)
F.J. Dwyer, USGS, Columbia, MO
I.E. Greer, USGS, Columbia, MO
R.A. Hoke, SAIC, Hackensack, NJ
C.G. Ingersoll, USGS, Columbia, MO
N.E. Kemble, USGS, Columbia, MO
P.P. Landrum, NOAA, Ann Arbor, Ml
H. Lee, USEPA, Newport, OR
D.R. Mount, USEPA, Duluth, MN
T.J. Norberg-King, USEPA, Duluth, MN
P.K. Sibley, University of Guelph, Guelph, Ontario
P.V. Winger, USGS, Athens, GA
3. Section 16; Statistical Analysis
J. Heltshe, SAIC, Narragansett, Rl
R.A. Hoke, SAIC, Hackensack, NJ
H. Lee, USEPA, Newport, OR
T.J. Norberg-King, USEPA, Duluth, MN
C. Schlekat, SAIC, Narragansett, Rl
4. Section 17; Precision and Accuracy
G.T. Ankley, USEPA, Duluth, MN
G.A. Burton, Wright State University, Dayton, OH
M.S. Greenburg, Wright State University, Dayton, OH
C.G. Ingersoll, USGS, Columbia, MO
N.E. Kemble, USGS, Columbia, MO
T.J. Norberg-King, USEPA, Duluth, MN
C.D. Rowland, Wright State University, Dayton, OH
P.K. Sibley, University of Guelph, Guelph, Ontario
Review comments from the following individuals are gratefully acknowledged on the
first edition of this manual (USEPA, 1994a): T. Armitage, USEPA, OST, Washing-
ton, D.C.; J. Arthur, R. Spehar, and C. Stephan, USEPA, ORD, Duluth, MN; T.
Bailey, USEPA, OPP, Washington, D.C.; C. Philbrick Barr and P. Nolan, USEPA,
Region 1, Lexington, MA; S. Collyard, T. Dawson, J. Jenson, J. Juenemann, and J.
Thompson, ILS, Duluth, MN; P. Crocker and S. McKinney, USEPA, Region 6,
Dallas, TX; S. Ferraro and R. Swartz, ORD, Newport, OR; L. Cast, TAI, Newtown,
OH; G. Hanson, USEPA, OSW, Washington, D.C.; D. Klemm, EMSL, Newtown,
OH; D. Reed, USEPA, OWM, Washington, D.C.; C. Scheklat and J. Scott, SAIC,
Narragansett, Rl; F. Schmidt, USEPA, OWOW, Washington, D.C.; J. Smrchek,
USEPA, OPPT, Washington, D.C.
Review comments for the following individuals are gratefully acknowledged on the
second edition of the manual: P. Crocker, USEPA, Region 6, Dallas, TX; P. De Lisle,
Coastal Bioanalysts, Gloucester Point, VA; R. Haley, NCASI, Anacortes, WA; R.
Hoke, DuPont, Newark, DE; S. Kroner, USEPA, OSW, Washington, D.C.; P. Landrum,
NOAA, Ann Arbor, Ml; J. Lazorchak, USEPA, ORD, Cincinnati, OH; S. Lin, USEPA,
OWOW, Washington, D.C.; A. Samel, DuPont, Newark, DE; J. Smrchek, USEPA,
OPPT, Washington, D.C.; M. Thompson, USEPA, OST, Washington, D.C.; P.
Winger, USGS, Athens, GA.
XVIII
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Acknowledgements (continued)
Participation by the following laboratories in the round-robin testing is greatly
appreciated for the first edition of the manual (USEPA, 1994a): ABC Laboratories,
Columbia, MO; Environment Canada, Burlington, Ontario; EVS Consultants,
Vancouver, BC; Michigan State University, East Lansing, Ml; National Fisheries
Contaminant Research Center, Athens, GA; Midwest Science Center, Columbia,
MO; Center University of Mississippi, University, MS; University of Wisconsin-
Superior, Superior, WS; USEPA, Cincinnati, OH; USEPA, Duluth, MN; Washington
Department of Ecology, Manchester, WA; Wright State University, Dayton, OH.
Culturing support was supplied for USEPA Duluth by S. Collyard, J. Juenemann, J.
Jenson, and J. Denny.
Participation by the following laboratories in the round-robin testing is greatly
appreciated for this second edition of the manual: Aquatech Biological Sciences,
South Burlington, VT; AScI Environment, Duluth, MN; Arkansas State University,
State University, AR; Bayer, Stillwell, KS; Beak, Brampton, Ontario; Carolina
Ecotox, Durham, NC; Columbia Environmental Research Center, Columbia, MO;
Dell Engineering, Holland, Ml; EA Engineering, Sparks, MD; Great Lakes Research
Center, Traverse City, Ml; Michigan State University, East Lansing, Ml; NCASI,
Anacortes, WA; Patuxent Wildlife Research Center, Athens, GA; SAIC, Naragansett,
Rl; Springborn Laboratories, Wareham, MA; University of Mississippi, Oxford, MS;
Wright State University, Dayton, OH; USEPA, Duluth, MN; USEPA-Region 1,
Lexington, MA; Zeneca, Bracknell, Berks, United Kingdom.
USEPA's Office of Science and Technology provided support for the development of
this manual.
XIX
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XX
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Section 1
Introduction
1.1 Significance of Use
1.1.1 Sediment provides habitat for many aquatic organ-
isms and is a major repository for many of the more
persistent chemicals that are introduced into surface
waters. In the aquatic environment, most anthropogenic
chemicals and waste materials including toxic organic
and inorganic chemicals eventually accumulate in sedi-
ment. Mounting evidence exists of environmental degra-
dation in areas where USEPA Water Quality Criteria
(WQC; Stephan et al., 1985) are not exceeded, yet organ-
isms in or near sediments are adversely affected
(Chapman, 1989). The WQC were developed to protect
organisms in the water column and were not intended to
protect organisms in sediment. Concentrations of chemi-
cals in sediment may be several orders of magnitude
higherthan in the overlying water; however, bulk sediment
concentrations have not been strongly correlated to bio-
availability (Burton, 1991). Partitioning or sorption of a
compound between water and sediment may depend on
many factors, including aqueous solubility, pH, redox,
affinity for sediment organic carbon and dissolved organic
carbon, grain size of the sediment, sediment mineral
constituents (oxides of iron, manganese, and aluminum),
and the quantity of acid volatile sulfides in sediment (Di
Toroetal., 1990, 1991). Although certain chemicals are
highly sorbed to sediment, these compounds may still be
available to the biota. Contaminated sediments may be
directly toxic to aquatic life or can be a source of contami-
nants for bioaccumulation in the food chain.
1.1.2 Assessments of sediment quality have commonly
included sediment chemical analyses and surveys of
benthic community structure. Determination of sediment
chemical concentrations on a dry weight basis alone
offers little insight into predicting adverse biological ef-
fects because bioavailability may be limited by the intri-
cate partitioning factors mentioned above. Likewise,
benthic community surveys may be inadequate because
they sometimes fail to discriminate between effects of
contaminants and those that result from unrelated
non-contaminant factors, including water-quality fluctua-
tions, physical parameters, and biotic interactions. To
obtain a direct measure of sediment toxicity or bioaccu-
mulation, laboratory tests have been developed in which
surrogate organisms are exposed to sediments under
controlled conditions. Sediment toxicity tests have evolved
into effective tools that provide direct, quantifiable evi-
dence of biological consequences of sediment
contamination that can only be inferred from chemical or
benthic community analyses. To evaluate sediment qual-
ity nationwide, USEPA developed the National Sediment
Inventory (NSI), which is a compilation of existing sedi-
ment quality data and protocols used to evaluate the data.
The NSI was used to produce the first biennial report to
Congress on sediment quality in the United States as
required under the Water Resources Development Act of
1992 (USEPA, 1997a; 1997b; 1997c). USEPA's evalua-
tion of the data shows that sediment contamination exists
in every region and state of the country and various
waters throughout the United States contain sediment
that is sufficiently contaminated with toxic pollutants to
pose potential risks to fish and to humans and wildlife who
eat fish. The use of consistent sediment testing methods
described in this manual will provide high quality data
needed for the NSI, future reports to Congress, and
regulatory programs to prevent, remediate, and manage
contaminated sediments (USEPA, 1998).
1.1.3 The objective of a sediment test is to determine
whether chemicals in sediment are harmful to or are
bioaccumulated by benthic organisms. The tests can be
used to measure interactive toxic effects of complex
chemical mixtures in sediment. Furthermore, knowledge
of specific pathways of interactions among sediments
and test organisms is not necessary to conduct the tests
(Kemp and Swartz, 1988). Sediment tests can be used to
(1) determine the relationship between toxic effects and
bioavailability; (2) investigate interactions among chemi-
cals; (3) compare the sensitivities of different organisms;
(4) determine spatial and temporal distribution of contami-
nation; (5) evaluate dredged material; (6) measure toxicity
as part of product licensing or safety testing or chemical
approval; (7) rank areas for cleanup, and (8) set cleanup
goals and estimate the effectiveness of remediation or
management practices.
1.1.4 A variety of standard methods have been developed
for assessing the toxicity of contaminants associated with
sediments using amphipods, midges, polychaetes, oli-
gochaetes, mayflies, or cladocerans (i.e., ASTM,1999a;
ASTM,1999b; ASTM, 1999c; ASTM, 1999d; USEPA,
1994a; USEPA, 1994b; Environment Canada, 1997a;
Environment Canada, 1997b). Several endpoints are
suggested in these methods to measure effects of con-
taminants in sediment including survival, growth, behavior,
or reproduction; however, survival of test organisms in
1
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10-d exposures is the endpoint most commonly reported.
These short-term exposures which only measure effects
on survival can be used to identify high levels of contami-
nation, but may not be able to identify moderately contami-
nated sediments (Sibley et al., 1996; Sibley et al., 1997a;
Sibley et al., 1998; Benoit et al., 1997; Ingersoll et al.,
1998). Sublethal endpoints in sediment tests may also
prove to be better estimates of responses of benthic
communities to contaminants in the field (Kemble et al.,
1994) The first edition of this manual (USEPA, 1994a)
described 10-d toxicity tests with the amphipod Hyalella
azteca and midge Chironomus tentans (Section 11,12).
Thissecond edition of the manual now outlines approaches
for evaluating sublethal endpoints in longer-term sediment
exposures with these two species (Section 14,15). Guid-
ance is also presented in Section 13 regarding sediment
bioaccumulation testing with the oligochaete Lumbriculus
variegatus.
1.1.5 Results of toxicity tests on sediments spiked at
different concentrations of chemicals can be used to
establish cause and effect relationships between chemi-
cals and biological responses. Results of toxicity tests
with test materials spiked into sediments at different
concentrations may be reported in terms of an LC50
(median lethal concentration), an EC50 (median effect
concentration), an IC50 (inhibition concentration), or as a
NOEC (no observed effect concentration) or LOEC (low-
est observed effect concentration). In some cases, re-
sults of bioaccumulation tests may also be reported in
terms of a Biota-sediment Accumulation Factor (BSAF)
(Ankleyetal., 1992a; Ankley et al., 1992b).
1.1.6 Evaluating effect concentrations for chemicals in
sediment requires knowledge of factors controlling their
bioavailability. Similar concentrations of a chemical in
units of mass of chemical per mass of sediment dry
weight often exhibit a range in toxicity in different sedi-
ments (Di Toro et al., 1990; Di Toro et al., 1991). Effect
concentrations of chemicals in sediment have been corre-
lated to interstitial water concentrations, and effect con-
centrations in interstitial water are often similar to effect
concentrations in water-only exposures. The bioavailabil-
ity of nonionic organic compounds in sediment is often
inversely correlated with the organic carbon concentra-
tion. Whatever the route of exposure, these correlations
of effect concentrations to interstitial water concentra-
tions indicate that predicted or measured concentrations
in interstitial water can be used to quantify the exposure
concentration to an organism. Therefore, information on
partitioning of chemicals between solid and liquid phases
of sediment is useful for establishing effect concentra-
tions (Di Toro etal., 1991).
1.1.7 Field surveys can be designed to provide either a
qualitative reconnaissance of the distribution of sediment
contamination or a quantitative statistical comparison of
contamination among sites. Surveys of sediment toxicity
or bioaccumulation are usually part of more comprehen-
sive analyses of biological, chemical, geological, and
hydrographic data. Statistical correlations may be im-
proved and sampling costs may be reduced if subsamples
are taken simultaneously for sediment tests, chemical
analyses, and benthic community structure.
1.1.8 Table 1.1 lists several approaches the USEPA has
considered for the assessment of sediment quality
(USEPA, 1992c). These approaches include (1) equilibrium
partitioning, (2) tissue residues, (3) interstitial watertoxicity,
(4) benthic community structure, (5) whole-sediment toxic-
ity and sediment-spiking tests, (6) Sediment Quality Triad,
and (7) sediment quality guidelines (see Chapman, 1989
and USEPA, 1989a; USEPA, 1990a; USEPA, 1990b;
USEPA, 1992b for a critique of these methods). The
sediment assessment approaches listed in Table 1.1 can
be classified as numeric (e.g., equilibrium partitioning),
descriptive (e.g., whole-sediment toxicity tests), or a
combination of numeric and descriptive approaches (e.g.,
Effect Range Median; USEPA, 1992c). Numeric methods
can be used to derive chemical-specific equilibrium parti-
tioning sediment guidelines (ESGs) or other sediment
quality guidelines (SQGs). Descriptive methods such as
toxicity tests with field-collected sediment cannot be
used alone to develop numerical ESGs or other SQGs for
individual chemicals. Although each approach can be
used to make site-specific decisions, no one single ap-
proach can adequately address sediment quality. Overall,
an integration of several methods using the weight of
evidence is the most desirable approach for assessing
the effects of contaminants associated with sediment
(Long and Morgan, 1990; MacDonald etal., 1996; Ingersoll
et al., 1996; 1997). Hazard evaluations integrating data
from laboratory exposures, chemical analyses, and benthic
community assessments provide strong complementary
evidence of the degree of pollution-induced degradation in
aquatic communities (Chapman etal., 1992; Chapman et
al., 1997; Burton, 1991).
1.2 Program Applicability
1.2.1 The USEPA has authority under a variety of
statutes to manage contaminated sediments (Table 1.2
and USEPA, 1990e). USEPA's Contaminated Sediment
Management Strategy (USEPA, 1998) establishes the
following four goals for contaminated sediments and de-
scribes actions that the Agency intends to take to accom-
plish these goals: (1) to prevent further contamination of
sediments that may cause unacceptable ecological or
human health risks; (2) when practical, to clean up exist-
ing sediment contamination that adversely affects the
Nation's waterbodies or their uses, or that causes other
significant effects on human health or the environment;
(3) to ensure that sediment dredging and the disposal of
dredged material continue to be managed in an environ-
mentally sound manner; and (4) to develop and consis-
tently apply methodologies for analyzing contaminated
sediments. The Agency plans to employ its pollution
prevention and source control programs to address the
first goal. To accomplish the second goal, USEPA will
consider a range of risk management alternatives to
reduce the volume and effects of existing contaminated
sediments, including in-situ containment and contaminated
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Table 1.1 Sediment Quality Assessment Procedures1
Type
Method
Numeric Descriptive Combination
Approach
Equilibrium Partitioning
Tissue Residues
Interstitial Water Toxicity
Benthic Community
Structure
Whole-sediment Toxicity
and Sediment Spiking
Sediment Quality Triad
Sediment Quality Guidelines
A sediment quality value for a given contaminant is determined by
calculating the sediment concentration of the contaminant that
corresponds to an interstitial water concentration equivalent to the
USEPA water-quality criterion for the chemical.
Safe sediment concentrations of specific chemicals are established
by determining the sediment chemical concentration that results in
acceptable tissue residues.
Toxicity of interstitial water is quantified and identification
evaluation procedures are applied to identify and quantify chemical
components responsible for sediment toxicity.
Environmental degradation is measured by evaluating alterations
in benthic community structure.
Test organisms are exposed to sediments that may contain
known or unknown quantities of potentially toxic chemicals. At the
end of a specified time period, the response of the test organisms
is examined in relation to a specified endpoint. Dose-response
relationships can be established by exposing test organisms to
sediments that have been spiked with known amounts of chemicals
or mixtures of chemicals.
Sediment chemical contamination, sediment toxicity, and benthic
community structure are measured on the same sediment sample.
Correspondence between sediment chemistry, toxicity, and field
effects is used to determine sediment concentrations that
discriminate conditions of minimal, uncertain, and major biological
effects.
The sediment concentration of contaminanants associated with
toxic responses measured in laboratory exposures or in field
assessments (i.e., Apparent Effect Threshold (AET), Effect Range
Median (ERM), Probable Effect Level (PEL)).
Modified from USEPA (1992c)
sediment removal. Finally, the Agency is developing
tools for use in pollution prevention, source control,
remediation, and dredged material management to meet
the collective goals. These tools include national invento-
ries of sediment quality and environmental releases of
contaminants, numerical assessment guidelines to evalu-
ate contaminant concentrations, and standardized bioas-
says to evaluate the bioaccumulation and toxicity poten-
tial of sediment samples.
1.2.2 The Clean Water Act (CWA) is the single most
important law dealing with environmental quality of sur-
face waters in the United States. The objective of the
CWA is to restore and maintain the chemical, physical,
and biological integrity of the Nation's waters (CWA,
Section 101). Federal and state monitoring programs
traditionally have focused on evaluating water column
problems caused by point source dischargers. Findings
in the National Sediment Quality Survey, Volume I of the
first biennial report to Congress on sediment quality in the
U.S., indicate that this focus needs to be expanded to
include sediment quality impacts (Section 1.1.2 and
USEPA, 1997a).
1.2.3 The Office of Water (OW), the Office of Preven-
tion, Pesticides, and Toxic Substances (OPPTS), the
Office of Solid Waste (OSW), and the Office of Emer-
gency and Remedial Response (OERR) are all committed
to the principle of consistent tiered testing described in
the Contaminated Sediment Management Strategy
(USEPA, 1998). Agency-wide consistent testing is desir-
able because all USEPA programs will use standard
methods to evaluate health risk and produce comparable
data. It will also provide the basis for uniform cross-
program decision-making within the USEPA. Each pro-
gram will, however, retain the flexibility of deciding whether
identified risks would trigger regulatory actions.
1.2.4 Tiered testing refers to a structured, hierarchical
procedure for determining data needs relative to decision-
making that consists of a series of tiers, or levels, of
investigative intensity. Typically, increasing tiers in a
tiered testing framework involve increased information
and decreased uncertainty (USEPA, 1998). Each EPA
program office intends to develop guidance for interpret-
ing the tests conducted within the tiered framework and to
explain how information within each tier would trigger
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Law2
Table 1.2 Statutory Needs for Sediment Quality Assessment1
Area of Need
CERCLA • Assessment of need for remedial action with contaminated sediments; assessment of degree of cleanup required,
disposition of sediments
CWA • National Pollutant Discharge Elimination System (NPDES) permitting, especially under Best Available Technology
(BAT) in water-quality-limited water
Section 403(c) criteria for ocean discharges; mandatory additional requirements to protect marine environment
Section 301 (g) waivers for publicly owned treatment works (POTWs) discharging to marine waters
Section 404 permits for dredge and fill activities (administered by the U.S. Army Corps of Engineers [USAGE])
FIFRA • Reviews of uses for new and existing chemicals
Pesticide labeling and registration
MPRSA • Permits for ocean dumping
NEPA • Preparation of environmental impact statements for projects with surface water discharges
TSCA • Section 5: Premanufacture notification reviews for new industrial chemicals
Sections 4, 6, and 8: Reviews for existing industrial chemicals
RCRA • Assessment of suitability (and permitting of) on-land disposal or beneficial use of contaminated sediments considered
"hazardous"
1 Modified from Dickson et al., 1987 and Southerland et al., 1992.
2 CERCLA Comprehensive Environmental Response, Compensation and Liability Act (Superfund).
CWA Clean Water Act.
FIFRA Federal Insecticide, Fungicide, and Rodenticide Act.
MPRSA Marine Protection, Resources and Sanctuary Act.
NEPA National Environmental Policy Act.
TSCA Toxic Substances Control Act.
RCRA Resource Conservation and Recovery Act.
regulatory action. Depending on statutory and regulatory
requirements, the program specific guidance will describe
decisions based on a weight of evidence approach, a
pass-fail approach, or comparison to a reference site.
The following two approaches are currently being used by
USEPA: (1) the Office of Water-U.S. Army Corps of
Engineers dredged material testing framework and (2) the
OPPTS ecological risk assessment tiered testing frame-
work. USEPA-USACE (1998a) describes the dredged
material testing framework and Smrchek and Zeeman
(1998) summarizes the OPPTS testing framework. A
tiered testing framework has not yet been chosen for
Agency-wide use, but some of the components have
been identified to be standardized. These components
include toxicity tests, bioaccumulation tests, sediment
quality guidelines, and other measurements that may
have ecological significance, including benthic commu-
nity structure evaluation, colonization rate, and in situ
sediment testing within a mesocosm (USEPA, 1992a).
1.3 Scope and Application
1.3.1 A variety of standard methods have been previously
developed for assessing the toxicity of chemicals in
sediments using amphipods, midges, polychaetes, oli-
gochaetes, mayflies, or cladocerans (USEPA, 1994a;
USEPA, 1994b; ASTM, 1999a; ASTM, 1999b; ASTM;
1999c; ASTM, 1999d; Environment Canada, 1997a; Envi-
ronment Canada, 1997b). Several endpoints are suggested
in these methods to measure effects of chemicals in
sediment including survival, growth, behavior, or repro-
duction; however, survival of test organ isms in 10-d expo-
sures is the endpoint most commonly reported. These
short-term exposures which only measure effects on
survival can be used to identify high levels of contamina-
tion, but might not be able to identify moderate levels of
contamination in sediments (Benoit et al., 1997; Ingersoll
et al., 1998; Sibley et al., 1996; Sibley et al., 1997a;
Sibley et al., 1997b; Sibley et al., 1998).
1.3.2 Procedures described in Sections 11 and 12 for
conducting 10-d sediment toxicity tests with the amphi-
pod H. azteca (measuring survival) and the midge
C. tentans (measuring survival and growth) were de-
scribed in the first edition of the manual (USEPA, 1994a).
Section 14 of this second edition of the manual now
describes a method for determining potential sublethal
effects of contaminants associated with sediment on
H. azteca, including effects on reproduction based on a
procedure described by Ingersoll et al. (1998). Section 15
of this second edition of the manual now describes a
method for determining sublethal endpoints in sediment
tests based on a life-cycle test with C. tentans described
by Benoit et al. (1997), Sibley et al. (1996), and Sibley et
al. (1997a). Procedures are primarily described fortesting
freshwater sediments; however, estuarine sediments (up
to 15%o salinity) can also be tested in 10-d sediment tests
with H. azteca.
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1.3.2.1 The decision to conduct 10-d or long-term toxicity
tests with H. azteca or C. tentans depends on the goal of
the assessment. In some instances, sufficient informa-
tion may be gained by measuring sublethal endpoints in
10-d tests. In other instances, the 10-d tests could be
used to screen samples for toxicity before long-term tests
are conducted. While the long-term tests are needed to
determine direct effects on reproduction, measurement of
growth in these toxicity tests may serve as an indirect
estimate of reproductive effects of chemicals associated
with sediments (Section 14.4.5 and 15.4.6.2). Additional
studies are ongoing to more thoroughly evaluate the
relative sensitivity between lethal and sublethal endpoints
measured in 10-d tests and between sublethal endpoints
measured in the long-term tests. Results of these studies
and additional applications of the methods described in
Sections 14 and 15 will provide data that can be used to
assist in determining where application of long-term tests
will be most appropriate.
1.3.2.2 Use of sublethal endpoints for assessment of
contaminant risk is not unique to toxicity testing with
sediments. Numerous regulatory programs require the use
of sublethal endpoints in the decision-making process
(Pittinger and Adams, 1997) including: (1) Water Quality
Criteria (and State Standards); (2) National Pollution Dis-
charge Elimination System (NPDES) effluent monitoring
(including chemical-specific limits and sublethal endpoints
in toxicity tests); (3) Federal Insecticide, Rodenticide and
Fungicide Act (FIFRA) and the Toxic Substances Control
Act (TSCA; tiered assessment includes several sublethal
endpoints with fish and aquatic invertebrates); (4) Super-
fund Comprehensive Environmental Response, Compen-
sation and Liability Act (CERCLA); (5) Organization of
Economic Cooperation and Development (OECD; suble-
thal toxicity testing with fish and invertebrates); (6) Euro-
pean EconomicCommunity (EC; sublethal toxicity testing
with fish and invertebrates); and (7) the Paris Commission
(behavioral endpoints).
1.3.3 Guidance for conducting 28-d bioaccumulation
tests with the oligochaete Lumbriculus variegatus is also
provided in this manual (Section 13). Overlying water is
renewed daily and organisms are not fed during bioaccu-
mulation tests. Methods are also described for
determining bioaccumulation kinetics of different classes
of compounds during 28-d exposures with L. variegatus.
1.3.4 Additional research and methods development are
now in progress to (1) refine sediment Toxicity Identifica-
tion Evaluation (TIE) procedures (Ankley and Thomas,
1992), (2) refine sediment spiking procedures, (3) develop
in situ toxicity tests to assess sediment toxicity and
bioaccumulation under field conditions, (4) evaluate rela-
tive sensitivity of endpoints measured in toxicity tests,
(5) develop methods for additional species, (6) evaluate
relationships between toxicity and bioaccumulation, and
(7) produce additional data on confirmation of responses
in laboratory tests with natural populations of benthic
organisms. This information will be described in future
editions of this manual or other USEPA manuals.
1.3.4.1 This methods manual serves as a companion to
the marine sediment testing method manuals (USEPA,
1994b; USEPA, 1999).
1.3.5 Procedures described in this manual are based on
the following documents: ASTM (1999a), ASTM (1999b),
ASTM (1999c), ASTM (1999d), Ankley et al. (1993),
Phipps et al. (1993), Call et al. (1994), USEPA (1991 a),
USEPA (1994a), USEPA (1994b), Ingersoll et al. (1995),
Ingersoll et al. (1998), Sibley et al. (1996), Sibley et al.
(1997a), Sibley et al. (1997b), and Benoit et al. (1997).
This manual outlines specific test methods for evaluating
the toxicity of sediments in 10-d exposures with H. azteca
and C. tentans. The manual also outlines general guid-
ance on procedures for evaluating the effects of sediment
contaminants in long-term exposures with H. azteca and
C. tentans and bioaccumulation of contaminants in
sediment with L. variegatus. Some issues that may be
considered in interpretation of test results are the subject
of continuing research, including the influence of feeding
on bioavailability, nutritional requirements of the test or-
ganisms, additional performance criteria for organism
health, and confirmation of responses in laboratory tests
with natural benthic populations. As additional research is
completed on these and other test species, the results
will be incorporated into future editions of this manual.
See Section 4 for additional details.
1.3.6 General procedures described in this manual might
be useful for conducting tests with other aquatic organ-
isms; however, modifications may be necessary. Altering
the procedures described in this manual may alter bio-
availability and produce results that are not directly com-
parable with results of acceptable procedures. Compari-
son of results obtained using modified versions of these
procedures might provide useful information concerning
new concepts and procedures for conducting sediment
tests with aquatic organisms (e.g., Diporeiaspp., Tubifex
tubifex, Hexagenia spp.). If tests are conducted with
procedures different from those described in this manual,
additional tests are required to determine comparability of
results.
1.3.6.1 Methods have been described for culturing and
testing indigenous species that may be as sensitive or
more sensitive than the species recommended in this
manual. However, the USEPA currently allows the use of
indigenous species only where state regulations require
their use or prohibit importation of the recommended
species. Where state regulations prohibit importation or
use of the recommended test species, permission should
be requested from the appropriate regulatory agency be-
fore using indigenous species.
1.3.6.2 Where states have developed culturing and test-
ing methods for indigenous species other than those
recommended in this manual, data comparing the sensi-
tivity of the substitute species and one or more of the
recommended species must be obtained with sediments
or reference toxicants to ensure that the species selected
are at least as sensitive and appropriate as the recom-
mended species.
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1.3.7 Selection of Test Organisms
1.3.7.1 The choice of a test organism has a major
influence on the relevance, success, and interpretation of
a test. Test organism selection should be based on both
environmental relevance and practical concerns (DeWitt
et al., 1989; Swartz, 1989). Ideally, a test organism
should (1) have a toxicological database demonstrating
relative sensitivity and discrimination to a range of chemi-
cals of concern in sediment; (2) have a database for
interlaboratory comparisons of procedures (e.g., round-robin
studies); (3) be in contact with sediment (e.g., water
column vs. benthic organism); (4) be readily available
through culture or from field collection; (5) be easily
maintained in the laboratory; (6) be easily identified; (7) be
ecologically or economically important; (8) have a broad
geographical distribution, be indigenous (either present or
historical) to the site being evaluated, or have a niche
similarto organisms of concern (e.g., similar feeding guild
or behavior to the indigenous organisms); (9) be tolerant
of a broad range of sediment physico-chemical character-
istics (e.g., grain size); and (10) be compatible with
selected exposure methods and endpoints (Table 1.3,
ASTM, 1998d). The method should also be (11) peer
reviewed (e.g., journal articles, ASTM guides) and (12)
confirmed with responses with natural populations of
benthic organisms (Sections 1.3.7.9 and 1.3.8.5).
1.3.7.2 Of these criteria (Table 1.3), a database demon-
strating relative sensitivity to chemicals, contact with
sediment, ease of culture in the laboratory, interlaboratory
comparisons, tolerance to varying sediment physico-
chemical characteristics, and confirmation with responses
of natural benthic populations were the primary criteria
used for selecting H. azteca, C. tentans, and L. variegatus
for the current edition of this manual. Many organisms
that might be appropriate for sediment testing do not now
meet these selection criteria because historically little
emphasis has been placed on developing standardized
testing procedures for benthic organisms. A similar data-
base must be developed in order for other organisms to be
included in future editions of this manual (e.g., mayflies
[Hexagen/aspp.], other midges [C. riparius], other amphi-
pods [Diporeia spp.], cladocerans [Daphnia magna,
Ceriodaphnia dubia], or mollusks).
1.3.7.3 An important consideration in the selection of
specific species for test method development is the
existence of information concerning relative sensitivity of
the organisms both to single chemicals and complex
mixtures. A number of studies have evaluated the sensi-
tivity of H. azteca, C. tentans and L. variegatus, relative
to one another, as well as other commonly tested fresh-
water species. For example, Ankley et al. (1991 b) found
H. azteca to be as, or slightly more, sensitive than
Ceriodaphnia dubia to a variety of sediment elutriate and
pore-water samples. In that study, L. variegatus were less
sensitive to the samples than either the amphipod or the
cladoceran. West et al. (1993) found the rank sensitivity
of the three species to the lethal effects of copper in
sediments could be ranked (from greatest to least): H.
azteca > C. tentans > L. variegatus. In short-term (48 to
96 h) exposures, L. variegatus generally was less sensi-
tive than H. azteca, C. dubia, or Pimephales promelas
to cadmium, nickel, zinc, copper, and lead
(Schubauer-Berigan et al., 1993). Of the latter three
species, no one was consistently the most sensitive to all
five metals.
1.3.7.3.1 In a study of Great Lakes sediment, H. azteca,
C. tentans, and C. riparius were among the most sensitive
and discriminatory of 24 organisms tested (Burton and
Ingersoll, 1994; Burton et al., 1996a; Ingersoll et al.,
1993). Kemble et al. (1994) found the rank sensitivity of
fourspeciesto metal-contaminated sediments to be (from
greatest to least): H. azteca > C. riparius > Oncorhynchus
mykiss (rainbow trout) > Daphnia magna. The relative
sensitivity of the three endpoints evaluated in the H. azteca
test with Clark Fork River sediments was (from greatest
to least): length > sexual maturation > survival.
1.3.7.3.2 In 10-d water-only and whole-sediment tests, H.
azteca and C. tentans were more sensitive than D. magna
to fluoranthene (Suedel et al., 1993).
1.3.7.3.3 Water-only tests also have been conducted for
10 d with a number of chemicals using the three species
described in this manual (Phippset al., 1995; Table 1.4).
All tests were flow-through exposures using a soft natural
water (Lake Superior) with measured chemical concentra-
tions that, other than the absence of sediment, were
conducted under conditions (e.g..temperature, photope-
riod, feeding) similar to those being described for the
standard 10-d sediment test. In general, H. azteca was
more sensitive to copper, zinc, cadmium, nickel and lead
than either C. tentans or L. variegatus. Chironomus ten-
tans and H. azteca exhibited a similar sensitivity to
several of the pesticides tested. Lumbriculus variegatus
was not tested with several of the pesticides; however, in
other studies with whole sediments contaminated by DDT
and associated metabolites, and in short-term (96-h)
experiments with organophosphate insecticides (diazinon,
chlorpyrifos), L. variegatus has proven to be far less
sensitive than either H. azteca or C. tentans. These
results highlight two important points germane to the
methods in this manual. First, neither of the two test
species selected for estimating sediment toxicity
(H. azteca, C. tentans) was consistently more sensitive
to all chemicals, indicating the importance of using mul-
tiple test organisms when performing sediment assess-
ments. Second, L. variegatus appears to be relatively
insensitive to most of the test chemicals, which perhaps
is a positive attribute for an organism used in bioaccumu-
lation tests.
1.3.7.3.4 Using the data from Table 1.4, sensitivity of
H. azteca, C. tentans and L. variegatus can be evaluated
relative to other freshwater species. For this analysis,
acute and chronic toxicity data from water quality criteria
(WQC) documents for copper, zinc, cadmium, nickel,
lead, DDT, dieldrin and chlorpyrifos, and toxicity informa-
tion from the AQUIRE database (AQUIRE, 1992) for ODD
and DDE, were compared to assay results for the three
species (Phippset al., 1995). The sensitivity of H. azteca
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Table 1.3 Rating of Selection Criteria for Freshwater Sediment Toxicity Testing Organisms1
Criterion Hyalella Diporeia Chironomus Chironomus Lumbriculus Tubifex Hexagenia Mollusks Daphnia spp. and
azteca spp. tentans riparius variegatus tubifex spp. Ceriodaphnia spp.
Relative
sensitivity
toxicity +
database
Round-robin
studies +
conducted
Contact with + +
sediment
Laboratory +
culture
Taxonomic +/- +/-
identification
Ecological + +
importance
Geographical + +/-
distribution
Sediment
physico- + +
chemical
tolerance
Response
confirmed + +
with benthic
populations
Peer reviewed + +
Endpoints2 S, G, M, R S, B, A
monitored
NA
S, G, E, R
S, G, E
B, S, R
S, R
S, G
S, G, R
1 A "+" or"-" rating indicates a positive or negative attribute
2 S = Survival, G = Growth, B = Bioaccumulation, A = Avoidance, R = Reproduction, M = Maturation, E = Emergence, NA = not applicable
Table 1.4
Chemical
Water-only, 10-d LC50 (ug/L) Values for Hyalella
azteca, Chironomus tentans, and Lumbriculus
variegatus 1
H. azteca
C. tentans L. variegatus
Copper
Zinc
Cadmium
Nickel
Lead
p,p'-DDT
p,p'-DDD
p,p'-DDE
Dieldrin
Chlorpyrifos
35
73
2.83
780
<16
0.07
0.17
1.39
7.6
0.086
54
1,1 252
NT4
NT
NT
1.23
0.18
3.0
1.1
0.07
35
2,984
158
12,160
794
NT
NT
>3.3
NT
NT
Chemicals tested at ERL-Duluth in soft water—hardness 45 mg/L
as CaCO3 at pH 7.8 to 8.2 (Phipps et al., 1995).
50% mortality at highest concentration tested.
70% mortality at lowest concentration tested.
NT = not tested.
to metals and pesticides, and C. tentans to pesticides
was comparable to chronic toxicity data generated for
other test species. This was not completely unexpected
given that the 10-d exposures used for these two species
are likely more similar to chronic partial life-cycle tests
than the 48- to 96-h exposures traditionally defined as
acute in WQC documents. Interestingly, in some in-
stances (e.g., dieldrin, Chlorpyrifos), LC50 data generated
for H. azteca or C. tentans were comparable to or lower
than any reported for other freshwater species in the WQC
documents. This observation likely is a function not only
of the test species, but of the test conditions; many of the
tests on which early WQC were based were static, rather
than flow-through, and utilized unmeasured contaminant
concentrations.
1.3.7.4 Relative species sensitivity frequently varies
among chemicals; consequently, a battery of tests in-
cluding organisms representing different trophic levels
may be needed to assess sediment quality (Craig, 1984;
-------
Williams et al., 1986a; Long et al., 1990; Ingersoll et al.,
1990; Burton and Ingersoll, 1994; Burton et al., 1996a;
USEPA, 1989c). For example, Reish (1988) reported the
relative toxicity of six metals (As, Cd, Cr, Cu, Hg, and Zn)
to crustaceans, polychaetes, pelecypods, and fishes and
concluded that no single species or group of test organ-
isms was the most sensitive to all of the metals.
1.3.7.5 Measurable concentrations of ammonia are com-
mon in the pore water of many sediments and have been
found to be a common cause of toxicity in pore water
(Jones and Lee, 1988; Ankley et al., 1990; Schubauer-
Berigan and Ankley, 1991). Acute toxicity of ammonia to
H. azteca, C. tentans, and L. variegatus has been evalu-
ated in several studies. As has been found for many
other aquatic organisms, the toxicity of ammonia to
C. tentans and L. variegatus has been shown to be de-
pendent on pH. Four-day LC50 values for L. variegatus in
water-column (no sediment) exposures ranged from 6.6 to
390 mg/L total ammonia as pH was increased from 6.3 to
8.6 (Schubauer-Berigan et al., 1995). For C. tentans, 4-d
LC50 values ranged from 82 to 370 mg/L total ammonia
over a similar pH range (Schubauer-Berigan et al., 1995).
Ankley et al. (1995) reported that the toxicity of ammonia
to H. azteca (also in water-only exposures) showed differ-
ing degrees of pH-dependence in different test waters.
Toxicity was not pH dependent in soft reconstituted wa-
ter, with 4-d LC50 values of about 20 mg/L at pH ranging
from 6.5 to 8.5. In contrast, ammonia toxicity in hard
reconstituted water exhibited substantial pH dependence
with LC50 values decreasing from >200 to 35 mg/L total
ammonia over the same pH range. Borgmann and
Borgmann (1997) later showed that the variation in ammo-
nia toxicity across these waters could be attributed to
differences in sodium and potassium content, which ap-
pear to influence the toxicity of ammonia to H. azteca.
1.3.7.5.1 Although these studies provide benchmark
concentrations that may be of concern in sediment pore
waters, additional studies by Whiteman et al. (1996)
indicated that the relationship between water-only LC50
values and those measured in sediment exposures differs
among organisms. In sediment exposures, the 10-d LC50
for L. variegatus and C. tentans occurred when sediment
pore water reached about 150% of the LC50 determined
from water-only exposures. However, experiments with
H. azteca showed that the 10-d LC50 was not reached
until pore water concentrations were nearly 10 times the
water-only LC50, at which time the ammonia concentra-
tion in the overlying water was equal to the water-only
LC50. The authors attribute this discrepancy to avoid-
ance of sediment by H. azteca. Thus, although it appears
that water-only LC50 values may provide suitable screen-
ing values for potential ammonia toxicity, higher concen-
trations may be necessary to actually induce ammonia
toxicity in sediment exposures, particularly for H. azteca.
Further, these data underscore the importance of measur-
ing the pH of pore water when ammonia toxicity may be of
concern. Ankley and Schubauer-Berigan (1995) and Besser
et al. (1998) describe procedures for conducting toxicity
identification evaluations (TIEs) for pore-water or whole-
sediment samples to determine whether ammonia is
contributing to the toxicity of sediment samples.
1.3.7.6 Sensitivity of a species to chemicals is also
dependant on the duration of the exposure and the end-
points evaluated. Sections 14.4 and 15.4 describe
results of studies which demonstrate the utility of measur-
ing sublethal endpoints in sediment toxicity tests with H.
azteca and C. tentans.
1.3.7.7 The sensitivity of an organism to chemicals
should be balanced with the concept of discrimination
(Burton and Ingersoll, 1994; Burton et al., 1996). The
response of a test organism should provide discrimination
between different levels of contamination.
1.3.7.8 The sensitivity of an organism is related to the
route of exposure and biochemical response to chemi-
cals. Sediment-dwelling organisms can receive exposure
from three primary sources: interstitial water, sediment
particles, and overlying water. Food type, feeding rate,
assimilation efficiency, and clearance rate will control the
dose of chemicals from sediment. Benthic invertebrates
often selectively consume different particle sizes (Harkey
et al., 1994) or particles with higher organic carbon con-
centrations, which may have higher chemical concentra-
tions. Grazers and other collector-gatherers that feed on
aufwuchs, or surface films, and detritus may receive
most of their body burden directly from materials attached
to sediment or from actual sediment ingestion. In amphi-
pods (Landrum, 1989) and clams (Boese et al., 1990),
uptake through the gut can exceed uptake across the gills
of certain hydrophobic compounds. Organisms in direct
contact with sediment may also accumulate chemicals
by direct adsorption to the body wall or by absorption
through the integument (Knezovich etal., 1987).
1.3.7.9 Despite the potential complexities in estimating
the dose that an animal receives from sediment, the
toxicity and bioaccumulation of many chemicals in sedi-
ment such as Kepone®, fluoranthene, organochlorines,
and metals have been correlated with either the concen-
tration of these chemicals in interstitial water or, in the
case of nonionic organic chemicals, in sediment on an
organic-carbon normalized basis (DiToro etal., 1990; Di
Toroetal., 1991). The relative importance of whole sedi-
ment and interstitial water routes of exposure depends on
the test organism and the specific chemical (Knezovich
et al., 1987). Because benthic communities contain a
diversity of organisms, many combinations of exposure
routes can be important. Therefore, behavior and feeding
habits of a test organism can influence its ability to
accumulate chemicals from sediment and should be con-
sidered when selecting test organisms for sediment
testing.
1.3.7.10 The response of H. azteca and C. tentans in
laboratory toxicity studies has been compared with the
response of natural benthic populations.
1.3.7.10.1 Chironomids were not found in sediment
samples that decreased growth of C. tentans by 30% or
-------
more in 10-d laboratory toxicity tests (Giesyetal., 1988).
Wentsel et al. (1977a, 1977b, 1978) reported a correlation
between responses of C. tentans in laboratory tests and
the abundance of C. tentans in metal-contaminated sedi-
ments.
1.3.7.10.2 Canfield et al. (1994, 1996, 1998) evaluated
the composition of benthic invertebrate communities in
sediments for the following areas: (1) three Great Lakes
Areas of Concern (AOC; Buffalo River, NY; Indiana
Harbor, IN; Saginaw River, Ml), (2) the upper Mississippi
River, and (3) the Clark Fork River located in Montana.
Results of these benthic community assessments were
compared to sediment chemistry and toxicity (28-d sedi-
ment exposures with H. azteca which monitored effects
on survival, growth, and sexual maturation). Good con-
cordance was evident between measures of laboratory
toxicity, sediment contamination, and benthic inverte-
brate community composition in extremely contaminated
samples. However, in moderately contaminated samples,
less concordance was observed between the composition
of the benthic community and either laboratory toxicity
test results orsediment contaminant concentration. Labo-
ratory sediment toxicity tests better identified chemical
contamination in sediments compared to many of the
commonly used measures of benthic invertebrate com-
munity composition. Benthic measures may reflect other
factors such as habitat alteration in addition to responding
to contaminants. Canfield et al. (1994, 1996, 1998)
identified the need to better evaluate noncontaminant
factors (i.e., TOC, grain size, water depth, habitat alter-
ation) in order to better interpret the response of benthic
invertebrates to sediment contamination.
1.3.7.10.3 The results from laboratory sediment toxicity
tests were compared to colonization of artificial sub-
strates exposed in situ to Great Lakes sediment (Burton
and Ingersoll, 1994; Burton et al., 1996a). Survival or
growth of/-/, azteca and C. tentans in 10- to 28-d labora-
tory exposures were negatively correlated to percent chi-
ronomids and percent tolerant taxa colonizing artificial
substrates in the field. Schlekat et al. (1994) reported
generally good agreement between sediment tests with H.
azteca and benthic community responses in the Anacostia
River, Washington, D.C.
1.3.7.10.4 Sediment toxicity to amphipods in 10-d toxic-
ity tests, field contamination, and field abundance of
benthic amphipods were examined along a sediment con-
tamination gradient of DDT (Swartzetal., 1994). Survival
of Eohaustorius estuarius, Rhepoxynius abronius, and H.
azteca in laboratory toxicity tests was positively corre-
lated to abundance of amphipods in the field and, along
with the survival of/-/, azteca, was negatively correlated
to DDT concentrations. The threshold for 10-d sediment
toxicity in laboratory studies was about 300 u,g DDT
(+metabolites)/g organic carbon. The threshold for abun-
dance of amphipods in the field was about 100 u,g DDT
(+metabolites)/g organic carbon. Therefore, correlations
between toxicity, contamination, and field populations
indicate that short-term sediment toxicity tests can pro-
vide reliable evidence of biologically adverse sediment
contamination in the field, but may be underprotective of
sublethal effects.
1.3.8 Selection of Organisms for Sediment
Bioaccumulation Testing
1.3.8.1 Several studies have demonstrated that hydro-
phobic organic compounds are bioaccumulated from sedi-
ment by freshwater infaunal organisms, including larval
insects (C. tentans, Adams et al., 1985; Adams, 1987;
Hexagenia limbata, Gobas et al., 1989), oligochaetes
(Tubifextubifexand Limnodrilus hoffmeisteri, Oliver, 1984;
Oliver, 1987; Connell et al., 1988), and by marine organ-
isms (polychaetes, Nephtys incisa; mollusks, Mercenaria
mercenaria, Yoldia limatula; Lake et al., 1990). Consum-
ers of these benthic organisms may bioaccumulate or
biomagnify chemicals. Therefore, in addition to sediment
toxicity, it may be important to examine the uptake of
chemicals by aquatic organisms from contaminated sedi-
ments.
1.3.8.2 Various species of organisms have been sug-
gested for use in studies of chemical bioaccumulation
from aquatic sediments. Several criteria should be con-
sidered before a species is adopted for routine use in
these types of studies (Ankley et al., 1992a; Call et al.,
1994). These criteria include (1) availability of organisms
throughout the year, (2) known chemical exposure his-
tory, (3) adequate tissue mass for chemical analyses, (4)
ease of handling, (5) tolerance of a wide range of sedi-
ment physico-chemical characteristics (e.g., particle size),
(6) low sensitivity to chemicals associated with sediment
(e.g., metals, organics), (7) amenability to long-term ex-
posures without adding food, (8) and ability to accurately
reflect concentrations of chemicals in field-exposed or-
ganisms (e.g., exposure is realistic). With these criteria in
mind, the advantages and disadvantages of several po-
tential freshwater taxa for bioaccumulation testing are
discussed below.
1.3.8.3 Freshwater clams provide an adequate tissue
mass, are easily handled, and can be used in long-term
exposures. However, few non-exotic freshwater species
are available for testing. Exposure of clams is uncertain
because of valve closure. Furthermore, clams are filter
feeders and may accumulate lower concentrations of
chemicals compared with detritivores (Lake et al., 1990).
Chironomids can be readily cultured, are easy to handle,
and reflect appropriate routes of exposure. However, their
rapid life cycle makes it difficult to perform long-term
exposures with hydrophobic compounds; also, chironomids
can readily biotransform organic compounds such as
benzo[a]pyrene (Harkey et al., 1994). Larval mayflies
reflect appropriate routes of exposure, have adequate
tissue mass for residue analysis, and can be used in
long-term tests. However, mayflies cannot be continuously
cultured in the laboratory and consequently are not always
available for testing. Furthermore, the background
concentrations of chemicals and health of field-collected
individuals may be uncertain. Amphipods (e.g., H. azteca)
can be cultured in the laboratory, are easy to handle, and
reflect appropriate routes of exposure. However, their size
-------
may be insufficient for residue analysis and H. azteca are
sensitive to chemicals in sediment. Fish (e.g., fathead
minnows) provide an adequate tissue mass, are readily
available, are easy to handle, and can be used in long-term
exposures. However, the route of exposure is not
appropriate for evaluating the bioavailability of
sediment-associated chemicals to benthic organisms.
1.3.8.4 Oligochaetes are infaunal benthic organisms that
meet many of the test criteria listed above. Certain oli-
gochaete species are easily handled and cultured, pro-
vide reasonable biomass for residue analyses, and are
tolerant of varying sediment physical and chemical char-
acteristics. Oligochaetes are exposed to chemicals via all
appropriate routes of exposure, including pore water and
ingestion of sediment particles. Oligochaetes need not be
fed during long-term bioaccumulation exposures (Phipps
et al., 1993). Various oligochaete species have been used
in toxicity and bioaccumulation evaluations (Chapman et
al., 1982a, Chapman et al., 1982b; Wiederholm, 1987;
Kielty et al., 1988a; Kielty et al., 1988b; Phipps et al.,
1993), and field populations have been used as indicators
of the pollution of aquatic sediments (Brinkhurst, 1980;
Spencer, 1980; Oliver, 1984; Lauritsen, 1985; Robbinset
al., 1989; Ankley et al., 1992b; Brunson et al., 1993;
Brunson et al., 1998). An additional desirable characteris-
tic of Lumbriculus variegatus in bioaccumulation tests is
that this species does not biotransform PAHs (Harkey et
al., 1994).
1.3.8.5 The response of L. variegatus in laboratory
bioaccumulation studies has been confirmed with natural
populations of Oligochaetes.
1.3.8.5.1 Total PCB concentrations in laboratory-exposed
L. variegatus were similar to concentrations measured in
field-collected Oligochaetes from the same sites (Ankley
et al., 1992b). PCB homologue patterns also were similar
between laboratory-exposed and field-collected Oligocha-
etes. The more highly chlorinated PCBs tended to have
greater bioaccumulation in the field-collected organisms.
In contrast, total PCBs in laboratory-exposed (Pimephales
promelas) and field-collected (Ictalurus melas) fish re-
vealed poor agreement in bioaccumulation relative to the
sediment concentrations at the same sites.
1.3.8.5.2 Chemical concentrations measured in
L. variegatus after 28-d exposures to sediment in the
laboratory were compared to chemical concentrations in
field-collected Oligochaetes from the 13 pools of the upper
Mississippi River where these sediments were collected
(Brunson et al., 1998). Chemical concentrations were
relatively low in sediments and tissues from the pools
evaluated. Only polycyclic aromatic hydrocarbons (PAHs)
and total polychlorinated biphenyls (PCBs) were frequently
measured above detection limits. A positive correlation
was observed between lipid-normalized concentrations of
PAHs detected in laboratory-exposed L. variegatus and
field-collected Oligochaetes across all sampling locations.
Rank correlations for concentrations of individual com-
pounds between laboratory-exposed and field-collected
Oligochaetes were strongest for benzo(e)pyrene, perylene,
benzo(b,k)-fluoranthene, and pyrene (Spearman rank cor-
relations > 0.69). About 90% of the paired PAH concen-
trations in laboratory-exposed and field-collected Oligocha-
etes were within a factor of three of one another indicating
laboratory results could be extrapolated to the field with a
reasonable degree of certainty.
1.4 Performance-based Criteria
1.4.1 USEPA's Environmental Monitoring Manage-
ment Council (EMMC) recommended the use of
performance-based methods in developing chemical
analytical standards (Williams, 1993). Performance-based
methods were defined by EMMC as a monitoring approach
that permits the use of appropriate methods that meet
pre-established demonstrated performance standards
(Section 9.2).
1.4.2 The USEPA Office of Water's Office of Science
and Technology and Office of Research and Development
held a workshop on September 16-18,1992 in Washing-
ton, DC to provide an opportunity for experts in the field of
sediment toxicology and staff from USEPA's Regional
and Headquarters program offices to discuss the develop-
ment of standard freshwater and marine sediment testing
procedures (USEPA, 1992a; USEPA, 1994a). Workgroup
participants reached a consensus on several culturing
and testing methods. In developing guidance for culturing
fresh water test organisms to be included in the USEPA
methods manual for sediment tests, it was agreed that no
single method should be required to culture organisms.
However, the consensus at the workshop was that since
the success of a test depends on the health of the
cultures, having healthy test organisms of known quality
and age for testing was the key consideration. A
performance-based criteria approach was selected as the
preferred method through which individual laboratories
should evaluate culture methods rather than by
control-based criteria. This method was chosen to allow
each laboratory to optimize culture methods and minimize
effects of test organism health on the reliability and
comparability of test results. See Tables 11.3,12.3,13.4,
14.3, and 15.3 for a listing of performance criteria for
culturing and testing.
10
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Section 2
Summary of Method
2.1 Method Description and
Experimental Design
2.1.1 Method Description
2.1.1.1 This manual describes procedures for testing
freshwater organisms in the laboratory to evaluate the
potential toxicity or bioaccumulation of chemicals associ-
ated with whole sediments. Sediments may be collected
from the field or spiked with compounds in the laboratory.
Toxicity methods are outlined for two organisms, the
amphipod Hyalella azteca and the midge Chironomus
tentans. Methods are described for conducting 10-d
toxicity tests with amphipods (Section 11) or midges
(Section 12). Toxicity tests are conducted for 10 d in
300-mL chambers containing 100 ml of sediment and 175
ml of overlying water. Overlying water is added daily and
test organisms are fed during the toxicity tests. The
endpoints in the 10-d toxicity test with H. azteca and C.
tentans are survival and growth. Procedures are primarily
described for testing freshwater sediments; however, es-
tuarine sediments (up to 15 %o salinity) can also be tested
in 10-d toxicity tests with H. azteca.
2.1.1.2 Guidance is also described in the manual for
conducting long-term sediment toxicity tests with
H. azteca (Section 14) and C. tentans (Section 15). The
long-term sediment exposures with H. azteca are started
with 7- to 8-d-old amphipods. On Day 28, amphipods are
isolated from the sediment and placed in water-only cham-
bers where reproduction is measured on Day 35 and 42.
Endpoints measured in the long-term amphipod test in-
clude survival (Day 28, 35, and 42), growth (Day 28 and
42), and reproduction (number of young per female pro-
duced from Day 28 to 42). The long-term sediment
exposures with C. tentans start with newly hatched larvae
(<24-h old) and continues through emergence, reproduc-
tion, and hatching of the F1 generation (about 60-d expo-
sures). Survival and growth are determined at 20 d.
Starting on Day 23 to the end of the test, emergence and
reproduction of C. tentans are monitored daily. The
number of eggs per female is determined for each egg
mass, which is incubated for 6 d to determine hatching
success.
2.1.1.3 Guidance for conducting 28-d bioaccumulation
tests with the oligochaete Lumbriculus variegatus is also
provided in the manual. The overlying water is added daily
and the test organisms are not fed during bioaccumulation
tests. Section 13 also describes procedures for determin-
ing bioaccumulation kinetics of different classes of com-
pounds during 28-d exposures with L. variegatus.
2.1.2 Experimental Design
The following section is a general summary of experimen-
tal design. See Section 16 for additional detail.
2.1.2.1 Control and Reference Sediment
2.1.2.1.1 Sediment tests include a control sediment
(sometimes called a negative control). A control sedi-
ment is a sediment that is essentially free of contami-
nants, is used routinely to assess the acceptability of a
test, and is not necessarily collected near the site of
concern. Any contaminants in control sediment are thought
to originate from the global spread of pollutants and do
not reflect any substantial input from local or nonpoint
sources (ASTM, 1999c). A control sediment provides a
measure of test acceptability, evidence of test organism
health, and a basis for interpreting data obtained from the
test sediments. A reference sediment is typically col-
lected near an area of concern (e.g., a disposal site) and
is used to assess sediment conditions exclusive of
materials) of interest. Testing a reference sediment pro-
vides a site-specific basis for evaluating toxicity.
2.1.2.1.1.1 In general, the performance of test organisms
in the negative control is used to judge the acceptability
of a test, and either the negative control or reference
sediment may be used to evaluate performance in the
experimental treatments, depending on the purpose of
the study. Any study in which organisms in the negative
control do not meet performance criteria must be consid-
ered questionable because it suggests that adverse fac-
tors affected the test organisms. Key to avoiding this
situation is using only control sediments that have a
demonstrated record of performance using the same test
procedure. This includes testing of new collections from
sediment sources that have previously provided suitable
control sediment.
2.1.2.1.1.2 Because of the uncertainties introduced by
poor performance in the negative control, such studies
should be repeated to insure accurate results. However,
the scope or sampling associated with some studies may
make it difficult or impossible to repeat a study. Some
researchers have reported cases where performance in
11
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the negative control is poor, but performance criteria are
met in a reference sediment included in the study design.
In these cases, it might be reasonable to infer that other
samples that show good performance are probably not
toxic; however, any samples showing poor performance
should not be judged to have shown toxicity, since it is
unknown whether the adverse factors that caused poor
control performance might have also caused poor perfor-
mance in the test treatments.
2.1.2.1.2 Natural geomorphological and physico-chemi-
cal characteristics such as sediment texture may influ-
ence the response of test organisms (DeWitt et al., 1988).
The physico-chemical characteristics of test sediment
must be within the tolerance limits of the test organism.
Ideally, the limits of a test organism should be determined
in advance; however, controls for factors such as grain
size and organic carbon can be evaluated if the recom-
mended limits are approached or exceeded in a test
sediment. See Section 10.1 for information on physico-
chemical requirements of test organisms. If the physico-
chemical characteristics of a test sediment exceed the
tolerance limits of the test organism, it may be desirable
to include a control sediment that encompasses those
characteristics. The effects of some sediment character-
istics (e.g., grain size ortotal organic carbon) on sediment
test results may be addressed with regression equations
(DeWitt et al., 1988; Ankley et al., 1994a). The use of
formulated sediment can also be used to evaluate physico-
chemical characteristics of sediment on test organisms
(Walsh et al., 1991; Suedel and Rodgers, 1994; Kemble et
al.,1999; USEPA, 1998).
2.1.2.2 The experimental design depends on the purpose
of the study. Variables that need to be considered include
the number and type of control sediments, the number of
treatments and replicates, and water-quality characteris-
tics.
2.1.2.2.1 The purpose of the study might be to determine
a specific endpoint such as an LC50 and may include a
control sediment, a positive control, a solvent control, and
several concentrations of sediment spiked with a chemi-
cal (see Section 8.3.2).
2.1.2.2.2 The purpose of the study might be to determine
whether field-collected sediments are toxic, and may
include controls, reference sediments, and test sedi-
ments. Controls are used to evaluate the acceptability of
the test (Tables 11.3, 12.3, 13.4, 14.3, 15.3) and might
include a control sediment, a formulated sediment (Sec-
tion 7.2), a sand substrate (for C. tentans; Section 12.2,
15.2), or water-only exposures (for H. azteca; Section
14.3.7.8). Testing a reference sediment provides a
site-specific basis for evaluating toxicity of the test sedi-
ments. Comparisons of test sediments to multiple refer-
ence or control sediments representative of the physical
characteristics of the test sediment (i.e., grain size, or-
ganic carbon) may be useful in these evaluations. A
summary of field sampling design is presented by Green
(1979). See Section 16 for additional guidance on experi-
mental design and statistics.
2.1.2.3 If the purpose of the study is to conduct a
reconnaissance field survey to identify contaminated sites
for further investigation, the experimental design might
include only one sample from each site to allow for
maximum spatial coverage. The lack of replication at a
site usually precludes statistical comparisons (e.g., analy-
sis of variance [ANOVA]) among sites, but these surveys
can be used to identify contaminated sites for further
study or may be evaluated using regression techniques
(Sokal and Rohlf, 1981; Steel and Torrie, 1980).
2.1.2.4 In other instances, the purpose of the study might
be to conduct a quantitative sediment survey of chemis-
try and toxicity to determine statistically significant differ-
ences between effects among control and test sediments
from several sites. The number of replicates per site
should be based on the need for sensitivity or power
(Section 16). In a quantitative survey, replicates (sepa-
rate samples from different grabs collected at the same
site) would need to be taken at each site. Chemical and
physical characterizations of each of these grabs would
be required for each of these replicates used in sediment
testing. Separate subsamples might be used to determine
within-sample variability or to compare test procedures
(e.g., comparative sensitivity among test organisms), but
these subsamples cannot be considered to be true field
replicates for statistical comparisons among sites (ASTM,
1999a).
2.1.2.5 Sediments often exhibit high spatial and temporal
variability (Stemmer et al., 1990a). Therefore, replicate
samples may need to be collected to determine variance
in sediment characteristics. Sediments should be col-
lected with as little disruption as possible; however,
subsampling, compositing, or homogenization of sedi-
ment samples may be necessary for some experimental
designs.
2.1.2.6 Site locations might be distributed along a known
pollution gradient, in relation to the boundary of a disposal
site, or at sites identified as being contaminated in a
reconnaissance survey. Both spatial and temporal com-
parisons can be made. In pre-dredging studies, a sam-
pling design can be prepared to assess the contamination
of samples representative of the project area to be dredged.
Such a design should include subsampling of cores taken
to the project depth.
2.1.2.7 The primary focus of the physical and experimen-
tal test design, and statistical analysis of the data, is the
experimental unit. The experimental unit is defined as the
smallest physical entity to which treatments can be inde-
pendently assigned (Steel and Torrie, 1980) and to which
air and water exchange between test chambers is kept to
a minimum. As the number of test chambers per treat-
ment increases, the number of degrees of freedom and
the power of a significance test increase, and therefore,
the width of the confidence interval on a point estimate,
such as an LC50, decreases (Section 16). Because of
factors that might affect test results, all test chambers
should be treated as similarly as possible. Treatments
should be randomly assigned to individual test chamber
12
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locations. Assignment of test organisms to test cham-
bers should be impartial (Davis etal., 1998).
2.2 Types of Tests
2.2.1 Methods for conducting 10-d toxicity tests are
outlined fortwo organisms, the amphipod H. azteca (Sec-
tion 11) and the midge C. tentans (Section 12). The
manual primarily describes methods for testing freshwa-
ter sediments; however, the methods described can also
be used for testing H. azteca in estuarine sediments in
10-d tests (up to 15%o salinity).
2.2.2 Guidance for conducting long-term toxicity tests is
also outlined for H. azteca (Section 14) and C. tentans
(Section 15).
2.2.3 Guidance for conducting 28-d bioaccumulation
tests with the oligochaete L. variegatus is described in
Section 13. Procedures are also described for determin-
ing bioaccumulation kinetics of different classes of com-
pounds during 28-d exposures with L variegatus.
2.3 Test Endpoints
2.3.1 Endpoints measured in the 10-d toxicity tests are
survival and growth. Length or weight is reported as the
average of the surviving organisms at the end of the test
(Sections 11 and 12). From these data, biomass can also
be calculated (dry weight of surviving organisms divided by
the initial numberof organisms). The rationale forevaluat-
ing biomass in toxicity testing is as follows: small differ-
ences in either growth or survival may not be statistically
significantly different from the control; however, a com-
bined estimate of biomass may increase the statistical
power of the test. Although USEPA (1994c, d) describes
procedures for reporting biomass as a measure of growth
in effluent toxicity tests, the approach has not yet been
routinely applied to sediment testing. Therefore, biomass
is not listed as a primary endpoint in the methods described
in Sections 11,12,14, and 15.
2.3.2 Endpoints measured in the long-term H. azteca
exposures include survival (Day 28, 35, and 42), growth
(Day 28 and 42), and reproduction (number of young per
female produced from Day 28 to 42). The long-term
sediment exposures with C. tentans start with newly
hatched larvae (<24-h old) and continue through emer-
gence, reproduction, and hatching of the F1 generation
(about 60-d exposures). Survival is determined at 20 d.
Starting on Day 23 to the end of the test, emergence and
reproduction of C. tentansare monitored daily. The number
of eggs perfemale is determined for each egg mass, which
is incubated for 6 d to determine hatching success.
2.3.2.1 The long-term toxicity test methods for Hyalella
azteca and Chironomus tentans (Sections 14 and 15) can
be used to measure effects on reproduction as well as
long-term survival and growth. Reproduction is a key
variable influencing the long-term sustainability of popula-
tions (Rees and Crawley, 1989) and has been shown to
provide valuable and sensitive information in the assess-
ment of sediment toxicity (Derr and Zabik, 1972; Wentsel
et al., 1978; Williams et al., 1987; Postma et al., 1995;
Sibleyetal., 1996,1997a; Ingersolletal., 1998). Further,
as concerns have emerged regarding the environmental
significance of chemicals that can act directly or indirectly
on reproductive endpoints (e.g., endocrine disrupting com-
pounds), the need forcomprehensive reproductive toxicity
tests has become increasingly important. Reproductive
endpoints measured in sediment toxicity tests with H.
azteca and C. tentans tend to be more variable compared
with those for survival or growth (Section 14.4.6 and
15.4.6). Hence, additional replicates would be required to
achieve the same statistical power as for survival and
growth endpoints (Section 16). The procedures described
in Sections 14 and 15 include measurement of a variety of
lethal and sublethal endpoints; minor modifications of the
basic methods can be used in cases where only a subset
ofthese endpoints is of interest (Sections 14.1.3 and 15.1.2).
2.3.3 Endpoints measured in bioaccumulation tests are
tissue concentrations of contaminants and for some types
of studies, lipid content. Behavior of test organisms should
be qualitatively observed daily in all tests (e.g., avoidance
of sediment).
13
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Section 3
Definitions
3.1 Terms
The following terms were defined in Lee (1980), NRC
(1989), USEPA (1989c), USEPA-USACE (1991),
USEPA-USACE (1998a), ASTM (1999a), ASTM (1999b),
orASTM(1999h).
3.1.1 Technical Terms
3.1.1.1 Bioaccumulation. The net accumulation of a
substance by an organism as a result of uptake from all
environmental sources.
3.1.1.2 Bioaccumulation factor. Ratio of tissue residue
to contaminant source concentration at steady state.
3.1.1.3 Bioaccumulation potential. Qualitative assess-
ment of whether a contaminant is bioavailable.
3.1.1.4 Bioconcentration. The net assimilation of a
substance by an aquatic organism as a result of uptake
directly from aqueous solution.
3.1.1.5 Bioconcentration factor (BCF). Ratio of tissue
residue to water contaminant concentration at steady
state.
3.1.1.6 Biota-sediment accumulation factor (BSAF).
The ratio of tissue residue to source concentration (e.g.,
sediment at steady state normalized to lipid and sediment
organic carbon).
3.1.1.7 Clean. Denotes a sediment or waterthat does not
contain concentrations of test materials which cause
apparent stress to the test organisms or reduce their
survival.
3.1.1.8 Concentration. The ratio of weight or volume of
test materials) to the weight or volume of sediment or
water.
3.1.1.9 Contaminated sediment. Sediment containing
chemical substances at concentrations that pose a known
or suspected threat to environmental or human health.
3.1.1.10 Control sediment. A sediment that is essen-
tially free of contaminants and is used routinely to assess
the acceptability of a test. Any contaminants in control
sediment may originate from the global spread of pollut-
ants and do not reflect any substantial input from local or
nonpoint sources. Comparing test sediments to control
sediments is a measure of the toxicity of a test sediment
beyond inevitable background contamination. Control
sediment is also called a negative control because no
toxic effects are anticipated in this treatment.
3.1.1.11 Depuration. Loss of a substance from an
organism as a result of any active (e.g., metabolic break-
down) or passive process when the organism is placed
into an uncontaminated environment. Contrast with Elimi-
nation.
3.1.1.12 Effect concentration (EC). The toxicant con-
centration that would cause an effect in a given percent-
age of the test population. Identical to LC when the
observable adverse effect is death. For example, the
EC50 is the concentration of toxicant that would cause a
specified effect in 50% of the test population.
3.1.1.13 Elimination. General term for the loss of a
substance from an organism that occurs by any active or
passive means. The term is applicable either in a con-
taminated environment (e.g., occurring simultaneously
with uptake) or in a clean environment. Contrast with
Depuration.
3.1.1.14 Equilibrium partitioning sediment guide-
lines (ESGs). Numerical concentrations of chemical
contaminants in sediment at or below which direct lethal
or sublethal toxic effects on benthic organisms are not
expected. ESGs are based on the theory that an equilib-
ria exists among contaminant concentration in sediment
pore water, contaminant associated with a binding phase
in sediment, and biota. ESGs are derived by assigning a
protective water-only effects concentration to the pore
water (such as a Final Chronic Value), and expressing the
associated equilibrium sediment concentration in terms of
the principal binding phase that limits contaminant bio-
availability (e.g., total organic carbon for nonionic organ-
ics or acid volatile sulfides for metals).
3.1.1.15 Formulated sediment. Mixtures of materials
used to mimic the physical components of a natural
sediment.
3.1.1.16 Inhibition concentration (1C). The toxicant
concentration that would cause a given percent reduction
in a non-quantal measurement for the test population. For
14
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example, the IC25 is the concentration of toxicant that
would cause a 25% reduction in growth for the test
population, and the IC50 is the concentration of toxicant
that would cause a 50% reduction.
3.1.1.17 Interstitial water or pore water. Water occupy-
ing space between sediment or soil particles.
3.1.1.18 /fr. Uptake rate coefficient from the aqueous
phase, with units of g-waterxg-tissue-1 xtime-1. Contrast
with k.
3.1.1.19
time'1.
Elimination rate constant, with units of
3.1.1.20 ks. Sediment uptake rate coefficient from the
sediment phase, with units of g-sediment x g-tissue'1 x
time'1. Contrast with k,.
3.1.1.21
cient.
K . Organic carbon-water partitioning coeffi-
3.1.1.22 Kow. Octanol-water partitioning coefficient.
3.1.1.23 Kinetic Bioaccumulation Model. Any model
that uses uptake and/or elimination rates to predict tissue
residues.
3.1.1.24 Lethal concentration (LC). The toxicant con-
centration that would cause death in a given percentage
of the test population. Identical to EC when the observ-
able adverse effect is death. For example, the LC50 is the
concentration of toxicant that would cause death in 50%
of the test population.
3.1.1.25 Lowest observed effect concentration (LOEC).
The lowest concentration of a toxicant to which organ-
isms are exposed in a test that causes an adverse effect
on the test organisms (i.e., where a significant difference
exists between the value for the observed response and
that for the controls).
3.1.1.26 Wo observed effect concentration (NOEC).
The highest concentration of a toxicant to which organ-
isms are exposed in a test that causes no observable
adverse effect on the test organisms (i.e., the highest
concentration of a toxicant in which the value for the
observed response is not statistically significantly differ-
ent from the controls).
3.1.1.27 Overlying water. The water placed over sedi-
ment in a test chamber during a test.
3.1.1.28 Reference sediment. A whole sediment near an
area of concern used to assess sediment conditions
exclusive of material(s) of interest. The reference sedi-
ment may be used as an indicator of localized sediment
conditions exclusive of the specific pollutant input of
concern. Such sediment would be collected nearthesite
of concern and would represent the background condi-
tions resulting from any localized pollutant inputs as well
as global pollutant input. This is the manner in which
reference sediment is used in dredged material evaluations.
3.1.1.29 Reference-toxicity test. A test conducted with
reagent-grade reference chemical to assess the sensitiv-
ity of the test organisms. Deviations outside an estab-
lished normal range may indicate a change in the sensitiv-
ity of the test organism population. Reference-toxicity
tests are most often performed in the absence of sedi-
ment.
3.1.1.30 Sediment. Particulate material that usually lies
below water. Formulated particulate material that is in-
tended to lie below water in a test.
3.1.1.31 Spiked sediment. A sediment to which a
material has been added for experimental purposes.
3.1.1.32 Steady state. An equilibrium or "constant" tissue
residue resulting from the balance of the flux of compound
into and out of the organism. Operationally determined by
no statistically significant difference in tissue residue
concentrations from three consecutive sampling periods.
3.1.1.33 Whole sediment. Sediment and associated
pore waterthat have had minimal manipulation. The term
bulk sediment has been used synonymously with whole
sediment.
3.1.2 Grammatical Terms
The words "must," "should," "may," "can," and "might"
have very specific meanings in this manual.
3.1.2.1 "Must" is used to express an absolute require-
ment, that is, to state that a test ought to be designed to
satisfy the specified conditions, unless the purpose of the
test requires a different design. "Must" is only used in
connection with the factors that directly relate to the
acceptability of a test.
3.1.2.2 "Should" is used to state that the specified
condition is recommended and ought to be met if pos-
sible. Although a violation of one "should" is rarely a
serious matter, violation of several will often render the
results questionable.
3.1.2.3 Terms such as "is desirable," "is often desirable,"
and "might be desirable" are used in connection with less
important factors.
3.1.2.4 "May" is used to mean "is (are) allowed to," "can"
is used to mean "is (are) able to," and "might" is used to
mean "could possibly." Thus, the classic distinction be-
tween "may" and "can" is preserved, and "might" is never
used as a synonym for either "may" or "can."
15
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Section 4
Interferences
4.1 General Introduction
4.1.1 Interferences are characteristics of a sediment or
sediment test system, aside from those related to
sediment-associated chemicals of concern, that can po-
tentially affect test organism survival, growth, or repro-
duction. These interferences can potentially confound
interpretation of test results in two ways: (1) false-positive
response, i.e., toxicity is observed in the test when
contamination is not present at concentrations known to
elicit a response, or there is more toxicity than expected;
and (2) false-negative response, i.e., no toxicity or
bioaccumulation is observed when contaminants are
present at concentrations known to elicit a response, or
there is less toxicity or bioaccumulation than expected.
4.1.2 There are three categories of interfering factors that
can cause false-negative or false-positive responses:
(1) those characteristics of sediments affecting survival
independent of chemical concentration (i.e.,
noncontaminant factors), (2) changes in chemical
bioavailability as a function of sediment manipulation or
storage, and (3) the presence of indigenous organisms.
Although test procedures and test organism selection
criteria were developed to minimize these interferences,
this section describes the nature of these interferences.
4.1.3 Because of the heterogeneity of natural sediments,
extrapolation from laboratory studies to the field can
sometimes be difficult (Table 4.1; Burton, 1991). Sedi-
ment collection, handling, and storage procedures may
alter bioavailability and concentration of chemicals of
concern by changing the physical, chemical, or biological
characteristics of the sediment. Maintaining the integrity
of a field-collected sediment during removal, transport,
mixing, storage, and testing is extremely difficult and may
complicate the interpretation of effects. Direct compari-
sons of organisms exposed in the laboratory and in the
field would be useful to verify laboratory results. However,
spiked sediment may not be representative of contami-
nated sediment in the field. Mixing time (Stemmer et al.,
1990a), aging (Word etal., 1987; Landrum, 1989; Landrum
and Faust, 1992) and the chemical form of the material
can affect responses of test organisms in spiked sedi-
ment tests.
4.1.4 Laboratory testing with field-collected sediments
may be useful in estimating cumulative effects and
interactions of multiple chemicals in a sample. Tests with
Table 4.1 Advantages and Disadvantages for Use of
Sediment Tests1
Advantages
• Sediment tests measure bioavailable fraction of
contaminant(s).
• Sediment tests provide a direct measure of benthic effects,
assuming no field adaptation or amelioration of effects.
• Limited special equipment is required for testing.
• Ten-day toxicity test methods are rapid and inexpensive.
• Legal and scientific precedence exists for use; ASTM standard
guides are available.
• Sediment tests measure unique information relative to
chemical analyses or benthic community analyses.
• Tests with spiked chemicals provide data on cause-effect
relationships.
• Sediment toxicity tests can be applied to all chemicals of
concern.
• Tests applied to field samples reflect cumulative effects of
contaminants and contaminant interactions.
• Toxicity tests are amenable to confirmation with natural
benthos populations.
Disadvantages
• Sediment collection, handling, and storage may alter bioavail-
ability.
• Spiked sediment may not be representative of field contami-
nated sediment.
• Natural geochemical characteristics of sediment may affect
the response of test organisms.
• Indigenous animals may be present in field-collected sedi-
ments.
• Route of exposure may be uncertain and data generated in
sediment toxicity tests may be difficult to interpret if factors
controlling the bioavailability of contaminants in sediment are
unknown.
• Tests applied to field samples may not discriminate effects of
individual chemicals.
• Few comparisons have been made of methods or species.
• Only a few chronic methods for measuring sublethal effects
have been developed or extensively evaluated.
• Laboratory tests have inherent limitations in predicting
ecological effects.
1 Modified from Swartz (1989)
field samples usually cannot discriminate between effects
of individual chemicals. Most sediment samples contain
a complex matrix of inorganic and organic chemicals with
many unidentified compounds. The use of Toxicity
16
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Identification Evaluations (TIE) in conjunction with
sediment tests with spiked chemicals may provide
evidence of causal relationships and can be applied to
many chemicals of concern (Ankley and Thomas, 1992;
Adams et al., 1985; USEPA, 1996b). Sediment spiking
can also be used to investigate additive, antagonistic, or
synergistic effects of specific chemical mixtures in a
sediment sample (Swartzetal., 1988).
4.1.5 Spiked sediment may not be representative of
contaminated sediment in the field. Mixing time (Stemmer
et al., 1990b) and aging (Word et al., 1987; Landrum,
1989; and Landrum and Faust, 1992) of spiked sediment
can affect responses of organisms.
4.1.6 Most assessments of contaminated sediment rely
on short-term-lethality testing methods (e.g., ^10 d;
USEPA-USACE, 1977; USEPA-USACE, 1991; Sections
11 and 12). Short-term-lethality tests are useful in identi-
fying "hot spots" of sediment contamination but may not
be sensitive enough to evaluate moderately contaminated
areas. Sediment quality assessments using sublethal
responses of benthic organisms, such as effects on
growth and reproduction, have been used to successfully
evaluate moderately contaminated areas (Scott, 1989;
Kemble et al., 1994; Ingersoll et al., 1998; Sections 14
and 15).
4.1.7 Despite the interferences discussed in this section,
existing sediment test methods that include measure-
ment of sublethal endpoints may be used to provide a
rapid and direct measure of effects of contaminants on
benthic communities (e.g., Canfield etal., 1996). Labora-
tory tests with field-collected sediment can also be used
to determine temporal, horizontal, or vertical distribution
of contaminants in sediment. Most tests can be com-
pleted within two to four weeks. Legal and scientific
precedents exist for use of toxicity and bioaccumulation
tests in regulatory decision-making (e.g., USEPA, 1986a).
Furthermore, sediment tests with complex contaminant
mixtures are important tools for making decisions about
the extent of remedial action for contaminated aquatic
sites and for evaluating the success of remediation activi-
ties.
4.2 Noncontaminant Factors
4.2.1 Results of sediment tests can be used to predict
effects that may occur with aquatic organisms in the field
as a result of exposure under comparable conditions. Yet
motile organisms might avoid exposure in the field. Pho-
toinduced toxicity caused by ultraviolet (UV) light may be
important for some compounds associated with sediment
(e.g., polycyclic aromatic hydrocarbons (PAHs); Daven-
port and Spacie, 1991; Ankley etal., 1994b). Fluorescent
light does not contain UV light, but natural sunlight does.
Lighting can therefore affect toxicological responses and
is an important experimental variable for photoactivated
chemicals. However, lighting typically used to conduct
laboratory tests does not include the appropriate spec-
trum of ultraviolet radiation to photoactivate compounds
(Oris and Giesy, 1985; Ankley et al., 1994b). Therefore,
laboratory tests may not account for toxicity expressed
by this mode of action.
4.2.2 Natural geomorphological and physico-chemical
characteristics such as sediment texture may influence
the response of test organisms (DeWitt et al., 1988). The
physico-chemical characteristics of test sediment need
to be within the tolerance limits of the test organism.
Ideally, the limits of the test organism should be deter-
mined in advance; however, control samples reflecting
differences in factors such as grain size and organic
carbon can be evaluated if the limits are exceeded in the
test sediment (Section 10.1). The effects of sediment
characteristics can also be addressed with regression
equations (DeWitt et al., 1988; Ankley et al., 1994a).
Effects of physico-chemical characteristics of sediment
on test organisms can also be evaluated by using formu-
lated sediment for testing (Section 7.2; Walsh et al.,
1991; Suedel and Rodgers, 1994; Kemble etal., 1999).
See Sections 11.4, 12.4, 13.4, 14.4, and 15.4 for a
discussion of the relationships between grain size of
sediment and responses of test organisms.
4.2.3 A weak relationship was evident between mean
reproduction of/-/, azteca in the 42-d test and grain size
(Section 14.4.3; Ingersoll etal., 1998). Additional study is
needed to better evaluate potential relationships between
reproduction of/-/, azteca and the physical characteristics
of the sediment. The weak relationship between grain
size of sediment and reproduction may have been due to
the fact that some of the samples with higher amounts of
sand also had higher concentrations of organic chemicals
compared with other samples (Ingersoll et al., 1998).
Hyalella azteca tolerated a wide range in sediment par-
ticle size and organic matter in 10- to 28-d tests measur-
ing effects on survival or growth (Ankley et al., 1994a;
Suedel and Rodgers, 1994; Ingersoll et al., 1996; Ingersoll
etal., 1998; Kemble etal., 1999; Section 14.4.3).
4.2.3.1 Until additional studies have been conducted
which substantiate this lack of a correlation between
physical characteristics of sediment and reproduction
measured in the 42-d H. azteca test, it would be desirable
to test control or reference sediments which are represen-
tative of the physical characteristics of field-collected
sediments. Formulated sediments could be used to
bracket the ranges in physical characteristics expected in
the field-collected sediments being evaluated (Section
7.2). Addition of YCT should provide a minimum amount
of food needed to support adequate survival, growth, and
reproduction of H. azteca in sediments low in organic
matter (Section 14.2). Without addition of food, H. azteca
can starve during exposures (McNulty et al., 1999) mak-
ing it impossible to differentiate effects of chemicals from
other sediment characteristics.
4.2.4 Additional potential interferences of tests are de-
scribed in Sections 11.4, 12.4, 13.4,14.4, and 15.4.
17
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4.3 Changes in Bioavailability
4.3.1 Sediment toxicity tests are meant to serve as an
indicator of contaminant-related toxicity that might be
expected under field or natural conditions. Some studies
have indicated differences between results of laboratory
testing and results of field testing of sediments using in
situ exposures (Sasson-Brickson and Burton, 1991).
4.3.2 Sediment collection, handling, and storage proce-
dures may alter contaminant bioavailability and concen-
tration by changing the physical, chemical, or biological
characteristics of the sediment. Manipulations such as
mixing, homogenization, and sieving may temporarily
disrupt the equilibrium of organic compounds in sediment.
Similarly, oxidation of anaerobic sediments increases the
availability of certain metals (Di Toro et al., 1990). Be-
cause the availability of contaminants can be a function
of the degree of manipulation, this manual recommends
that handling, storage, and preparation of the sediment for
testing be as consistent as possible. If sieving is per-
formed, it is done primarily to remove predatory organ-
isms and large debris. This manipulation most likely
results in a worst-case condition of heightened bioavail-
ability yet eliminates predation as a factor that might
confound test results. When sediments are sieved, it may
be desirable to take samples before and after sieving
(e.g., pore-water metals or DOC, AVS, TOC) to document
the influence of sieving on sediment chemistry. USEPA
does not recommend sieving freshwater sediments on a
routine basis. See USEPA (1999) and ASTM (1999b).
4.3.3 Testing sediments at temperatures different from
the field might affect contaminant solubility, partitioning
coefficients, or other physical and chemical characteris-
tics. Interaction between sediment and overlying water
and the ratio of sediment to overlying water can influence
bioavailability (Stemmer et al., 1990b).
4.3.4 The addition of food, water, or solvents to the test
chambers might obscure the bioavailability of contami-
nants in sediment or might provide a substrate for bacte-
rial or fungal growth (Harkeyetal., 1997). Without addition
of food, the test organisms may starve during exposures
(Ankley et al., 1994a; McNulty et al., 1999). However, the
addition of food may alter the availability of the contami-
nants in the sediment (Wiederholmetal., 1987, Harkey et
al., 1994) depending on the amount of food added, its
composition (e.g., TOC), and the chemical(s) of interest.
4.3.5 Depletion of aqueous and sediment-sorbed con-
taminants resulting from uptake by an organism or
absorption to a test chamber can also influence availabil-
ity. In most cases, the organism is a minor sink for
contaminants relative to the sediment. However, within
the burrow of an organism, sediment desorption kinetics
might limit uptake rates. Within minutes to hours, a major
portion of the total chemical can be inaccessible to the
organisms because of depletion of available residues.
The desorption of a particular compound from sediment
may range from easily reversible (labile; within minutes)
to irreversible (non-labile; within days or months; Karickhoff
and Morris, 1985). Interparticle diffusion oradvection and
the quality and quantity of sediment organic carbon can
also affect sorption kinetics.
4.3.6 The route of exposure may be uncertain, and data
from sediment tests may be difficult to interpret if factors
controlling the bioavailability of contaminants in sediment
are unknown. Bulk-sediment chemical concentrations may
be normalized to factors other than dry weight. For ex-
ample, concentrations of nonionic organic compounds
might be normalized to sediment organic-carbon content
(USEPA, 1992c) and certain metals normalized to acid
volatile sulfides (Di Toro et al., 1990). Even with the
appropriate normalizing factors, determination of toxic
effects from ingestion of sediment or from dissolved
chemicals in the interstitial water can still be difficult
(Lamberson and Swartz, 1988).
4.4 Presence of Indigenous Organisms
4.4.1 Indigenous organisms may be present in
field-collected sediments. An abundance of the same
organism or organisms taxonomically similar to the test
organism in the sediment sample may make interpreta-
tion of treatment effects difficult. For example, growth of
amphipods, midges, or mayflies may be reduced if high
numbers of oligochaetes are in a sediment sample
(Reynoldson et al., 1994). Previous investigators have
inhibited the biological activity of sediment with sieving,
heat, mercuric chloride, antibiotics, or gamma irradiation
(see ASTM, 1999b). However, further research is needed
to determine effects on contaminant bioavailability or
other modifications of sediments from treatments such as
those used to remove or destroy indigenous organisms.
18
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Section 5
Health, Safety, and Waste Management
5.1 General Precautions
5.1.1 Development and maintenance of an effective
health and safety program in the laboratory requires an
ongoing commitment by laboratory management and in-
cludes (1) the appointment of a laboratory health and
safety officer with the responsibility and authority to de-
velop and maintain a safety program, (2) the preparation
of a formal written health and safety plan, which is pro-
vided to each laboratory staff member, (3) an ongoing
training program on laboratory safety, and (4) regular
safety inspections.
5.1.2 This manual addresses procedures that may in-
volve hazardous materials, operations, and equipment,
but it does not purport to address all of the safety prob-
lems associated with their use. It is the responsibility of
the user to establish appropriate safety and health prac-
tices, and determine the applicability of regulatory limita-
tions before use. While some safety considerations are
included in this manual, it is beyond the scope of this
manual to encompass all safety requirements necessary
to conduct sediment tests.
5.1.3 Collection and use of sediment may involve sub-
stantial risks to personal safety and health. Contaminants
in field-collected sediment may include carcinogens, mu-
tagens, and other potentially toxic compounds. Inasmuch
as sediment testing is often begun before chemical analy-
ses can be completed, worker contact with sediment
needs to be minimized by (1) using gloves, laboratory
coats, safety glasses, face shields, and respirators as
appropriate, (2) manipulating sediment under a ventilated
hood or in an enclosed glove box, and (3) enclosing and
ventilating the exposure system. Personnel collecting
sediment samples and conducting tests should take all
safety precautions necessary for the prevention of bodily
injury and illness that might result from ingestion or
invasion of infectious agents, inhalation or absorption of
corrosive or toxic substances through skin contact, and
asphyxiation because of lack of oxygen or presence of
noxious gases.
5.1.4 Before beginning sample collection and laboratory
work, personnel should determine that all required safety
equipment and materials have been obtained and are in
good condition.
5.2 Safety Equipment
5.2.1 Personal Safety Gear
5.2.1.1 Personnel should use appropriate safety equip-
ment, such as rubber aprons, laboratory coats, respira-
tors, gloves, safety glasses, face shields, hard hats, and
safety shoes.
5.2.2 Laboratory Safety Equipment
5.2.2.1 Each laboratory should be provided with safety
equipment such as first aid kits, fire extinguishers, fire
blankets, emergency showers, and eye wash stations.
5.2.2.2 All laboratories should be equipped with a tele-
phone to enable personnel to summon help in case of
emergency.
5.3 General Laboratory and Field
Operations
5.3.1 Laboratory personnel should be trained in proper
practices for handling and using chemicals that are en-
countered during procedures described in this manual.
Routinely encountered chemicals include acids, organic
solvents, and standard materials for reference-toxicity
tests. Special handling and precautionary guidance in
Material Safety Data Sheets should be followed for re-
agents and other chemicals purchased from supply houses.
5.3.2 Work with some sediment may require compliance
with rules pertaining to the handling of hazardous materi-
als. Personnel collecting samples and performing tests
should not work alone.
5.3.3 It is advisable to wash exposed parts of the body
with bactericidal soap and water immediately after collect-
ing or manipulating sediment samples.
5.3.4 Strong acids and volatile organic solvents should
be used in a fume hood or under an exhaust canopy over
the work area.
5.3.5 An acidic solution should not be mixed with a
hypochlorite solution because hazardous vapors might be
produced.
19
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5.3.6 To prepare dilute acid solutions, concentrated acid
should be added to water, not vice versa. Opening a bottle
of concentrated acid and adding concentrated acid to
water should be performed only under a fume hood.
5.3.7 Use of ground-fault systems and leak detectors is
strongly recommended to help prevent electrical shocks.
Electrical equipment or extension cords not bearing the
approval of Underwriter Laboratories should not be used.
Ground-fault interrupters should be installed in all "wet"
laboratories where electrical equipment is used.
5.3.8 All containers should be adequately labeled to
identify their contents.
5.3.9 Good housekeeping contributes to safety and
reliable results.
5.4 Disease Prevention
5.4.1 Personnel handling samples that are known or
suspected to contain human wastes should be given the
opportunity to be immunized against hepatitis B, tetanus,
typhoid fever, and polio. Thorough washing of exposed
skin with bactericidal soap should follow handling these
samples.
5.5 Safety Manuals
5.5.1 For further guidance on safe practices when han-
dling sediment samples and conducting toxicity tests,
check with the permittee and consult general industrial
safety manuals including USEPA (1986b) and Walters
and Jameson (1984).
5.6 Pollution Prevention, Waste Manage-
ment, and Sample Disposal
5.6.1 It is the laboratory's responsibility to comply with
the federal, state, and local regulations governing the
waste management, particularly hazardous waste identifi-
cation rules and land disposal restrictions, and to protect
the air, water and land by minimizing and controlling all
releases from fume hoods and bench operations. Also,
compliance is required with any sewage discharge per-
mits and regulations. For further information on waste
management, consult "The Waste Management Manual
for Laboratory Personnel" available from the American
Chemical Society's Department of Government Relations
and Science Policy, 1155 16th Street N.W., Washington,
D.C. 20036.
5.6.2 Guidelines for the handling and disposal of hazard-
ous materials should be strictly followed. The federal
government has published regulations for the manage-
ment of hazardous waste and has given the states the
option of either adopting those regulations or developing
their own. If states develop their own regulations, they are
required to be at least as stringent as the federal regula-
tions. As a handler of hazardous materials, it is a
laboratory's responsibility to know and comply with the
applicable state regulations. Refer to The Bureau of
National Affairs Inc., (1986) for the citations of the federal
requirements.
5.6.3 Substitution of nonhazardous chemicals and re-
agents should be encouraged and investigated whenever
possible. For example, use of a nonhazardous compound
for a positive control in reference-toxicity tests is advis-
able. Reference-toxicity tests with copper can provide
appropriate toxicity at concentrations below regulated
levels.
20
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Section 6
Facilities, Equipment, and Supplies
6.1 General
6.1.1 Before a sediment test is conducted in any test
facility, it is desirable to conduct a "nontoxicant" test with
each test species in which all test chambers contain a
control sediment (sometimes called the negative control)
and clean overlying water. Survival, growth, or reproduc-
tion of the test organisms will demonstrate whether facili-
ties, water, control sediment, and handling techniques are
adequate to result in acceptable species-specific control
numbers. Evaluations may also be made on the magni-
tude of between-chamber variance in a test. See
Section 9.14.
6.2 Facilities
6.2.1 The facility must include separate areas for cultur-
ing test organisms and sediment testing to reduce the
possibility of contamination by test materials and other
substances, especially volatile compounds. Holding, ac-
climation, and culture chambers should not be in a room
where sediment tests are conducted, stock solutions or
sediments are prepared, or equipment is cleaned. Test
chambers may be placed in a temperature-controlled
recirculating water bath, environmental chamber, or equiva-
lent facility with temperature control. An enclosed test
system is desirable to provide ventilation during tests to
limit exposure of laboratory personnel to volatile sub-
stances.
6.2.2 Light of the quality and luminance normally ob-
tained in the laboratory is adequate (about 100 to 1000 lux
using wide-spectrum fluorescent lights; e.g., cool-white or
daylight) has been used successfully to culture and test
organisms. Lux is the unit selected for reporting lumi-
nance in this manual. Multiply units of lux by 0.093 to
convert to units of foot candles. Multiply units of lux by
6.91 x 10~3 to convert to units of u,E/m2/s (assuming an
average wavelength of 550 nm (u,mol ~2 s~1 = W m x ^[nm]
x 8.36 x 10-3); ASTM, 1999g). Luminance should be
measured at the surface of the water in test chambers. A
uniform photoperiod of 16L:8D can be achieved in the
laboratory or in an environmental chamber using auto-
matic timers.
6.2.3 During phases of rearing, holding, and testing, test
organisms should be shielded from external disturbances
such as rapidly changing light or pedestrian traffic.
6.2.4 The test facility should be well ventilated and free of
fumes. Laboratory ventilation systems should be checked
to ensure that return air from chemistry laboratories or
sample handling areas is not circulated to culture or
testing rooms, or that air from testing rooms does not
contaminate culture rooms. Air pressure differentials
between rooms should not result in a net flow of poten-
tially contaminated air to sensitive areas through open or
loose-fitting doors. Air used for aeration must be free of
oil and fumes. Oil-free air pumps should be used where
possible. Filters to remove oil, water, and bacteria are
desirable. Particles can be removed from the air using
filters such as BALSTON® Grade BX (Balston, Inc.,
Lexington, MA) or equivalent, and oil and other organic
vapors can be removed using activated carbon filters
(e.g., BALSTON® C-1 filter), or equivalent.
6.3 Equipment and Supplies
6.3.1 Equipment and supplies that contact stock solu-
tions, sediment, or overlying water should not contain
substances that can be leached or dissolved in amounts
that adversely affect the test organisms. In addition,
equipment and supplies that contact sediment or water
should be chosen to minimize sorption of test materials
from water. Glass, type 316 stainless steel, nylon,
high-density polyethylene, polypropylene, polycarbonate,
and fluorocarbon plastics should be used whenever pos-
sible to minimize leaching, dissolution, and sorption. Con-
crete and high-density plastic containers may be used for
holding and culture chambers, and in the water-supply
system. These materials should be washed in detergent,
acid rinsed, and soaked in flowing water for a week or
more before use. Cast-iron pipe should not be used in
water-supply systems because colloidal iron will be added
to the overlying water and strainers will be needed to
remove rust particles. Copper, brass, lead, galvanized
metal, and natural rubber must not contact overlying
water or stock solutions before or during a test. Items
made of neoprene rubber and other materials not men-
tioned above should not be used unless it has been
shown that their use will not adversely affect survival,
growth, or reproduction of the test organisms.
6.3.2 New lots of plastic products should be tested for
toxicity by exposing organisms to them under ordinary
test conditions before general use.
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6.3.3 General Equipment
6.3.3.1 Environmental chamber or equivalent facility with
photoperiod and temperature control (20°C to 25°C).
6.3.3.2 Water purification system capable of producing at
least 1 mega-ohm water (USEPA, 1991 a).
6.3.3.3 Analytical balance capable of accurately weigh-
ing to 0.01 mg.
6.3.3.4 Reference weights, Class S—for documenting
the performance of the analytical balance(s). The balance(s)
should be checked with reference weights that 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 taking the last weight of a series.
6.3.3.5 Volumetric flasks and graduated cylinders—
Class A, borosilicate glass or nontoxic plastic labware,
10 to 1000 ml for making test solutions.
6.3.3.6 Volumetric pipets—Class A, 1 to 100 ml.
6.3.3.7 Serological pipets—1 to 10 ml, graduated.
6.3.3.8 Pipet bulbs and fillers—PROPIPET® or equiva-
lent.
6.3.3.9 Droppers, and glass tubing with fire polished
edges, 4- to 6-mm ID—for transferring test organisms.
6.3.3.10 Wash bottles—for rinsing small glassware, in-
strument electrodes and probes.
6.3.3.11 Glass or electronic thermometers—for measur-
ing water temperature.
6.3.3.12 National Bureau of Standards Certified ther-
mometer (see USEPA Method 170.1; USEPA, 1979b).
6.3.3.13 Dissolved oxygen (DO), pH/selective ion, and
specific conductivity meters and probes for routine physi-
cal and chemical measurements are needed. Unless a
test is being conducted to specifically measure the effect
of DO or conductivity, a portable field-grade instrument is
acceptable.
6.3.3.14 See Table 6.1 fora list of additional equipment
and supplies. Appendix C outlines additional equipment
and supplies needed for conducting the long-term expo-
sures with C. tentans.
6.3.4 Water-delivery System
6.3.4.1 The water-delivery system used in water-renewal
testing can be one of several designs (Appendix A). The
system should be capable of delivering water to each
replicate test chamber. Mount and Brungs (1967) diluters
have been successfully modified for sediment testing.
Other diluter systems have also been useful (Ingersoll
and Nelson, 1990; Maki, 1977; Benoit et al., 1993; Zumwalt
et al., 1994; Brunson et al., 1998). The water-delivery
system should be calibrated before the test by determin-
ing the flow rate of the overlying water. The general
operation of the system should be visually checked daily
throughout the length of the test. If necessary, the
water-delivery system should be adjusted during the test.
At any particular time during the test, flow rates through
any two test chambers should not differ by more than 10%.
6.3.4.2 The overlying water can be replaced manually
(e.g., siphoning); however, manual systems take more
time to maintain during a test. In addition, automated
systems generally result in less suspension of sediment
compared to manual renewal.
6.3.5 Test Chambers
6.3.5.1 Test chambers may be constructed in several
ways and of various materials, depending on the experi-
mental design and the contaminants of interest. Clear
silicone adhesives, suitable for aquaria, sorb some or-
ganic compounds that might be difficult to remove. There-
fore, as little adhesive as possible should be in contact
with the test material. Extra beads of adhesive should be
on the outside of the test chambers rather than on the
inside. To leach potentially toxic compounds from the
adhesive, all new test chambers constructed using sili-
cone adhesives should be held at least 48 h in overlying
water before use in a test.
6.3.5.2 Test chambers for specific tests are described in
Sections 11,12,13,14, and 15.
6.3.6 Cleaning
6.3.6.1 All nondisposable sample containers, test cham-
bers, and other equipment that have come in contact with
sediment should be washed after use in the manner
described below to remove surface contaminants.
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). Hexane might also be used as a solvent for
removing nonionic organic compounds. However,
acetone is preferable if only one organic solvent is
used to clean equipment.
6. Rinse three times with deionized water.
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Table 6.1 Equipment and Supplies for Culturing and Testing Specific Test Organisms1
A. Biological Supplies
Brood stock of test organisms
Active dry yeast (HA)
Cerophyl® (dried cereal leaves; HA)
Trout food pellets (HA)
Tetrafin® or Tetramin® goldfish food (CT)
Trout starter (LV)
Helisoma sp. snails (optional; LV)
Algae (e.g., Selenastrum capricornutum, Chlorella; CT)
Diatoms (e.g., Navicula sp; HA)
B. Glassware
Culture chambers
Test chambers (300-mL high-form lipless beaker; HA and CT)
Test chambers (15.8- x 29.3- x 11.7-cm, W x L x H; LV)
Juvenile holding beakers (e.g., 1 L; HA)
Crystallizing dishes or beakers (200 to 300 mL; CT)
Erlenmeyer flasks (250 and 500 mL; CT)
Larval rearing chambers (e.g., 19 L capacity; CT)
1/4" glass tubing (for aspirating flask; CT)
Glass bowls (20-cm diameter; LV)
Glass vials (10 mL; LV)
Wide-bore pipets (4- to 6-mm ID)
Glass disposable pipets
Burettes (for hardness and alkalinity determinations)
Graduated cylinders (assorted sizes, 10 mL to 2 L)
C. Instruments and Equipment
Dissecting microscope
Stainless-steel sieves (e.g., U.S. Standard No. 25, 30
35, 40, 50 mesh)
Delivery system for overlying water (See Appendix B for a
listing of equipment needed for water delivery systems)
Photoperiod timers
Light meter
Temperature controllers
Thermometer
Continuous recording thermometers
Dissolved oxygen meter
pH meter
Ion-specific meter
Ammonia electrode (or ammonia test kit)
Specific-conductance meter
Drying oven
Desiccator
Balance (0.01 mg sensitivity)
C. Instruments and Equipment
Blender
Refrigerator
Freezer
Light box
Hemacytometer (HA)
Paper shredder, cutter, or scissors (CT, LV)
Tissue homogenizer (LV)
Electric drill with stainless steel auger (diameter 7.6 cm,
overall length 38 cm, auger bit length 25.4 cm (Section 8.3)
D. Miscellaneous
Ventilation system for test chambers
Air supply and airstones (oil free and regulated)
Cotton surgical gauze or cheese cloth (HA)
Stainless-steel screen (no. 60 mesh, for test chambers)
Glass hole-cutting bits
Silicon adhesive caulking
Plastic mesh (110-um mesh opening; Nytex® 110; HA)
Aluminum weighing pans (Sigma Chemical Co., St. Louis, MO)
Fluorescent light bulbs
Nalgene bottles (500 mL and 1000 mL for food preparation and
storage)
Deionized water
Airline tubing
White plastic dish pan
"Coiled-web material" (3-M, St. Paul, MN; HA)
White paper toweling (for substrate; CT)
Brown paper toweling (for substrate; LV)
Screening material (e.g., Nitex® (110 mesh), window screen,
or panty hose; CT)
Water squirt bottle
Dissecting probes (LV)
Dental picks (LV)
Shallow pans (plastic (light-colored), glass, stainless steel)
E. Chemicals
Detergent (nonphosphate)
Acetone (reagent grade)
Hexane (reagent grade)
Hydrochloric acid (reagent grade)
Chloroform and methanol (LV)
Copper Sulfate, Potassium Chloride
Reagents for reconstituting water
Formalin (or Notox®)
Sucrose
HA = Hyalella azteca
CT = Chironomus tentans
LV = Lumbriculus variegatus
1 Appendix C outlines additional equipment and supplies for the long-term exposures with C. tentans.
6.3.6.2 All test chambers and equipment should be
thoroughly rinsed or soaked with the dilution water imme-
diately before use in a test.
6.3.6.3 Many organic solvents (e.g., methylene chloride)
leave a film that is insoluble in water. A dichromate-sulfuric
acid cleaning solution can be used in place of both the
organic solvent and the acid (see ASTM, 1999e), but the
solution might attack silicone adhesive and leave chro-
mium residues on glass. An alternative to use of
dichromate-sulfuric acid could be to heat glassware for
8hat450°C.
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Section 7
Water, Formulated Sediment, Reagents, and Standards
7.1 Water
7.1.1 Requirements
7.1.1.1 Water used to test and culture organisms should
be uniform in quality. Acceptable water should allow
satisfactory survival, growth, or reproduction of the test
organisms. Test organisms should not show signs of
disease or apparent stress (e.g., discoloration, unusual
behavior). If problems are observed in the culturing or
testing of organisms, it is desirable to evaluate the char-
acteristics of the water. See USEPA (1991 a) and ASTM
(1999a) for a recommended list of chemical analyses of
the water supply.
7.1.2 Source
7.1.2.1 A natural water is considered to be of uniform
quality if monthly ranges of the hardness, alkalinity, and
specific conductance are less than 10% of their respec-
tive averages and if the monthly range of pH is less than
0.4. Natural waters should be obtained from an uncon-
taminated well orspring, if possible, orfrom a surface-water
source. If surface water is used, the intake should be
positioned to (1) minimize fluctuations in quality and
contamination, (2) maximize the concentration of dis-
solved oxygen, and (3) ensure low concentrations of
sulfideand iron. Municipal water supplies may be variable
and may contain unacceptably high concentrations of
materials such as copper, lead, zinc, fluoride, chlorine, or
chloramines. Chlorinated water should not be used for
culturing or testing because residual chlorine and
chlorine-produced oxidants are toxic to many aquatic
organisms. Use of tap water is discouraged unless it is
dechlorinated and passed through a deionizer and carbon
filter (USEPA, 1991 a).
7.1.2.2 For site-specific investigations, it is desirable to
have the water-quality characteristics of the overlying
water as similar as possible to the site water. For certain
applications the experimental design might require use of
water from the site where sediment is collected.
7.1.2.3 Waterthat might be contaminated with facultative
pathogens may be passed through a properly maintained
ultraviolet sterilizer equipped with an intensity meter and
flow controls or passed through a filter with a pore size of
0.45 urn or less.
7.1.2.4 Water might need aeration using air stones,
surface aerators, or column aerators. Adequate aeration
will stabilize pH, bring concentrations of dissolved oxygen
and other gases into equilibrium with air, and minimize
oxygen demand and concentrations of volatiles. Exces-
sive aeration may reduce hardness and alkalinity of hard
water (e.g., 280 mg/L hardness as CaCO3; E.L. Brunson,
USGS, Columbia, MO, personal communication). The
concentration of dissolved oxygen in source water should
be between 90 to 100% saturation to help ensure that
dissolved oxygen concentrations are acceptable in test
chambers.
7.1.3 Reconstituted Water
7.1.3.1 Ideally, reconstituted watershould be prepared by
adding specified amounts of reagent-grade chemicals to
high-purity distilled or deionized water (ASTM, 1999e;
USEPA, 1991 a). Problems have been observed with use
of reconstituted water in long-term exposures with
H. azteca (Section 7.1.3.4.3). In some applications,
acceptable high-purity water can be prepared using deion-
ization, distillation, or reverse-osmosis units (Section
6.3.3.2; USEPA, 1991 a). In some applications, test water
can be prepared by diluting natural water with deionized
water (Kemble et al., 1994) or by adding salts to relatively
dilute natural waters.
7.1.3.2 Deionized watershould be obtained from a sys-
tem capable of producing at least 1 mega-ohm water. 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 mixed-bed
water treatment system. Some investigators have ob-
served that holding reconstituted water prepared from
deionized water for several days before use in sediment
tests may improve performance of test organisms.
7.1.3.3 Conductivity, pH, hardness, dissolved oxygen,
and alkalinity should be measured on each batch of
reconstituted water. The reconstituted water should be
aerated before use to adjust pH and dissolved oxygen to
the acceptable ranges (e.g., Section 7.1.3.4.1). USEPA
(1991 a) recommends using a batch of reconstituted water
for two weeks.
24
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7.1.3.4 Reconstituted Fresh Wafer (Smith et al., 1997)
7.1.3.4.1 To prepare 100 L of reconstituted fresh water,
use the reagent-grade chemicals as follows:
1. Place about 75 L of deionized water in a properly
cleaned container.
2. Add 5 g of CaSO4 and 5 g of CaCI2 to a 2-L aliquot of
deionized water and mix (e.g., on a stir plate) for 30
min or until the salts dissolve.
3. Add 3 g of MgSO4, 9.6 g NaHCO3, and 0.4 g KCI to a
second 2-L aliquot of deionized water and mix on a
stir plate for 30 min.
4. Pour the two 2-L aliquots containing the dissolved
salts into the 75 L of deionized water and fill the
carboy to 100 L with deionized water.
5. Aerate the mixture for at least 24 h before use.
6. The water quality of the reconstituted water should be
approximately the following: hardness, 90 to 100mg/L
as CaCO3, alkalinity 50 to 70 mg/L as CaCO3, con-
ductivity 330 to 360 mS/cm, and pH 7.8 to 8.2.
7.1.3.4.2 This reconstituted fresh water (reformulated
moderately hard reconstituted water) described by Smith
et al. (1997) and described in the first edition of this
manual (USEPA, 1994a) has been used successfully in
10-d round-robin testing with H. azteca, C. tentans, and
C. riparius (Section 17). This reconstituted water has a
higher proportion of chloride to sulfate compared to the
reconstituted waters described in ASTM (1999e) and
USEPA (1991a).
7.1.3.4.3 McNulty et al. (1999) and Kemble et al. (1998,
1999) observed poor survival of H. azteca in tests con-
ducted 14 to 28 d using a variety of reconstituted waters
including the reconstituted water described by Smith et al.
(1997). Borgmann (1996) described a reconstituted water
that was used successfully to maintain H. azteca in
culture; however, some laboratories have not had suc-
cess with reproduction of the H. azteca when using this
reconstituted water in the 42-d test (T.J. Norberg-King,
USEPA, Duluth, MN, personal communication). Research
is ongoing to develop additional types of reconstituted
waters suitable for H. azteca. Until an acceptable recon-
stituted water has been developed for long-term expo-
sures with H. azteca, a natural water demonstrated to
support adequate survival, growth, and reproduction of
amphipods is recommended for use in long-term H. az-
teca exposures (Section 14.2; Ingersoll et al., 1998;
Kemble etal., 1998, 1999).
7.1.3.5 Synthetic Seawater
7.1.3.5.1 Reconstituted salt water can be prepared by
adding commercial sea salts, such as FORTY FATH-
OMS®, HW MARINEMIX®, INSTANT OCEAN®, or
equivalent to deionized water.
7.1.3.5.2 A synthetic seawater formulation called GP2 is
prepared with reagent grade chemicals that can be diluted
with deionized water to the desired salinity (USEPA,
1994d).
7.1.3.5.3 Ingersoll et al. (1992) describe procedures for
culturing H. azteca at salinities up to 15%o. Reconstituted
salt water was prepared by adding INSTANT OCEAN®
salts to a 25:75 (v/v) mixture of freshwater (hardness
283 mg/L as CaCO3) and deionized water that was held at
least two weeks before use. Synthetic seawater was
conditioned by adding 6.2 mL of Frit-zyme® #9 nitrifying
bacteria (Nitromonassp. and Nitrobactersp.; Fritz Chemi-
cal Company, Dallas, TX) to each liter of water. The
cultures were maintained by using renewal procedures;
25% of the culture water was replaced weekly. Hyalella
azteca have been used to evaluate the toxicity of estua-
rine sediments up to 15 %o salinity in 10-d exposures
(Nebeker and Miller, 1988; Roach et al., 1992; Winger et
al., 1993; Ingersoll et al., 1996).
7.2 Formulated Sediment
7.2.1 General Requirements
7.2.1.1 Formulated sediments are mixtures of materials
that mimic the physical components of natural sedi-
ments. Formulated sediments have not been routinely
applied to evaluate sediment contamination. A primary
use of formulated sediment could be as a control sedi-
ment. Formulated sediments allow for standardization of
sediment testing or provide a basis for conducting sedi-
ment research. Formulated sediment provides a basis by
which any testing program can assess the acceptability
of their procedures and facilities. In addition, formulated
sediment provides a consistent measure evaluating
performance-based criteria necessary for test acceptabil-
ity. The use of formulated sediment eliminates interfer-
ences caused by the presence of indigenous organisms.
For toxicity tests with sediments spiked with specific
chemicals, the use of a formulated sediment eliminates or
controls the variation in sediment physico-chemical char-
acteristics and provides a consistent method for evaluat-
ing the fate of chemicals in sediment. See USEPA (1999)
and ASTM (1999b) for additional detail regarding uses of
formulated sediment.
7.2.1.2 A formulated sediment should (1) support the
survival, growth, or reproduction of a variety of benthic
invertebrates, (2) provide consistent acceptable biological
endpoints for a variety of species, and (3) be composed of
materials that have consistent characteristics. Consis-
tent material characteristics include (1) consistency of
materials from batch to batch, (2) contaminant concentra-
tions below concentrations of concern, and (3) availability
to all individuals and facilities (Kemble et al., 1999).
7.2.1.3 Physico-chemical characteristics that might be
considered when evaluating the appropriateness of a
formulated sediment include percent sand, percent clay,
percent silt, organic carbon content, cation exchange
25
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capacity (CEC), oxidation reduction potential (redox), pH,
and carbon:nitrogen:phosphorus ratios.
7.2.2 Sources of Materials
7.2.2.1 A variety of methods describe procedures for
making formulated sediments. These procedures often
use similar constituents; however, they often include
either a component or a formulation step that would result
in variation from test facility to test facility. In addition,
most of the procedures have not been subjected to stan-
dardization and consensus approval or round-robin (ring)
testing. The procedure outlined by Kemble et al. (1999)
below was evaluated in round-robin testing with Hyalella
azteca and Chironomus tentans (Section 17.6).
7.2.2.2 Most formulated sediments include sand and
clay/silt that meet certain specifications; however, some
may be quite different. For example, three sources of clay
and silt include Attagel® 50, ASP® 400, and ASP®
400P. Table 7.1 summarizes the characteristics of these
materials. The percentage of clay ranges from 56.5 to
88.5 and silt ranges from 11.5 to 43.5. These characteris-
tics should be evaluated when considering the materials
to use in a formulated sediment.
Table 7.1
Characteristics of Three Sources of Clays and
Silts Used in Formulated Sediments
Characteristic Attagel® 50 ASP® 400
ASP® 400P
% Sand
% Clay
% Silt
Soil class
0.0
88.50
11.50
Clay
0.01
68.49
31.50
Clay
0.0
56.50
43.50
Silty clay
Note: Table 7.3 lists suppliers for these materials.
7.2.2.3 A critical component of formulated sediment is
the source of organic carbon. Many procedures have
used peat as the source of organic carbon. Othersources
of organic carbon listed in Table 7.2 have been evaluated
including humus, potting soil, maple leaves, composted
Table 7.2. Carbon, Nitrogen, Phosphorus Levels for
Various Sources of Organic Carbon (Kemble et
al., 1998a)
Organic carbon
Source
Peat
Maple leaves 1
Maple leaves 2
Cow manure
Rabbit chow
Humic acid
Cereal leaves
Chlorella
Trout chow
Tetramin®
Tetrafin®
Alpha cellulose
Carbon
(%)
47
42
47
30
40
40
47
40
43
37
36
30
Nitrogen
(mg/g)
4
6
3
11
18
3
4
41
36
45
29
0.7
Phosphorus
(ug/g)
0.4
1.3
1.7
8.2
0.2
ND1
0.4
5.7
11.0
9.6
8.6
ND
cow manure, rabbit chow, cereal leaves, chlorella, trout
chow, Tetramin®, Tetrafin®, and alpha cellulose. Only
peat, humus, potting soil, composted cow manure, and
alpha cellulose have been used successfully without
fouling the overlying water in sediment testing (Kemble et
al., 1999). The other sources of organic carbon listed in
Table 7.2 caused dissolved oxygen concentrations to fall
to unacceptable levels (Kemble et al., 1999). Kemble et
al. (1999) reported that conditioning of formulated sedi-
ment was not necessary when alpha cellulose was used
as a source of organic carbon to prepare sediment for use
as a negative control. In addition, alpha cellulose is a
consistent source of organic carbon that is relatively
biologically inactive and low in concentrations of chemi-
cals of concern. It is one of three forms of cellulose
(alpha, beta, and gamma) that differ in their degree of
polymerization. Alpha cellulose has the highest degree of
polymerization and is the chief constituent of paper pulp.
The beta and gamma forms have a much lower degree of
polymerization and are known as hemicellulose. Hence,
compared with other sources of organic carbon, alpha
cellulose would not serve as a food source, but would
serve as an organic carbon constituent for sediment to
add texture or to provide a partitioning compartment for
chemicals. Using alpha cellulose as a source of organic
carbon for sediment-spiking studies has not been ad-
equately evaluated. Recent work conducted byJ. Besser
(USGS, Columbia, MO, unpublished data) indicated that
using alpha cellulose as a source or organic carbon in 21 -
d studies resulted in some generation of sulfide in the
pore water, which may affect the bioavailability of metals
spiked in sediment.
7.2.2.4 An important consideration in the selection of an
organic carbon source may be the ratio of carbon to
nitrogen to phosphorus. As demonstrated in Table 7.2,
percentage carbon ranged from 30 to 47, nitrogen ranged
from 0.7 to 45 mg/g, and phosphorus ranged from below
detection to 11 ug/g for several different carbon sources.
These characteristics should be evaluated when consid-
ering the materials to use in a formulated sediment.
7.2.3 Procedure
7.2.3.1 A summary of various procedures that have been
used to formulate sediment are listed below. Suppliers of
various components are listed in Table 7.3.
1. Walsh etal. (1981): (1) Wash sand (Mystic White No.
85, 45, and 18—New England Silica Inc.; Note: Mys-
tic White sands are no longer available. Kemble et al.
(1999) found White Quartz sand to be an acceptable
substitute; Table 7.3) and sieve into three grain sizes:
coarse (500 to 1500 mm); medium (250 to 499 mm);
and fine (63 to 249 mm). (2) Obtain clay and silt from
Engelhard Corp. (3) Mill and sieve peat moss through
an 840-mm screen. (4) Mix constituents dry in the
following quantities: coarse sand (0.6%); medium
sand (8.7%); fine sand (69.2%); silt (10.2%); clay
(6.4%); and organic matter (4.9%).
Not detected.
26
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Component
Table 7.3 Sources of Components Used in Formulated Sediments
Sources
Sand • White Quartz sand #1 dry, #2, #3—New England Silica, Inc., South Windsor, CT (Note: Mystic White sands are no
longer available. Kemble et al. (1999) found White Quartz sand to be an acceptable substitute).
• Product No. 33094, BDH Chemical, Ltd., Poole, England
Kaolinite • ASP 400, ASP 400P, ASP 600, ASP 900—Englehard Corporation, Edison, NJ
• Product No. 33059, BDH Chemical, Ltd., Poole, England
Montmorillonite • W.D. Johns, Source Clays, University of Missouri, Columbia, MO
Clay • Lewiscraft Sculptor's Clay, available in hobby and artist supply stores
Humus • Sims Bark Co., Inc., Tuscumbia, AL
Alpha cellulose • Sigma Co., St. Louis, MO
Peat • D.L. Browning Co., Mather, Wl
• Joseph Bentley, Ltd., Barrow-on-Humber, South Humberside, England
• Mellinger's, North Lima, OH
Potting soil • Zehr's No Name Potting Soil, Mississauga, Ontario
Humic acid • Aldrich Chemical Co, Milwaukee, Wl
Cow manure • A.H. Hoffman, Inc., Landisville, PA
Dolomite • Ward's Natural Science Establishment, Inc., Rochester, NY
2. Harrahy and Clements (1997): (1) Rinse peat moss
then soak for 5 d in deionized water renewing water
daily. (2) After acclimation for 5 d, remove all water
and spread out to dry. (3) Grind moss and sieve using
the following sieve sizes: 1.18 mm (discard these
particles); 1.00 mm (average size 1.09 mm); 0.85 mm
(average size 0.925); 0.60 (average size 0.725); 0.425
mm (average size 0.5125 mm); retainer (average size
0.2125 mm). (4) Use a mixture of sizes that provides
an average particle size of 840 mm. (5) Wash me-
dium quartz sand and dry. (6) Obtain clay and silt
using ASP 400 (Englehard Corp). (7) Mix constituents
dry in the following quantities: sand (850 g); silt and
clay (150 g); dolomite (0.5 g); sphagnum moss (22 g);
and humicacid (0.1g). (8) Mix sediment for an hour on
a rolling mill and store dry until ready for use.
3. Hanes et al. (1991): (1) Sieve sand and retain two
particle sizes (90 to 180 urn and 180 to 250 urn) which
are mixed in a ratio of 2:1. (2) Dry potting soil for24 h
at room temperature and sieve through a 1-mm screen.
Clay is commercially available sculptors clay. (3)
Determine percent moisture of clay and soil after
drying for 24 h at 60 to 100°C (correct for percent
moisture when mixing materials). (4) Mix constituents
by weight in the following ratios: sand mixture (42%);
clay (42%); and soil (16%). (5) Autoclave after mixing
in a foil-covered container for 20 min. Mixture can be
stored indefinitely if kept covered after autoclaving.
4. Naylor(1993): (1) Sieve acid-washed sand to obtain a
40-to 100-mm size. (2) Obtain clay as kaolin light. (3)
Grind and sieve peat moss using a 2-mm screen
(peat moss which is allowed to dry out will not rehy-
drate and will float on the water surface). (4) Adjust for
the use of moist peat moss by determining moisture
content (dry 5 samples of peat at 60°C until constant
weight is achieved). (5) Mix constituents by weight in
the following percentages: sand (69%); kaolin (20%);
peat (10% [adjust for moisture content]); and CaCO3
(1%). (6) Mix for 2 h in a soil shaker and store in
sealed containers.
5. Suedel and Rodgers (1994): (1) Sieve sand (Mystic
White #18 and 90; Note: Mystic White sands are no
longer available. Kemble et al. (1999) found White
Quartz sand to be an acceptable substitute; Table 7.3)
to provide three different size fractions: coarse (2.0 to
0.5 mm), medium (0.5 to 0.25 mm) and fine (0.25 to
0.05 mm). (2) Ash silt (ASP 400), clay (ASP 600 and
900), montmorillonite clay, and dolomite at 550°C for
1 h to remove organic matter. (3) Dry humus (70°C)
and mill to 2.0 mm. (4) Add dolomite as 1 % of the silt
requirement. (5) Age materials for 7 d in flowing water
before mixing. (6) Mix constituents to mimic the
desired characteristics of the sediment of concern.
6. Kemble et al. (1999) describe procedures for making
a variety of formulated sediments ranging in grain
size and organic carbon. A sediment with 19% sand
and 2% organic carbon was produced by combining:
(1) 219 grams of sand (White Quartz #1 dry), (2)1242
grams of a silt-clay mixture (ASP 400), (3) 77.3
grams of alpha cellulose, (4) 0.15 grams of humic
27
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acid, and (5) 7.5 grams of dolomite (the dolomite is a
source of bicarbonate buffering that occurs naturally
in soils and sediments). Steps for processing the
sand before use include: (1) rinsing sand with gentle
mixing in well water (hardness 283 mg/L as CaCO3,
alkalinity 255 mg/L as CaCO3, pH 7.8) until the water
runs clear, (2) rinsing the sand for 5 min with deion-
ized water, and (3) air drying the sand. Constituents
are mixed for 1 h on a rolling mill and stored dry until
ready for use (i.e., no conditioning required). When
formulated sediments are made with a high silt-clay
content, the alkalinity and hardness of the pore water
may drop due to cation exchange. Gentle mixing of
the formulated sediment with overlying water before
use in testing reduces this change in the water quality
characteristics of the pore water.
7.3 Reagents
7.3.1 Data sheets should be followed for reagents and
other chemicals purchased from supply houses. The test
materials) should be at least reagent grade, unless a test
using a formulated commercial product, technical-grade,
or use-grade material is specifically needed. Reagent
containers should be dated when received from the sup-
plier, and the shelf life of the reagent should not be
exceeded. Working solutions should be dated when pre-
pared and the recommended shelf life should not be
exceeded.
7.4 Standards
7.4.1 Appropriate standard methods for chemical and
physical analyses should be used when possible. For
those measurements for which standards do not exist or
are not sensitive enough, methods should be obtained
from other reliable sources.
28
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Section 8
Sample Collection, Storage, Manipulation, and Characterization
8.1 Collection
8.1.1 Before the preparation or collection of sediment, a
procedure should be established for the handling of sedi-
ment that might contain unknown quantities of toxic chemi-
cals (Section 5).
8.1.2 Sediments are spatially and temporally variable
(Stemmer et al., 1990a). Replicate samples should be
collected to determine variance in sediment characteris-
tics. Sediment should be collected with as little disruption
as possible; however, subsampling, compositing, or ho-
mogenization of sediment samples might be necessary
for some experimental designs. Sampling can cause loss
of sediment integrity, change in chemical speciation, or
disruption of chemical equilibrium (ASTM, 1999b). A
benthic grab or core should be used rather than a dredge
to minimize disruption of the sediment sample. Sediment
should be collected from a depth that will represent ex-
pected exposure. For example, oligochaetes may burrow
4 to 15 cm into sediment. Samples collected for evalua-
tions of dredged material should include sediment cores
to the depth of removal. Surveys of the toxicity of surficial
sediment are often based on cores of the upper 2 cm
sediment depth.
8.1.3 Exposure to direct sunlight during collection should
be minimized, especially if the sediment contains pho-
tolytic compounds. Sediment samples should be cooled
to 4°C in the field before shipment (ASTM, 1999b). Dry ice
can be used to cool samples in the field; however, sedi-
ments should never be frozen. Monitors can be used to
measure temperature during shipping (e.g., TempTale
Temperature Monitoring and Recording System, Sensitech,
Inc., Beverly, MA).
8.1.4 For additional information on sediment collection
and shipment see USEPA (1999) and ASTM (1999b).
8.2 Storage
8.2.1 Since the contaminants of concern and influencing
sediment characteristics are not always known, it is
desirable to hold the sediments after collection in the dark
at 4°C. Traditional convention has held that toxicity tests
should be started as soon as possible following collection
from the field, although actual recommended storage
times range from two weeks (ASTM, 1999b) to less than
eight weeks (USEPA-USACE, 1998a). Discrepancies in
recommended storage times reflected a lack of data
concerning the effects of long-term storage on the physi-
cal, chemical, and toxicological characteristics of the
sediment. However, numerous studies have recently
been conducted to address issues related to sediment
storage (Dillon etal., 1994; Becker and Ginn, 1995;Carr
and Chapman, 1995; Moore et al., 1996; Sarda and
Burton, 1995; Sijmetal., 1997; DeFoe and Ankley, 1998).
The conclusions and recommendations offered by these
studies vary substantially and appear to depend primarily
upon the type or class of contaminants) present. Consid-
ered collectively, these studies suggest that the recom-
mended guidance that sediments be tested sometime
between the time of collection and 8 weeks storage is
appropriate. Additional guidance is provided below.
8.2.2 Extended storage of sediments that contain high
concentrations of labile chemicals (e.g., ammonia, vola-
tile organics) may lead to a loss of these chemicals and a
corresponding reduction in toxicity. Under these circum-
stances, the sediment should be tested as soon as
possible after collection, but not later than within two
weeks (Sarda and Burton, 1995). Sediments that exhibit
low-level to moderate toxicity can exhibit considerable
temporal variability in toxicity, although the direction of
change is often unpredictable (Carrand Chapman, 1995;
Moore etal., 1996; DeFoe and Ankley, 1998). Forthese
types of sediments, the recommended storage time of <8
weeks may be most appropriate. In some situations, a
minimum storage period for low-to-moderately contami-
nated sediments may help reduce variability. For ex-
ample, DeFoe and Ankley (1998) observed high variability
in survival during early testing periods (e.g., <2 weeks) in
sediments with low toxicity. DeFoe and Ankley (1998)
hypothesized that this variability partially reflected the
presence of indigenous predators that remained alive
during this relatively short storage period. Thus, if preda-
tory species are known to exist, and the sediment does
not contain labile contaminants, it may be desirable to
store the sediment for a short period before testing (e.g., 2
weeks) to reduce potential for interferences from indig-
enous organisms. Sediments that contain comparatively
stable compounds (e.g., high molecularweight compounds
such as PCBs) or which exhibit a moderate-to-high level
of toxicity, typically do not vary appreciably in toxicity in
relation to storage duration (Moore et al., 1996; DeFoe
and Ankley, 1998). Forthese sediments, long-term stor-
age (e.g., >8 weeks) can be undertaken.
29
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8.2.3 Researchers may wish to conduct additional char-
acterizations of sediment to evaluate possible effects of
storage. Concentrations of chemicals of concern could
be measured periodically in pore water during the storage
period and at the start of the sediment test (Kemble et al.,
1994). Ingersoll et al. (1993) recommend conducting a
toxicity test with pore water within two weeks from sedi-
ment collection and at the start of the sediment test.
Freezing might further change sediment properties such
as grain size or chemical partitioning and should be
avoided (ASTM, 1999b; Schuytema et al., 1989). Sedi-
ment should be stored with no air overthe sealed samples
(no head space) at 4°C before the start of a test (Shuba et
al.,1978). Sediment may be stored in containers con-
structed of suitable materials as outlined in Section 6.
8.3 Manipulation
8.3.1 Homogenization
8.3.1.1 Samples tend to settle during shipment. As a
result, water above the sediment should not be discarded
but should be mixed back into the sediment during ho-
mogenization. Sediment samples should not be sieved to
remove indigenous organisms unless there is a good
reason to believe indigenous organisms may influence
the response of the test organism. However, large indig-
enous organisms and large debris can be removed using
forceps. Reynoldson et al. (1994) observed reduced growth
of amphipods, midges, and mayflies in sediments with
elevated numbers of oligochaetes and recommended siev-
ing sediments suspected to have high numbers of indig-
enous oligochaetes. If sediments must be sieved, it may
be desirable to analyze samples before and after sieving
(e.g., pore-water metals, DOC, AVS, TOC) to document
the influence of sieving on sediment chemistry.
8.3.1.2 If sediment is collected from multiple field samples,
the sediment can be pooled and mixed by stirring or using
a rolling mill, feed mixer, or other suitable apparatus (see
ASTM, 1999b). Homogenization of sediment can be ac-
complished using a variable-speed hand-held drill outfit-
ted with a stainless-steel auger (diameter 7.6 cm, overall
length 38 cm, auger bit length 25.4 cm; Part No. 800707,
Augers Unlimited, Exton, PA; Kemble etal., 1994).
8.3.2 Sediment Spiking
8.3.2.1 Test sediment can be prepared by manipulating
the properties of a control sediment. Mixing time (Stemmer
et al., 1990a) and aging (Word et al., 1987; Landrum,
1989; Landrum and Faust, 1992) of spiked sediment can
affect bioavailability of chemicals in sediment. Many
studies with spiked sediment are often started only a few
days afterthe chemical has been added to the sediment.
This short time period may not be long enough for sedi-
ments to equilibrate with the spiked chemicals (Section
8.3.2.2.3). Consistent spiking procedures should be fol-
lowed in orderto make interlaboratory comparisons. See
USEPA (1999) and ASTM (1999b) for additional detail
regarding sediment spiking.
8.3.2.1.1 The cause of sediment toxicity and the magni-
tude of interactive effects of chemicals can be estimated
by spiking a sediment with chemicals or complex waste
mixtures (Lamberson and Swartz, 1992). Sediments spiked
with a range of concentrations can be used to generate
either point estimates (e.g., LC50) or a minimum concen-
tration at which effects are observed (lowest observed
effect concentration; LOEC). Results of tests may be
reported in terms of a BSAF (Ankley et al., 1992b). The
influence of sediment physico-chemical characteristics
on chemical toxicity can also be determined with
sediment-spiking studies (Adams et al., 1985).
8.3.2.2 The test material(s) should be at least reagent
grade, unless a test using a formulated commercial prod-
uct, technical-grade, or use-grade material is specifically
needed. Before a test is started, the following should be
known about the test material: (1) the identity and concen-
tration of major ingredients and impurities, (2) water solu-
bility in test water, (3) log Kow, BCF (from other test
species), persistence, hydrolysis, and photolysis rates of
the test substances, (4) estimated toxicity to the test
organism and to humans, (5) if the test concentration(s)
are to be measured, the precision and bias of the analyti-
cal method at the planned concentration^) of the test
material, and (6) recommended handling and disposal
procedures. Addition of test material(s) to sediment may
be accomplished using various methods, such as a
(1) rolling mill, (2) feed mixer, or (3) hand mixing (ASTM,
1999b; USEPA, 1999). Modifications of the mixing tech-
niques might be necessary to allow time for a test mate-
rial to equilibrate with the sediment. Mixing time of spiked
sediment should be limited from minutes to a few hours,
and temperature should be kept low to minimize potential
changes in the physico-chemical and microbial character-
istics of the sediment (ASTM, 1999b). Duration of contact
between the chemical and sediment can affect partition-
ing and bioavailability (Word etal., 1987). Care should be
taken to ensure that the chemical is thoroughly and
evenly distributed in the sediment. Analyses of sediment
subsamples are advisable to determine the degree of
mixing homogeneity (Ditsworth et al., 1990). Moreover,
results from sediment-spiking studies should be com-
pared to the response of test organisms to chemical
concentrations in natural sediments (Lamberson and
Swartz, 1992).
8.3.2.2.1 Organic chemicals have been added: (1) di-
rectly in a dry (crystalline) form; (2) coated on the inside
walls of the container (Ditsworth et al., 1990); or (3) coated
onto silica sand (e.g., 5% w/w of sediment) which is
added to the sediment (D.R. Mount, USEPA, Duluth, MN,
personal communication). In techniques 2 and 3, the
chemical is dissolved in solvent, placed in a glass spiking
container (with or without sand), then the solvent is slowly
evaporated. The advantage of these three approaches is
that no solvent is introduced to the sediment, only the
chemical being spiked. When testing spiked sediments,
procedural blanks (sediments that have been handled in
the same way, including solvent addition and evaporation,
but contain no added chemical) should be tested in addi-
tion to regular negative controls.
30
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8.3.2.2.2 Metals are generally added in an aqueous
solution (ASTM, 1999b; Carlson et al., 1991; Di Toro et
al., 1990). Ammonia has also been successfully spiked
using aqueous solutions (Besser et al., 1998). Inclusion
of spiking blanks is recommended.
8.3.2.2.3 Sufficient time should be allowed after spiking
for the spiked chemical to equilibrate with sediment com-
ponents. For organic chemicals, it is recommended that
the sediment be aged at least one month before starting a
test. Two months or more may be necessary for chemi-
cals with a high log Kow (e.g., >6; D.R. Mount, USEPA,
Duluth, MN, personal communication). For metals, shorter
aging times (1 to 2 weeks) may be sufficient. Periodic
monitoring of chemical concentrations in pore water dur-
ing sediment aging is highly recommended as a means to
assess the equilibration of the spiked sediments. Moni-
toring of pore water during spiked sediment testing is also
recommended.
8.3.2.3 Direct addition of a solvent (other than water) to
the sediment should be avoided if possible. Addition of
organic solvents may dramatically influence the concen-
tration of dissolved organic carbon in pore water. If an
organic solvent is to be used, the solvent should be at a
concentration that does not affect the test organism.
Further, both solvent control and negative control sedi-
ments must be included in the test. The solvent control
must contain the highest concentration of solvent present
and must be from the same batch used to make the stock
solution (see ASTM, 1999e).
8.3.2.4 If the test contains both a negative control and a
solvent control, the survival, growth, or reproduction of
the organisms tested should be compared. If a statisti-
cally significant difference is detected between the two
controls, only the solvent control may be used for meeting
the acceptability of the test and as the basis for calculat-
ing results. The negative control might provide additional
information on the general health of the organisms tested.
If no statistically significant difference is detected, the
data from both controls should be used for meeting the
acceptability of the test and as the basis for calculating
the results (ASTM, 1999f). If performance in the solvent
control is markedly different from that in the negative
control, it is possible that the data are compromised by
experimental artifacts and may not accurately reflect the
toxicity of the chemical in natural sediments.
8.3.3 Test Concentration(s) for Laboratory
Spiked Sediments
8.3.3.1 If a test is intended to generate an LC50, a
toxicant concentration series (0.5 or higher) should be
selected that will provide partial mortalities at two or more
concentrations of the test chemical. The LC50 of a
particular compound may vary depending on physical and
chemical sediment characteristics. It may be desirable to
conduct a range-finding test in which the organisms are
exposed to a control and three or more concentrations of
the test material that differ by a factor often. Results from
water-only tests could be used to establish concentrations
to be tested in a whole-sediment test based on predicted
pore-water concentrations (Di Toro et al., 1991).
8.3.3.2 Bulk-sediment chemical concentrations might be
normalized to factors other than dry weight. For example,
concentrations of nonpolar organic compounds might be
normalized to sediment organic-carbon content, and si-
multaneously extracted metals might be normalized to
acid volatile sulfides (Di Toro et al., 1990; Di Toro et al.,
1991).
8.3.3.3 In some situations it might be necessary to
simply determine whether a specific concentration of test
material is toxic to the test organism, or whether adverse
effects occur above or below a specific concentration.
When there is interest in a particular concentration, it
might only be necessary to test that concentration and
not to determine an LC50.
8.4 Characterization
8.4.1 All sediments should be characterized and at least
the following determined: pH and ammonia of the pore
water, organic carbon content (total organic carbon, TOC),
particle size distribution (percent sand, silt, clay), and
percent water content (ASTM, 1999a; Plumb, 1981). See
Section 8.4.4.7 for methods to isolate pore water.
8.4.2 Other analyses on sediments might include biologi-
cal oxygen demand, chemical oxygen demand, cation
exchange capacity, Eh, total inorganic carbon, total vola-
tile solids, acid volatile sulfides, metals, synthetic organic
compounds, oil and grease, petroleum hydrocarbons, as
well as interstitial water analyses for various physico-
chemical parameters.
8.4.3 Macrobenthos may be evaluated by subsampling
the field-collected sediment. If direct comparisons are to
be made, subsamples for toxicity testing should be col-
lected from the same sample to be used for analysis of
sediment physical and chemical characterizations. Quali-
tative descriptions of the sediment can include color,
texture, and presence of macrophytes or animals. Moni-
toring the odor of sediment samples should be avoided
because of potential hazardous volatile chemicals.
8.4.4 Analytical Methodology
8.4.4.1 Chemical and physical data should be obtained
using appropriate standard methods whenever possible.
For those measurements for which standard methods do
not exist or are not sensitive enough, methods should be
obtained from other reliable sources.
8.4.4.2 The precision, accuracy, and bias of each analyti-
cal method used should be determined in the appropriate
matrix: that is, sediment, water, tissue. Reagent blanks
and analytical standards should be analyzed, and recov-
eries should be calculated.
8.4.4.3 Concentration of spiked test material(s) in sedi-
ment, interstitial water, and overlying water should be
31
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measured as often as practical during a test. If possible,
the concentration of the test material in overlying water,
interstitial water and sediments should be measured at
the start and end of a test. Measurement of test materials)
degradation products might also be desirable.
8.4.4.4 Separate chambers should be set up at the start
of a test and destructively sampled during and at the end
of the test to monitor sediment chemistry. Test organ-
isms and food should be added to these extra chambers.
8.4.4.5 Measurement of test material(s) concentration in
water can be accomplished by pipeting water samples
from about 1 to 2 cm above the sediment surface in the
test chamber. Overlying water samples should not con-
tain any surface debris, any material from the sides of the
test chamber, or any sediment.
8.4.4.6 Measurement of test material(s) concentration in
sediment at the end of a test can be taken by siphoning
most of the overlying water without disturbing the surface
of the sediment, then removing appropriate aliquots of the
sediment for chemical analysis.
8.4.4.7 Interstitial water
8.4.4.7.1 Interstitial water (pore water), defined as the
water occupying the spaces between sediment or soil
particles, is often isolated to provide either a matrix for
toxicity testing or to provide an indication of the concen-
tration or partitioning of contaminants within the sediment
matrix. Draft USEPA equilibrium partitioning sediment
guidelines (ESGs) are based on the presumption that the
concentration of chemicals in the interstitial water are
correlated directly to their bioavailability and, therefore,
their toxicity (Di Toro et al., 1991). Of additional impor-
tance is contaminants in interstitial waters can be trans-
ported into overlying waters through diffusion, bioturbation,
and resuspension processes (Van Rees et al., 1991).
The usefulness of interstitial water sampling for determin-
ing chemical contamination or toxicity will depend on the
study objectives and nature of the sediments at the study
site.
8.4.4.7.2 Isolation of sediment interstitial water can be
accomplished by a wide variety of methods, which are
based on eitherphysical separation oron diffusion/equilib-
rium. The common physical-isolation procedures can be
categorized as: (1) centrifugation, (2) compression/squeez-
ing, or (3) suction/vacuum. Diffusion/equilibrium proce-
dures rely on the movement (diffusion) of pore-water
constituents across semipermeable membranes into a
collecting chamber until an equilibrium is established. A
description of the materials and procedures used in the
isolation of pore wateris included in the reviews by Bufflap
and Allen (1995a), ASTM (1999b), and USEPA (1999).
8.4.4.7.3 When relatively large volumes of water are
required (>20 ml) fortoxicitytesting orchemical analyses,
appropriate quantities of sediment are generally collected
with grabs or corers forsubsequent isolation of the intersti-
tial water. Several isolation procedures, such as centrifu-
gation (An kley and Scheubauer-Berigan, 1994), squeezing
(Carrand Chapman, 1995) and suction (Wingerand Lasier,
1991; Wingeret al., 1998), have been used successfully to
obtain adequate volumes for testing purposes. Peepers
(dialysis) generally do not produce sufficient volumes for
most analyses; however, larger sized peepers (500-mL
volume) have been used for collecting interstitial water in
situ for chemical analyses and organism exposures (Bur-
ton, 1992; Sarda and Burton, 1995).
8.4.4.7.4 There is no one superior method forthe isolation
of interstitial water used fortoxicity testing and associated
chemical analyses. Factors to consider in the selection of
an isolation procedure may include: (1) volume of pore
waterneeded, (2) ease of isolation (materials, preparation
time, and time required for isolation), and (3) artifacts in the
pore water caused by the isolation procedure. Each ap-
proach has unique strengths and limitations (Bufflap and
Allen, 1995a,1995b; Wingeret al., 1998), which vary with
sediment characteristics, chemicals of concern, toxicity
test methods, and desired test resolution (i.e., data quality
objectives). Forsuctionorcompressionseparation,which
uses a filter or a similar surface, there may be changes to
the characteristics of the interstitial water compared with
separation using centrifugation (Ankley etal., 1994; Horowitz
et al., 1996). For most toxicity test procedures, relatively
large volumes of interstitial water (e.g., liters) are frequently
needed forstatic or renewal exposures with the associated
water chemistry analyses. Although centrifugation can be
used to generate large volumes of interstitial water, it is
difficult to use centrifugation to isolate water from coarser
sediment. If smaller volumes of interstitial water are
adequate and logistics allow, it may be desirable to use
peepers, which establish an equilibrium with the pore water
through a permeable membrane. If logistics do not allow
placement of peeper samplers, an alternative procedure
could be to collect cores that can be sampled using side
port suctioningorcentrifugation(G.A. Burton, Wright State
University, personal communication). However, if larger
samples of interstitial water are needed, it would be
necessary to collect multiple cores as quickly as possible
using an inert environment and to centrifuge samples at
ambient temperatures. See USEPA (1999) and ASTM
(1999b) foradditional detail regarding isolation of interstitial
water.
32
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Section 9
Quality Assurance and Quality Control
9.1 Introduction
9.1.1 Developing and maintaining a laboratory quality
assurance (QA) program requires an ongoing commit-
ment by laboratory management and also includes the
following: (1) appointment of a laboratory quality assur-
ance officer with the responsibility and authority to de-
velop and maintain a QA program, (2) preparation of a
Quality Assurance Project Plan with Data Quality Objec-
tives, (3) preparation of written descriptions of laboratory
Standard Operating Procedures (SOPs) fortest organism
culturing, testing, instrument calibration, sample
chain-of-custody, laboratory sample tracking system, and
(4) provision of adequate, qualified technical staff and
suitable space and equipment to assure reliable data.
Additional guidance for QA can be obtained in USEPA
(1989d) and in USEPA (1994e).
9.1.2 QA practices within a testing laboratory should
address all activities that affect the quality of the final
data, such as (1) sediment sampling and handling, (2) the
source and condition of the test organisms, (3) condition
and operation of equipment, (4) test conditions, (5) instru-
ment calibration, (6) replication, (7) use of reference
toxicants, (8) record keeping, and (9) data evaluation.
9.1.3 Quality control (QC) practices, on the other hand,
consist 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
testing see FDA (1978), USEPA (1979a), USEPA (1980a),
USEPA (1980b), USEPA (1991 a), USEPA (1994c),
USEPA (1994d), USEPA (1995), DeWoskin (1984), and
Taylor (1987).
9.2 Performance-based Criteria
9.2.1 USEPA Environmental Monitoring Management Coun-
cil (EMMC) recommended the use of performance-based
methods in developing standards for chemical ana-
lytical methods (Williams, 1993). Performance-based
methods were defined by EMMC as a monitoring
approach that permits the use of appropriate meth-
ods that meet pre-established demonstrated performance
standards. Minimum required elements of performance,
such as precision, reproducibility, bias, sensitivity, and
detection limits should be specified, and the method
should be demonstrated to meet the performance
standards.
9.2.2 Participants at a September 1992 USEPA sedi-
ment toxicity workshop arrived at a consensus on several
culturing and testing methods for freshwater organisms
(Appendix A of USEPA, 1994a). In developing guidance
for culturing test organisms to be included in this manual
for sediment tests, it was generally agreed that no single
method must be used to culture organisms. Success of a
test relies on the health of the culture from which organ-
isms are taken for testing. Having healthy organisms of
known quality and age for testing is the key consideration
relative to culture methods. Therefore, a performance-based
criteria approach is the preferred method through which
individual laboratories should evaluate culture health rather
than using control-based criteria. Performance-based cri-
teria were chosen to allow each laboratory to optimize
culture methods while providing organisms that produce
reliable and comparable test results. See Tables 11.3,
12.3, 13.4, 14.3 and 15.3 for a listing of performance
criteria for culturing and testing.
9.3 Facilities, Equipment, and Test
Chambers
9.3.1 Separate areas for test organism culturing and
testing must be provided to avoid loss of cultures due to
cross-contamination. Ventilation systems should be de-
signed and operated to prevent recirculation or leakage of
air from chemical analysis laboratories or sample storage
and preparation areas into test organism culturing or
sediment testing areas, and from sediment testing labora-
tories and sample preparation areas into culture rooms.
9.3.2 Equipment for temperature control should be ad-
equate to maintain recommended test-water tempera-
tures. Recommended materials should be used in the
fabricating of the test equipment that comes in contact
with the sediment or overlying water.
9.3.3 Before a sediment test is conducted in a new
facility, a "noncontaminant" test should be conducted in
which all test chambers contain a control sediment and
overlying water. This information is used to demonstrate
that the facility, control sediment, water, and handling
procedures provide acceptable responses of test organ-
isms (See Section 9.14).
33
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9.4 Test Organisms
9.4.1 The organisms should appear healthy, behave
normally, feed well, and have low mortality in cultures,
during holding (e.g., <20% for 48 h before the start of a
test), and in test controls. The species of test organisms
should be positively identified to species.
9.5 Water
9.5.1 The quality of water used for organism culturing and
testing is extremely important. Overlying water used in
testing and water used in culturing organisms should be
uniform in quality. Acceptable water should allow satis-
factory survival, growth, or reproduction of the test organ-
isms. Test organisms should not show signs of disease
or apparent stress (e.g., discoloration, unusual behavior).
See Section 7 for additional details.
9.6 Sample Collection and Storage
9.6.1 Sample holding times and temperatures should
conform to conditions described in Section 8.
9.7 Test Conditions
9.7.1 It is desirable to measure temperature continuously
in at least one chamber during each test. Temperatures
should be maintained within the limits specified for each
test. Dissolved oxygen, alkalinity, water hardness, con-
ductivity, ammonia, and pH should be checked as pre-
scribed in Sections 11.3,12.3, 13.3, 14.3 and 15.3.
9.8 Quality of Test Organisms
9.8.1 It may be desirable for laboratories to periodically
perform 96-h water-only reference-toxicity tests to assess
the sensitivity of culture organisms (Section 9.16). Data
from these reference-toxicity tests could be used to as-
sess genetic strain or life-stage sensitivity to select chemi-
cals. The requirement in the first edition of this manual for
laboratories to conduct monthly reference-toxicity tests
(USEPA, 1994a) has not been included as a requirement
in this second edition for testing sediments because of
the inability of reference-toxicity tests to identify stressed
populations of test organisms (McNulty et al., 1999).
Physiological measurements such as lipid content might
also provide useful information regarding the health of the
cultures.
9.8.2 It is desirable to determine the sensitivity of test
organisms obtained from an outside source. The supplier
should provide data with the shipment describing the
history of the sensitivity of organisms from the same
source culture. The supplier should also certify the spe-
cies identification of the test organisms and provide the
taxonomic references or name(s) of the taxonomic expert(s)
consulted.
9.8.3 All organisms in a test must be from the same
source (Section 10.2.2). Organisms may be obtained
from laboratory cultures orfrom commercial orgovernment
sources (Table 10.1). The test organisms used should be
identified using an appropriate taxonomic key, and verifi-
cation should be documented (Pennak, 1989; Merritt and
Cummins, 1996). Obtaining organisms from wild popula-
tions should be avoided unless organisms are cultured
through several generations in the laboratory. In addition,
the ability of the wild population of sexually reproducing
organisms to cross breed with the existing laboratory
population should be determined (Duan et al.,1997). Sen-
sitivity of the wild population to select chemicals (e.g.,
Table 1.4) should also be documented.
9.9 Quality of Food
9.9.1 Problems with the nutritional suitability of the food
will be reflected in the survival, growth, or reproduction of
the test organisms in cultures or in sediment tests.
9.9.2 Food used to culture organisms used in bioaccumu-
lation tests must be analyzed for compounds to be mea-
sured in the bioaccumulation tests.
9.10 Test Acceptability
9.10.1 Tables 11.3, 12.3, 13.4, 14.3 and 15.3 outline
requirements for acceptability of the tests. An individual
test may be conditionally acceptable if temperature, dis-
solved oxygen, and other specified conditions fall outside
specifications, depending on the degree of the departure
and the objectives of the tests (see test condition sum-
maries in Tables 11.1, 12.1, 13.1, 14.1, and 15.1). The
acceptability of a 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 should be noted when reporting
data from a test.
9.11 Analytical Methods
9.11.1 All routine chemical and physical analyses for
culture and testing water, food, and sediment should
include established quality assurance practices outlined
in USEPA methods manuals (USEPA, 1979a; USEPA,
1979b; USEPA, 1991 a; USEPA, 1994b).
9.11.2 Reagent containers should be dated when re-
ceived from the supplier, and the shelf life of the reagent
should not be exceeded. Working solutions should be
dated when prepared and the recommended shelf life
should not be exceeded.
9.12 Calibration and Standardization
9.12.1 Instruments used for routine measurements of
chemical and physical characteristics such as pH, dis-
solved oxygen, temperature, and conductivity should be
calibrated before use each day according to the instru-
ment manufacturer's procedures as indicated in the gen-
eral section on quality assurance (see USEPA Methods
150.1, 360.1,170.1, and 120.1; USEPA, 1979b). Calibra-
tion data should be recorded in a permanent log.
34
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9.12.2 A known-quality water should be included in the
analyses of each batch of water samples (e.g., water
hardness, alkalinity, conductivity). It is desirable to in-
clude certified standards in the analysis of water samples.
9.13 Replication and Test Sensitivity
9.13.1 The sensitivity of sediment tests will depend in
part on the number of replicates/treatment, the signifi-
cance level selected, and the type of statistical analysis.
If the variability remains constant, the sensitivity of a 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 (Section 16).
9.14 Demonstrating Acceptable
Performance
9.14.1 Intralaboratory precision, expressed as a coeffi-
cient of variation (CV) of the range in response for each
type of test to be used in a laboratory, can 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 (e.g.,
the same test duration, type of water, age of test organ-
isms, feeding) and the same data analysis methods. This
should be done to gain experience for the toxicity tests
and to serve as a point of reference for future testing. A
reference-toxicity concentration series (0.5 or higher)
should be selected that will provide partial mortalities at
two or more concentrations of the test chemical
(Section 8.3.3). Information from previous tests can be
used to improve the design of subsequent tests to opti-
mize the dilution series selected for future testing.
9.14.2 Before conducting tests with potentially contami-
nated sediment, it is strongly recommended that the
laboratory conduct the tests with control sediment(s)
alone. Results of these preliminary studies should be
used to determine if use of the control sediment and other
test conditions (i.e., water quality) result in acceptable
performance in the tests as outlined in Tables 11.1,12.1,
13.1,14.1,and15.1.
9.14.3 Laboratories should demonstrate that their person-
nel are able to recover an average of at least 90% of the
organisms from whole sediment. For example, test organ-
isms could be added to control sediment or test sedi-
ments and recovery could be determined after 1 h
(Tomasovicet al., 1994).
9.15 Documenting Ongoing Laboratory
Performance
9.15.1 Outliers, which are data falling outside the control
limits, and trends of increasing or decreasing sensitivity
are readily identified. If the reference-toxicity results from
a given test fall outside the "expected" range (e.g., +2
SD), the sensitivity of the organisms and the credibility of
the test results may be suspect. In this case, the test
procedure should be examined for defects and should be
repeated with a different batch of test organisms
(Section 16).
9.15.2 A sediment test may be acceptable if specified
conditions of a reference-toxicity test fall outside the
expected ranges (Section 9.10.2). Specifically, a sedi-
ment test should not be judged unacceptable if the LC50
for a given reference-toxicity test falls outside the ex-
pected range or if mortality in the control of the reference-
toxicity test exceeds 10%. All the performance criteria
outlined in Tables 11.3, 12.3, 13.4, 14.3, and 15.3 must
be considered when determining the acceptability of a
sediment test. The acceptability of the sediment test
would depend on the experience and judgment of the
investigator and the regulatory authority.
9.15.3 Performance should improve with experience, and
the control limits should gradually narrow, as the statis-
tics stabilize. However, control limits of+2 SD, by defini-
tion, will be exceeded 5% of the time, regardless of how
well a laboratory performs. Forthis reason, good laborato-
ries that develop very narrow control limits may be penal-
ized if a test result that falls just outside the control limits
is rejected cte facto. The width of the control limits should
be considered in decisions regarding rejection of data
(Section 17).
9.16 Reference Toxicants
9.16.1 Historically, reference-toxicity testing has been
thought to provide three types of information relevant to
the interpretation of toxicity test data: (1) an indication of
the relative "health" of the organisms used in the test;
(2) a demonstration that the laboratory can perform the
test procedure in a reproducible manner; and (3) informa-
tion to indicate whether the sensitivity of the particular
strain or population in use at a laboratory is comparable to
those in use in other facilities. With regard to the first type
of information, recent work by McNulty et al. (1999)
suggests that reference-toxicity tests may not be effec-
tive in identifying stressed populations of test organisms.
In addition, reference-toxicity tests recommended for use
with sediment toxicity tests are short-term, water column
tests, owing in part to the lack of a standard sediment for
reference-toxicity testing. Because the test procedures
for reference-toxicity tests are not the same as for the
sediment toxicity tests of interest, the applicability of
reference-toxicity tests to demonstrate ability to repro-
ducibly perform the sediment test procedures is greatly
reduced. Particularly for the long-term sediment toxicity
tests with H. azteca and C. tentans, performance of
control organisms overtime may be a better indicator of
success in handling and testing these organisms (Sec-
tions 14 and 15).
9.16.2 Although the requirement for monthly testing has
been removed in this second edition of the manual,
periodic reference-toxicity testing should still be con-
ducted as an indication of overall comparability of results
among laboratories (at a minimum, sixtests over a 3-year
period should be conducted to evaluate potential differences
in life stage or genetic strain of test organisms). In
35
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particular, reference-toxicity tests should be performed
more frequently when organisms are obtained from out-
side sources, when there are changes in culture prac-
tices, or when brood stock from an outside source is
incorporated into a laboratory culture.
9.16.3 Reference toxicants such as sodium chloride
(NaCI), potassium chloride (KCI), cadmium chloride (CdCI2),
and copper sulfate (CuSO4) are suitable for use. No one
reference toxicant can be used to measure the sensitivity
of test organisms with respect to another toxicant with a
different mode of action (Lee, 1980). However, it may be
unrealistic to test more than one ortwo reference toxicants
routinely. KCI has been used successfully in round-robin
water-only exposures with H. azteca and C. tentans
(Section 17).
9.16.4 Test conditions for conducting reference-toxicity
tests with H. azteca, C. tentans, and L. variegatus are
outlined in Tables 9.1 and 9.2. Reference-toxicity tests
can be conducted using one organism/chamber or mul-
tiple organisms in each chamber. Some laboratories have
observed low control survival when more than one midge/
chamber is tested in water-only exposures.
9.17 Record Keeping
9.17.1 Section 16.1 outlines recommendations for record
keeping (i.e., data files, chain-of-custody).
Table 9.1 Recommended Test Conditions for Conducting Reference-toxicity Tests with One Organism/Chamber
Parameter Conditions
1. Test type:
2. Dilution series:
3. Toxicant:
4. Temperature:
5. Light quality:
6. Illuminance:
7. Photoperiod:
8. Renewal of water:
9. Age of organisms:
10. Test chamber:
11. Volume of water:
12. Number of organisms/chamber:
13. Number of replicate chambers/treatment:
14. Feeding:
15. Substrate:
16. Aeration:
17. Dilution water:
18. Test chamber cleaning:
19. Water quality:
20. Test duration:
21. Endpoint:
22. Test acceptability:
Water-only test
Control and at least 5 test concentrations (0.5 dilution factor)
NaCI, KCI, Cd, or Cu
23 ± 1 °C
Wide-spectrum fluorescent lights
About 100 to 1000 lux
16L8D
None
H. azteca: 7- to 14-d old (1- to 2-d range in age)
C. tentans: second- to third-instar larvae (about 10-d-old larvae)1
L. variegatus: adults
30-mL plastic cups (covered with glass or plastic)
20 ml
1
10 minimum
H. azteca: 0.1 ml YCT (1800 mg/L stock) on Day 0 and 2
C. tentans: 0.25 ml Tetrafin® (4 g/L stock) on Day 0 and 2
L. variegatus: not fed
H. azteca: Nitex® screen (110 mesh)
C. tentans: sand (monolayer)
L. variegatus: no substrate
None
Culture water, well water, surface water, site water, or reconstituted water
None
Hardness, alkalinity, conductivity, dissolved oxygen, and pH at the beginning and
end of a test. Temperature daily.
96 h
Survival (LC50)
90% control survival
Age requirement: All animals must be third or second instar with at least 50% of the organisms at third instar.
36
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Table 9.2 Recommended Test Conditions for Conducting Reference-toxicity Tests with More Than One
Organism/Chamber
1.
2.
3.
4.
5.
6.
7.
8.
Parameter
Test type:
Dilution series:
Toxicant:
Temperature:
Light quality:
Illuminance:
Photoperiod:
Renewal of water:
Conditions
Water-only test
Control and at least 5 test concentrations (0.5 dilution factor)
NaCI, KCI, Cd, or Cu
23 ± 1 °C
Wide-spectrum fluorescent lights
About 100 to 1000 lux
16L8D
None
9. Age of organisms1
10. Test chamber:
11. Volume of water:
12. Number of organisms/chamber:
13. Number of replicate chambers/treatment:
14. Feeding:
15. Substrate:
16. Aeration:
17. Dilution water:
18. Test chamber cleaning:
19. Water quality:
20. Test duration:
21. Endpoint:
22. Test acceptability:
H. azteca: 7- to 14-d old (1- to 2-d range in age)
C. tentans: second to third instar (about 10-d-old larvae)1
L. variegatus: adults
250-mL glass beaker (covered with glass or plastic)
100 ml (minimum)
10 minimum
3 minimum
H. azteca: 0.5 ml YCT (1800 mg/L stock) on Day 0 and 2
C. tentans: 1.25 ml Tetrafin® (4 g/L stock) on Day 0 and 2
L. variegatus: not fed
H. azteca: Nitex® screen (110 mesh)
C. tentans: sand (monolayer)
L. variegatus: no substrate
None
Culture water, well water, surface water, site water or
reconstituted water
None
Hardness, alkalinity, conductivity, dissolved oxygen, and pH
at the beginning and end of a test. Temperature daily.
96 h
Survival (LC50)
90% control survival
1 Age requirement: All animals must be third or second instar with at least 50% of the organisms at third instar.
37
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Section 10
Collecting, Culturing, and Maintaining Test Organisms
10.1 Life Histories
10.1.1 Hyalella azteca
10.1.1.1 Hyalella azteca inhabit permanent lakes, ponds,
and streams throughout North and South America (de
March, 1981; Pennak, 1989). Occurrence of/-/, azteca is
most common in warm (20°C to 30°C for much of the
summer) mesotrophic or eutrophic lakes that support
aquatic plants. These amphipods are also found in ponds,
sloughs, marshes, rivers, ditches, streams, and springs,
but in lower numbers. Hyalella azteca have achieved
densities of >10,000/m2 in preferred habitats (de March,
1981).
10.1.1.2 Hyalella azteca are epibenthic detritivores that
burrow into the sediment surface. Margrave (1970a) re-
ported that H. azteca selectively ingest bacteria and
algae. The behavior and feeding habits of/-/, azteca make
them excellent test organisms for sediment assessments.
10.1.1.3 Reproduction by H. azteca is sexual. The adult
males are larger than females and have larger second
gnathopods (de March, 1981). Males pair with females by
grasping the females (amplexus) with their gnathopods
while on the backs of the females. After feeding together
for 1 to 7 d the female is ready to molt and the two
organisms separate for a short time while the female
sheds her old exoskeleton. Once the exoskeleton is
shed, the two organisms reunite and copulation occurs.
The male places sperm nearthe marsupium of the female
and her pleopods sweep the sperm into the marsupium.
The organisms again separate and the female releases
eggs from her oviducts into the marsupium where they are
fertilized. Hyalella azteca average about 18 eggs/brood
(Pennak, 1989) with larger organisms having more eggs
(Cooper, 1965).
10.1.1.4 The developing embryos and newly hatched
young are kept in the marsupium until the next molt. At
24°C to 28°C, hatching ranges from 5 to 10 d after
fertilization (Embody, 1911; Bovee, 1950; Cooper, 1965).
The time between molts for females is 7 to 8 d at 26°C to
28°C (Bovee, 1950). Therefore, about the time embryos
hatch, the female molts and releases the young. Hyalella
azteca average 15 broods in 152 d (Pennak, 1989).
Pairing of the sexes is simultaneous with embryo incubation
of the previous brood in the marsupium. Hyalella azteca
have a minimum of nine instars (Geisler, 1944). There are
5 to 8 pre-reproductive instars (Cooper, 1965) and an
indefinite number of post-reproductive instars. The first
five instars form the juvenile stage of development, instar
stages 6 and 7 form the adolescent stage when sexes
can be differentiated, instar stage 8 is the nuptial stage,
and all later instars are the adult stages of development
(Pennak, 1989).
10.1.1.5 Hyalella azteca have been successfully cultured
at illuminance of about 100 to 1000 lux (Ingersoll and
Nelson, 1990; Ankley etal., 1991 a; Ankley etal., 1991b).
Hyalella azteca avoid bright light, preferring to hide under
litter and feed during the day.
10.1.1.6 Temperatures tolerated by H. azteca range from
0 to 33°C (Embody, 1911; Bovee, 1949; Sprague, 1963).
At temperatures less than 10°C the organisms rest and
are immobile (de March, 1977; de March, 1978). At tem-
peratures of 10°C to 18°C, reproduction can occur. Juve-
niles grow more slowly at colder temperatures and be-
come larger adults. Smaller adults with higher reproduc-
tion are typical when organisms are grown at 18°C to
28°C. The highest rates of reproduction occur at 26°C to
28°C (de March, 1978) while lethality occurs at 33°C to
37°C (Bovee, 1949; Sprague, 1963).
10.1.1.7 Hyalella azteca are found in waters of widely
varying types. Hyalella azteca can inhabit saline waters
up to 29 %0; however, their distribution in these saline
waters has been correlated to water hardness (Ingersoll et
al., 1992). Hyalella azteca inhabit water with high Mg
concentrations at conductivities up to 22,000 uS/cm, but
only up to 12,000 uS/cm in Na-dominated waters (Ingersoll
etal., 1992). De March (1981) reported H. azteca were not
collected from locations where calcium was less than
7 mg/L. Hyalella azteca have been cultured in reconsti-
tuted salt water with a salinity up to 15%o (Ingersoll etal.,
1992; Winger and Lasier, 1993). In laboratory studies,
Sprague (1963) reported a 24-h LC50 for dissolved oxy-
gen at 20°C of 0.7 mg/L. Pennak and Rosine (1976)
reported similar findings. Nebekeret al. (1992) reported
48-h and 30-d LC50s for H. azteca of less than 0.3 mg/L
dissolved oxygen. Weight and reproduction of/-/, azteca
were reduced after 30-d exposure to 1.2 mg/L dissolved
oxygen.
38
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10.1.1.8 Hyalella azteca tolerate a wide range of sub-
strates. Ingersoll et al. (1996) reported that H. azteca
tolerated sediments ranging from more than 90% silt- and
clay-sized particles to 100% sand-sized particles without
detrimental effects on either survival or growth. Hyalella
azteca tolerated a wide range in grain size and organic
matter in 10- to 42-d tests with formulated sediment
(Suedel and Rodgers, 1994; Ingersoll et al., 1998). Ankley
et al. (1994a) evaluated the effects of natural sediment
physico-chemical characteristics on the results of 10-d
laboratory toxicity tests with H. azteca, C. tentans, and
L. variegatus. Tests were conducted with and without the
addition of exogenous food. Survival of organisms was
decreased in tests without added food. Physico-chemical
sediment characteristics including grain size and TOC
were not significantly correlated to the response of
H. azteca in either fed or unfed tests. See Sections 4.2.3
and 14.4 for additional detail regarding studies of the
influence of grain size in long-term sediment toxicity tests
with H. azteca.
10.1.2 Chironomus tentans
10.1.2.1 Chironomus tentans have a holarctic distribution
(Townsend et al., 1981) and are commonly found in
eutrophic ponds and lakes (Flannagan, 1971; Driver, 1977).
Midge larvae are important in the diet offish and waterfowl
(Sadler, 1935; Siegfried, 1973; Driveretal., 1974; McLarney
et al., 1974). Larvae of C. tentans usually penetrate a few
cm into sediment. In both lotic and lentic habitats with
soft bottoms, about 95% of the chironomid larvae occur in
the upper 10 cm of substrates, and very few larvae are
found below 40 cm (Townsend et al., 1981). Larvae were
found under the following conditions in British Columbia
lakes by Topping (1971): particle size <0.15 mm to 2.Omm,
temperature 0 to 23.3°C, dissolved oxygen 0.22 to
8.23 mg/L, pH 8.0 to 9.2, conductivity 481 to
4,136 umhos/cm, and sediment organic carbon 1.9 to
15.5%. Larvae were absent from lakes if hydrogen sulfide
concentration in overlying water exceeded 0.3 mg/L. Abun-
dance of larvae was positively correlated with conductiv-
ity, pH, amount of food, percentages of particles in the
0.59 to 1.98 mm size range, and concentrations of Na, K,
Mg, Cl, SO4, and dissolved oxygen. Others (e.g., Curry,
1962; Oliver, 1971) have reported a temperature range of
0 to 35°C and a pH range of 7 to 10.
10.1.2.2 Chironomus tentans are aquatic during the larval
and pupal stages. The life cycle of C. tentans can be
divided into four distinct stages: (1) an egg stage, (2) a
larval stage, consisting of four instars, (3) a pupal stage,
and (4) an adult stage. Mating behavior has been de-
scribed by Sadler (1935) and others (ASTM, 1999a).
Males are easily distinguished from females because
males have large, plumose antennae and a much thinner
abdomen with visible genitalia. The male has paired geni-
tal claspers on the posteriortip of the abdomen (Townsend
et al., 1981). The adult female weighs about twice as
much as the male, with about 30% of the female weight
contributed by the eggs. After mating, adult females
oviposit a single transparent, gelatinous egg mass di-
rectly into the water. At the USEPA Office of Research
and Development Laboratory (Duluth, MN), the females
oviposit eggs within 24 h after emergence. Egg cases
contain a variable number of eggs from about 500 to 2000
eggs/eggcase (J. Jenson, ILS, Duluth, MN, personal
communication) and will hatch in 2 to 4 d at 23°C. Under
optimal conditions larvae will pupate and emerge as adults
after about 21 d at 23°C. Larvae begin to construct tubes
(or cases) on the second or third day after hatching. The
cases lengthen and enlarge as the larvae grow with the
addition of small particles bound together with threads
from the mouths of larvae (Sadler, 1935). The larvae draw
food particles inside the tubes and also feed in the
immediate vicinity of either end of the open-ended tubes
with their caudal extremities anchored within the tube.
The four larval stages are followed by a black-colored
pupal stage (lasting about 3 d) and emergence to a
terrestrial adult (imago) stage. The adult stage lasts for
3 to 5 d, during which time the adults mate during flight
and the females oviposit their egg cases (2 to 3 d post-
emergence; Sadler, 1935).
10.1.2.3 Grain size tolerance of C. tentans in sediment
testing is described in Section 12.4.3 for 10-d exposures
and in Section 15.4.3 for long-term exposures.
10.1.3 Lumbriculus variegatus
10.1.3.1 Lumbriculus variegatus inhabit a variety of
sediment types throughout the United States and Europe
(Chekanovskaya, 1962; Cook, 1969; Spencer, 1980;
Brinkhurst, 1986). Lumbriculus variegatus typically tunnel
in the upper aerobic zone of sediments of reservoirs,
rivers, lakes, ponds, and marshes. When not tunneling,
they bury their anterior portion in sediment and undulate
their posterior portion in overlying water for respiratory
exchange.
10.1.3.2 Adults of L. variegatus can reach a length of
40 to 90 mm, diameter of 1.0 to 1.5 mm, and wet weight of
5 to 12 mg (Call etal., 1991; Phipps et al., 1993). Lipid
content is about 1.0% (wet weight, Ankley et al., 1992b;
Brunson et al., 1993; Brunson et al., 1998). Lumbriculus
variegatus most commonly reproduce asexually, although
sexual reproduction has been reported (Chekanovskaya,
1962). Newly hatched worms have not been observed in
cultures (Call etal., 1991; Phipps etal., 1993). Cultures
consist of adults of various sizes. Populations of labora-
tory cultures double (number of organisms) every 10 to
14 d at 20°C (Phipps et al., 1993).
10.1.3.3 Lumbriculus variegatus tolerate a wide range of
substrates. Ankley et al. (1994a) evaluated the effects of
natural sediment physico-chemical characteristics on the
results of 10-d laboratory toxicity tests with H. azteca,
C. tentans, and L. variegatus. Tests were conducted with
and without the addition of exogenous food. Survival and
reproduction of organisms was decreased in tests without
added food. Physico-chemical sediment characteristics
including grain size and TOC were not significantly corre-
lated to reproduction or growth of L. variegatus in either
fed or unfed tests.
39
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10.2 General Culturing Procedures
10.2.1 Acceptability of a culturing procedure is based in
part on performance of organisms in culture and in the
sediment test (Section 1.4 and 9.2). No single technique
for culturing test organisms is required. What may work
well for one laboratory may not work as well for another
laboratory. While a variety of culturing procedures are
outlined in Section 10.3 for/-/, azteca, in Section 10.4 for
C. tentans, and in Section 10.5 for L. variegatus, organ-
isms must meet the test acceptability requirements listed
in Tables 11.3, 12.3,13.4, 14.3, and 15.3.
10.2.2 All organisms in a test must be from the same
source. Organisms may be obtained from laboratory cul-
tures or from commercial or government sources
(Table 10.1). The test organism used should be identified
using an appropriate taxonomic key, and verification should
be documented. Obtaining organisms from wild popula-
tions should be avoided unless organisms are cultured
through several generations in the laboratory. In addition,
Table 10.1 Sources of Starter Cultures of Test Organisms
Source Species
U.S. Environmental Protection Agency H. azteca
Mid-Continent Ecological Division C. tentans
6201 Congdon Boulevard L. variegatus
Duluth, MN 55804
Teresa Norberg-King (218/529-5163, fax -5003)
email: norberg-king.teresa@epa.gov
U.S. Environmental Protection Agency H. azteca
Environmental Monitoring System Laboratory L. variegatus
26 W. Martin Luther Dr.
Cincinnati, OH 45244
Jim Lazorchak (513/569-7076, fax -7609)
email: lazorchak.jim@epa.gov
Columbia Environmental Research Center H. azteca
U.S. Geological Survey C. tentans
4200 New Haven Road L. variegatus
Columbia, MO 65201
Eugene Greer (573/876-1820, fax -1896)
email: eugene_greer@usgs.gov
Great Lakes Environmental Research L. variegatus
Laboratory, NOAA
2205 Commonwealth Boulevard
Ann Arbor, Ml 48105-1593
Peter Landrum (313/741-2276, fax -2055)
email: landrum@glerl.noaa.gov
Wright State University H. azteca
Institute for Environmental Quality C. tentans
Dayton, OH 45435 L. variegatus
Allen Burton (937/775-2201, fax -4997)
email: aburton@wright.edu
Michigan State University H. azteca
Department of Fisheries and Wildlife C. tentans
No. 13 Natural Resources Building L. variegatus
East Lansing, Ml 48824-1222
John Giesy (517/353-2000, fax 517/432-1984)
email: jgiesy@aol.com
the ability of the wild population of sexually reproducing
organisms to crossbreed with the existing laboratory popu-
lation should be determined (Duan et al. ,1997). Sensitiv-
ity of the wild population to select chemicals (e.g., Table
1.4) should also be documented.
10.2.3 Test organisms obtained from commercial sources
should be shipped in well-oxygenated water in insulated
containers to maintain temperature during shipment. Tem-
perature and dissolved oxygen of the water in the shipping
containers should be measured on arrival to determine if
the organisms might have been subjected to low dis-
solved oxygen or temperature fluctuations. The tempera-
ture of the shipped water should be gradually adjusted to
the desired culture temperature at a rate not exceeding
2°C per 24 h. Additional reference-toxicity testing is sug-
gested if organisms are not cultured at the testing labora-
tory (Section 9.16).
10.2.4 A group of organisms should not be used for a test
if they appear to be unhealthy, discolored, or otherwise
stressed (e.g., >20% mortality for48 h before the start of
a test). If the organisms fail to meet these criteria, the
entire batch should be discarded and a new batch should
be obtained. All organisms should be as uniform as
possible in age and life stage. Test organisms should be
handled as little as possible. When handling is necessary,
it should be done as gently, carefully, and as quickly as
possible.
10.2.5 H. azteca, C. tentans, and L. variegatus can be
cultured in a variety of waters. Water of a quality sufficient
to culture fathead minnows (Pimephales promelas) or
cladocerans will generally be adequate.
10.2.5.1 Variable success has been reported using re-
constituted waters to culture or test H. azteca in long-term
exposures (i.e., >10d; See Section 7.1.3 for details).
10.2.5.2 Organisms can be cultured using either static or
renewal procedures. Renewal of water is recommended to
limit loss of the culture organisms from a drop in dis-
solved oxygen or a buildup of waste products. In renewal
systems, there should be at least one volume addition/d
of culture water to each chamber. In static systems, the
overlying water volume should be changed at least weekly
by siphoning down to a level just above the substrate and
slowly adding fresh water. Extra care should be taken to
ensure that proper water quality is maintained in static
systems. For example, aeration is needed in static sys-
tems to maintain dissolved oxygen at >2.5 mg/L.
10.2.5.3 A recirculating system using an under-gravel
filter has been used to culture amphipods and midges
(P.V. Winger, USGS, Athens, GA, personal communica-
tion). The approach for using a recirculating system to
culture organisms has been described by New et al.
(1974), Crandall etal. (1981), and Rottmann and Campton
(1989). Under-gravel filters can be purchased from
aquarium suppliers and consist of an elevated plate with
holes that fit on the bottom of an aquarium. The plate has
a standpipe to which a pump can be attached. Gravel or
40
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an artificial substrate (e.g., plastic balls or multi-plate
substrates) is placed on the plate. The substrates provide
surface area for microorganisms that use nitrogenous
compounds. A simple example of a recirculating system
is two aquaria positioned one above the other with a total
volume of 120 L. The bottom aquarium contains the
under-gravel filter system, gravel, or artificial substrate,
and a submersible pump. The top aquarium is used for
culture of animals and has a hole in the bottom with a
standpipe for returning overflow water to the bottom
aquarium. Water lost to evaporation is replaced weekly,
and water is replaced at one- to two-month intervals.
Cultures fed foods such as Tetramin® or Tetrafin® should
include limestone gravel to help avoid depression in pH.
Recirculating systems require less maintenance than static
systems.
10.2.6 Cultures should be maintained at 23°C with a
16L8D photoperiod at an illuminance of about 100 to 1000
lux (USEPA, 1994a; ASTM, 1999a). Cultures should be
observed daily. Water temperature should be measured
daily or continuously, and dissolved oxygen should be
measured weekly. It may be desirable for laboratories to
periodically perform 96-h water-only reference-toxicity tests
to assess the sensitivity of culture organisms (Section
9.16.2). Data from these reference-toxicity tests could be
used to assess genetic strain or life-stage sensitivity to
select chemicals. The previous requirement for laborato-
ries to conduct monthly reference-toxicity tests (USEPA
1994a) has not been included as a requirement in this
second edition fortesting sediments due to the inability of
reference-toxicity tests to identify stressed populations of
test organisms (Section 9.16; McNulty et al., 1999).
Culture water hardness, alkalinity, ammonia, and pH should
be measured at least quarterly. If amphipods are cultured
using static conditions, it is desirable to measure water
quality more frequently. If reconstituted water is used to
culture organisms, water quality should be measured on
each batch of reconstituted water. Culture procedures
should be evaluated and adjusted as appropriate to re-
store or maintain the health of the culture.
10.3 Culturing Procedures for Hyalella
azteca
10.3.1 The culturing procedures described below are
based on methods described in USEPA (1991 a), Ankley
et al. (1994a), Call et al. (1994), Tomasovic et al. (1994),
Greer (1993), Ingersoll and Nelson (1990), Ingersoll et al.
(1998), ASTM (1999a) and USEPA (1994a). The culturing
procedure must produce 7- to 14-d-old amphipods to start
a 10-d sediment test (Table 11.3). The 10-d test with H.
azteca should start with a narrow range in size or age of
H. azteca (1- to 2-d range in age) to reduce potential
variability in growth at the end of the 10-d test. This
narrower range would be easiest to obtain using known-
age organisms (i.e., Section 10.3.2, 10.3.4) instead of
sieving the cultures (Section 10.3.5) to obtain similar-
sized amphipods (i.e., amphipods within a range of 1-to
2-d old will be more uniform in size than organisms within
the range of 7 d). The culturing procedure must produce
7- to 8-d-old amphipods to start a long-term test with H.
azteca (Table 14.3).
10.3.2 The following procedure described by Call et al.
(1994) and USEPA (1991 a) can be used to obtain known-
age amphipods to start a test. Mature amphipods
(50 organisms >30-d old at 23°C) are held in 2-L glass
beakers containing 1 L of aerated culture water and cotton
gauze as a substrate. Amphipods are fed 10 ml of a
yeast-Cerophyl®-trout chow (YCT) mixture (Appendix B)
and 10 ml of the green algae Selenastrum capricornutum
(about 3.5 x107cells/ml_). Five ml of each food is added
to each culture daily, except for renewal days, when
10 ml of each food is added.
10.3.2.1 Water in the culture chambers is changed
weekly. Survival of adults and juveniles and production of
young amphipods should be measured at this time. The
contents of the culture chambers are poured into a trans-
lucent white plastic or white enamel pan. Afterthe adults
are removed, the remaining amphipods will range in age
from <1 - to 7-d old. Young amphipods are transferred with
a pipet into a 1 -L beaker containing culture water and are
held for one week before starting a toxicity test. Organ-
isms are fed 10 ml of YCT and 10 ml of green algae on
start-up day, and 5 ml of each food each following day
(Appendix B). Survival of young amphipods should be
>80% during this one-week holding period. Records should
be kept on the number of surviving adults, number of
breeding pairs, and young production and survival. This
information can be used to develop control charts that are
useful in determining whether cultures are maintaining a
vigorous reproductive rate indicative of culture health.
Some of the adult amphipods can be expected to die in
the culture chambers, but mortality greater than about
50% should be cause for concern. Reproductive rates in
culture chambers containing 60 adults can be as high as
500 young per week. A decrease in reproductive rate may
be caused by a change in water quality, temperature, food
quality, or brood stock health. Adult females will continue
to reproduce for several months.
10.3.3 A second procedure for obtaining known-age
amphipods is described by Borgmann et al. (1989). Known-
age amphipods are cultured in 2.5-L chambers containing
about 1 L of culture water and between 5 and 25 adult
H. azteca. Each chamber contains pieces of cotton gauze
presoaked in culture water. Once a week the test organ-
isms are isolated from the gauze and collected using a
sieve. Amphipods are then rinsed into petri dishes where
the young and adults are sorted. The adults are returned
to the culture chambers containing fresh water and food.
10.3.4 A third procedure for obtaining known-age amphi-
pods is described by Greer (1993), Tomasovic et al.
(1994), and Ingersoll et al. (1998). Mass cultures of
mixed-age amphipods are maintained in 80-L glass aquaria
containing about 50 L of water (Ingersoll and Nelson,
1990). A flaked food (e.g., Tetrafin®) is added to each
culture chamber receiving daily water renewals to provide
about 20 g dry solids/50 L of water twice weekly in an 80-L
culture chamber. Additional flaked food is added when
41
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most of the flaked food has been consumed. Laboratories
using static systems should develop lower feeding rates
specific to their systems. Each culture chamber has a
substrate of maple leaves and artificial substrates (six
20-cm diameter sections per 80-L aquaria of nylon
"coiled-web material"; 3-M, St. Paul, MN). Before use,
leaves are soaked in 30%o salt water for about 30 d to
reduce the occurrence of planaria, snails, or other organ-
isms in the substrate. The leaves are then flushed with
water to remove the salt water and residuals of naturally
occurring tannic acid before placement in the cultures.
10.3.4.1 To obtain known-age amphipods, a U.S. Stan-
dard Sieve #25 (710-um mesh) is placed underwater in a
chamber containing mixed-age amphipods. A #25 sieve
will retain mature amphipods, and immature amphipods
will pass through the mesh. Two or three pieces of
artificial substrate (3-M coiled-web material) or a mass of
leaves with the associated mixed-age amphipods are
quickly placed into the sieve. The sieve is brought to the
top of the water in the culture chamber keeping all but
about 1 cm of the sieve underwater. The artificial sub-
strates or leaves are then shaken under water several
times to dislodge the attached amphipods. The artificial
substrates or leaves are taken out of the sieve and placed
back in the culture chamber. The sieve is agitated in the
waterto rinse the smaller amphipods back into the culture
chamber. The larger amphipods remaining in the sieve are
transferred with a pipet into a dish and then placed into a
shallow glass pan (e.g., pie pan) where immature amphi-
pods are removed. The remaining mature amphipods are
transferred using a pipet into a second #25 sieve which is
held in a glass pan containing culture water.
10.3.4.2 The mature amphipods are left in the sieve in the
pan overnight to collect any newborn amphipods that are
released. After 24 h, the sieve is moved up and down
several times to rinse the newborn amphipods (<24-h old)
into the surrounding water in the pan. The sieve is re-
moved from the pan, and the mature amphipods are
placed back into their culture chamber or placed in a
second pan containing culture water if additional organ-
isms are needed fortesting. The newborn amphipods are
moved with a pipet and placed in a culture chamber with
flowing water during a grow-out period. The newborn am-
phipods should be counted to determine if adequate num-
bers have been collected for the test.
10.3.4.3 Isolation of about 1500 (750 pairs) adults in
amplexus provided about 800 newborn amphipods in 24 h
and required about six man-hours of time. Isolation of
about 4000 mixed-age adults (some in amplexus and
others not in amplexus) provided about 800 newborn
amphipods in 24 h and required less than one man-hour of
time. The newborn amphipods should be held for 6 to 13 d
to provide 7- to 14-d-old organisms to start a 10-d test
(Section 11) or should be held for 7 d to provide 7- to
8-d-old organisms to start a long-term test (Section 14).
The neonates are held in a 2-L beaker for 6 to 13 d before
the start of a test. On the first day of isolation, the
neonates are fed 10 ml of YCT (1800 mg/L stock solu-
tion) and 10 ml of Selenastrum capricornutum (about
3.5x 107 cells/ml). On the third, fifth, seventh, ninth,
eleventh, and thirteenth days after isolation, the amphi-
pods are fed 5 ml of both YCT and S. capricornutum.
Amphipods are initially fed a higher volume to establish a
layer of food on the bottom of the culture chamber. If
dissolved oxygen drops below 4 mg/L, about 50% of the
water should be replaced (Ingersolletal., 1998).
10.3.5 Laboratories that use mixed-age amphipods for
testing must demonstrate that the procedure used to
isolate amphipods will produce test organisms that are
7-to 14-d old. For example, amphipods passing through a
U.S. Standard #35 sieve (500 urn), but stopped by a
#45 sieve (355 urn) averaged 1.54 mm (SD 0.09) in length
(P.V. Winger, USGS, Athens, GA, unpublished data). The
mean length of these sieved organisms corresponds to
that of 6-d-old amphipods (Figure 10.1). After holding for
3 d before testing to eliminate organisms injured during
sieving, these amphipods would be about 9 d old (length
1.84 mm, SD 0.11) at the start of a toxicity test.
10.3.5.1 Ingersoll and Nelson (1990) describe the follow-
ing procedure for obtaining mixed-age amphipods of a
similar size to start a test. Smaller amphipods are iso-
lated from larger amphipods using a stack of U.S. Stan-
dard sieves: #30 (600 urn), #40 (425 urn), and #60 (250 urn).
Sieves should be held under water to isolate the amphi-
pods. Amphipods may float on the surface of the water if
they are exposed to air. Artificial substrate or leaves are
placed in the #30 sieve. Culture water is rinsed through
the sieves and small amphipods stopped by the #60 sieve
are washed into a collecting pan. Larger amphipods in the
#30 and #40 sieves are returned to the culture chamber.
The smaller amphipods are then placed in 1-L beakers
containing culture water and food (about 200 amphipods
per beaker) with gentle aeration.
10.3.5.2 Amphipods should be held and fed at a rate
similar to the mass cultures for at least 2 d before the
start of a test to eliminate animals injured during handling.
10.3.6 See Section 10.2.6 for procedures used to evalu-
ate the health of cultures.
10.4 Culturing Procedures for
Chironomus tentans
10.4.1 The culturing methods described below are based
on methods described in USEPA (1991 a), Ankley et al.
(1994a), Call et al. (1994), Greer (1993), ASTM (1999a),
and USEPA (1994a). A C. tentans 10-d survival and
growth test must be started with second- to third-instar
larvae (about 10-d-old larvae; Section 12; Figure 10.2). At
a temperature of 23°C, larvae should develop to the third
instar by 9 to 11 d after hatching (about 11 to 13 d
post-oviposition). The instar of midges at the start of a
test can be determined based on head capsule width
(Table 10.2) or based on weight or length at sediment test
initiation. Average length of midge larvae should be 4 to 6
mm, while average dry weight should be 0.08 to 0.23 mg/
individual. A C. tentans long-term test must be started
with larvae less than 24 h old (see Section 15.3 for a
42
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4
E
E
O)
c
CD
0
Sizu retained on 555 p-n siuve ci*te' passiny 500 jrf sieve
01 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16
Day
Figure 10.1 Mean length (+/- 2SD) and relative age of Hya lei la azteca collected by sieving in comparison with length of
known-age organisms. P.V. Winger, USGS, Athens, GA, unpublished data.
Thoracic
Segments
Table 10.2
Chironomus tentans Instar and Head Capsule
Widths1
Head
Capsule
Figure 10.2. Chironomus tentans larvae. Note thoracic segments
which are used to measure instars. (Reprinted from
Clifford, 1991 with kind permission from the Univer-
sity of Alberta Press.)
description of an approach for obtaining C. tentans larvae
less than 24 h old).
10.4.2 Historically, third-instarC. tentans we re frequently
referred to as the second instar in the published literature.
When C. tentans larvae were measured daily, the
C. tentans raised at 22°C to 24°C were third instar, not
second instar, by 9 to 11 d after hatching (T.J.
Norberg-King, USEPA, Duluth, MN, unpublished data).
10.4.3 Both silica sand and shredded paper toweling
have been used as substrates to culture C. tentans.
Either substrate may be used if a healthy culture can be
maintained. Greer(1993) used sand or paper toweling to
culture midges; however, sand was preferred due to the
Instar
First
Second
Third
Fourth
Days after
hatching
1 to 4.4
4.4 to 8.5
8.5 to 12.5
>12.5
Mean (mm)
0.10
0.20
0.38
0.67
Range (mm)
0.09 to 0.1 3
0.1 8 to 0.23
0.33 to 0.45
0.63 to 0.71
1 T.J. Norberg-King, USEPA, Duluth, MN, unpublished data.
ease in removing larvae for testing. Sources of sand are
listed in Section 7.
10.4.3.1 Papertowels are prepared according to a proce-
dure adapted from Batac-Catalan and White (1982). Plain
white kitchen papertowels are cut into strips. Cut toweling
is loosely packed into a blender with culture water and
blended for a few seconds. Small pieces should be
available to the organism; blending fortoo long will result
in a fine pulp that will not settle in a culture tank. Blended
towels can then be added directly to culture tanks, elimi-
nating any conditioning period for the substrate. A mass
of the toweling sufficient to fill a 150-mL beaker is placed
into a blender containing 1 L of deionized water, and
blended for 30 sec or until the strips are broken apart in
the form of a pulp. The pulp is then sieved using a 710-um
43
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sieve and rinsed well with deionized water to remove the
shortest fibers.
10.4.3.2 Dry shredded papertoweling loosely packed into
a 2-L beaker will provide sufficient substrate for about ten
19-L chambers (USEPA, 1991 a). The shredded toweling
placed in a 150-mL beaker produces enough substrate for
one 19-L chamber. Additional substrate can be frozen in
deionized water for later use.
10.4.4 Five egg cases will provide a sufficient number of
organisms to start a new culture chamber. Egg cases
should be held at 23°C in a glass beaker or crystallizing
dish containing about 100 to 150 ml of culture water
(temperature change should not exceed 2°C perd). Food
is not added until the embryos start to hatch (in about 2 to
4dat23°C)to reduce the risk of oxygen depletion. About
200 to 400 larvae are then placed into each culture
chamber. Crowding of larvae will reduce growth. See
Section 10.4.5.1 or10.4.6.1 for a description of feeding
rates. Larvae should reach the third instar by about 10 d
after median hatch (about 12 to 14 d after the time the
eggs were laid; Table 10.2).
10.4.5 Chironomus tentans are cultured in soft water at
the USEPA laboratory in Duluth (USEPA, 1993c) in glass
aquaria (19.0-L capacity, 36x21 x26 cm high). A water
volume of about 6 to 8 L in these flow-through chambers
can be maintained by drilling an overflow hole in one end
11 cm from the bottom. The top of the aquarium is
covered with a mesh material to trap emergent adults.
Pantyhose with the elasticized waist is positioned around
the chamber top and the legs are cut off. Fiberglass-
window screen glued to a glass strip (about 2- to 3-cm
wide) rectangle placed on top of each aquarium has also
been used by Call et al. (1994). About 200 to 300 ml of
40-mesh silica sand is placed in each chamber.
10.4.5.1 The stocking density of the number of C.
tentans eggs should be about 600 eggs per 6 to 8 L of
water. Dawson et al. (1999) found that the cultures in 15-
L aquaria and 7 L of waterwere self-regulating in density
regardless of the initial number of eggs stocked in each
tank. However, tanks with a higher initial stocking
density (i.e., 1400 eggs/tank) increased the time of peak
adult emergence to 30 to 33 d, whereas tanks with lower
stocking densities (600 or 1000 eggs/tank) had peak
emergence at 22 to 25 d after hatching.
10.4.5.2 Fish food flakes (i.e., Tetrafin®) are added to
each culture chamber to provide a final food concentra-
tion of about 0.04 mg dry solids/mL of culture water. A
stock suspension of the solids is blended with distilled
water to form an initial slurry. It is then filtered through
a 200-micron Nitex screen and diluted with distilled water
to form a 56 g dry solids/L final slurry (Appendix B). The
larvae in each tank are fed 2.5 ml of slurry (140 mg of
Tetrafin per day) from Day 0 to Day 7 and 5 ml of slurry
(280 mg Tetrafin per day) from Day 8 on. Feeding is done
afterthe water renewal process is completed. The stock
suspension should be well mixed immediately before
removing an aliquot for feeding. Each batch of food
should be refrigerated and can be used for up to two
weeks (Appendix B). Laboratories using static systems
should develop lower feeding rates specific to their
systems.
10.4.6 Chironomus tentans are cultured by Greer (1993)
in Rubbermaid® 5.7-L polyethylene cylindrical containers.
The containers are modified by cutting a semicircle into
the lid 17.75 cm across by 12.5 cm. Stainless-steel
screen (20 mesh/0.4 cm) is cut to size and melted to the
plastic lid. The screen provides air exchange, retains
emerging adults, and is a convenient way to observe the
culture. Two holes about 0.05 cm in diameter are drilled
through the uncut portion of the lid to provide access for
an air line and to introduce food. The food access hole is
closed with a No. 00 stopper. Greer (1993) cultures midges
under static conditions with moderate aeration, and about
90% of the water is replaced weekly. Each 5.7-L culture
chamber contains about 3 L of water and about 25 mL of
fine sand. Eight to 10 chambers are used to maintain
the culture.
10.4.6.1 Midges in each chamber are fed 6 mL/d of a
100 g/L suspension offish food flakes (e.g., Tetrafin®) on
Tuesday, Wednesday, Thursday, Friday, and Sunday. A
6-mL chlorella suspension (deactivated "Algae-Feast®
Chlorella," Earthrise Co., Callpatria, CA) is added to each
chamber on Saturday and on Monday. The chlorella sus-
pension is prepared by adding 5 g of dry chlorella
powder/L of water. The mixture should be refrigerated and
can be used for up to two weeks.
10.4.6.2 The water should be replaced more often if
animals appear stressed (e.g., at surface or pale color at
the second instar) or if the water is cloudy. Water is
replaced by first removing emergent adults with an aspira-
tor. Any growth on the sides of the chamber should be
brushed off before water is removed. Care should be
taken not to pour or siphon out the larvae when removing
the water. Larvae will typically stay near the bottom;
however, a small-mesh sieve or nylon net can be used to
catch any larvae that float out. Afterthe chambers have
been cleaned, temperature-adjusted culture water is poured
back into each chamber. The water should be added
quickly to stir up the larvae. Using this procedure, the
approximate size, number, and the general health of the
culture can be observed.
10.4.7 Adult emergence will begin about three weeks
after hatching at 23°C. Once adults begin to emerge, they
can be gently siphoned into a dry aspirator flask on a daily
basis. An aspirator can be made using a 250- or 500-mL
Erlenmeyer flask, a two-hole stopper, some short sec-
tions of 0.25-inch glass tubing, and Tygon® tubing for
collecting and providing suction (Figure 10.3). Adults
should be aspirated with short inhalations to avoid injuring
the organisms. The mouthpiece on the aspirator should
be replaced or disinfected between use. Sex ratio of the
adults should be checked to ensure that a sufficient
number of males are available for mating and fertilization.
One male may fertilize more than one female. However, a
44
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Tygon Tubing
500 ml Erlenmeyer
Mesh Cover
Nitex Screen
Water
Figure 10.3 Aspirator chamber (A) and reproduction and oviposit chamber (B) for adult midges.
45
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ratio of one male to three females improves fertilization
success.
10.4.7.1 A reproduction and oviposit chamber may be
prepared in several different ways (Figure 10.3). Culture
water (about 50 to 75 ml) can be added to the aspiration
flask in which the adults were collected (Figure 10.3;
Batac-Catalan and White, 1982). The USEPA Office of
Research and Development Laboratory (Duluth, MN;
USEPA, 1991 a) uses a 500-mL collecting flask with a
length of Nitex® screen positioned vertically and extend-
ing into the culture water (Figure 10.3). The Nitex® screen
is used by the females to position themselves just above
the water during oviposition. The two-hole stopper and
tubing of the aspirator should be replaced by screened
material or a cotton plug for good air exchange in the
oviposition chamber.
10.4.7.2 Greer (1993) uses an oviposition box to hold
emergent adults. The box is constructed of a 5.7-L cham-
ber with a 20-cm tall cylindrical chamber on top. The top
chamber is constructed of stainless-steel screen (35 mesh/
2.54 cm) melted onto a plastic lid with a 17.75-cm hole. A
5-cm hole is cut into the side of the bottom chamber and a
#11 stopper is used to close the hole. Egg cases are
removed by first sliding a piece of plexiglass between the
top and bottom chambers. Adult midges are then aspi-
rated from the bottom chamber. The top chamber with
plexiglass is removed from the bottom chamber and a
forceps is used to remove the egg cases. The top cham-
ber is put back on top of the bottom chamber, the plexiglass
is removed, and the aspirated adults are released from
the aspirator into the chamberthrough the 5-cm hole.
10.4.8 About two to three weeks before the start of a test,
at least 3 to 5 egg cases should be isolated for hatching
using procedures outlined in Section 10.4.4.
10.4.9 Records should be kept on the time to first
emergence and the success of emergence for each cul-
ture chamber. It is also desirable to monitor growth and
head capsule width periodically in the cultures. See Sec-
tion 10.2.6 for additional detail on procedures for evaluat-
ing the health of the cultures.
10.5 Culturing Procedures for
Lumbriculus variegatus
10.5.1 The culturing procedures described below are
based on methods described in Phipps et al. (1993),
USEPA (1991 a), Call et al. (1994), Brunson et al. (1998),
and USEPA (1994a). Bioaccumulation tests are started
with adult organisms.
10.5.2 Lumbriculus variegatus are generally cultured with
daily renewal of water (57- to 80-L aquaria containing 45 to
50 L of water).
10.5.3 Paper towels can be used as a substrate for
culturing L. variegatus (Phipps et al., 1993). Substrate is
prepared by cutting unbleached brown paper towels into
strips either with a paper shredder or with scissors. Cut
toweling is loosely packed into a blender with culture
water and blended for a few seconds. Small pieces
should be available to the organisms; blending fortoo long
will result in a fine pulp that will not settle in culture tanks.
Blended towels can then be added directly to culture
tanks, eliminating any conditioning period forthe substrate.
The papertowel substrate is renewed with blended towels
when thin or bare areas appear in the cultures. The
substrate in the chamber will generally last for about two
months.
10.5.4 Oligochaetes probably obtain nourishment from
ingesting the organic matter in the substrate (Pennak,
1989). Lumbriculus variegatus in each of the culture
chambers are fed a 10-mL suspension of 6 g of trout
starter 3 times/week. The particles will temporarily disperse
on the surface film, break through the surface tension,
and settle out over the substrate. Laboratories using
static systems should develop lower feeding rates spe-
cific to their systems. Food and substrate used to culture
oligochaetes should be analyzed for compounds to be
evaluated in bioaccumulation tests. If the concentration
of the test compound is above the detection level and the
food is not measured, the test may be invalidated. Recent
studies in other laboratories, for example, have indicated
elevated concentrations of PCBs in substrate and/or food
used for culturing the oligochaete (J. Amato, AScI Corpo-
ration, Duluth, MN, personal communication).
10.5.5 Phipps et al. (1993) recommend starting a new
culture with 500 to 1000 worms. Conditioned paper towel-
ing should be added when the substrate in a culture
chamber is thin.
10.5.6 On the day before the start of a test, oligochaetes
can be isolated by transferring substrate from the cultures
into a beaker using a fine-mesh net. Additional organisms
can be removed using a glass pipet (20-cm long, 5-mm
i.d.; Phipps etal., 1993). Water can be slowly trickled into
the beaker. The oligochaetes will form a mass and most
of the remaining substrate will be flushed from the beaker.
On the day the test is started, organisms can be placed in
glass or stainless-steel pans. A gentle stream of water
from the pipet can be used to spread out clusters of
oligochaetes. The remaining substrate can be siphoned
from the pan by allowing the worms to reform in a cluster
on the bottom of the pan. For bioaccumulation tests,
aliquots of worms to be added to each test chamber can
be transferred using a blunt dissecting needle or dental
pick. Excess water can be removed during transfer by
touching the mass of oligochaetes to the edge of the pan.
The mass of oligochaetes is then placed in a tared weigh
boat, quickly weighed, and immediately introduced into
the appropriate test chamber. Organisms should not be
blotted with a papertowel to remove excess water (Brunson
etal.,1998).
10.5.7 The culture population generally doubles (number
of organisms) in about 10 to 14d. See Section 10.2.6 for
additional detail on procedures for evaluating the health of
the cultures.
46
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Section 11
Test Method 100.1
Hyalella azteca 10-d Survival and Growth Test for Sediments
11.1 Introduction
11.1.1 Hyalella azteca (Saussure) have many desirable
characteristics of an ideal sediment toxicity testing organ-
ism including relative sensitivity to contaminants associ-
ated with sediment, short generation time, contact with
sediment, ease of culture in the laboratory, and tolerance
to varying physico-chemical characteristics of sediment.
Their response has been evaluated in interlaboratory studies
and has been confirmed with natural benthic populations.
Many investigators have successfully used H. azteca to
evaluate the toxicity of freshwater sediments (e.g., Nebeker
et al., 1984a; Borgmann and Munwar, 1989; Ingersoll and
Nelson, 1990; Ankley et al., 1991 a; Ankley et al., 1991b;
Burton etal., 1989; Winger and Lasier, 1993; Kemble et
al., 1994). H. azteca has been used for a variety of
sediment assessments (Ankley et al., 1991; West et al.,
1993; Hokeetal., 1994,1995; West etal., 1994). Hyalella
azteca can also be used to evaluate the toxicity of
estuarine sediments (up to 15 %o salinity; Nebeker and
Miller, 1988; Roach et al., 1992; Winger et al., 1993).
Endpoints typically monitored in 10-d sediment toxicity
tests with H. azteca include survival and growth.
11.1.2 A test method for conducting a 10-d sediment
toxicity test is described in Section 11.2 for H. azteca.
Methods outlined in Appendix A of USEPA(1994a) and in
Section 11.1.1 were used for developing test method
100.1. Results of tests using procedures different from
the procedures described in Section 11.2 may not be
comparable, and these different procedures may alter
contaminant bioavailability. Comparison of results ob-
tained using modified versions of these procedures might
provide useful information concerning new concepts and
procedures for conducting sediment tests with aquatic
organisms. If tests are conducted with procedures differ-
ent from the procedures described in this manual, addi-
tional tests are required to determine comparability of
results (Section 1.3).
11.2 Recommended Test Method for
Conducting a 10-d Sediment Toxicity
Test with Hyalella azteca
11.2.1 Recommended conditions for conducting a 10-d
sediment toxicity test with H. azteca are summarized in
Table 11.1. A general activity schedule is outlined in
Table 11.2. Decisions concerning the various aspects of
experimental design, such as the number of treatments,
number of test chambers/treatment, and water-quality
characteristics should be based on the purpose of the test
and the methods of data analysis (Section 16). The
number of replicates and concentrations tested depends
in part on the significance level selected and the type of
statistical analysis. When variability remains constant,
the sensitivity of a test increases as the number of
replicates increase.
11.2.2 The recommended 10-d sediment toxicity test
with H. azteca must be conducted at 23°C with a 16L8D
photoperiod at an illuminance of about 100 to 1000 lux
(Table 11.1). Test chambers are 300-mL high-form lipless
beakers containing 100 ml of sediment and 175 ml of
overlying water. Ten 7- to 14-d-old amphipods are used to
start a test. The 10-d test should start with a narrow range
in size or age of/-/, azteca (i.e., 1-to 2-d range in age) to
reduce potential variability in growth at the end of a 10-d
test (Section 10.3.1). The number of replicates/treatment
depends on the objective of the test. Eight replicates are
recommended for routine testing (Section 16). Amphipods
in each test chamber are fed 1.0 ml of YCT food daily
(Appendix B). The first edition of the manual (USEPA,
1994a) recommended a feeding level of 1.5 ml of YCT
daily; however, this feeding level was revised to 1.0 ml to
be consistent, with the feeding level in the long-term test
with H. azteca (Section 14). Each chamber re-
ceives 2 volume additions/d of overlying water. Water
renewals may be manual or automated. Appendix A
describes water-renewal systems that can be used to
deliver overlying water. Overlying water can be culture
water, well water, surface water, site water, or reconsti-
tuted water. For site-specific evaluations, the characteris-
tics of the overlying water should be as similar as pos-
sible to the site where sediment is collected. Require-
ments fortest acceptability are summarized in Table 11.3.
11.3 General Procedures
11.3.1 Sediment into Test Chambers
11.3.1.1 The day before the sediment test is started
(Day -1) each sediment should be thoroughly homog-
enized and added to the test chambers (Section 8.3.1).
Sediment should be visually inspected to judge the de-
gree of homogeneity. Excess water on the surface of the
sediment can indicate separation of solid and liquid
components. If a quantitative measure of homogeneity is
47
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Parameter
Table 11.1 Test Conditions for Conducting a 10-d Sediment Toxicity Test with Hyalella azteca
Conditions
1. Test type:
2. Temperature:
3. Light quality:
4. Illuminance:
5. Photoperiod:
6. Test chamber:
7. Sediment volume:
8. Overlying water volume:
9. Renewal of overlying water:
10. Age of organisms:
11. Number of organisms/chamber:
12. Number of replicate chambers/treatment:
13. Feeding:
14. Aeration:
15. Overlying water:
16. Test chamber cleaning:
17. Overlying water quality:
18. Test duration:
19. Endpoints:
20. Test acceptability:
Whole-sediment toxicity test with renewal of overlying water
23 ± 1 °C
Wide-spectrum fluorescent lights
About 100 to 1000 lux
16L8D
300-mL high-form lipless beaker
100mL
175 ml
2 volume additions/d (Appendix A); continuous or intermittent (e.g., 1 volume
addition every 12 h)
7- to 14-d old at the start of the test (1- to 2-d range in age)
10
Depends on the objective of the test. Eight replicates are recommended for routine
testing (see Section 16).
YCT food, fed 1.0 ml daily (1800 mg/L stock) to each test chamber. The first
edition of the manual (USEPA, 1994a) recommended a feeding level of 1.5 ml of
YCT daily; however, this feeding level was revised to 1.0 ml to be consistent with
the feeding level in the long-term tests with H. azteca (Section 14).
None, unless dissolved oxygen in overlying water drops below 2.5 mg/L.
Culture water, well water, surface water, site water, or reconstituted water
If screens become clogged during a test, gently brush the outside of the screen
(Appendix A).
Hardness, alkalinity, conductivity, pH, and ammonia at the beginning and end of a
test. Temperature and dissolved oxygen daily.
10d
Survival and growth
Minimum mean control survival of 80% and measurable growth of test organisms in
the control sediment. Additional performance-based criteria specifications are
outlined in Table 11.3.
Day
Table 11.2 General Activity Schedule for Conducting a 10-d Sediment Toxicity Test with Hyalella azteca 1
Activity
-7 Separate known-age amphipods from the cultures and place in holding chambers. Begin preparing food for the test. There
should be a 1- to 2-d range in age of amphipods used to start the test.
-6 to -2 Feed and observe isolated amphipods (Section 10.3), monitor water quality (e.g., temperature and dissolved oxygen).
-1 Feed and observe isolated amphipods (Section 10.3), monitor water quality. Add sediment into each test chamber, place
chambers into exposure system, and start renewing overlying water.
0 Measure total water quality (pH, temperature, dissolved oxygen, hardness, alkalinity, conductivity, ammonia). Transfer 10
7- to 14-day-old amphipods into each test chamber. Release organisms under the surface of the water. Add 1.0 ml of YCT
into each test chamber. Archive 20 test organisms for length determination or archive 80 test organisms for dry weight
determination. Observe behavior of test organisms.
1 to 8 Add 1.0 ml of YCT food to each test chamber. Measure temperature and dissolved oxygen. Observe behavior of test
organisms.
9 Measure total water quality.
10 Measure temperature and dissolved oxygen. End the test by collecting the amphipods with a sieve (Section 11.3.7.1).
Count survivors and prepare organisms for weight or length measurements.
1 Modified from Call etal., 1994
48
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Table 11.3 Test Acceptability Requirements for a 10-d Sediment Toxicity Test with Hyalella azteca
A. It is recommended for conducting a 10-d test with Hyalella azteca that the following performance criteria be met:
1. Age of H. azteca at the start of the test must be between 7- to 14-d old. The 10-d test should start with a narrow range in size or
age of H. azteca (i.e., 1- to 2-d range in age) to reduce potential variability in growth at the end of a 10-d test (Section 10.3.1).
2. Average survival of H. azteca in the control sediment must be greater than or equal to 80% at the end of the test. Growth of test
organisms should be measurable in the control sediment at the end of the 10-d test (i.e., relative to organisms at the start of the
test).
3. Hardness, alkalinity, and ammonia in the overlying water typically should not vary by more than 50% during the test, and dissolved
oxygen should be maintained above 2.5 mg/L in the overlying water.
B. Performance-based criteria for culturing H. azteca include the following:
1. It may be desirable for laboratories to periodically perform 96-h water-only reference-toxicity tests to assess the sensitivity of
culture organisms (Section 9.16.2). Data from these reference-toxicity tests could be used to assess genetic strain or life-stage
sensitivity of test organisms to select chemicals.
2. Laboratories should track parental survival in the cultures and record this information using control charts if known-age cultures are
maintained. Records should also be kept on the frequency of restarting cultures and the age of brood organisms.
3. Laboratories should record the following water-quality characteristics of the cultures at least quarterly: pH, hardness, alkalinity, and
ammonia. Dissolved oxygen in the cultures should be measured weekly. Temperature of the cultures should be recorded daily. If
static cultures are used, it may be desirable to measure water quality more frequently.
4. Laboratories should characterize and monitor background contamination and nutrient quality of food if problems are observed in
culturing or testing organisms.
5. Physiological measurements such as lipid content might provide useful information regarding the health of the cultures.
C. Additional requirements:
1. All organisms in a test must be from the same source.
2. Storage of sediments collected from the field should follow guidance outlined in Section 8.2.
3. All test chambers (and compartments) should be identical and should contain the same amount of sediment and overlying water.
4. Negative-control sediment and appropriate solvent controls must be included in a test. The concentration of solvent used must not
adversely affect test organisms.
5. Test organisms must be cultured and tested at 23°C (±1°C).
6. The daily mean test temperature must be within ±1°C of 23°C. The instantaneous temperature must always be within ±3°C of 23°C.
7. Natural physico-chemical characteristics of test sediment collected from the field should be within the tolerance limits of the test
organisms.
required, replicate subsamples should betaken from the than 10%. Hardness, alkalinity and ammonia
sediment batch and analyzed forTOC, chemical concen- concentrations in the water above the sediment, within a
trations, and particle size. treatment, typically should not vary by more than 50%
during the test. Mount and Brungs (1967) diluters have
11.3.1.2 Each test chamber should contain the same been modified for sediment testing, and other automated
amount of sediment, determined either by volume or by water-delivery systems have also been used (Maki, 1977;
weight. Overlying water is added to the chambers on Ingersoll and Nelson, 1990; Benoitetal., 1993; Zumwalt
Day-1 in a manner that minimizes suspension of sedi- et al., 1994; Brunson et al., 1998; Wall et al., 1998;
ment. This can be accomplished by gently pouring water Leppanen and Maier, 1998). The water-delivery system
along the sides of the chambers or by pouring water onto should be calibrated before a test is started to verify that
a baffle (e.g., a circular piece of Teflon® with a handle the system is functioning properly. Renewal of overlying
attached) placed above the sediment to dissipate the water is started on Day -1 before the addition of test
force of the water. A test begins when the organisms are organisms or food on Day 0. Appendix A describes
added to the test chambers (Day 0). water-renewal systems that can be used for conducting
sediment tests.
11.3.2 Renewal of Overlying Water
11.3.2.2 In water-renewal tests with one to four volume
11.3.2.1 Renewal of overlying water is required during a additions of overlying water/d, water-quality characteris-
test. At any particular time during the test, flow rates tics generally remain similarto the inflowing water (Ingersoll
through any two test chambers should not differ by more and Nelson, 1990; Ankley et al., 1993); however, in static
49
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tests, water quality may change profoundly during the
exposure (Shuba et al., 1978). For example, in static
whole-sediment tests, the alkalinity, hardness, and
conductivity of overlying water more than doubled in
several treatments during a four-week exposure (Ingersoll
and Nelson, 1990). Additionally, concentrations of meta-
bolic products (e.g., ammonia) may also increase during
static exposures, and these compounds can either be
directly toxic to the test organisms or may contribute to
the toxicity of the contaminants in the sediment.
Furthermore, changes in water-quality characteristics such
as hardness may influence the toxicity of many inorganic
(Gauss et al., 1985) and organic (Mayer and Ellersieck,
1986) contaminants. Although contaminant concentra-
tions are reduced in the overlying water in water-renewal
tests, organisms in direct contact with sediment generally
receive a substantial proportion of a contaminant dose
directly from either the whole sediment or from the
pore water.
11.3.3 Acclimation
11.3.3.1 Test organisms must be cultured and tested at
23°C. Ideally, test organisms should be cultured in the
same water that will be used in testing. However, acclima-
tion of test organisms to the test water is not required.
11.3.3.2 Culturing of organisms and toxicity assessment
are typically conducted at 23°C. However, occasionally
there is a need to perform evaluations at temperatures
different than that recommended. Under these circum-
stances, it may be necessary to acclimate organisms to
the desired test temperature to prevent thermal shock
when moving immediately from the culture temperature to
the test temperature (ASTM, 1999a). Acclimation can be
achieved by exposing organisms to a gradual change in
temperature; however, the rate of change should be rela-
tively slow to prevent thermal shock. A change in tem-
perature of 1 °C every 1 to 2 h has been used successfully
in some studies (P.K. Sibley, University of Guleph, Guelph,
Ontario, personal communication; APHA, 1989). Testing
at temperatures other than 23°C needs to be preceded by
studies to determine expected performance under alter-
nate conditions.
11.3.4 Placing Organisms in Test Chambers
11.3.4.1 Test organisms should be handled as little as
possible. Amphipods should be introduced into the overly-
ing water below the air-water interface. Test organisms
can be pipetted directly into overlying water. The size of
the test organisms at the start of the test should be
measured using the same measure (length orweight) that
will be used to assess their size at the end of the test. For
length, a minimum of 20 organisms should be measured.
Forweight measurement, a larger sample size (e.g., 80)
may be desirable because of the relative small mass of
the organisms. This information can be used to determine
consistency in the size of the organisms used to start a
test.
11.3.5 Feeding
11.3.5.1 For each beaker, 1.0 ml of YCT is added from
Day 0 to Day 9. Without addition of food, the test
organisms may starve during exposures. However, the
addition of the food may alter the availability of the
contaminants in the sediment (Wiederholm et al., 1987;
Harkey et al., 1994). Furthermore, if too much food is
added to the test chamber or if the mortality of test
organisms is high, fungal or bacterial growth may develop
on the sediment surface. Therefore, the amount of food
added to the test chambers is kept to a minimum.
11.3.5.2 Suspensions of food should be thoroughly mixed
before aliquots are taken. If excess food collects on the
sediment, a fungal or bacterial growth may develop on the
sediment surface, in which case feeding should be sus-
pended for one or more days. A drop in dissolved oxygen
below 2.5 mg/L during a test may indicate that the food
added is not being consumed. Feeding should be sus-
pended for the amount of time necessary to increase the
dissolved oxygen concentration (ASTM, 1999a). If feed-
ing is suspended in one treatment, it should be sus-
pended in all treatments. Detailed records of feeding rates
and the appearance of the sediment surface should be
made daily.
11.3.6 Monitoring a Test
11.3.6.1 All chambers should be checked daily and
observations made to assess test organism behavior
such as sediment avoidance. However, monitoring ef-
fects on burrowing activity of test organisms may be
difficult because the test organisms are often not visible
during the exposure. The operation of the exposure sys-
tem should be monitored daily.
11.3.6.2 Measurement of Overlying Water-quality
Characteristics
11.3.6.2.1 Conductivity, hardness, pH, alkalinity, and
ammonia should be measured in all treatments at the
beginning and end of a test. Overlying water should be
sampled just before water renewal from about 1 to 2 cm
above the sediment surface using a pipet. It may be
necessary to composite water samples from individual
replicates. The pipet should be checked to make sure no
organisms are removed during sampling of overlying water.
Water quality should be measured on each batch of water
prepared for the test.
11.3.6.2.2 Dissolved oxygen should be measured daily
and should be maintained at a minimum of 2.5 mg/L. If a
probe is used to measure dissolved oxygen in overlying
water, it should be thoroughly inspected between samples
to make sure that organisms are not attached and should
be rinsed between samples to minimize cross contamina-
tion. Aeration can be used to maintain dissolved oxygen
in the overlying water above 2.5 mg/L (i.e., about 1
bubble/second in the overlying water). Dissolved oxygen
and pH can be measured directly in the overlying water
with a probe.
50
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11.3.6.2.3 Temperature should be measured at least
daily in at least one test chamber from each treatment.
The temperature of the water bath orthe exposure cham-
ber should be continuously monitored. The daily mean
test temperature must be within ±1°C of 23°C. The
instantaneous temperature must always be within ±3°C
of23°C.
11.3.7 Ending a Test
11.3.7.1 Any of the surviving amphipods in the water
column or on the surface of the sediment can be pipetted
from the beaker before sieving the sediment. Immobile
organisms isolated from the sediment surface or from
sieved material should be considered dead. A #40 sieve
(425-um mesh) can be used to remove amphipods from
sediment. Alternatively, Kemble et al. (1994) suggest
sieving of sediment using the following procedure: (1) pour
about half of the overlying waterthrough a #50- (300-um)
U.S. standard mesh sieve, (2) swirl the remaining waterto
suspend the upper 1 cm of sediment, (3) pour this slurry
through the #50-mesh sieve and wash the contents of the
sieve into an examination pan, (4) rinse the coarser
sediment remaining in the test chamber through a #40-
(425-um) mesh sieve and wash the contents of this
second sieve into a second examination pan. Surviving
test organisms are removed from the two pans and counted.
If growth (length) is to be measured (Ingersoll and Nelson,
1990), the organisms can be preserved in 8% sugar
formalin solution. The sugarformalin solution is prepared
by adding 120 g of sucrose to 80 ml of formalin, which is
then brought to a volume of 1 L using deionized water.
This stock solution is mixed with an equal volume of
deionized water when used to preserve organisms.
NoTox® (Earth Safe Industries, Belle Mead, NJ) can be
used as a substitute for formalin (Ungeret al., 1993).
11.3.7.2 A consistent amount of time should be taken to
examine sieved material for recovery of test organisms
(e.g., 5 min/replicate). Laboratories should demonstrate
that their personnel are able to recover an average of at
least 90% of the organisms from whole sediment. For
example, test organisms could be added to control ortest
sediments, and recovery could be determined after 1 h
(Tomasovicet al., 1994).
11.3.8 Test Data
11.3.8.1 Survival and growth are measured attheendof
the 10-d sediment toxicity test with H. azteca. Growth of
amphipods is often a more sensitive toxicity endpoint
compared to survival (Burton and Ingersoll, 1994; Kemble
et al., 1994; Becker et al., 1995; Ingersoll et al., 1996;
Ingersoll etal., 1998; Steevens and Benson, 1998). The
duration of the 10-d test starting with 7- to 14-d-old
amphipods is not long enough to determine sexual matu-
ration or reproductive effects. The 42-d test (Section 14)
is designed to evaluate additional sublethal endpoints in
sediment toxicity tests with H. azteca. See Section
14.4.5.3 for a discussion of measuring dry weight vs.
length of/-/, azteca.
1st Antenna
2nd Antenna
/A
3rd Uropod
2nd Uropod
1st Uropod
Figure 11.1 Hyalella azteca. (A) denotes the uropods; (B) denotes
the base of the first antennae; (C) denotes the
gnathopod used for grasping females. Meaurement
of length is made from base of the 3rd uropod (A) to
(B). Females are recognized by the presence of egg
cases or the absence of an enlarged gnathopod.
(Reprinted from Cole and Watkins, 1997 with kind
permission from Kluwer Academic Publishers.)
11.3.8.2 Amphipod body length (±0.1 mm) can be mea-
sured from the base of the first antenna to the tip of the
third uropod along the curve of the dorsal surface
(Figure 11.1). Ingersoll and Nelson (1990) describe the
use of a digitizing system and microscope to measure
lengths of H. azteca. Kemble et al. (1994) also photo-
graphed invertebrates (at a magnification of 3.5X) and
measured length using a computer-interfaced digitizing
tablet. Antennal segment number can also be used to
estimate length or weight of amphipods (E.L. Brunson,
USGS, Columbia, MO, personal communication). Wet or
dry weight measurements have also been used to esti-
mate growth of/-/, azteca (ASTM, 1999a). If test organ-
isms are to be used for an evaluation of bioaccumulation,
it is not advisable to dry the sample before conducting the
residue analysis. If conversion from wet weight to dry
weight is necessary, aliquots of organisms can be weighed
to establish wet to dry weight conversion factors. A
consistent procedure should be used to remove the ex-
cess water from the organisms before measuring wet
weight.
11.3.8.3 Dry weight of amphipods should be determined
by pooling all living organisms from a replicate and drying
the sample at about 60°C to 90°C to a constant weight.
The sample is brought to room temperature in a desicca-
tor and weighed to the nearest 0.01 mg to obtain mean
weight per surviving organism per replicate (see Section
14.3.7.6) The first edition of this manual (USEPA,
1994a) recommended dry weight as a measure of growth
for both H. azteca and C. tentans. For C. tentans, this
recommendation was changed in the current edition to
ash-free dry weight (AFDW) instead of dry weight, with the
intent of reducing bias introduced by gut contents (Sibley
et al., 1997a). However, this recommendation was not
extended to include H. azteca. Studies by Dawson et al.
(personal communication, T.D. Dawson, Integrated Labo-
ratory Systems, Duluth, MN) have indicated that the ash
content of H. azteca is not greatly decreased by purging
organisms in clean water before weighing, suggesting that
sediment does not comprise a large portion of the overall
dry weight. In addition, using AFDW further decreases an
51
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already small mass, potentially increasing measurement
error. For this reason, dry weight continues to be the
recommended endpoint for estimating growth of/-/, azteca
via weight (growth can also be determined via length).
11.4 Interpretation of Results
11.4.1 Section 16 describes general information for inter-
pretation of test results. The following sections describe
species-specific information that is useful in helping to
interpret the results of sediment toxicity tests with
H. azteca.
11.4.2 Age Sensitivity
11.4.2.1 The sensitivity of H. azteca appears to be
relatively similar up to at least 24- to 26-d-old organisms
(Collyard etal., 1994). For example, the toxicity of diazinon,
Cu, Cd, and Zn was similar in 96-h water-only exposures
starting with 0- to 2-d-old organisms through 24- to 26-d-
old organisms (Figure 11.2). The toxicity of alkylphenol
ethoxylate (a surfactant) tended to increase with age. In
general, this suggests that tests started with 7- to 14-d-
old amphipods would be representative of the sensitivity
of H. azteca up to at least the adult life stage.
11.4.3 Grain Size
11.4.3.1 Hyalella azteca are tolerant of a wide range of
substrates. Physico-chemical characteristics (e.g., grain
size orTOC) of sediment were not significantly correlated
to the response of H. azteca in toxicity tests in which
organisms were fed (Section 10.1.1.8; Ankley etal.,
1994a).
11.4.4 Isolating Organisms at the End of a Test
11.4.4.1 Quantitative recovery of young amphipods (e.g.,
0- to 7-d old) is difficult given their small size (Figure 11.3,
Tomasovic et al., 1994). Recovery of older and larger
amphipods (e.g., 21-d old) is much easier. This was a
primary reason for deciding to start 10-d tests with 7- to
14-d-old amphipods (organisms are 17- to 24-d old at the
end of the 10-d test).
11.4.5 Influence of Indigenous Organisms
11.4.5.1 Survival of H. azteca in 28-d tests was not
reduced in the presence of oligochaetes in sediment
samples (Reynoldson et al., 1994). However, growth of
amphipods was reduced when high numbers of
oligochaetes were placed in a sample. Therefore, it is
important to determine the number and biomass of indig-
enous organisms in field-collected sediment in order to
better interpret growth data (Reynoldson et al., 1994;
DeFoe and Ankley, 1998). Furthermore, presence of preda-
tors may also influence the response of test organisms in
sediment (Ingersoll and Nelson, 1990).
11.4.6 Ammonia toxicity
11.4.6.1 Section 1.3.7.5 addresses interpretative guid-
ance for evaluating toxicity associated with ammonia in
sediment.
52
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67
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Section 12
Test Method 100.2
Chironomus tentans 10-d Survival and Growth Test for Sediments
12.1 Introduction
12.1.1 Chironomus tentans (Fabricius) have many desir-
able characteristics of an ideal sediment toxicity testing
organism including relative sensitivity to contaminants
associated with sediment, contact with sediment, ease of
culture in the laboratory, tolerance to varying physico-
chemical characteristics of sediment, and short genera-
tion time. Their response has been evaluated in interlabo-
ratory studies and has been confirmed with natural benthic
populations. Many investigators have successfully used
C. tentans to evaluate the toxicity of freshwater sedi-
ments (e.g., Wentsel etal., 1977; Nebekeret al., 1984a;
Nebeker et al., 1988; Adams et al., 1985; Giesy et al.,
1988; Hokeetal., 1990; West et al., 1993; Ankley et al.,
1993; Ankley et al., 1994a; Ankley et al.,1994b).
C. tentans has been used for a variety of sediment
assessments (Westetal., 1993; Hoke etal., 1994,1995;
West et al., 1994; Ankley et al., 1994c). Endpoints typi-
cally monitored in 10-d sediment toxicity tests with
C. tentans include survival and growth (ASTM, 1999a).
12.1.2 A specific test method for conducting a 10-d
sediment toxicity test is described in Section 12.2 for
C. tentans. Methods outlined in Appendix A of USEPA
(1994a) and in Section 12.1.1 were used for developing
test method 100.2. Results of tests using procedures
different from the procedures described in Section 12.2
may not be comparable and these different procedures
may alter contaminant bioavailability. Comparison of re-
sults obtained using modified versions of these proce-
dures might provide useful information concerning new
concepts and procedures for conducting sediment tests
with aquatic organisms. If tests are conducted with proce-
dures different from the procedures described in this
manual, additional tests are required to determine compa-
rability of results (Section 1.3).
12.2 Recommended Test Method for
Conducting a 10-d Sediment Toxicity
Test with Chironomus tentans
12.2.1 Recommended conditions for conducting a 10-d
sediment toxicity test with C. tentans are summarized in
Table 12.1. A general activity schedule is outlined in
Table 12.2. Decisions concerning the various aspects of
experimental design, such as the number of treatments,
number of test chambers/treatment, and water-quality
characteristics should be based on the purpose of the test
and the methods of data analysis (Section 16). The
number of replicates and concentrations tested depends
in part on the significance level selected and the type of
statistical analysis. When variability remains constant,
the sensitivity of a test increases as the number of
replicates increases.
12.2.2 The recommended 10-d sediment toxicity test
with C. tentans must be conducted at 23°C with a 16L8D
photoperiod at an illuminance of about 100 to 1000 lux
(Table 12.1). Test chambers are 300-mL high-form lipless
beakers containing 100 ml of sediment and 175 ml of
overlying water. Ten second- to third-instar midges (about
10-d old) are used to start a test (Section 10.4.1). The
number of replicates/treatment depends on the objective
of the test. Eight replicates are recommended for routine
testing (see Section 16). Midges in each test chamber are
fed 1.5 ml of a 4-g/L Tetrafin® suspension daily. Each
test chamber receives 2 volume additions/d of overlying
water. Water renewals may be manual or automated.
Appendix A describes water-renewal systems that can be
used to deliver overlying water. Overlying water can be
culture water, well water, surface water, site water, or
reconstituted water. For site-specific evaluations, the char-
acteristics of the overlying water should be as similar as
possible to the site where sediment is collected. Require-
ments fortest acceptability are summarized in Table 12.3.
12.3 General Procedures
12.3.1 Sediment into Test Chambers
The day before the sediment test is started (Day-1) each
sediment should be thoroughly homogenized and added
to the test chambers (Section 8.3.1). Sediment should be
visually inspected to judge the extent of homogeneity.
Excess water on the surface of the sediment can indicate
separation of solid and liquid components. If a quantita-
tive measure of homogeneity is required, replicate sub-
samples should be taken from the sediment batch and
analyzed forTOC, chemical concentrations, and particle
size.
12.3.1.1 Each test chamber should contain the same
amount of sediment, determined either by volume or by
weight. Overlying water is added to the chambers in a
mannerthat minimizes suspension of sediment. This can
be accomplished by gently pouring water along the sides
55
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Table 12.1 Recommended Test Conditions
Parameter
for Conducting a 10-d Sediment Toxicity Test with Chironoinus tentans
Conditions
1. Test type:
2. Temperature:
3. Light quality:
4. Illuminance:
5. Photoperiod:
6. Test chamber:
7. Sediment volume:
8. Overlying water volume:
9. Renewal of overlying water:
10. Age of organisms:
11. Number of organisms/chamber:
12. Number of replicate chambers/treatment:
13. Feeding:
14. Aeration:
15. Overlying water:
16. Test chamber cleaning:
17. Overlying water quality:
18. Test duration:
19. Endpoints:
20. Test acceptability:
Whole-sediment toxicity test with renewal of overlying water
23 ± 1 °C
Wide-spectrum fluorescent lights
About 100 to 1000 lux
16L8D
300-mL high-form lipless beaker
100mL
175 ml
2 volume additions/d (Appendix A); continuous or intermittent (e.g., one volume
addition every 12 h)
Second- to third-instar larvae (about 10-d-old larvae; all organisms must be third
instar or younger with at least 50% of the organisms at third instar; Section 10.4.1)
10
Depends on the objective of the test. Eight replicates are recommended for routine
testing (see Section 16).
Tetrafin® goldfish food, fed 1.5 ml daily to each test chamber (1.5 ml contains
6.0 mg of dry solids)
None, unless dissolved oxygen in overlying water drops below 2.5 mg/L.
Culture water, well water, surface water, site water, or reconstituted water
If screens become clogged during a test, gently brush the outside of the screen
(Appendix A).
Hardness, alkalinity, conductivity, pH, and ammonia at the beginning and end of a
test. Temperature and dissolved oxygen daily.
10d
Survival and growth (ash-free dry weight, AFDW)
Minimum mean control survival must be 70%, with minimum mean weight/surviving
control organism of 0.48 mg AFDW. Performance-based criteria specifications are
outlined in Table 12.3.
of the chambers or by pouring water onto a baffle (e.g., a
circular piece of Teflon with a handle attached) placed
above the sediment to dissipate the force of the water.
Renewal of overlying water is started on Day -1. A test
begins when the organisms are added to the test cham-
bers (Day 0).
12.3.2 Renewal of Overlying Water
12.3.2.1 Renewal of overlying water is required during a
test. At any particular time during the test, flow rates
through any two test chambers should not differ by more
than 10%. Hardness, alkalinity and ammonia concentra-
tions in the water above the sediment, within a treatment,
typically should not vary by more than 50% during the
test. Mount and Brungs (1967) diluters have been modi-
fied for sediment testing, and other automated water-
delivery systems have also been used (Maki, 1977;
Ingersoll and Nelson, 1990; Benoit et al., 1993; Zumwalt
et al., 1994; Brunson et al., 1998; Wall et al., 1998;
Leppanen and Maier, 1998). Each water-delivery system
should be calibrated before a test is started to verify that
the system is functioning properly. Renewal of overlying
water is started on Day -1 before the addition of test
organisms or food on Day 0. Appendix A describes
water-renewal systems that can be used for conducting
sediment tests.
12.3.2.2 In water-renewal tests with one to four volume
additions of overlying water/d, water-quality characteris-
tics generally remain similarto the inflowing water (Ingersoll
and Nelson, 1990; Ankley et al., 1993); however, in static
tests, water quality may change profoundly during the
exposure (Shuba et al., 1978). For example, in static
whole-sediment tests, the alkalinity, hardness, and con-
ductivity of overlying water more than doubled in several
treatments during a four-week exposure (Ingersoll and
Nelson, 1990). Additionally, concentrations of metabolic
products (e.g., ammonia) may also increase during static
exposures, and these compounds can either be directly
toxic to the test organisms or may contribute to the
toxicity of the contaminants in the sediment. Furthermore,
changes in water-quality characteristics such as hardness
may influence the toxicity of many inorganic (Gauss et
56
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Day
Table 12.2 General Activity Schedule for Conducting a 10-d Sediment Toxicity Test with Chironomus tentans 1
Activity
-14 Isolate adults for production of egg cases.
-13 Place newly deposited egg cases into hatching dishes.
-12 Prepare a larval rearing chamber with new substrate.
-11 Examine egg cases for hatching success. If egg cases have hatched, transfer first-instar larvae and any remaining unhatched
embryos from the crystallizing dishes into the larval rearing chamber. Feed organisms.
-10 Same as Day -11.
-9 to -2 Feed and observe midges (Section 10.4). Measure water quality (e.g., temperature and dissolved oxygen).
-1 Add food to each larval rearing chamber and measure temperature and dissolved oxygen. Add sediment into each test chamber,
place chamber into exposure system, and start renewing overlying water.
0 Measure total water quality (temperature, pH, hardness, alkalinity, dissolved oxygen, conductivity, ammonia). Remove third-instar
larvae from the culture chamber substrate. Add 1.5 ml of Tetrafin® (4.0 g/L) into each test chamber. Transfer 10 larvae into each
test chamber. Release organisms under the surface of the water. Archive 20 test organisms for instar determination and weight
or length determination. Observe behavior of test organisms.
1 to 8 Add 1.5 ml of food to each test chamber. Measure temperature and dissolved oxygen. Observe behavior of test organisms.
9 Measure total water quality.
10 Measure temperature and dissolved oxygen. End the test by collecting the midges with a sieve. Measure weight or length of
surviving larvae.
1 Modified from Call etal., 1994
al., 1985) and organic (Mayer and Ellersieck, 1986) con-
taminants. Although contaminant concentrations are re-
duced in the overlying water in water-renewal tests, organ-
isms in direct contact with sediment generally receive a
substantial proportion of a contaminant dose directly from
either the whole sediment or from the interstitial water.
12.3.3 Acclimation
12.3.3.1 Test organisms must be cultured and tested at
23°C. Ideally, test organisms should be cultured in the
same water that will be used in testing. However, acclima-
tion of test organisms to the test water is not required.
12.3.3.2 Culturing of organisms and toxicity assessment
are typically conducted at 23°C. However, occasionally
there is a need to perform evaluations at temperatures
different than that recommended. Under these circum-
stances, it may be necessary to acclimate organisms to
the desired test temperature to prevent thermal shock
when moving immediately from the culture temperature to
the test temperature (ASTM, 1999a). Acclimation can be
achieved by exposing organisms to a gradual change in
temperature; however, the rate of change should be rela-
tively slow to prevent thermal shock. A change in tem-
perature of 1 °C every 1 to 2 h has been used successfully
in some studies (P.K. Sibley, University of Guelph, Guelph,
Ontario, personal communication; APHA, 1989). Testing
at temperatures other than 23°C needs to be preceded by
studies to determine expected performance under alter-
nate conditions.
12.3.4 Placing Organisms in Test Chambers
12.3.4.1 Test organisms should be handled as little as
possible. Midges should be introduced into the overlying
water below the air-water interface. Test organisms can
be pipetted directly into overlying water. Developmental
stage of the test organisms should be documented from a
subset of at least 20 organisms used to start the test
(Section 10.4.1). Developmental stage can be deter-
mined from head capsule width (Table 10.2), length (4 to 6
mm), or dry weight (0.08 to 0.23 mg/individual). It is
desirable to measure size at test initiation using the same
measure as will be used to assess growth at the end of
the test.
12.3.5 Feeding
12.3.5.1 For each beaker, 1.5 ml of Tetrafin® is fed from
Day 0 to Day 9. Without addition of food, the test
organisms may starve during exposures. However, the
addition of the food may alter the availability of the
contaminants in the sediment (Wiederholm et al., 1987;
Harkey et al., 1994). Furthermore, if too much food is
added to the test chamber or if the mortality of test
organisms is high, fungal or bacterial growth may develop
on the sediment surface. Therefore, the amount of food
added to the test chambers is kept to a minimum.
12.3.5.2 Suspensions of food should be thoroughly mixed
before aliquots are taken. If excess food collects on the
sediment, a fungal or bacterial growth may develop on the
sediment surface, in which case feeding should be sus-
pended for one or more days. A drop in dissolved oxygen
below 2.5 mg/L during a test may indicate that the food
57
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Table 12.3 Test Acceptability Requirements for a 10-d Sediment Toxicity Test with Chironomus tentans
A. It is recommended for conducting a 10-d test with C. tentans that the following performance criteria be met:
1. Tests must be started with second- to third-instar larvae (about 10-d-old larvae; see Section 10.4.1).
2. Average survival of C. tentans in the control sediment must be greater than or equal to 70% at the end of the test.
3. Average size of C. tentans in the control sediment must be at least 0.48 mg AFDW at the end of the test.
4. Hardness, alkalinity, and ammonia in the overlying water typically should not vary by more than 50% during the test, and dissolved
oxygen should be maintained above 2.5 mg/L in the overlying water.
B. Performance-based criteria for culturing C. tentans include the following:
1. It may be desirable for laboratories to periodically perform 96-h water-only reference-toxicity tests to assess the sensitivity of
culture organisms (Section 9.16.2). Data from these reference-toxicity tests could be used to assess genetic strain or life-stage
sensitivity of test organisms to select chemicals.
2. Laboratories should keep a record of time to first emergence for each culture and record this information using control charts.
Records should also be kept on the frequency of restarting cultures.
3. Laboratories should record the following water-quality characteristics of the cultures at least quarterly: pH, hardness, alkalinity, and
ammonia. Dissolved oxygen in the cultures should be measured weekly. Temperature of the cultures should be recorded daily. If
static cultures are used, it may be desirable to measure water quality more frequently.
4. Laboratories should characterize and monitor background contamination and nutrient quality of food if problems are observed in
culturing or testing organisms.
5. Physiological measurements such as lipid content might provide useful information regarding the health of the cultures.
C. Additional requirements:
1. All organisms in a test must be from the same source.
2. Storage of sediments collected from the field should follow guidance outlined in Section 8.2.
3. All test chambers (and compartments) should be identical and should contain the same amount of sediment and overlying water.
4. Negative-control sediment and appropriate solvent controls must be included in a test. The concentration of solvent used must not
adversely affect test organisms.
5. Test organisms must be cultured and tested at 23°C (±1°C).
6. The daily mean test temperature must be within ±1°C of 23°C. The instantaneous temperature must always be within ±3°C of 23°C.
7. Natural physico-chemical characteristics of test sediment collected from the field should be within the tolerance limits of the test
organisms.
added is not being consumed. Feeding should be sus- sampled just before water renewal from about 1 to 2 cm
pended for the amount of time necessary to increase the above the sediment surface using a pipet. It may be
dissolved oxygen concentration (ASTM, 1999a). If feeding necessary to composite water samples from individual
is suspended in one treatment, it should be suspended in replicates. The pipet should be checked to make sure no
all treatments. Detailed records of feeding rates and the organisms are removed during sampling of overlying wa-
appearance of the sediment surface should be made ter. Water quality should be measured on each batch of
daily. water prepared for the test.
12.3.6 Monitoring a Test 12.3.6.2.2 Water-only exposures evaluating the tolerance
of C. tentans larvae to depressed DO have indicated that
12.3.6.1 All chambers should be checked daily and significant reductions in weight occurred after 10-d expo-
observations made to assess test organism behavior sure to 1.1 mg/L DO, but not at 1.5 mg/L (V. Mattson,
such as sediment avoidance. However, monitoring ef- USEPA, Duluth, MN, personal communication). This
fects on burrowing activity of test organisms may be finding concurs with the observations during method
difficult because the test organisms are often not visible development at the USEPA laboratory in Duluth that
during the exposure. The operation of the exposure sys- excursions of DO as low as 1.5 mg/L did not seem to
tern should be monitored daily. have an effect on midge survival and development (P.K.
Sibley, University of Guelph, Guelph, Ontario, personal
72.3.6.2 Measurement of Overly ing Water-Quality communication). Based on these findings, it appears that
Characteristics periodic depressions of DO below 2.5 mg/L (but not below
1.5 mg/L) are not likely to adversely affect test results,
12.3.6.2.1 Conductivity, hardness, pH, alkalinity, and and thus shou|d not be a reason to discarc| test data.
ammonia should be measured in all treatments at the Nonetheless tests should be managed toward a goal of
beginning and end of a test. Overlying water should be DO > 2.5 mg/L to insure satisfactory performance. If the
58
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DO level of the water falls below 2.5 mg/L for any one
treatment, aeration is encouraged and should be done in
all replicates for the duration of the test. Occasional
brushing of screens on outside of beakers will help main-
tain the exchange of water during renewals using the
exposure system described by Benoit et al. (1993). If a
probe is used to measure DO in overlying water, it should
be thoroughly inspected between samples to make sure
that organisms are not attached and should be rinsed
between samples to minimize cross contamination. Aera-
tion can be used to maintain dissolved oxygen in the
overlying water above 2.5 mg/L (i.e., about 1 bubble/
second in the overlying water).
12.3.6.2.3 Temperature should be measured at least
daily in at least one test chamber from each treatment.
The temperature of the water bath orthe exposure cham-
ber should be continuously monitored. The daily mean
test temperature must be within ±1 °C of 23°C. The instan-
taneous temperature must always be within ±3°C of 23°C.
12.3.7 Ending a Test
12.3.7.1 Immobile organisms isolated from the sediment
surface or from sieved material should be considered
dead. A #40 sieve (425-urn mesh) can be used to remove
midges from sediment. Alternatively, Kemble et al. (1994)
suggest sieving of sediment using the following proce-
dure: (1) pour about half of the overlying water through a
#50- (300-um) U.S. standard mesh sieve, (2) pour about
half of the sediment through the #50-mesh sieve and
wash the contents of the sieve into an examination pan,
(3) rinse the coarser sediment remaining in the test cham-
ber through a #40- (425-um) mesh sieve and wash the
contents of this second sieve into a second examination
pan. Surviving midges can then be isolated from these
pans. See Section 12.3.8.1 and 12.3.8.2 for the proce-
dures for measuring weight or length of midges.
12.3.7.2 A consistent amount of time should be taken to
examine sieved material for recovery of test organisms
(e.g., 5 min/replicate). Laboratories should demonstrate
that their personnel are able to recover an average of at
least 90% of the organisms from whole sediment. For
example, test organisms could be added to control sedi-
ment and recovery could be determined after 1 h
(Tomasovicet al., 1994).
12.3.8 Test Data
12.3.8.1 Ash-free dry weight (AFDW) and survival are the
endpoints measured at the end of the 10-d sediment
toxicity test with C. tentans. The 10-d method for C. tentans
in the first edition of this manual (USEPA, 1994a), as well
as most previous research, has used dry weight as a
measure of growth. However, Sibleyetal. (1997b) found
that the grain size of sediments influences the amount of
sediment that C. tentans larvae ingest and retain in their
gut. As a result, in finer-grain sediments, a substantial
portion of the measured dry weight may be comprised of
sediment ratherthan tissue. While this may not represent
a strong bias in tests with identical grain size distributions
in all treatments, most field assessments are likely to
have varying grain size among sites. This will likely
create differences in dry weight among treatments that
are not reflective of true somatic growth. Forthis reason,
weight of midges should be measured as ash-free dry
weight (AFDW) instead of dry weight. AFDW will more
directly reflect actual differences in tissue weight by
reducing the influence of sediment in the gut. The dura-
tion of the 10-d test starting with third-instar larvae is not
long enough to determine emergence of adults. Average
size of C. tentans in the control sediment must be at least
0.6 mg at the end of the test (0.48 mg AFDW) (Ankley et
al., 1993; ASTM, 1999a; Section 17.5). If test organisms
are to be used for an evaluation of bioaccumulation, it is
not advisable to dry the sample before conducting the
residue analysis. If conversion from wet weight to dry
weight is necessary, aliquots of organisms can be weighed
to establish wet to dry weight conversion factors. A
consistent procedure should be used to remove the ex-
cess water from the organisms before measuring wet
weight.
12.3.8.2 For determination of AFDW, first pool all living
larvae in each replicate and dry the sample to a constant
weight (e.g., 60°C for 24 h). Note that the weigh boats
should be ashed before use to eliminate weighing errors
due to the pan oxidizing during ashing. The sample is
brought to room temperature in a dessicator and weighed
to the nearest 0.01 mg to obtain mean weights per surviv-
ing organism per replicate. The dried larvae in the pan are
then ashed at 550°C for 2 h. The pan with the ashed
larvae is then reweighed and the tissue mass of the larvae
is determined as the difference between the weight of the
dried larvae plus pan and the weight of the ashed larvae
plus pan. In rare instances where preservation is re-
quired, an 8% sugar formalin solution can be used to
preserve samples (USEPA, 1994a), but the effects of
preservation on the weights and lengths of the midges
have not been sufficiently studied. Pupae or adult organ-
isms must not be included in the sample to estimate ash-
free dry weight. If head capsule width is to be measured,
it should be measured on surviving midges at the end of
the test before ash-free dry weight is determined.
12.3.8.3 Measurement of length is optional. Separate
replicate beakers should be set up to sample lengths of
midges at the end of an exposure. An 8% sugar formalin
solution can be used to preserve samples for length
measurements (Ingersoll and Nelson, 1990). The sugar
formalin solution is prepared by adding 120 g of sucrose
to 80 mL of formalin, which is then brought to a volume of
1 L using deionized water. This stock solution is mixed
with an equal volume of deionized water when used to
preserve organisms. NoTox® (Earth Safe Industries,
Belle Mead, NJ) can be used as a substitute for formalin
(Unger et al., 1993). Midge body length (±0.1 mm) can be
measured from the anterior of the labrum to the posterior
of the last abdominal segment (Smock, 1980). Kemble et
al. (1994) photographed midges at magnification of 3.5X
and measured the images using a computer-interfaced
digitizing tablet. A digitizing system and microscope can
59
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also be used to measure length (Ingersoll and Nelson,
1990).
12.4 Interpretation of Results
12.4.1 Section 16 describes general information for inter-
pretation of test results. The following sections describe
species-specific information that is useful in helping to
interpret the results of sediment toxicity tests with
C. tentans.
12.4.2 Age Sensitivity
12.4.2.1 Midges are perceived to be relatively insensitive
organisms in toxicity assessments (Ingersoll, 1995). This
conclusion is based on measuring survival of fourth-instar
larvae in short-term water-only exposures, a procedure
that may underestimate the sensitivity of midges to toxi-
cants. The first and second instars of chironomids are
more sensitive to contaminants than the third or fourth
instars. For example, first-instar C. tentans larvae were
6 to 27 times more sensitive than fourth-instar larvae to
acute copper exposure (Nebeker et al., 1984b; Gauss et
al., 1985; Figure 12.1) and first-instar C. riparius larvae
were 127 times more sensitive than second-instar larvae
to acute cadmium exposure (Williams et al., 1986b;
Figure 12.1). In chronic tests with first-instar larvae, midges
were often as sensitive as daphnids to inorganic and
organic compounds (Ingersoll et al., 1990). Sediment
tests should be started with uniform age and size midges
because of the dramatic differences in sensitivity of
midges by age. Whereas third-instar midges are not as
sensitive as younger organisms, the larger larvae are
easier to handle and isolate from sediment at the end of a
test.
12.4.2.2 DeFoe and Ankley (1998) studied a variety of
contaminated sediments and showed that the sensitivity
of C. tentans 10-d tests is greatly increased by measure-
ment of growth in addition to survival. Growth of midges
in 10-d sediment tests was found to be a more sensitive
endpoint than survival of Hyalella azteca (DeFoe and
Ankley, 1998). In cases where sensitivity of organisms
before the third instar is of interest, the long-term sedi-
ment exposures can be used, since they begin with newly
hatched larvae (Section 15).
12.4.3 Physical characteristics of sediment
12.4.3.1 Grain Size
12.4.3.1.1 Larvae of C. tentans appear to be tolerant of a
wide range of particle size conditions in substrates. Sev-
eral studies have shown that survival is not affected by
particle size in natural sediments, sand substrates, or
formulated sediments in both 10-d and long-term expo-
sures (Ankley et al., 1994; Suedel and Rodgers, 1994;
Sibley et al., 1997b, 1998). Ankley et al. (1994a) found
that growth of C. tentans larvae was weakly correlated
with sediment grain size composition, but not organic
carbon, in 10-d tests using 50 natural sediments from the
Great Lakes. However, Sibley et al. (1997b) found that
the correlation between grain size and larval growth disap-
peared after accounting for inorganic material contained
within larval guts and concluded that growth of C. tentans
was not related to grain size composition in either natural
sediments or sand substrates. Avoiding confounding
influences of gut contents on weight is the impetus for
recommending ash-free dry weight (instead of dry weight)
as the index of growth in the 10-day and long-term
C. tentans tests. Failing to do so could lead to erroneous
conclusions regarding the toxicity of the test sediment
(Sibley et al., 1997b). Procedures for correcting for gut
contents are described in Section 12.3.8. Emergence,
reproduction (mean eggs/female), and hatch success
were also not affected by the particle size composition of
substrates in long-term tests with C. tentans (Sibley et
al., 1998; Section 15).
12.4.3.2 Organic Matter
12.4.3.2.1 Based on 10-d tests, the content of organic
matter in sediments does not appear to affect survival of
C. tentans larvae in natural and formulated sediments, but
maybe important with respect to larval growth. Ankley et
al. (1994a) found no relationship between sediment or-
ganic content and survival or growth in 10-d bioassays
with C. tentans in natural sediments. Suedel and Rodgers
(1994) observed reduced survival in 10-d tests with a
formulated sediment when organic matter was <0.91%;
however, supplemental food was not supplied in this
study, which may influence these results relative to the
10-d test procedures described in this manual. Lacey et
al. (1999) found that survival of C. tentans larvae was
generally not affected in 10-d tests by eitherthe quality or
quantity of synthetic (alpha-cellulose) or naturally derived
(peat, maple leaves) organic material spiked into a formu-
lated sediment, although a slight reduction in survival
below the acceptability criterion (70%) was observed in a
natural sediment diluted with formulated sediment at an
organic matter content of 6%. In terms of larval growth,
Lacey et al. (1999) did not observe any systematic rela-
tionship between the level of organic material (e.g., food
quantity) and larval growth for each carbon source. Al-
though a significant reduction in growth was observed at
the highest concentration (10%) of the leaf treatment in
the food quantity study, significantly higher larval growth
was observed in this treatment when the different carbon
sources were compared at about equal concentrations
(effect of food quality). In the latter study, the following
gradient of larval growth was established in relation to the
source of organic carbon: peat< natural sediment < alpha-
cellulose < leaves. Since all of the treatments received a
supplemental source of food, these data suggest that
both the quality and quantity of organic carbon in natural
and formulated sediments may represent an important
confounding factor for the growth endpoint in tests with
C. tentans (Lacey et al., 1999). However, it is important
to note that these data are based on 10-d tests; the
applicability of these data to long-term testing has not
been evaluated (Section 15).
60
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A. Chironomus riparius: Cadmium
z.o
2
o 15
O
X
4 1
CN
0.5
0
-
•
I , I
1
1
1
I.
CD
CD
0.5
0
1st
1st
2nd 3rd
INSTAR
B. Chironomus tentans: Copper
4th
Williams etal. (1985)
I
2nd 3rd
INSTAR
4th
Nebekeretal. (1964)
Figure 12.1 Lifestage sensitivity of chironomids.
61
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12.4.4 Isolating Organisms at the End of a Test 12.4.6. Sexual Dimorphism
12.4.4.1 Quantitative recovery of larvae at the end of a
10-d sediment test should not be a problem. The larvae
are red and typically greaterthan 5 mm long.
12.4.5 Influence of Indigenous Organisms
12.4.5.1 The influence of indigenous organisms on the
response of C. tentans in sediment tests has not been
reported. Survival of a closely related species, C. riparius
was not reduced in the presence of oligochaetes in sedi-
ment samples (Reynoldson etal., 1994). However, growth
of C. riparius was reduced when high numbers of oli-
gochaetes were placed in a sample. Therefore, it is
important to determine the number and biomass of indig-
enous organisms in field-collected sediment in order to
better interpret growth data (Reynoldson et al., 1994;
DeFoe and Ankley, 1998). Furthermore, presence of
predators may also influence the response of test organ-
isms in sediment (Ingersoll and Nelson, 1990).
12.4.6.1 Differences in size between males and females
of a closely related midge species (Chironomus riparius)
had little effect on interpretation of growth-related effects
in sediment tests (<3% probability of making a Type I
error [nontoxic sample classified as toxic] due to sexual
dimorphism; Day et al., 1994). Therefore, sexual dimor-
phism will probably not be a confounding factor when
interpreting growth results measured in sediment tests
with C. tentans.
12.4.7 Ammonia Toxicity
12.4.7.1 Section 1.3.7.5 addresses interpretative guid-
ance for evaluating toxicity associated with ammonia in
sediment.
62
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Section 13
Test Method 100.3
Lumbriculus variegatus Bioaccumulation Test for Sediments
13.1 Introduction
13.1.1 Lumbriculus variegatus (Oligochaeta) have many
desirable characteristics of an ideal sediment bioaccumu-
lation testing organism including contact with sediment,
ease of culture in the laboratory, and tolerance to varying
physico-chemical characteristics of sediment. The re-
sponse of L variegatus in laboratory exposures has been
confirmed with natural benthic populations. Many investi-
gators have successfully used L. variegatus in toxicity or
bioaccumulation tests. Toxicity studies have been con-
ducted in water-only tests (Bailey and Liu, 1980; Hornig,
1980; Ewell et al., 1986; Nebeker et al., 1989; Ankley et
al., 1991 a; Ankley etal., 1991b), in effluent tests (Hornig,
1980), and in whole-sediment tests (Nebeker etal., 1989;
Ankley et al., 1991 a; Ankley et al., 1991b; Ankley et al.,
1992a; Call et al., 1991; Carlson etal., 1991; Phipps et
al., 1993; West et al., 1993). Several studies have re-
ported the use of L variegatus to examine bioaccumula-
tion of chemicals from sediment (Schuytema etal., 1988;
Nebeker et al., 1989; Ankley et al., 1991b; Call et al.,
1991; Carlson etal., 1991; Ankley etal., 1993; Kukkonen
and Landrum, 1994; and Brunson et al., 1993, 1998).
However, interlaboratory studies have not yet been con-
ducted with L. variegatus.
13.1.2 Additional research is needed on the standardiza-
tion of bioaccumulation procedures with sediment. There-
fore, Section 13.2 describes general guidance for con-
ducting a 28-d sediment bioaccumulation test with
L. variegatus. Methods outlined in Appendix A of USEPA
(1994a) and in Section 13.1.1 were used for developing
this general guidance. Results of tests using procedures
different from the procedures described in Section 13.2
may not be comparable, and these different procedures
may alter bioavailability. Comparison of results obtained
using modified versions of these procedures might pro-
vide useful information concerning new concepts and
procedures for conducting sediment tests with aquatic
organisms. If tests are conducted with procedures differ-
ent from the procedures described in this manual, addi-
tional tests are required to determine comparability of
results (Section 1.3).
13.2 Procedure for Conducting Sediment
Bioaccumulation Tests with
Lumbriculus variegatus
13.2.1 Recommended test conditions for conducting a
28-d sediment bioaccumulation test with L. variegatus are
summarized in Table 13.1. Table 13.2 outlines proce-
dures for conducting sediment toxicity tests with L varie-
gatus. A general activity schedule is outlined in Table 13.3.
Decisions concerning the various aspects of experimen-
tal design, such as the number of treatments, number of
test chambers/treatment, and water-quality characteris-
tics should be based on the purpose of the test and the
methods of data analysis (Section 16). The number of
replicates and concentrations tested depends in part on
the significance level selected and the type of statistical
analysis. When variability remains constant, the sensitiv-
ity of a test increases as the number of replicates increases.
13.2.2 The recommended 28-d sediment bioaccumula-
tion test with L. variegatus can be conducted with adult
oligochaetes at 23°C with a 16L8D photoperiod at a
illuminance of about 100 to 1000 lux (Table 13.1). Test
chambers can be 4 to 6 L that contain 1 to 2 L of sediment
and 1 to 4 L of overlying water. The number of replicates/
treatment depends on the objective of the test. Five
replicates are recommended for routine testing
(Section 16). To minimize depletion of sediment contami-
nants, the ratio of total organic carbon in sediment to dry
weight of organisms should be about 50:1. A minimum of
1 g/replicate with up to 5 g/replicate should be tested.
Oligochaetes are not fed during the test. Each chamber
receives 2 volume additions/d of overlying water. Appen-
dix A and Brunson et al., (1998) describe water-renewal
systems that can be used to deliver overlying water.
Overlying water can be culture water, well water, surface
water, site water, or reconstituted water. For site-specific
evaluations, the characteristics of the overlying water
should be as similar as possible to the site where sedi-
ment is collected. Requirements fortest acceptability are
outlined in Table 13.4.
13.2.2.1 Before starting a 28-d sediment bioaccumulation
test with L. variegatus, a toxicity screening test can be
conducted for at least 4 d using procedures outlined in
Table 13.2 (Brunson etal., 1993). The preliminary toxicity
screening test is conducted at 23°C with a 16L8D photo-
period at an illuminance of about 100 to 1000 lux. Test
chambers are 300-mL high-form lipless beakers containing
63
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Table 13.1 Recommended Test Conditions for Conducting a 28-d Sediment Bioaccumulation Test with Luinbriculus variegatus
Parameter Conditions
1. Test type:
2. Temperature:
3. Light quality:
4. Illuminance:
5. Photoperiod:
6. Test chamber:
7. Sediment volume:
8. Overlying water volume:
9. Renewal of overlying water:
10. Age of test organisms:
11. Loading of organisms in chamber:
12. Number of replicate chambers/treatment:
13. Feeding:
14. Aeration:
15. Overlying water:
16. Test chamber cleaning:
17. Overlying water quality:
18. Test duration:
19. Endpoint:
20. Test acceptability:
Whole-sediment bioaccumulation test with renewal of overlying water
23 ± 1 °C
Wide-spectrum fluorescent lights
About 100 to 1000 lux
16L8D
4- to 6-L aquaria with stainless steel screens or glass standpipes
1 L or more depending on TOC
1 L or more depending on TOC
2 volume additions/d (Appendix A); continuous or intermittent (e.g., one volume
addition every 12 h)
Adults
Ratio of total organic carbon in sediment to organism dry weight should be no less
than 50:1. Minimum of 1 g/replicate. Preferably 5 g/replicate.
Depends on the objective of the test. Five replicates are recommended for routine
testing (see Section 16).
None
None, unless DO in overlying water drops below 2.5 mg/L
Culture water, well water, surface water, site water, or reconstituted water
If screens become clogged during the test, gently brush the outside of the screen
(Appendix A).
Hardness, alkalinity, conductivity, pH, and ammonia at the beginning and end of a
test. Temperature and dissolved oxygen daily.
28 d
Bioaccumulation
Performance-based criteria specifications are outlined in Table 13.4.
100 ml of sediment and 175 ml of overlying water. Ten
adult oligochaetes/replicate are used to start a test. Four
replicates are recommended fortoxicity screening tests.
Oligochaetes are not fed during the test. Each chamber
receives 2 volume additions/d of overlying water. Appen-
dix A and Brunson et al. (1998) describe water-renewal
systems that can be used to deliver overlying water.
Overlying water should be similar to the water to be used
in the bioaccumulation test. Endpoints monitored at the
end of a toxicity test are number of organisms and
behavior. Numbers of L. variegatus in the toxicity screen-
ing test should not be significantly reduced in the test
sediment relative to the control sediment. Test organisms
should burrow into test sediment. Avoidance of test sedi-
ment by L. variegatus may decrease bioaccumulation.
13.3 General Procedures
13.3.1 Sediment into Test Chambers
13.3.1.1 The day before the sediment test is started
(Day -1) each sediment should be thoroughly homog-
enized and added to the test chambers (Section 8.3.1).
Sediment should be visually inspected to judge the extent
of homogeneity. Excess water on the surface of the
sediment can indicate separation of solid and liquid com-
ponents. If a quantitative measure of homogeneity is
required, replicate subsamples should be taken from the
sediment batch and analyzed for TOC, chemical concen-
trations, and particle size.
13.3.1.2 Each test chamber should contain the same
amount of sediment, determined either by volume or by
weight. Overlying water is added to the chambers in a
mannerthat minimizes suspension of sediment. This can
be accomplished by gently pouring water along the sides
of the chambers or by pouring water onto a baffle (e.g., a
circular piece of Teflon® with a handle attached) placed
above the sediment to dissipate the force of the water.
Renewal of overlying water is started on Day -1. A test
begins when the organisms are added to the test cham-
bers (Day 0).
13.3.2 Renewal of Overlying Water
13.3.2.1 Renewal of overlying water is recommended
during a test. At any particular time during the test, flow
rates through any two test chambers should not differ by
more than 10%. Hardness, alkalinity and ammonia
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Table 13.2 Recommended Test Conditions for Conducting a Preliminary 4-d Sediment Toxicity Screening Test with
Lumbriculus variegatus
Parameter
Conditions
1. Test type:
2. Temperature:
3. Light quality:
4. Illuminance:
5. Photoperiod:
6. Test chamber:
7. Sediment volume:
8. Overlying water volume:
9. Renewal of overlying water:
10. Age of test organisms:
11. Number of organisms/chamber:
12. Number of replicate chambers/treatment:
13. Feeding:
14. Aeration:
15. Overlying water:
16. Test chamber cleaning:
17. Overlying water quality:
18. Test duration:
19. Endpoints:
20. Test acceptability:
4-d whole-sediment toxicity test with renewal of overlying water
23 ± 1 °C
Wide-spectrum fluorescent lights
About 100 to 1000 lux
16L8D
300-mL high-form lipless beaker
100 ml
175 ml
2 volume additions/d (Appendix A); continuous or intermittent (e.g., one volume
addition every 12 h)
Adults
10
4 minimum
None
None, unless DO in overlying water drops below 2.5 mg/L
Culture water, well water, surface water, site water, or reconstituted water
If screens become clogged during the test, gently brush the outside of the screen.
Hardness, alkalinity, conductivity, pH, and ammonia at the beginning and end of
a test. Temperature and dissolved oxygen daily.
4d (minimum; up to 10 d)
Number of organisms and behavior. There should be no significant reduction in
number of organisms in a test sediment relative to the control.
Performance-based criteria specifications are outlined in Table 13.4.
concentrations in the water above the sediment, within a
treatment, should not vary by more than 50% during the
test. Mount and Brungs (1967) diluters have been modi-
fied for sediment testing, and other automated water-
delivery systems have also been used (Maki, 1977;
Ingersoll and Nelson, 1990; Benoit et al., 1993; Zumwalt
et al., 1994; Brunson et al., 1998; Wall et al., 1998;
Leppanen and Maier, 1998). Each water-delivery system
should be calibrated before a test is started to verify that
the system is functioning properly. Renewal of overlying
water is started on Day -1 before the addition of test
organisms on Day 0 (Appendix A).
13.3.2.2 In water-renewal tests with one to four volume
additions of overlying water/d, water-quality characteris-
tics generally remain similarto the inflowing water (Ingersoll
and Nelson, 1990; Ankley et al., 1993); however, in static
tests, water quality may change profoundly during the
exposure (Shuba et al., 1978). For example, in static
whole-sediment tests, the alkalinity, hardness, and con-
ductivity of overlying water more than doubled in several
treatments during a four-week exposure (Ingersoll and
Nelson, 1990). Additionally, concentrations of metabolic
products (e.g., ammonia) may also increase during static
exposures, and these compounds can either be directly
toxic to the test organisms or may contribute to the
toxicity of the contaminants in the sediment. Further-
more, changes in water-quality characteristics such as
hardness may influence the toxicity of many inorganic
(Gauss et al., 1985) and organic (Mayer and Ellersieck,
1986) contaminants. Although contaminant concentra-
tions are reduced in the overlying water in water-renewal
tests, organisms in direct contact with sediment generally
receive a substantial proportion of a contaminant dose
directly from either the whole sediment or from the inter-
stitial water.
13.3.3 Acclimation
13.3.3.1 Test organisms must be cultured and tested at
23°C. Ideally, test organisms should be cultured in the
same water that will be used in testing. However, acclima-
tion of test organisms to the test water is not required.
13.3.3.2 Culturing of organisms and toxicity assessment
are typically conducted at 23°C. However, occasionally
there is a need to perform evaluations at temperatures
different than that recommended. Under these circum-
stances, it may be necessary to acclimate organisms to
the desired test temperature to prevent thermal shock
65
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Table 13.3 General Activity Schedule for Conducting a 28-d Sediment Bioaccumulation Test with Lumbriculus variegatus
A. Conducting a 4-d Toxicity Screening Test (conducted before the 28-d bioaccumulation test)
Day Activity
-1 Isolate worms for conducting toxicity screening test. Add sediment into each test chamber, place chambers into exposure system,
and start renewing overlying water.
0 Measure total water quality (pH, temperature, dissolved oxygen, hardness, alkalinity, conductivity, ammonia). Transfer 10 worms
into each test chamber. Measure weight of a subset of 20 organisms used to start the test. Observe behavior of test organisms.
1 to 2 Measure temperature and dissolved oxygen. Observe behavior of test organisms.
3 Same as Day 1. Measure total water quality.
4 Measure temperature and dissolved oxygen. End the test by collecting the oligochaetes with a sieve and determine weight of
survivors. Bioaccumulation tests should not be conducted with L. variegatus if a test sediment significantly reduces number of
oligochaetes relative to the control sediment or if oligochaetes avoid the sediment.
B. Conducting a 28-d Bioaccumulation Test
Day Activity
-1 Isolate worms for conducting bioaccumulation test. Add sediment into each test chamber, place chambers into exposure system,
and start renewing overlying water.
0 Measure total water quality (pH, temperature, dissolved oxygen, hardness, alkalinity, conductivity, ammonia). Transfer
appropriate amount of worms (based on weight) into each test chamber. Sample a subset of worms used to start the test for residue
analyses. Observe behavior of test organisms.
1 to 6 Measure temperature and dissolved oxygen. Observe behavior of test organisms.
7 Same as Day 1. Measure total water quality.
8 to 13 Same as Day 1
14 Same as Day 7
15 to 20 Same as Day 1
21 Same as Day 7
22 to 26 Same as Day 1
27 Measure total water quality.
28 Measure temperature and dissolved oxygen. End the uptake by collecting the worms with a sieve. Separate any indigenous
organisms from L. variegatus. Determine the weight of survivors. Eliminate the gut contents of surviving worms in water for 6
to 8 h. Longer purging periods (not to exceed 24 hours) may be used if all target analytes have Log Kow>5 (Section 13.3.7.3).
when moving immediately from the culture temperature to weights should be measured on a subset of at least 100
the test temperature (ASTM, 1999a). Acclimation can be organisms used to start the test. The ratio of total organic
achieved by exposing organisms to a gradual change in carbon in sediment to dry weight of organisms at the start
temperature; however, the rate of change should be rela- of the test should be no less than 50:1.
tively slow to prevent thermal shock. A change in tem-
perature of 1 °C every 1 to 2 h has been used successfully 13.3.4.2 Oligochaetes added to each replicate should not
in some studies (P.K. Sibley, University of Guelph,Guelph, be blotted to remove excess water (Section 10.5.6).
Ontario, personal communication). Testing at tempera- Oligochaetes can be added to each replicate at about
tures other than 23°C needs to be preceded by studies to 1.33X of the target stocking weight (Brunson et al.,
determine expected performance under alternate 1998). This additional 33% should account forthe excess
conditions. weight from water in the sample of nonblotted oligocha-
etes at the start of the test.
13.3.4 Placing Organisms in Test Chambers
13.3.5 Feeding
13.3.4.1 Isolate oligochaetes for starting a test as de-
scribed in Section 10.5.6. A subset of L. variegatus at the 13.3.5.1 Lumbriculus variegatus should not be fed during
start of the test should be sampled to determine starting a bioaccumulation test.
concentrations of chemicals of concern. Mean group
66
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Table 13.4 Test Acceptability Requirements for a 28-d Sediment Bioaccumulation Test with Lumbriculus variegatus
A. It is recommended for conducting a 28-d test with L. variegatus that the following performance criteria be met:
1. Numbers of L. variegatus in a 4-d toxicity screening test should not be significantly reduced in the test sediment relative to the
control sediment.
2. Test organisms should burrow into test sediment. Avoidance of test sediment by L. variegatus may decrease bioaccumulation.
3. Hardness, alkalinity, and ammonia in the overlying water typically should not vary by more than 50% during the test, and dis-
solved oxygen should be maintained above 2.5 mg/L in the overlying water.
B. Performance-based criteria for culturing L. variegatus include the following:
1. It may be desirable for laboratories to periodically perform 96-h water-only reference toxicity tests to assess the sensitivity of
culture organisms (Section 9.16.2). Data from these reference-toxicity tests could be used to assess genetic strain or life-stage
sensitivity of test organisms to select chemicals.
2. Laboratories should monitor the frequency with which the population is doubling in the culture (number of organisms) and record
this information using control charts (doubling rate would need to be estimated on a subset of animals from a mass culture).
Records should also be kept on the frequency of restarting cultures. If static cultures are used, it may be desirable to measure
water quality more frequently.
3. Food used to culture organisms should be analyzed before the start of a test for compounds to be evaluated in the bioaccumula-
tion test.
4. Laboratories should record the following water-quality characteristics of the cultures at least quarterly and the day before the start
of a sediment test: pH, hardness, alkalinity, and ammonia. Dissolved oxygen in the cultures should be measured weekly.
Temperature of the cultures should be recorded daily.
5. Laboratories should characterize and monitor background contamination and nutrient quality of food if problems are observed in
culturing or testing organisms.
6. Physiological measurements such as lipid content might provide useful information regarding the health of the cultures.
C. Additional requirements:
1. All organisms in a test must be from the same source.
2. Storage of sediments collected from the field should follow guidance outlined in Section 8.2.
3. All test chambers (and compartments) should be identical and should contain the same amount of sediment and overlying water.
4. Negative-control sediment and/or the appropriate solvent controls must be included in a test. The concentration of solvent used
must not affect test organisms adversely.
5. Test organisms must be cultured and tested at 23°C (±1°C).
6. The daily mean test temperature must be within ±1°C of 23°C. The instantaneous temperature must always be within ±3°C of 23°C
7. Natural physico-chemical characteristics of test sediment collected from the field should be within the tolerance limits of the test
organisms.
13.3.6 Monitoring a Test necessary to composite water samples from individual
replicates. The pipet should be checked to make sure no
13.3.6.1 All chambers should be checked daily and organisms are removed during sampling of overlying water.
observations made to assess test organism behavior Waterquality should be measured on each batch of water
such as sediment avoidance. However, monitoring ef- prepared for the test.
fects on burrowing activity of test organisms may be
difficult because the test organisms are often not visible 13.3.6.2.2 Dissolved oxygen should be measured daily
during the exposure. The operation of the exposure sys- and should be above 2.5 mg/L. If a probe is used to
tern should be monitored daily. measure dissolved oxygen in overlying water, it should be
thoroughly inspected between samples to make sure that
73.3.6.2 Measurement of Overlying Water-quality organisms are not attached and should be rinsed between
Characteristics samples to minimize cross contamination. Aeration can
be used to maintain dissolved oxygen in the overlying
13.3.6.2.1 Conductivity, hardness, pH, alkalinity, and water above 2.5 mg/L (i.e., about 1 bubble/second in the
ammonia should be measured in all treatments at the overlying water). Dissolved oxygen and pH can be mea-
beginning and end of a test. Overlying water should be sured directly in the overlying waterwith a probe.
sampled just before water renewal from about 1 to 2 cm
above the sediment surface using a pipet. It may be
67
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13.3.6.2.3 Temperature should be measured at least
daily in at least one test chamber from each treatment.
The temperature of the water bath orthe exposure chamber
should be continuously monitored. The daily mean test
temperature must be within ±1°C of 23°C. The instanta-
neous temperature must always be within ±3°C of 23°C.
13.3.7 Ending a Test
13.3.7.1 Sediment at the end of the test can be sieved
through a fine-meshed screen sufficiently small to retain
the oligochaetes (e.g., U.S. standard sieve #40 (425-um
mesh) or#60 (250-um mesh). The sieved material should
be quickly transferred to a shallow pan to keep oligocha-
etes from moving through the screen. Immobile organ-
isms should be considered dead.
13.3.7.2 The sediment contribution to the body weight of
Lumbriculus variegatus is reported to be about 20% of the
wet weight and the contribution to chemical concentra-
tions ranges from 0 to 11% in two laboratory studies
(Kukkonen and Landrum, 1994; 1995). Analyses by
Mount et al. (1998) suggest that under certain conditions
substantially larger errors may occur if gut contents are
included in samples for tissue analysis. Accordingly,
after separating the organisms from the sediment, test
animals are held in clean water to allow the worms to
purge their guts of sediment. To initiate gut purging, live
oligochaetes are transferred from the sieved material to a
1 -L beaker containing overlying water only. Oligochaetes
should not be placed in clean sediment to eliminate gut
contents. Clean sediment can add to the dry weight of the
oligochaetes, which would result in a dilution of chemical
concentrations on a dry weight basis. Further, purging in
clean sediment is thought to accelerate depuration of
chemical from tissues (Kukkonen and Landrum, 1994).
The elimination beakers may need to be aerated to main-
tain dissolved oxygen above 2.5 mg/L.
13.3.7.3 The first edition of this manual (USEPA, 1994a)
specified a 24-h holding period for gut purging, based on
the findings of Call et al. (1991) who reported that
L. variegatus clear more than 90% of their gut contents in
24 h. Kukkonen and Landrum (1995) reported L. variega-
tus will purge out the intestinal contents in 10 h in water,
and more recently, Mount et al. (1999) found that gut
purging of L. variegatus was essentially complete in
only 6 h . Shorter purging periods may be preferable to
reduce depuration of chemical from tissue during holding
in clean water, particularly for compounds with log Kow
< 5 (Figure 13.1). Mount et al. (1999) estimated that aftera 6-h
purging period, compounds with log Kow > 3.85 would
remain at >90% of their initial concentrations, but after
24 h, only compounds with log Kow > 5 would be at >90%
of the initial concentration in tissue. Forthis reason, it is
recommended that the purging period last 6 to 8 h. Longer
purging periods (not to exceed 24 hours) may be used if
all target analytes have log Kow > 5.
13.3.7.4 Field-collected sediments may include indig-
enous oligochaetes. The behavior and appearance of
indigenous oligochaetes are usually different from L. var-
iegatus. It may be desirable to test extra chambers
without the addition of L. variegatus to check for the
presence of indigenous oligochaetes in field-collected
sediment (Phippsetal., 1993). Bioaccumulation of chemi-
cals by indigenous oligochaetes exposed in the same
chamber with introduced L. variegatus in a 28-d test has
been evaluated (Brunson etal., 1993). Peak concentrations
of select PAHs and DDT in this study were similar in the
indigenous oligochaetes and L. variegatus exposed in the
same chamber for 28 d.
13.3.7.5 Care should be taken to isolate at least the
minimum amount of tissue mass from each replicate
chamber needed for analytical chemistry.
13.3.8 Test Data
13.3.8.1 Sensitivity of tissue analyses is dependent
largely on the mass of tissue available and the sensitivity
of the analytical procedure. To obtain meaningful results
from bioaccumulation tests, it is essential that desired
detection limits be established before testing, and that the
test design allow for sufficient tissue mass. Tissue
masses required for various analyses at selected lower
limits of detection are listed in Table 13.5. Detection
limits for individual PAHs in tissue are listed in Table 13.6.
For most chemicals, a minimum mass of 1 g/replicate
(wet weight) and preferably 5 g/replicate (wet weight)
should be tested. Again, however, to insure results will be
meaningful, required masses for analytes of interest to
the study should be specifically evaluated before the
study is designed.
13.3.8.2 If an estimate of dry weight is needed, a
subsample should be dried to a constant weight at about
60 to 90°C. The sample is brought to room temperature in
a desiccator and weighed to the nearest 0.01 mg. Lum-
briculus variegatus typically contain about 1 % lipid (wet
weight). It may be desirable to determine ash-free dry
weight (AFDW) of oligochaetes instead of dry weight.
Measurement of AFDW is recommended over dry weight
for C. tentans due to the contribution of sediment in the
gut to the weight of midge (Section 12.3.8; Sibley et al.,
1997b). Additional data are needed to determine the
contribution of sediment in the gut of L variegatusio body
weight before a definitive recommendation can be made
to measure AFDW of oligochaetes routinely.
13.3.8.3 Depending on specific study objectives, total
lipids can be measured on a subsample of the total tissue
mass of each thawed replicate sample. Gardner et al.
(1985) describe procedures for measuring lipids in 1 mg of
tissue. Different methods of lipid analysis can yield differ-
ent results (Randall et al., 1991). The analytical method
used for lipid analysis should be calibrated against the
chloroform-methanol extraction method described by Folch
et al. (1957) and Bligh and Dyer (1959).
13.3.8.3.1 A number of studies have demonstrated that
lipids are the major storage site for organic chemicals in a
variety of organisms (Roberts et al., 1977; Oliver and
Niimi, 1983; de Boer, 1988). Because of the importance of
68
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0
S»
—j
Q-
jr-
"*
m
0 .
0)1
13 •
Q.
r- 1
A» •
CD .
s»l
3
D_ |
i •
CO 1
0
i"!
CL •
x:
C^J I
•^ '
0 '
|>i
a. i
CO
•*- i
0
E>
Q_
x:
i
CN
CO
- N.
O
D)
O
- 10
O
CM
O
O
O
00
O
CD
o
CN
Figure 13.1 Predicted depuration of nonionic organic chemicals from tissue of Lumbriculus variegatus as a function of Ko and
duration of depuration, assuming no contribution of sediment in the gut. Shaded area represents tlO% of tissue
concentration at the beginning of the depuration period (Mount et al., 1999).
69
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Table 13.5 Grams of Lumbriculus variegatus Tissue (Wet
Weight) Required
for Various Analytes at
Selected Lower Limits of
Analyte
PCBs
PCB (total1)
PCB (congener2)
Level of chlorination
mono-trichloro
tetra-hexachloro
hepta-octachloro
nona-decachloro
Organochlorine pesticides 1
p,p' DDE
p,p' - ODD
p,p' - DDT
o,p' - DDE
o,p' ODD
o,p' DDT
Alpha-chlordane
Gamma-chlordane
Dieldrin
Endrin
Heptachlorepoxide
Oxychlordane
Mi rex
Trans - nonachlor
Toxaphene
PAHs 3
PAHs
Dioxins 4
TCDD (ng/g)
Inorganic 5
Cadmium
Copper
Lead
Zinc
1 Schmitt etal., 1990
2 USEPA, 1990c
3 \ /~,»~»~;i~,i-n»~ «* -.1 -i noo
1.0
Lower
0.600
0.025
0.050
0.075
0.125
0.050
0.050
0.050
0.050
0.050
0.050
0.050
0.050
0.050
0.050
0.050
0.050
0.050
0.050
0.600
0.012
0.020
0.005
0.005
0.005
0.005
Detection
Grams of Tissue
2.0
Limit of Detection
0.300
0.0125
0.025
0.0375
0.0625
0.025
0.025
0.025
0.025
0.025
0.025
0.025
0.025
0.025
0.025
0.025
0.025
0.025
0.025
0.300
0.006
0.010
0.0025
0.0025
0.0025
0.0025
5.0
(ug/g)
0.120
0.005
0.010
0.015
0.025
0.010
0.010
0.010
0.010
0.010
0.010
0.010
0.010
0.010
0.010
0.010
0.010
0.010
0.010
0.120
0.002
0.004
0.001
0.001
0.001
0.001
Table 13.6 Detection Limits (ng) of Individual PAHs by
HPLC-FD1
Analyte Detection Limit (ng)
Benzo(a)pyrene 0.01
Pyrene 0.03
Benzo(k)fluoranthene 0.03
Dibenz(a,h)anthracene 0.03
Anthracene 0.10
Benz(a)anthracene 0.10
Benzo(e)pyrene 0.10
Benzo(b)fluoranthene 0.10
Benzo(g,h,i)perylene 0.10
3-Methyleholanthrene 0.10
1 Obana etal., 1981
lipids, it may be desirable to normalize bioaccumulated
concentrations of nonpolar organics to the tissue lipid
concentration. Lipid concentration is one of the factors
required in deriving the BSAF (Section 16). However, the
difficulty with using this approach is that each lipid method
generates different lipid concentrations (see Kates (1 986)
for discussion of lipid methodology). The differences in
lipid concentrations directly translate to a similar variation
in the lipid-normalized chemical concentrations or BSAF.
13.3.8.3.2 For comparison of lipid-normalized tissue
residues or BASFs, it is necessary to either promulgate a
standard lipid technique or to intercalibrate the various
techniques. Standardization of a single method is difficult
because the lipid methodology is often intimately tied in
with the extraction procedure for chemical analysis. As an
interim solution, the Bligh-Dyer lipid method (Bligh and
Dyer, 1 959) is recommended as a temporary "intercalibration
standard" (ASTM, 1999c).
13.3.8.3.3 The potential advantages of Bligh-Dyer in-
clude its ability to extract neutral lipids not extracted by
many other solvent systems and the wide use of this
method (or the same solvent system) in biological and
toxicological studies (e.g., Roberts et al., 1977; Oliver
and Niimi, 1 983; de Boer, 1 988; Landrum, 1 989). Because
the technique is independent of any particular analytical
extraction procedure, it will not change when the extrac-
tion technique is changed. Additionally, the method can
be modified for small tissue sample sizes as long as the
solvent ratios are maintained (Herbes and Allen, 1983;
Gardner etal., 1985).
4 USEPA, 1990d
5 Schmitt and Finger, 1987
13.3.8.3.4 If the Bligh-Dyer method is not the primary
lipid method used, the chosen lipid analysis method
should be compared with Bligh-Dyer for each tissue type.
The chosen lipid method can then be converted to
"Bligh-Dyer" equivalents and the lipid-normalized tissue
70
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residues reported in "Bligh-Dyer equivalents." In the in-
terim, it is suggested that extra tissue of each species be
frozen for future lipid analysis in the event that a different
technique proves more advantageous (ASTM, 1999c).
13.4 Interpretation of Results
13.4.1 Section 16 describes general information for inter-
pretation of test results. The following sections describe
species-specific information that is useful in helping to
interpret the results of sediment bioaccumulation tests
with L. variegatus.
13.4.2 Duration of Exposure
13.4.2.1 Because data from bioaccumulation tests often
will be used in ecological or human health risk assess-
ments, the procedures are designed to generate quantita-
tive estimates of steady-state tissue residues. Eighty
percent of steady state is used as the general criterion
(ASTM, 1999c). Because results from a single or few
species often will be extrapolated to other species, the
procedures are designed to maximize exposure to
sediment-associated chemicals so as not to systemati-
cally underestimate residues in untested species.
13.4.2.2 A kinetic study can be conducted to estimate
steady-state concentrations instead of conducting a 28-d
bioaccumulation test (e.g..sample on Day 1, 3, 7,14,28;
Brunson et al., 1993; USEPA-USACE, 1991). A kinetic
test conducted under the same test conditions outlined
above, can be used when 80% of steady state will not be
obtained within 28 d or when more precise estimates of
steady-state tissue residues are required. Exposures
shorter than 28 d may be used to determine whether
compounds are bioavailable (i.e., bioaccumulation
potential).
13.4.2.3 DDT reportedly reached 90% of steady state by
Day 14 of a 56-d exposure with L variegatus. However,
low molecular weight PAHs (e.g., acenaphthylene, fluo-
rene, phenanthrene) generally peaked at Day 3 and tended
to decline to Day 56 (Brunson et al., 1993). In general,
concentrations of high molecular weight PAHs (e.g.,
benzo[b]fluoranthene, benzo[e]pyrene, indeno-
[1,2,3-c,d]pyrene) either peaked at Day 28 or continued to
increase during the 56-d exposure.
13.4.3 Influence of Indigenous Organisms
13.4.3.1 Field-collected sediments may include indig-
enous oligochaetes. Phipps et al. (1993) recommend test-
ing extra chambers without the addition of L variegatusio
check for the presence of indigenous oligochaetes in
field-collected sediment.
13.4.4 Sediment Toxicity in Bioaccumulation
Tests
13.4.4.1 Toxicity or altered behavior of organisms in a
sample may not preclude use of bioaccumulation data;
however, information on adverse effects of a sample
should be included in the report.
13.4.4.2 Grain Size.
13.4.4.2.1 Lumbriculus variegatus are tolerant of a wide
range of substrates. Physico-chemical characteristics (e.g.,
grain size) of sediment were not significantly correlated to
the growth or reproduction of L. variegatus in 10-d toxicity
tests (see Section 10.1.3.3; Ankley etal., 1994a).
13.4.4.3 Sediment Organic Carbon
13.4.4.3.1 Reduced growth of L variegatus may result
from exposure to sediments with low organic carbon con-
centrations (G.T. Ankley, USEPA, Duluth, MN, personal
communication). Forthis reason, reduced growth observed
in bioaccumulation tests could be caused by either direct
toxicity or insufficient nutrition of the sediment. Testing
additional replicate chambers with supplemental food could
be used to help make this distinction, although the effect
of added food on accumulation of chemicals would need to
be considered in the test interpretation.
13.4.4.4 Ammonia Toxicity
13.4.4.4.1 Section 1.3.7.5 addresses interpretative guid-
ance for evaluating toxicity associated with ammonia in
sediment.
71
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Section 14
Test Method 100.4
Hyalella azteca 42-d Test for Measuring the Effects of Sediment-
associated Contaminants on Survival, Growth, and Reproduction
14.1 Introduction
14.1.1 Hyalella azteca are routinely used to assess the
toxicity of chemicals in sediment (Section 11; Nebekeret
al., 1984; Dillon and Gibson,1986; Burton et al., 1989;
Burton et al., 1992; Ingersoll and Nelson, 1990; Borgmann
and Munawar, 1989; Ankley et al., 1994; Winger and
Lazier, 1994; Suedeland Rodgers, 1994; Dayetal., 1995;
Kubitz et al.,1996). Test duration and endpoints recom-
mended in previously developed standard methods for
sediment testing with H. azteca include 10-d survival
(Section 11; USEPA, 1994a) and 10- to 28-d survival and
growth (ASTM, 1999a; Environment Canada, 1998a). Short-
term exposures which only measure effects on survival
can be used to identify high levels of contamination, but
may not be able to identify marginally contaminated sedi-
ments. The method described in this section can be used
to evaluate potential effects of contaminated sediment on
survival, growth, and reproduction of H. azteca in a
42-d test.
14.1.2 Section 14.2 describes general guidance for con-
ducting a 42-d test with H. azteca that can be used to
evaluate the effects of contaminants associated with
sediments on survival, growth and reproduction. Refine-
ments of these methods may be described in future
editions of this manual after additional laboratories have
successfully used the method (Section 17.6). The 42-d
test with H. azteca has not been adequately evaluated in
water with elevated salinity (Section 1.3.2).
14.1.3 The procedure outlined in Section 14.2 is based
on procedures described in Ingersoll et al. (1998). The
sediment exposure starts with 7- to 8-d-old amphipods.
On Day 28, amphipods are isolated from the sediment
and placed in water-only chambers where reproduction is
measured on Day 35 and 42. Typically, amphipods are
first in amplexus at about Day 21 to 28 with release of the
first brood between Day 28 to 42. Endpoints measured
include survival (Day 28, 35 and 42), growth (as length or
dry weight measured on Day 28 and 42), and reproduction
(number of young/female produced from Day 28 to 42).
The procedures described in Table 14.1 include measure-
ment of a variety of lethal and sublethal endpoints; minor
modifications of the basic methods can be used in cases
where only a subset of these endpoints is of interest.
14.1.3.1 Several designs were considered for measuring
reproduction in sediment exposures based on the repro-
ductive biology of/-/, azteca (Ingersoll et al., 1998). The
first design considered was a continuation of the 28-d
sediment exposures described in Ingersoll et al. (1996) for
an additional two weeks to determine the number of young
produced in the first brood. The limitation of this design is
the difficulty in quantitatively isolating young amphipods
from sediment (Tomasovicetal., 1995). A second design
considered was extension of the 28-d sediment exposure
for an additional month or longer until several broods are
released. These multiple broods could then be isolated
from the sediment. The limitation of this second design is
that specific effects on reproduction could not be differen-
tiated from reduced survival of offspring and it would still
be difficult to isolate the young amphipods from sediment.
A third design considered, and the one described in this
manual, was to expose amphipods in sediment until a few
days before the release of the first brood. The amphipods
could then be sieved from the sediment and held in water
to determine the number of young produced (Ingersoll et
al., 1998). This test design allows a quantitative measure
of reproduction. One limitation to this design is that
amphipods might recover from effects of sediment expo-
sure during this holding period in clean water (Landrum
and Scavia, 1983; Kane Driscoll et al., 1997); however,
amphipods are exposed to sediment during critical devel-
opmental stages before release of the first brood in clean
water.
14.1.4 The method has been used to evaluate a formu-
lated sediment and field-collected sediments with low to
moderate concentrations of contaminants (Ingersoll et al.,
1998). Survival of amphipods in these sediments was
typically >85% afterthe 28-d sediment exposures and the
14-d holding period in water to measure reproduction
(Ingersoll et al., 1998). The method outlined in 14.2 has
also been evaluated in round-robin testing with 8 to 12
laboratories (Section 17.6). Afterthe 28-d sediment expo-
sures in a control sediment (West Bearskin), survival was
>80% for >88% of the laboratories; length was >3.2 mm/
individual for >71% of the laboratories; and dry weight
was >0.15 mg/individual for >66% of the laboratories.
Reproduction from Day 28 to Day 42 was >2 young/
female for >71% of the laboratories participating in the
round-robin testing. Reproduction was more variable within
and among laboratories; hence, more replicates might be
72
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Table 14.1 Test Conditions for Conducting a 42-d
and ASTM 1999a).
Parameter
Sediment Toxicity Test with Hyalella azteca (modified from USEPA 1994a
Conditions
1. Test type:
2. Temperature:
3. Light quality:
4. Illuminance:
5. Photoperiod:
6. Test chamber:
7. Sediment volume:
8. Overlying water volume:
9. Renewal of overlying water:
10. Age of organisms:
11. Number of organisms/chamber:
12. Number of replicate chambers/treatment:
13. Feeding:
14. Aeration:
15. Overlying water:
16. Test chamber cleaning:
17. Overlying water quality:
18. Test duration:
19. Endpoints:
20. Test acceptability:
Whole-sediment toxicity test with renewal of overlying water
23 ± 1 °C
Wide-spectrum fluorescent lights
About 100 to 1000 lux
16L8D
300-mL high-form lipless beaker
100 ml
175 ml in the sediment exposure from Day 0 to Day 28 (175 to 275 ml in the water-
only exposure from Day 28 to Day 42)
2 volume additions/d (Appendix A); continuous or intermittent (e.g., one volume
addition every 12 h)
7- to 8-d old at the start of the test
10
12 (4 for 28-d survival and growth and 8 for 35- and 42-d survival, growth, and
reproduction). Reproduction is more variable than growth or survival; hence, more
replicates might be needed to establish statistical differences among treatments
(See Section 14.2.3).
YCT food, fed 1.0 ml (1800 mg/L stock) daily to each test chamber.
None, unless dissolved oxygen in overlying water drops below 2.5 mg/L.
Culture water, well water, surface water or site water. Use of reconstituted water
is not recommended.
If screens become clogged during a test, gently brush the outside of the screen
(Appendix A).
Hardness, alkalinity, conductivity, and ammonia at the beginning and end of a sediment
exposure (Day 0 and 28). Temperature daily. Conductivity weekly. Dissolved oxygen
(DO) and pH three times/week. Concentrations of DO should be measured more often
if DO drops more than 1 mg/L since the previous measurement.
42 d
28-d survival and growth; 35-d survival and reproduction; and 42-d survival, growth,
reproduction, and number of adult males and females on Day 42
Minimum mean control survival of 80% on Day 28. Additional performance-based
criteria specifications are outlined in Table 14.3 based on results of round-robin
testing (Sections 14.1.4 and 17.6).
needed to establish statistical differences among treat-
ments with this endpoint.
14.1.5 Growth of H. azteca in sediment tests often
provides unique information that can be used to
discriminate toxic effects of exposure to contaminants
(Brasher and Ogle, 1993; Borgmann, 1994; Kembleetal.,
1994; Ingersoll et al., 1996; Kubitz et al., 1996; Milan! et
al., 1996; Steevens and Benson, 1998). Either length or
weight can be measured in sediment tests with H. azteca.
However, additional statistical options are available if
length is measured on individual amphipods, such as
nested analysis of variance which can account for vari-
ance in length between replicates (Steevens and Benson,
1998). Ongoing water-only studies testing select
contaminants will provide additional data on the relative
sensitivity and variability of sublethal endpoints in toxicity
tests with H. azteca (Ingersoll et al., 1998).
14.1.6 Results of tests using procedures different from
the procedures described in Section 14.2 may not be
comparable, and these different procedures may alter
contaminant bioavailability. Comparisons of results ob-
tained using modified versions of these procedures might
provide useful information concerning new concepts and
procedures for conducting sediment tests with aquatic
organisms. If tests are conducted with procedures differ-
ent from the procedures described in this manual, addi-
tional tests are required to determine comparability of
results (Section 1.3).
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14.2 Procedure for Conducting a Hyalella nance of about 100 to 1000 lux (Table 14.1). Testcham-
azteca 42-d Test for Measuring the bers are 300-ml_ high-form lipless beakers containing
Effprte of <5pdimpnt a«5«5oriatpd 10° ml of sediment and 175 ml of overlying water. Ten
Effects ot bediment-associated amphipods in each test chamber are fed 1.0 ml of YCT
uontammants on survival, urowtn, daHy (Appendix B). Each test chamber receives 2 volume
and Reproduction additions/d of overlying water. Water renewals may be
manual or automated. Appendix A describes water-re-
14.2.1 Conditions for evaluating sublethal endpoints in a newa| systems that can be used to deliver overlying
sediment toxicity test with H. azteca are summarized in water. Overlying water should be a source of water that
Table 14.1. A general activity schedule is outlined in nas been demonstrated to support survival, growth, and
Table 14.2. Decisions concerning the various aspects of reproduction of H. azteca in culture. McNulty et al. (1999)
experimental design, such as the number of treatments, and Kemble et al. (1999) observed poor survival of
number of test chambers/treatment, and water-quality H azteca in tests conducted 14 to 28 d using a variety of
characteristics should be based on the purpose of the test reconstituted waters including the reconstituted water
and the methods of data analysis (Section 16). When (reformulated moderately hard reconstituted water) de-
variability remains constant, the sensitivity of a test scribed in Smith et a, (1997) and described in the first
increases as the number of replicates increase. edition of this manua, (USEPA, 1994a). Borgmann (1996)
described a reconstituted waterthat was used successfully
14.2.2 The 42-d sediment toxicity test with H. azteca is to maintain H. azteca in culture; however, some laborato-
conducted at 23°C with a 16L8D photoperiod at an illumi- ries have not had success when using this reconstituted
Table 14.2 General Activity Schedule for Conducting a 42-d Sediment Toxicity Test with Hyalella azteca
Day Activity
Pre-Test
-8 Separate known-age amphipods from the cultures and place in holding chambers. Begin preparing food for the test. The <24-h
amphipods are fed 10 ml of YCT (1800 mg/L stock solution) and 10 ml of Selenastrum capricornutum (about 3.Ox 107cells/mL)
on the first day of isolation and 5 ml of both YCT and S. capricornutum on the 3rd and 5th d after isolation.
-7 Remove adults and isolate <24-h-old amphipods (if procedures outlined in Section 10.3.4 are followed).
-6 to -2 Feed and observe isolated amphipods (Section 10.3), monitor water quality (e.g., temperature and dissolved oxygen).
-1 Feed and observe isolated amphipods (Section 10.3), monitor water quality. Add sediment into each test chamber, place chambers
into exposure system, and start renewing overlying water.
Sediment Test
0 Measure total water quality (pH, temperature, dissolved oxygen, hardness, alkalinity, conductivity, ammonia). Transfer ten 7- to
8-d-old amphipods into each test chamber. Release organisms under the surface of the water. Add 1.0 ml of YCT (1800 mg/L
stock) into each test chamber. Archive 20 test organisms for length determination or archive 80 test organisms for dry weight
determination. Observe behavior of test organisms.
1 to 27 Add 1.0 ml of YCT to each test beaker. Measure temperature daily, conductivity weekly, and dissolved oxygen (DO) and pH
three times/week. Observe behavior of test organisms.
28 Measure temperature, dissolved oxygen, pH, hardness, alkalinity, conductivity and ammonia. End the sediment-exposure portion
of the test by collecting the amphipods with a #40-mesh sieve (425-um mesh; U.S. standard size sieve). Use four replicates
for growth measurements: count survivors and preserve organisms in sugar formalin for growth measurements. Use eight
replicates for reproduction measurements: place survivors in individual replicate water-only beakers and add 1.0 ml of YCT to
each test beaker/d and 2 volume additions/d (Appendix A) of overlying water.
Reproduction Phase
29 to 35 Feed daily (1.0 ml of YCT). Measure temperature daily, conductivity weekly, and DO and pH three times a week. Measure
hardness and alkalinity weekly. Observe behavior of test organisms.
35 Record the number of surviving adults and remove offspring. Return adults to their original individual beakers and add food.
36 to 41 Feed daily (1.0 ml of YCT). Measure temperature daily, conductivity weekly, and DO and pH three times a week. Measure
hardness and alkalinity weekly. Observe behavior of test organisms.
41 Measure total water quality (pH, temperature, dissolved oxygen, hardness, alkalinity, conductivity, ammonia).
42 Record the number of surviving adults and offspring. Surviving adult amphipods on Day 42 are preserved in sugar formalin solution.
The number of adult males in each beaker is determined from this archived sample. This information is used to calculate the number
of young produced per female per replicate from Day 28 to Day 42.
74
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water in the 42-d test (T.J. Norberg-King, USEPA, Duluth, growth and survival endpoints and the other 8 replicates
MN, personal communication). For site-specific are used for measurement of survival and reproduction on
evaluations, the characteristics of the overlying water Day 35 and for measurement of survival, reproduction, or
should be as similar as possible to the site where sedi- growth on Day 42.
ment is collected. Requirements fortest acceptability are
summarized in Table 14.3. 14.3 General Procedures
14.2.3 The number of replicates and concentrations 14.3.1 Sediment into Test Chambers
tested depends in part on the significance level selected
and the type of statistical analysis. A total of 12 repli- 14.3.1.1 The day before the sediment test is started
cates, each containing ten 7- to 8-d-old amphipods, are (Day -1) each sediment should be thoroughly homog-
tested for each treatment. Starting the test with substan- enized and added to the test chambers (Section 8.3.1).
tially younger or older organisms may compromise the Sediment should be visually inspected to judge the de-
reproductive endpoint. For the total of 12 replicates the gree of homogeneity. Excess water on the surface of the
assignment of beakers is as follows: 12 replicates are set sediment can indicate separation of solid and liquid corn-
up on Day -1 of which 4 replicates are used for 28-d ponents. If a quantitative measure of homogeneity is
Table 14.3 Test Acceptability Requirements for a 42-d Sediment Toxicity Test with Hyalella azteca
A. It is recommended for conducting the 42-d test with H. azteca that the following performance criteria be met:
1. Age of H. azteca at the start of the test should be 7- to 8-d old. Starting a test with substantially younger or older organisms may
compromise the reproductive endpoint.
2. Average survival of H. azteca in the control sediment on Day 28 should be greater than or equal to 80%.
3. Laboratories participating in round-robin testing (Section 17.6) reported after 28-d sediment exposures in a control sediment
(West Bearskin), survival >80% for >88% of the laboratories; length >3.2 mm/individual for >71% of the laboratories; and dry
weight >0.15 mg/individual for >66% of the laboratories. Reproduction from Day 28 to Day 42 was >2 young/female for >71% of
the laboratories participating in the round-robin testing. Reproduction was more variable within and among laboratories; hence,
more replicates might be needed to establish statistical differences among treatments with this endpoint.
4. Hardness, alkalinity, and ammonia in the overlying water typically should not vary by more than 50% during the sediment
exposure, and dissolved oxygen should be maintained above 2.5 mg/L in the overlying water.
B. Performance-based criteria for culturing H. azteca include the following:
1. It may be desirable for laboratories to periodically perform 96-h water-only reference-toxicity tests to assess the sensitivity of
culture organisms (Section 9.16.2). Data from these reference-toxicity tests could be used to assess genetic strain or life-stage
sensitivity of test organisms to select chemicals.
2. Laboratories should track parental survival in the cultures and record this information using control charts if known-age cultures
are maintained. Records should also be kept on the frequency of restarting cultures and the age of brood organisms.
3. Laboratories should record the following water-quality characteristics of the cultures at least quarterly: pH, hardness, alkalinity,
and ammonia. Dissolved oxygen in the cultures should be measured weekly. Temperature of the cultures should be recorded
daily. If static cultures are used, it may be desirable to measure water quality more frequently.
4. Laboratories should characterize and monitor background contamination and nutrient quality of food if problems are observed in
culturing or testing organisms.
5. Physiological measurements such as lipid content might provide useful information regarding the health of the cultures.
C. Additional requirements:
1. All organisms in a test must be from the same source.
2. Storage of sediments collected from the field should follow guidance outlined in Section 8.2.
3. All test chambers (and compartments) should be identical and should contain the same amount of sediment and overlying water.
4. Negative-control sediment and appropriate solvent controls must be included in a test. The concentration of solvent used must not
adversely affect test organisms.
5. Test organisms must be cultured and tested at 23°C (±1°C).
6. The mean of the daily test temperature must be within ±1°C of 23°C. The instantaneous temperature must always be within ±3°C
of23°C.
7. Natural physico-chemical characteristics of test sediment collected from the field should be within the tolerance limits of the test
organisms.
75
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required, replicate subsamples should be taken from the
sediment batch and analyzed forTOC, chemical concen-
trations, and particle size.
14.3.1.2 Each test chamber should contain the same
amount of sediment, determined either by volume or by
weight. Overlying water is added to the chambers on
Day -1 in a manner that minimizes suspension of sedi-
ment. This can be accomplished by gently pouring water
along the sides of the chambers or by pouring water onto
a baffle (e.g., a circular piece of Teflon with a handle
attached) placed above the sediment to dissipate the
force of the water. Renewal of overlying water is started
on Day -1. A test begins when the organisms are added to
the test chambers (Day 0).
14.3.2 Renewal of Overlying Water
/
14.3.2.1 Renewal of overlying water is required during a
test. At any particular time during a test, flow rates
through any two test chambers should not differ by more
than 10%. Hardness, alkalinity and ammonia
concentrations in the water above the sediment, within a
treatment, typically should not vary by more than 50%
during the test. Mount and Brungs (1967) diluters have
been modified for sediment testing, and other automated
water-delivery systems have also been used (Maki, 1977;
Ingersoll and Nelson, 1990; Benoit et al., 1993; Zumwalt
et al., 1994; Brunson et al., 1998; Wall et al., 1998;
Leppanen and Maier, 1998). The water-delivery system
should be calibrated before a test is started to verify that
the system is functioning properly. Renewal of overlying
water is started on Day -1 before the addition of test
organisms or food on Day 0. Appendix A describes
water-renewal systems that can be used for conducting
sediment tests.
14.3.2.2 In water-renewal tests with one to four volume
additions of overlying water/d, water-quality characteristics
generally remain similar to the inflowing water (Ingersoll
and Nelson, 1990; Ankley et al., 1993); however, in static
tests, water quality may change profoundly during the
exposure (Shuba et al., 1978). For example, in static
whole-sediment tests, the alkalinity, hardness, and con-
ductivity of overlying water more than doubled in several
treatments during a four-week exposure (Ingersoll and
Nelson, 1990). Additionally, concentrations of metabolic
products (e.g., ammonia) may also increase during static
exposures, and these compounds can either be directly
toxic to the test organisms or may contribute to the
toxicity of the contaminants in the sediment. Further-
more, changes in water-quality characteristics such as
hardness may influence the toxicity of many inorganic
(Gauss et al., 1985) and organic (Mayer and Ellersieck,
1986) contaminants. Although contaminant concentra-
tions are reduced in the overlying water in water-renewal
tests, organisms in direct contact with sediment generally
receive a substantial proportion of a contaminant dose
directly from either the whole sediment or from the
pore water.
14.3.3 Acclimation
14.3.3.1 Test organisms must be cultured and tested at
23°C. Ideally, test organisms should be cultured in the
same water that will be used in testing. However, acclima-
tion of test organisms to the test water is not required.
14.3.3.2 Culturing of organisms and toxicity assessment
are typically conducted at 23°C. However, occasionally
there is a need to perform evaluations at temperatures
different than that recommended. Under these
circumstances, it may be necessary to acclimate organ-
isms to the desired test temperature to prevent thermal
shock when moving immediately from the culture tem-
perature to the test temperature (ASTM, 1999a). Accli-
mation can be achieved by exposing organisms to a
gradual change in temperature; however, the rate of change
should be relatively slow to prevent thermal shock. A
change in temperature of 1°C every 1 to 2 h has been
used successfully in some studies (P.K. Sibley, Univer-
sity of Guelph, Guelph, Ontario, personal communication;
APHA, 1989). Testing at temperatures other than 23°C
needs to be preceded by studies to determine expected
performance under alternate conditions.
14.3.4 Placing Organisms in Test Chambers
14.3.4.1 Test organisms should be handled as little as
possible. Amphipods should be introduced into the overly-
ing water below the air-water interface. Test organisms
can be pipetted directly into overlying water. The size of
the test organisms at the start of the test should be
measured using the same measure (length orweight) that
will be used to assess their size at the end of the test. For
length, a minimum of 20 organisms should be measured.
Forweight measurement, a larger sample size (e.g., 80)
may be desirable because of the relatively small mass of
the organisms. This information can be used to deter-
mine consistency in the size of the organisms used to
start a test.
14.3.5 Feeding
14.3.5.1 For each beaker, 1.0 ml of YCT is added from
Day 0 to Day 42. Without addition of food, the test
organisms may starve during exposures. However, the
addition of the food may alter the availability of the
contaminants in the sediment (Wiederholm et al., 1987;
Harkey et al., 1994). Furthermore, if too much food is
added to the test chamber, or if the mortality of test
organisms is high, fungal or bacterial growth may develop
on the sediment surface. Therefore, the amount of food
added to the test chambers is kept to a minimum.
14.3.5.2 Suspensions of food should be thoroughly mixed
before aliquots are taken. If excess food collects on the
sediment, a fungal or bacterial growth may develop on the
sediment surface, in which case feeding should be sus-
pended for one or more days. A drop in dissolved oxygen
below 2.5 mg/L during a test may indicate that the food
added is not being consumed. Feeding should be sus-
pended for the amount of time necessary to increase the
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dissolved oxygen concentration (ASTM, 1999a). If feed-
ing is suspended in one treatment, it should be sus-
pended in all treatments. Detailed records of feeding rates
and the appearance of the sediment surface should be
made daily.
14.3.6 Monitoring a Test
14.3.6.1 All chambers should be checked daily and
observations made to assess test organism behavior
such as sediment avoidance. However, monitoring ef-
fects on burrowing activity of test organisms may be
difficult because the test organisms are often not visible
during the exposure. The operation of the exposure sys-
tem should be monitored daily.
14.3.6.2 Measurement of Overly ing Water-quality
Characteristics
14.3.6.2.1 Conductivity, pH, DO, hardness, alkalinity,
and ammonia should be measured in all treatments at the
beginning and at the end of the sediment exposure portion
of the test. Water-quality characteristics should also be
measured at the beginning and end of the reproductive
phase (Day 29 to Day 42). Conductivity should be mea-
sured weekly, whereas pH and DO should be measured
three times/week (Section 14.3.6.2.2). Overlying water
should be sampled just before water renewal from about
1 to 2 cm above the sediment surface using a pipet. It
may be necessary to composite water samples from
individual replicates. The pipet should be checked to
make sure no organisms are removed during sampling of
overlying water.
14.3.6.2.2 Dissolved oxygen should be measured three
times/week and should be at a minimum of 2.5 mg/L. If a
probe is used to measure dissolved oxygen in overlying
water, it should be thoroughly inspected between samples
to make sure that organisms are not attached and should
be rinsed between samples to minimize cross contamina-
tion. Aeration can be used to maintain dissolved oxygen
in the overlying water above 2.5 mg/L (i.e., about 1
bubble/second in the overlying water). Dissolved oxygen
and pH can be measured directly in the overlying water
with a probe.
14.3.6.2.3 Temperature should be measured at least
daily in at least one test chamber from each treatment.
The temperature of the water bath orthe exposure cham-
ber should be continuously monitored. The daily mean
test temperature must be within ±1 °C of 23°C. The instan-
taneous temperature must always be within ±3°C of 23°C.
14.3.7 Ending a Test
14.3.7.1 Endpoints monitored include 28-d survival and
growth of amphipods and 35-d and 42-d survival, growth,
and reproduction (number of young/female) of amphipods.
Growth or reproduction of amphipods may be a more
sensitive toxicity endpoint compared to survival (Burton
and Ingersoll, 1994; Kembleetal., 1994; Ingersoll et al.,
1998).
14.3.7.2 On Day 28, 4 of the replicate beakers/sediment
are sieved with a #40-mesh sieve (425-um mesh; U.S.
standard size sieve) to remove surviving amphipods for
growth determinations. Any of the surviving amphipods in
the water column or on the surface of the sediment can be
pipetted from the beaker before sieving the sediment. The
sediment in each beaker should be sieved in two separate
aliquots (i.e., most of the amphipods will probably be
found in the surface aliquot). Immobile organisms isolated
from the sediment surface or from sieved material should
be considered dead. Surviving amphipods from these
4 replicates can be preserved in separate vials containing
8% sugar formalin solution if length of amphipods is to be
measured (Ingersoll and Nelson, 1990). The sugar forma-
lin solution is prepared by adding 120 g of sucrose to
80 ml of formalin which is then brought to a volume of 1 L
using deionized water. This stock solution is mixed with
an equal volume of deionized water when used to pre-
serve organisms. NoTox® (Earth Safe Industries, Belle
Mead, NJ) can be used as a substitute for formalin (Linger
etal.,1993).
14.3.7.3 A consistent amount of time should be taken to
examine sieved material for recovery of test organisms
(e.g., 5 min/replicate). Laboratories should demonstrate
that their personnel are able to recover an average of at
least 90% of the organisms from whole sediment. For
example, test organisms could be added to control ortest
sediments, and recovery could be determined after 1 h
(Tomasovicet al., 1994).
14.3.7.4 Growth of amphipods can be reported as either
length or weight; however, additional statistical options
are available if length is measured on individual organ-
isms (Section 14.4.5.3).
14.3.7.5 Amphipod body length (±0.1 mm) can be mea-
sured from the base of the first antenna to the tip of the
third uropod along the curve of the dorsal surface (Figure
11.1). Kembleetal. (1994) describe the use of a digitizing
system and microscope to measure lengths of/-/, azteca.
Kemble et al. (1994) also photographed invertebrates (at a
magnification of 3.5X) and measured length using a com-
puter-interfaced digitizing tablet.
14.3.7.6 Dry weight of amphipods in each replicate can
be determined on Day 28 and 42. If both weight and
length are to be determined, weight should be measured
after length on the preserved samples. Gaston et al.
(1995) and Duke et al. (1996) have shown that biomass or
length of several aquatic invertebrates did not signifi-
cantly change after two to four weeks of storage in 10%
formalin. If test organisms are to be used for an evalua-
tion of bioaccumulation, it is not advisable to dry the
sample before conducting the residue analysis. If conver-
sion from wet weight to dry weight is necessary, aliquots
of organisms can be weighed to establish wet to dry
weight conversion factors. A consistent procedure should
be used to remove the excess water from the organisms
before measuring wet weight.
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14.3.7.7 Dry weight of amphipods can be determined as
follows: (1) transferring the archived amphipods from a
replicate out of the sugar formalin solution into a crystal-
lizing dish; (2) rinsing amphipods with deionized water;
(3) transferring these rinsed amphipods to a preweighed
aluminum pan; (4) drying these samples for 24 h at60°C;
and (5) weighing the pan and dried amphipods on a
balance to the nearest 0.01 mg. Average dry weight of
individual amphipods in each replicate is calculated from
these data. Due to the small size of the amphipods,
caution should be taken during weighing (10 dried amphi-
pods after a 28-d sediment exposure may weigh less than
2.5 to 3.5 mg). Weigh pans need to be carefully handled
using powder-less gloves and the balance should be
calibrated with standard weights with each use. Use of
small aluminum pans (e.g., 7 x 22 x 7 mm, Sigma
Chemical Company, St. Louis, MO) will help reduce vari-
ability in measurements of dry weight. Weigh boats can
also be constructed from sheets of aluminum foil.
14.3.7.8 The first edition of this manual (USEPA, 1994a)
recommended dry weight as a measure of growth for both
H. azteca and C. tentans. For C. tentans, this recommen-
dation was changed in the current edition to ash-free dry
weight (AFDW) instead of dry weight, with the intent of
reducing bias introduced by gut contents (Sibley et al.,
1997a). However, this recommendation was not ex-
tended to include H. azteca. Studies by Dawson et al.
(personal communication, T.D. Dawson, Integrated Labo-
ratory Systems, Duluth, MN) have indicated that the ash
content of H. azteca is not greatly decreased by purging
organisms in clean water before weighing, suggesting that
sediment does not comprise a large portion of the overall
dry weight. In addition, using AFDW further decreases an
already small mass, potentially increasing measurement
error. For this reason, dry weight continues to be the
recommended endpoint for estimating growth of/-/, azteca
via weight (growth can also be determined via length).
14.3.7.9 On Day 28, the remaining 8 beakers/sediment
are also sieved and the surviving amphipods in each
sediment beaker are placed in 300-mL water-only beakers
containing 150 to 275 mL of overlying water and a 5-cm x
5-cm piece of Nitex screen (Nylon Bolting cloth; 44%
open area and 280-um aperture, Wildlife Supply Com-
pany, Saginaw, Ml; Ingersoll et al., 1998). In a subse-
quent study, improved reproduction of H. azteca was
observed when the Nitex screen was replaced with a 3-cm
x3-cm piece of the nylon "Coiled-web material" described
in Section 10.3.4 for use in culturing amphipods (T.J.
Norberg-King, USEPA, personal communication). Each
water-only beaker receives 1.0 mL of YCT stock solution
and about two volume additions of water daily.
14.3.7.10 Reproduction of amphipods is measured on
Day 35 and Day 42 in the water-only beakers by removing
and counting the adults and young in each beaker. On
Day 35, the adults are then returned to the same water-
only beakers. Adult amphipods surviving on Day 42 are
preserved in sugar formalin. The number of adult females
is determined by simply counting the adult males (mature
male amphipods will have an enlarged second gnathopod)
and assuming all otheradults are females (cf., Figure 11.1).
The number of females is used to determine number of
young/female/beaker from Day 28 to Day 42. Growth can
also be measured forthese adult amphipods.
14.4 Interpretation of Results
14.4.1 Data Analysis
14.4.1.1 Endpoints measured in the 42-d H. azteca test
include survival (Day 28, 35, and 42), growth (as length or
dry weight on Day 28 and 42), and reproduction (number
of young/female produced from Day 28 to 42). Section 16
describes general information regarding statistical analy-
sis of these data, including both point estimates (i.e.,
LC50s) and hypothesis testing (i.e., ANOVA). The follow-
ing sections describe species-specific information that is
useful in helping to interpret the results of 42-d sediment
toxicity tests with H. azteca.
14.4.2 Age Sensitivity
14.4.2.1 The sensitivity of H. azteca appears to be
relatively similar up to at least 24- to 26-d-old organisms
(Collyard etal., 1994). For example, the toxicity of diazinon,
Cu, Cd, and Zn was similar in 96-h water-only exposures
starting with 0- to 2-d-old organisms through 24- to 26-
-d-old organisms (Figure 11.2). The toxicity of alkylphenol
ethoxylate (a surfactant) tended to increase with age. In
general, this suggests that tests started with 7-d to 8-d-old
amphipods would be representative of the sensitivity of
H. azteca up to at least the adult life stage.
14.4.3 Grain Size
14.4.3.1 Hyalella azteca tolerate a wide range in sedi-
ment grain size and organic matter in 10- to 28-d tests
measuring effects on survival or growth (Ankley et al.,
1994; Suedel and Rodgers, 1994; Ingersoll et al., 1996;
Kemble et al., 1999). Using the method outlined in Sec-
tion 14.2, no significant correlations were observed be-
tween the survival, growth, or reproduction of/-/, azteca
and the physical characteristics of the sediment (grain
size ranging from predominantly silt to predominantly
sand), TOC (ranging from 0.3 to 9.6%), water content
(ranging from 19 to 81 %; Ingersoll et al., 1998). Addition-
ally, no significant correlations were observed between
these biological endpoints and the water-quality charac-
teristics (i.e., hardness, alkalinity, ammonia) of pore wa-
ter or overlying water in the sediments evaluated by
Ingersoll et al. (1998). Weak trends were observed be-
tween reproduction of amphipods and percent clay, per-
cent silt, and percent sand. Additional study is needed to
better evaluate potential relationships between reproduc-
tion of/-/, azteca and these physical characteristics of the
sediment. The weak relationship between the sediment
grain size and reproduction may have been due to the fact
that samples with higher amounts of sand also had higher
concentrations of organic contaminants compared to other
samples evaluated in Ingersoll et al. (1998).
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14.4.3.2 Until additional studies have been conducted
which substantiate this lack of a correlation between
physical characteristics of sediment and the reproductive
endpoints measured in the long-term sediment test with
H. azteca, it would be desirable to test control or refer-
ence sediments which are representative of the physical
characteristics of field-collected sediments. Formulated
sediments could be used to bracket the ranges in physi-
cal characteristics expected in the field-collected sedi-
ments being evaluated (Section 7.2). Addition of YCT
should provide a minimum amount of food needed to
support adequate survival, growth, and reproduction of
H. azteca in sediments low in organic matter. Without
addition of food, H. azteca can starve during exposures
(McNulty et al., 1999) making it impossible to differentiate
effects of contaminants from other sediment
characteristics.
14.4.4 Influence of Indigenous Organisms
14.4.4.1 Survival of H. azteca in 28-d tests was not
reduced in the presence of oligochaetes in sediment
samples (Reynoldson et al., 1994). However, growth of
amphipods was reduced when high numbers of oligo-
chaetes were placed in a sample. Therefore, it is impor-
tant to determine the number and biomass of indigenous
organisms in field-collected sediments in order to better
interpret growth data (Reynoldson et al., 1994; DeFoe and
Ankley, 1998). Furthermore, presence of predators may
also influence response of test organisms in sediment
(Ingersoll and Nelson, 1990).
14.4.5 Relationships between Growth and
Reproductive Endpoints
14.4.5.1 Natural or anthropogenic stressors that affect
growth of invertebrates may also affect reproduction,
because of a minimum size needed for reproduction
(Rees and Crawley, 1989; Ernsting et al., 1993; Moore
and Dillon, 1993; Enserink et al., 1995; Moore and Farrar,
1996; Sibley et al., 1996, 1997a). Ingersoll et al. (1998)
reported a significant correlation between reproduction
from Day 28 to 42 and length of/-/, azteca on Day 28 when
data are plotted by the mean of each treatment
(Figure 14.1 a; Spearman rank correlation of 0.59,
p=0.0001). Based on 28-d lengths, smaller amphipods
(<3.5 mm) tended to have lower reproduction and larger
amphipods (>4.3 mm) tended to have higher reproduction;
however, the range in reproduction was wide for amphi-
pods 3.5 to 4.3 mm in length. Based on 42-d lengths,
there was a weaker correlation between length and repro-
duction (i.e., reproduction and length measured in paired
replicates; Figure 14.1b, Spearman rank correlation of
0.49, p=0.0001). Similarly, plotting data by individual
replicates (data not shown) did not improve the relation-
ship between 42-d length and reproduction compared to
the plots by the mean of each treatment (Figure 14.1b;
Ingersoll etal., 1998).
14.4.5.2 Weaker relationships were observed between
reproduction and dry weight measured on Day 28
(Figure 14.2a, Spearman rank correlation of 0.44,
p = 0.0037, n = 42) or dry weight measured on Day 42
(Figure 14.2b, Spearman rank correlation 0.34, p = 0.0262,
n = 42). Round-robin studies (Section 17.6) have gener-
ated additional data that will be used to further evaluate
relationships between growth and reproduction of/-/, azteca
in sediment tests using the procedures outlined in
Section 14.2.
14.4.5.3 A significant correlation was evident between
length and dry weight of amphipods (Figure 14.3, Spearman
rank of 0.80, p=0.0001) indicating that either length or
weight could be measured in sediment tests with
H. azteca. However, additional statistical options are
available if length is measured on individual amphipods,
such as nested ANOVA which can account for variance in
length within replicates (Steevens and Benson, 1998).
Analyses are ongoing to evaluate the ability of length vs.
weight to discriminate between contaminated and uncon-
taminated samples in a database described in Ingersoll et
al. (1996).
14.4.5.4 The relatively variable relationship between
growth and reproduction probably reflects the fact that
most of these comparisons were made within a fairly
narrow range in length (3.5 to 5.0 mm; Figure 14.1) or dry
weight (0.25 to 0.50 mg; Figure 14.2). Other investigators
have reported a similar degree of variability in reproduc-
tion of/-/, azteca within a narrow range of length or weight,
with stronger correlations observed over wider ranges
(Hargrave, 1970b; Strong, 1972; Wen, 1993; Moore and
Farrar, 1996). The degree of correlation between growth
and reproduction may also be dependent on the genetic
strain of/-/, azteca evaluated (Strong, 1972; France, 1992).
14.4.5.5 The proportion of males to females within a
treatment or by replicate was not correlated to young
production, but may have contributed to a variation in
reproduction (Ingersoll et al., 1998). Wen (1993) reported
that when two or three males were placed in a beakerwith
one female H. azteca, the frequency of successful am-
plexus was reduced, possibly from aggression between
the males. Future study is needed to determine if increas-
ing the number of amphipods/beaker would result in a
more consistent proportion of males to females within a
beaker and would reduce variability in reproduction.
14.4.5.6 Reproduction was often more variable than
growth (Ingersoll et al., 1998). The coefficient of variation
(CV) was typically <10% for growth and >20% for repro-
duction. This difference in variation affects the statistical
power of the comparisons and the number of replicates
required for a test. For example, detection of a 20%
difference between treatment means at a statistical power
of 0.8 would require about 4 replicates at a CV of 10% and
14 replicates at a CV of 20% (Figure 16.5). Fewer repli-
cates would be required if detection of larger differences
among treatment means were of interest. Ongoing water-
only studies testing select contaminants will hopefully
provide additional data on the relative sensitivity and
variability of sublethal endpoints in toxicity tests with
H. azteca (Ingersoll etal., 1998).
79
-------
14 -
12 -
c
o
M—
o
CD
"E
14 -
12 -
10 -
8 -
6 -
4 -
2 -
0
b
O
O
O O
2.5 3.0 3.5 4.0 4.5 5.0
Length (mm, Day 42, by treatment)
5.5
Figure 14.1 Relationships between Hya lei la azteca length and reproduction by (a) treatment means for 28-d length
or (b) treatment means for 42-d length.
80
-------
_CD
03
E
75)
o
-------
co
o
-I—<
CD
1.0
0.9 -
0.8 -
0.7 -
0.6
0.5 -
0.4 -
0.3 -
0.2 -
0.1 -
0.0
OG
so
o
o
o
o
o
o
2.5 3.0 3.5 4.0 4.5 5.0 5.5
Length (mm, by replicate)
6.0
Figure 14.3 Relationship between Hyalella azteca length and dry weight. Triangles are data for Day 28 and circles are data for
Day 42 (Ingersoll et al.. 1998).
14.4.5.7 The 8-replicate design recommended in this
manual (Table 14.1) is a compromise between logistical
constraints and statistical considerations. Laboratories
experienced with this method have shown CVs of 25 to
50% (Ingersoll et al., 1998), though some higher values
were observed during the round-robin testing (Section
17.6), in which most labs had not previously performed
the test.
14.4.5.8 As discussed above, the number of replicates
can be adjusted according to the needs of a particular
study. For example, Kubitz et al. (1996) recommended a
two-step process for assessing growth in sediment tests
with H. azteca. Using this process, a limited number of
replicates would be tested in a screening step. Samples
identified as possibly affecting reproduction could then be
tested in a confirmatory step with additional replicates.
This two-step analysis conserves laboratory resources
and increases statistical power when needed to discrimi-
nate sublethal effects. A similar approach could be ap-
plied to evaluate reproductive effects of contaminants in
sediment where a limited number of replicates could be
initially tested to evaluate potential effects. Samples
identified as possibly toxic based on reproduction could
then be reevaluated using an increased number of repli-
cates. However, the use of sediments stored for extended
periods of time may introduce variability in results be-
tween the two studies (Section 8.2).
14.4.6 Relative Endpoint Sensitivity
14.4.6.1 Measurement of sublethal endpoints in sedi-
ment tests with H. azteca can provide unique information
that has been used to discriminate toxic effects of expo-
sure to contaminants. Table 14.4 compares the relative
sensitivity of survival and growth endpoints in 14- and
28-d tests with H. azteca (Ingersoll et al., 1996, 1998).
When 14-d and 28-d tests were conducted concurrently
measuring both survival and growth, both tests identified
34% of the samples as toxic and 53% of the samples as
not toxic (N=32). Both tests identified an additional 6% of
the samples as toxic. Survival or growth endpoints identi-
fied a similar percentage of samples as toxic in both the
14- and 28-d tests. However, the majority of the samples
used to make these comparisons were highly contami-
nated. Additional exposures conducted with moderately
contaminated sediment might exhibit a higher percentage
of sublethal effects in the 28-d test compared to the
14-d test.
14.4.6.2 When both survival and growth were measured
in 14-d tests (N=25), only 4% of the samples reduced
82
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Table 14.4 Percentage of Paired Tests or Paired Endpoints Identifying Samples as Toxic in Hyalella azteca 14-d or 28-d Tests.
See USEPA (1996a) and Ingersoll et al. (1996) for a description of this database.
Comparisons
Tox/tox1
Not/not2
Tox/not3
Not/tox4
N5
Survival or growth: 14 d/28 d
Survival: 14 d/28 d
Growth: 14 d/28 d
14 d: survival/growth
28 d: survival/growth
34
25
8
4
16
53
66
64
60
52
6
0
12
20
14
6
10
16
16
18
32
32
25
25
44
1 Tox/tox: samples toxic (significant reduction relative to the control p<0.05) with both tests (or both endpoints).
2 Not/not: samples not toxic with both tests (or both endpoints).
3 Tox/not: samples toxic to the first but not the second test (or endpoint).
4 Not/tox: samples not toxic to the first but toxic to the second test (or endpoint).
5 N: number of samples
both survival and growth; however, 20% reduced survival
only and 16% reduced growth only (60% did not reduce
survival orgrowth). Hence, if survival was the only endpoint
measured in 14-d tests, 16% of the toxic samples would
be incorrectly classified. Similar percentages are also
observed for the 28-d tests. When both survival and
growth were measured in the 28-d test (N=44), 16% of the
samples reduced both survival and growth, 14% reduced
survival only, 18% reduced growth only, and 52% did not
reduce survival orgrowth.
14.4.6.3 The endpoint comparisons in Table 14.4 repre-
sent only samples where both survival and growth could
be measured. If a sample was extremely toxic, it would
not be included in this comparison since growth could not
be measured. Moderately contaminated sediments that
did not severely reduce survival could have a reduced
growth. For example, in 28-d tests with sediments from
the Clark Fork River, growth was a more sensitive end-
point compared to survival or maturation. Only 13% of the
samples reduced survival and 20% of the samples re-
duced maturation; however, growth was reduced in 53%
of the samples (Kemble et al., 1994).
14.4.6.4 Other investigators have reported measurement
of growth in tests with H. azteca often provides unique
information that can help discriminate toxic effects of
exposure to contaminants in sediment (Kubitz et al.,
Milan! etal., 1996; Steevensand Benson, 1998) or water
(Brasher and Ogle, 1993; Borgmann, 1994). Similarly, in
sediment tests with the midge C. tentans, sublethal end-
points are often more sensitive than survival as indicators
of contaminant stress (Section 12 and 15). In contrast,
Borgmann et al. (1989) reported that growth or reproduc-
tion did not add additional information beyond measure-
ment of survival of/-/, azteca in water-only exposures with
cadmium or pentachlorophenol. Similarly, Dayetal. (1995)
reported that weight did not add additional information
beyond measurement of survival in 28-d tests with
H. azteca. Ramirez-Romero (1997) reported that repro-
duction of H. azteca was not affected by exposure to
sublethal concentrations of fluoranthene in sediment when
exposures were started with juvenile amphipods. Brasher
and Ogle (1993) started exposures with adult amphipods
and observed the sensitivity of reproduction compared to
survival of H. azteca was dependent on the chemical
tested (reproduction more sensitive to selenite and sur-
vival more sensitive to selenate in water-only exposures).
Long-term exposures starting with juvenile amphipods
would likely be more appropriate to assess effects of
contaminants on reproduction (i.e., Carr and Chapman,
1992; Nebekeretal., 1992).
14.4.7 Future Research
14.4.7.1 Additional studies are needed to further evaluate
the use of reconstituted water and ammonia on long-term
exposures with H. azteca. Section 1.3.8.5 addresses
interpretative guidance for evaluating toxicity associated
with ammonia in sediment. Ongoing water-only toxicity
tests with select chemicals (i.e., cadmium, ODD and
fluoranthene) should generate data that can be used to
better determine the relative sensitivity of survival, repro-
duction, and growth endpoints in tests with H. azteca
(Ingersoll et al., 1998). These water-only studies will also
be used to evaluate potential recovery of amphipods after
transfer into clean water to measure reproduction. In
addition to studies evaluating the relative sensitivity of
endpoints, research is also needed to evaluate the ability
of these laboratory endpoints to estimate responses of
benthic organisms exposed in the field to chemicals in
sediments (Canfield et al., 1996).
83
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Section 15
Test Method 100.5
Life-cycle Test for Measuring the Effects of Sediment-associated
Contaminants on Chironomus tentans
15.1 Introduction
15.1.1 The midge Chironomus tentans has been used
extensively in the short-term assessment of chemicals in
sediments (Wentsel et al., 1977; Nebeker et al., 1984;
Giesy et al., 1988; West et al., 1994), and standard
methods have been developed for testing with this midge
using 10-d exposures (Ingersoll et al., 1995; USEPA,
1994a; ASTM, 1999a). Chironomus tentans is a good
candidate for long-term toxicity testing because it nor-
mally completes its life cycle in a relatively short period of
time (25 to 30 d at 23°C), and a variety of developmental
(growth, survivorship) and reproductive (fecundity) end-
points can be monitored. In addition, emergent adults can
be readily collected so it is possible to transfer organisms
from the sediment test system to clean, overlying water
for direct quantification of reproductive success.
15.1.2 The long-term sediment toxicity test with the
midge, Chironomus tentans, is a life-cycle test in which
the effects of sediment exposure on survival, growth,
emergence, and reproduction are assessed (Benoit et al.,
1997). Procedures for conducting the long-term test
with C. tentans are described in Section 15.2. The test is
started with newly hatched larvae (<24-h old) and contin-
ues through emergence, reproduction, and hatching of the
F1 generation. Survival is determined at 20 d and at the
end of the test (about 50 to 65 d). Growth is determined at
20 d, which corresponds to the 10-d endpoint in the 10-d
C. tentans growth test started with 10-d-old larvae (Sec-
tion 12). From Day 23 to the end of the test, emergence
and reproduction are monitored daily. The number of
eggs is determined for each egg case, which is incubated
for 6 d to determine hatching success. Each treatment of
the life-cycle test is ended separately when no additional
emergence has been recorded for 7 consecutive days
(the 7-d criterion). When no emergence is recorded from a
treatment, ending of that treatment should be based on
the control sediment using this 7-d criterion. Appendix C
and Table 6.1 outline equipment and supplies needed to
conduct this test. The procedures described in Table
15.1 include measurement of a variety of lethal and
sublethal endpoints; minor modifications of the basic
methods can be used in cases where only a subset of
these endpoints is of interest.
15.1.3 The method outlined in Section 15.2 has been
evaluated in round-robin testing with 10 laboratories using
two clean sediments (Section 17.6). In the preliminary
round-robin with 1.5 ml of Tetrafin/d as a food source,
90% of labs met the survival criterion (>70%), 100% of
labs met the growth criterion (>0.48 mg AFDW), 70% of
labs met the emergence criterion (>50%), 90% of labs
met the reproduction criterion (>800 eggs/female), and
88% of labs met the percent hatch criterion (>80%).
Reproduction was generally more variable than growth or
survival within and among laboratories; hence, more repli-
cates might be needed to establish statistical signifi-
cance of small decreases in reproduction.
15.1.4 Growth and othersublethal endpoints in sediment
tests with C. tentans often provide unique information that
can be used to discriminate toxic effects of exposure to
contaminants. See Section 15.4.6 for additional details.
15.1.5 Results of tests using procedures different from
the procedures described in Section 15.2 may not be
comparable and these different procedures may alter
contaminant bioavailability. Comparison of results ob-
tained using modified versions of these procedures might
provide useful information concerning new concepts and
procedures for conducting sediment tests with aquatic
organisms. If tests are conducted with procedures differ-
ent from the procedures described in this manual, addi-
tional tests are required to determine comparability of
results (Section 1.3).
15.2 Procedure for Conducting a Life-
cycle Test for Measuring the Effects
of Sediment-associated
Contaminants on Chironomus
tentans
15.2.1 Conditions for conducting a long-term sediment
toxicity test with C. tentans are summarized in Table 15.1.
A general activity schedule is outlined in Table 15.2.
Decisions concerning the various aspects of experimental
design, such as the number of treatments, number of test
chambers/treatment, and water-quality characteristics
should be based on the purpose of the test and the
methods of data analysis (Section 16). When variability
84
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Table 15.1 Test Conditions for
Parameter
Conducting a Long-term Sediment Toxicity Test with Chironoinus tentans
Conditions
1. Test type:
2. Temperature:
3. Light quality:
4. Illuminance:
5. Photoperiod:
6. Test chamber:
7. Sediment volume:
8. Overlying water volume:
9. Renewal of overlying water:
10. Age of organisms:
11. Number of organisms/chamber:
12. Number of replicate chambers/treatment:
13. Feeding:
14. Aeration:
15. Overlying water:
16. Test chamber cleaning:
17. Overlying water quality:
18. Test duration:
19. Endpoints:
20. Test acceptability:
Whole-sediment toxicity test with renewal of overlying water
23±1°C
Wide-spectrum fluorescent lights
About 100 to 1000 lux
16L8D
300-mL high-form lipless beaker
100mL
175 ml
2 volume additions/d (Appendix A); continuous or intermittent (e.g., one volume
addition every 12 h)
< 24-h-old larvae
12
16 (12 at Day -1 and 4 for auxiliary males on Day 10)
Tetrafin® goldfish food, fed 1.5 ml daily to each test chamber starting Day -1
(1.0 ml contains 4.0 mg of dry solids)
None, unless dissolved oxygen in overlying water drops below 2.5 mg/L
Culture water, well water, surface water, site water, or reconstituted water
If screens become clogged during a test, gently brush the outside of the screen
(Appendix A).
Hardness, alkalinity, conductivity, and ammonia at the beginning, on Day 20, and
at the end of a test. Temperature daily (ideally continuously). Dissolved oxygen
(DO) and pH three times/week. Conductivity weekly. Concentrations of DO should
be measured more often if DO has declined by more than 1 mg/L since previous
measurement.
About 50 to 65 d; each treatment is ended separately when no additional emergence
has been recorded for seven consecutive days. When no emergence is recorded
from a treatment, termination of that treatment should be based on the control
sediment using this 7-d criterion.
20-d survival and weight; female and male emergence, adult mortality, the number
of egg cases oviposited, the number of eggs produced, and the number of hatched
eggs. Potential sublethal endpoints are listed in Table 15.4.
Average size of C. tentans in the control sediment at 20 d must be at least 0.6 mg/
surviving organism as dry weight or 0.48 mg/surviving organism as AFDW.
Emergence should be greater than or equal to 50%. Experience has shown that
pupae survival is typically >83% and adult survival is >96%. Time to death after
emergence is <6.5 d for males and <5.1 d for females. The mean number of eggs/
egg case should be greater than or equal to 800 and the percent hatch should be
greater than or equal to 80%. See Sections 15.1.3 and 17.6 for a summary of
performance in round-robin testing.
remains constant, the sensitivity of a test increases as
the number of replicates increases.
15.2.2 The long-term sediment toxicity test with C. ten-
tans is conducted at 23°C with a 16L8D photoperiod at an
illuminance of about 100 to 1000 lux (Table 15.1). Test
chambers are 300-mL high-form lipless beakers contain-
ing 100 ml of sediment and 175 ml of overlying water.
Each test chamber receives 2 volume additions/d of
overlying water. Water renewals may be manual or auto-
mated. Appendix A describes water-renewal systems that
can be used to deliver overlying water. Overlying water
should be a source of water that has been demonstrated
to support survival, growth, and reproduction of C. tentans
in culture. For site-specific evaluations, the characteris-
tics of the overlying water should be as similar as pos-
sible to the site where sediment is collected. Require-
ments for test acceptability are summarized in Table
15.3.
15.2.3 The number of replicates and concentrations
tested depends in part on the significance level selected
and the type of statistical analysis. For routine testing, a
total of 16 replicates, each containing 12, <24-h-old larvae
are tested for each treatment. For the total of 16 repli-
cates the assignment of beakers is as follows: initially,
12 replicates are set up on Day -1 of which 4 replicates
are used for 20-d growth and survival endpoints and 8
85
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Table 15.2 General Activity Schedule for Conducting a Long-term Sediment Toxicity Test with Chironomus tentans
Day Activity
Pre-Test
-4 Start reproduction flask with cultured adults (1:3 male:female ratio). For example for 15 to 25 egg cases, 10 males and 30 females
are typically collected. Egg cases typically range from 600 to 1500 eggs/case.
-3 Collect egg cases (a minimum of 6 to 8) and incubate at 23°C.
-2 Check egg cases for viability and development.
-1 1. Check egg cases for hatch and development.
2. Add 100 ml of homogenized test sediment to each replicate beaker and place in corresponding treatment holding tank. After
sediment has settled for at least 1 h, add 1.5 ml Tetrafin slurry (4g/L solution) to each beaker. Overlying water renewal begins
at this time.
Sediment Test
0 1. Transfer all egg cases to a crystallizing dish containing control water. Discard larvae that have already left the egg cases
in the incubation dishes. Add 1.5 ml food to each test beaker with sediment before the larvae are added. Add 12 larvae to each
replicate beaker (beakers are chosen by random block assignment). Let beakers sit (outside the test system) for 1 h following
addition of the larvae. After this period, gently immerse all beakers into their respective treatment holding tanks.
2. Measure temperature, pH, hardness, alkalinity, dissolved oxygen, conductivity and ammonia at start of test.
1-End On a daily basis, add 1.5 ml food to each beaker. Measure temperature daily. Measure the pH and dissolved oxygen three
times a week during the test. Measure conductivity weekly. If the DO has declined more than 1 mg/L since previous reading,
increase frequency of DO measurements and aerate if DO continues to be less than 2.5 mg/L. Measure hardness, alkalinity,
conductivity, ammonia, temperature, pH, and dissolved oxygen at the end of the test.
6 For auxiliary male production, start reproduction flask with culture adults (e.g., 10 males and 30 females; 1:3 male to female ratio).
7-10 Follow set-up schedule for auxiliary male beakers (4 replicates/treatment) described above for Day -3 to Day 0.
19 In preparation for weight determinations, ash weigh pans at 550°C for 2 h. Note that the weigh pans should be ashed before use
to eliminate weighing errors due to the pan oxidizing during ashing of samples.
20 1. Randomly select four replicates from each treatment and sieve the sediment to recover larvae for growth and survival
determinations. Pool all living larvae per replicate and dry the sample to a constant weight (e.g., 60°C for 24 h).
2. Install emergence traps on each of the remaining reproductive replicate beakers.
3. Measure temperature, pH, hardness, alkalinity, dissolved oxygen, conductivity and ammonia.
21 The sample with dried larvae is brought to room temperature in a dessicator and weighed to the nearest 0.01 mg . The dried
larvae in the pan are then ashed at 550°C for 2 h. The pan with the ashed larvae is then reweighed and the tissue mass of the
larvae determined as the difference between the weight of the dried larvae plus pan and the weight of the ashed larvae plus pan.
Chronic Measurements
23-End On a daily basis, record emergence of males and females, pupal, and adult mortality, and time to death for previously collected
adults. Each day, transfer adults from each replicate to a corresponding reproduction/oviposition (RIO) chamber. Transfer each
primary egg case from the R/O chamber to a corresponding petri dish to monitor incubation and hatch. Record each egg case
oviposited, number of eggs produced (using either the ring or direct count methods), and number of hatched eggs. If it is difficult
to estimate the number of eggs in an egg case, use a direct count to determine the number of eggs; however the hatchability data
will not be obtained for this egg case.
28 Place emergence traps on auxiliary male replicate beakers.
33-End Transfer males emerging from the auxiliary male replicates to individual inverted petri dishes. The auxiliary males are used for
mating with females from corresponding treatments from which most of the males had already emerged or in which no males
emerged.
40-End After 7 d of no recorded emergence in a given treatment, end the treatment by sieving the sediment to recover larvae, pupae,
or pupal exuviae. When no emergence occurs in a test treatment, that treatment can be ended once emergence in the control
sediment has ended using the 7-d criterion.
replicates for determination of emergence and reproduc- are stocked with 12, <24-h-old larvae 10 d following
tion. It is typical for males to begin emerging 4 to 7 d initiation of the test. Midges in each test chamber are fed
before females. Therefore, additional males, referred to 1.5 ml of a 4-g/L Tetrafin® suspension daily. Endpoints
as auxiliary males, need to be available during the prime monitored include 20-d survival and weight, emergence,
female emergence period for each respective chamber/ time to death (adults), reproduction, and egg hatchability.
sediment. To provide these males, 4 additional replicates
86
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Table 15.3 Test Acceptability Requirements for a Long-term Sediment Toxicity Test with Chironomus tentans
A. It is recommended for conducting a long-term test with C. tentans that the following performance criteria be met:
1. Tests must be started with less than 1-d- (<24-h) old larvae. Starting a test with substantially older organisms may compromise
the emergence and reproductive endpoint.
2. Average survival of C. tentans in the control sediment should be greater than or equal to 70% on Day 20 and greater than 65% at
the end of the test.
3. Average size of C. tentans in the control sediment at 20 d must be at least 0.6 mg/surviving organism as dry weight or 0.48 mg/
surviving organism as AFDW. Emergence should be greater than or equal to 50%. Experience has shown that pupae survival is
typically >83% and adult survival is >96%. Time to death after emergence is <6.5 d for males and <5.1 d for females. The mean
number of eggs/egg case should be greater than or equal to 800 and the percent hatch should be greater than or equal to 80%.
See Sections 15.1.3 and 17.6 for a summary of performance in round-robin testing.
4. Hardness, alkalinity, and ammonia in the overlying water typically should not vary by more than 50% during the test, and dissolved
oxygen should be maintained above 2.5 mg/L in the overlying water.
B. Performance-based criteria for culturing C. tentans include the following:
1. It may be desirable for laboratories to periodically perform 96-h water-only reference-toxicity tests to assess the sensitivity of
culture organisms (Section 9.16.2). Data from these reference-toxicity tests could be used to assess genetic strain or life-stage
sensitivity of test organisms to select chemicals.
2. Laboratories should keep a record of time to first emergence for each culture and record this information using control charts.
Records should also be kept on the frequency of restarting cultures.
3. Laboratories should record the following water-quality characteristics of the cultures at least quarterly: pH, hardness, alkalinity,
and ammonia. Dissolved oxygen in the cultures should be measured weekly. Temperature of the cultures should be recorded
daily. If static cultures are used, it may be desirable to measure water quality more frequently.
4. Laboratories should characterize and monitor background contamination and nutrient quality of food if problems are observed in
culturing or testing organisms.
5. Physiological measurements such as lipid content might provide useful information regarding the health of the cultures.
C. Additional requirements:
1. All organisms in a test must be from the same source.
2. Storage of sediments collected from the field should follow guidance outlined in Section 8.2.
3. All test chambers (and compartments) should be identical and should contain the same amount of sediment and overlying water.
4. Negative-control sediment and appropriate solvent controls must be included in a test. The concentration of solvent used must not
adversely affect test organisms.
5. Test organisms must be cultured and tested at 23°C (±1°C).
6. The daily mean test temperature must be within ±1°C of 23°C. The instantaneous temperature must always be within ±3°C of 23°C.
7. Natural physico-chemical characteristics of test sediment collected from the field should be within the tolerance limits of the test
organisms.
15.3 General Procedures 15.3.2 Hatching of Eggs
15.3.1 Collection of Egg Cases 15.3.2.1 Hatching of eggs should be complete by about
72 h. Hatched larvae remain with the egg case for about
15.3.1.1 Egg cases are obtained from adult midges held 24 h and appear to use the gelatinous component of the
in a sex ratio of 1:3 male:female. Ten males and egg case as an initial source of food (Sadler, 1935; Ball
30 females will produce between 15 to 25 egg cases, and Baker, 1995). After the first 24-h period with larvae
Adults should be collected fourdays before starting a test hatched, transferthe egg cases from the incubation petri
(Appendix C, Figure C.3). The day after collection of dish to another dish with clean test water. Larvae having
adults, 6 to 8 of the larger "C" shaped egg cases are already left the egg case in the incubation petri dish are
transferred to a petri dish with culture water and incubated discarded since their precise age and time away from the
at 23°C (Appendix C, Figure C.2). Hatching typically gelatinous food source is unknown. The action of trans-
begins around 48 h and larvae typically leave the egg ferring the egg case stimulates the remaining larvae to
case 24 h after the first hatch. The number of eggs in leave the egg case within a few hours. These are the
each egg case will vary, but typically ranges from 600 to larvae that are used to start the test.
1500 eggs. It should be noted that mating may have
occurred in culture tanks before males and females are
placed into flasks for collecting eggs.
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Table 15.4 Endpoints for a Long-term Sediment Toxicity Test with Chironomus tentans
Lethal
Survival
Larvae (20 d)
Larvae (End)
Pupae
Adults
Sublethal
Growth Emergence
Larvae Total/Percent
Cumulative (Rate)
Time to First
Time to Death
Reproduction
Sex Ratio
Time to Oviposition
Mean Eggs/Female
Egg Cases/Treatment
Egg Hatchability
15.3.3 Sediment into Test Chambers
15.3.3.1 The day before the sediment test is started
(Day -1) each sediment should be thoroughly homog-
enized and added to the test chambers (Section 8.3.1).
Sediment should be visually inspected to judge the extent
of homogeneity. Excess water on the surface of the
sediment can indicate separation of solid and liquid com-
ponents. If a quantitative measure of homogeneity is
required, replicate subsamples should be taken from the
sediment batch and analyzed for TOC, chemical con-
centrations, and particle size.
15.3.3.2 Each test chamber should contain the same
amount of sediment, determined either by volume or by
weight. Overlying water is added to the chambers in a
mannerthat minimizes suspension of sediment. This can
be accomplished by gently pouring water along the sides
of the chambers or by pouring water onto a baffle (e.g., a
circular piece of Teflon with a handle attached) placed
above the sediment to dissipate the force of the water.
Renewal of overlying water is started on Day -1. A test
begins when the organisms are added to the test cham-
bers (Day 0).
15.3.4 Renewal of Overlying Water
15.3.4.1 Renewal of overlying water is required during a
test. Two volume additions of overlying water (continuous
or intermittent) should be delivered to each test chamber
daily. At any particular time during the test, flow rates
through any two test chambers should not differ by more
than 10%. Hardness, alkalinity and ammonia concentra-
tions in the water above the sediment, within a treatment,
typically should not vary by more than 50% during the
test. Mount and Brungs (1967) diluters have been modi-
fied for sediment testing, and other automated water-
delivery systems have also been used (Maki, 1977;
Ingersoll and Nelson, 1990; Benoit et al., 1993; Zumwalt
et al., 1994; Brunson et al., 1998; Wall et al., 1998;
Leppanen and Maier, 1998). Each water-delivery system
should be calibrated before a test is started to verify that
the system is functioning properly. Renewal of overlying
water is started on Day -1 before the addition of test
organisms on Day 0. Appendix A describes water-renewal
systems that can be used for conducting sediment tests.
15.3.4.2 In water-renewal tests with one to four volume
additions of overlying water/d, water-quality characteris-
tics generally remain similarto the inflowing water (Ingersoll
and Nelson, 1990; Ankley et al., 1993); however, in static
tests, water quality may change profoundly during the
exposure (Shuba et al., 1978). For example, in static
whole-sediment tests, the alkalinity, hardness, and
conductivity of overlying water more than doubled in
several treatments during a four-week exposure (Ingersoll
and Nelson, 1990). Additionally, concentrations of meta-
bolic products (e.g., ammonia) may also increase during
static exposures, and these compounds can either be
directly toxic to the test organisms or may contribute to
the toxicity of the contaminants in the sediment. Further-
more, changes in water-quality characteristics such as
hardness may influence the toxicity of many inorganic
(Gauss et al., 1985) and organic (Mayer and Ellersieck,
1986) contaminants. Although contaminant concentra-
tions are reduced in the overlying water in water-renewal
tests, organisms in direct contact with sediment generally
receive a substantial proportion of a contaminant dose
directly from either the whole sediment or from the inter-
stitial water.
15.3.5 Acclimation
15.3.5.1 Test organisms must be cultured and tested at
23°C. Ideally, test organisms should be cultured in the
same water that will be used in testing. However, acclima-
tion of test organisms to the test water is not required.
15.3.5.2 Culturing of organisms and toxicity assessment
are typically conducted at 23°C. However, occasionally
there is a need to perform evaluations at temperatures
different than that recommended. Under these circum-
stances, it may be necessary to acclimate organisms to
the desired test temperature to prevent thermal shock
when moving immediately from the culture temperature to
the test temperature (ASTM, 1999a). Acclimation can be
achieved by exposing organisms to a gradual decline in
temperature; however, the rate of decline should be rela-
tively slow to prevent thermal shock. A decline in tem-
perature of 1 °C every 1 to 2 h has been used successfully
in some studies (P.K. Sibley, University of Guelph, Guelph,
Ontario, personal communication; APHA, 1989). Testing
at temperatures other than 23°C needs to be preceded by
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studies to determine expected performance under alter-
nate conditions.
15.3.6 Placing Organisms in Test Chambers
15.3.6.1 Test organisms should be handled as little as
possible. To start the test, larvae are collected with a
Pasteur pipet from the bottom of the incubation dish with
the aid of a dissecting microscope. Test organisms are
pipetted directly into overlying water and care should be
exercised to release them underthe surface of the water.
Transferring the larvae to exposure chambers within 4 h of
emerging from the egg case reportedly improves survival
(Benoit et al., 1997). Laboratory personnel should prac-
tice transferring first-instar midge larvae before tests with
sediment are conducted.
15.3.7 Feeding
15.3.7.1 Each beaker receives a daily addition of 1.5 ml
of Tetrafin® (4 mg/mL dry solids). Without addition of
food, the test organisms may starve during exposures.
However, the addition of the food may alterthe availability
of the contaminants in the sediment (Wiederholm et al.,
1987; Harkeyetal., 1994). Furthermore, if too much food
is added to the test chamber, or if the mortality of test
organisms is high, fungal or bacterial growth may develop
on the sediment surface. Therefore, the amount of food
added to the test chambers is kept to a minimum.
15.3.7.1 Suspensions of food should be thoroughly mixed
before aliquots are taken. If excess food collects on the
sediment, a fungal or bacterial growth may develop on the
sediment surface, in which case feeding should be sus-
pended for one or more days. A drop in dissolved oxygen
below 2.5 mg/L during a test may indicate that the food
added is not being consumed. Feeding should be sus-
pended for the amount of time necessary to increase the
dissolved oxygen concentration (ASTM, 1999a). If feed-
ing is suspended in one treatment, it should be sus-
pended in all treatments. Detailed records of feeding rates
and the appearance of the sediment surface should be
made daily.
15.3.8 Monitoring a Test
15.3.8.1 All chambers should be checked daily and
observations made to assess test organism behavior
such as sediment avoidance. However, monitoring ef-
fects on burrowing activity of test organisms may be
difficult because the test organisms are often not visible
during the exposure. The operation of the exposure sys-
tem should be monitored daily.
15.3.8.2 Measurement of Overlying Water-quality
Characteristics
15.3.8.2.1 Conductivity, hardness, alkalinity, and ammo-
nia should be measured in all treatments at the beginning
of the test, on Day 20, and at the end of the test.
Dissolved oxygen (DO) and pH measurements should be
taken at the beginning of a test and at least three times a
week until the end of the test. Conductivity should be
measured weekly. Overlying water should be sampled
just before water renewal from about 1 to 2 cm above the
sediment surface using a pipet. It may be necessary to
composite water samples from individual replicates. The
pipet should be checked to make sure no organisms are
removed during sampling of overlying water. Water quality
should be measured on each batch of water prepared for
the test.
15.3.8.2.2 Routine chemistries on Day 0 should be taken
before organisms are placed in the test beakers. Dis-
solved oxygen and pH can be measured directly in the
overlying water with a probe. However, for DO it is
important to allow the probe time to equilibrate in the
overlying water in an effort to accurately measure concen-
trations of DO. If a probe is used for measurements in
overlying water, it should be inspected between samples
to make sure that organisms are not attached and should
be rinsed between samples to minimize cross contamina-
tion.
15.3.8.2.3 Water-only exposures evaluating the tolerance
of C. tentans larva to depressed DO have indicated that
significant reductions in weight occurred after 10-d expo-
sure to 1.1 mg/L DO, but not at 1.5 mg/L (V. Mattson,
USEPA, Duluth, MN, personal communication). This
finding concurs with the observations during method de-
velopment at the USEPA laboratory in Duluth that excur-
sions of DO as low as 1.5 mg/L did not seem to have an
effect on midge survival and development (P.K. Sibley,
University of Guelph, Guelph, Ontario, personal commu-
nication). Based on these findings, periodic depressions
of DO below 2.5 mg/L (but not below 1.5 mg/L) are not
likely to adversely affect test results, and thus should not
be a reason to discard test data. Nonetheless, tests
should be managed toward a goal of DO >2.5 mg/L to
insure satisfactory performance. If the DO level of the
waterfalls below 2.5 mg/L for any one treatment, aeration
is encouraged and should be done in all replicates for the
duration of the test (i.e., about 1 bubble/second in the
overlying water). Occasional brushing of screens on
outside of beakers will help maintain the exchange of
water during renewals.
15.3.8.2.4 Temperature should be measured at least
daily in at least one test chamber from each treatment.
The temperature of the water bath orthe exposure cham-
ber should be continuously monitored. The daily mean
test temperature must be within ±1 °C of 23°C. The instan-
taneous temperature must always be within ±3°C of 23°C.
15.3.8.3 Monitoring Survival and Growth
15.3.8.3.1 At 20 d, 4 of the initial 12 replicates are
selected for use in growth and survival measurements.
Using a #40 sieve (425-um mesh) to remove larvae from
sediment, collect the C. tenfansand record data on record
sheet (Appendix D). Any immobile organisms isolated
from the sediment surface or from sieved material should
be considered dead. Often C. tentans larvae tend to lose
their coloration within 15 to 20 min of death and may
become rigidly elongate. Surviving larvae are kept sepa-
rated by replicate for weight measurements; if pupae are
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recovered (<1% occurrence at recommended testing
conditions), these organisms are included in survival data
but not included in the growth data. A consistent amount
of time should be taken to examine sieved material for
recovery of test organisms (e.g., 5 min/replicate).
15.3.8.3.2 The 10-d method for C. tentans in the first
edition of this manual (USEPA, 1994a), as well as most
previous research, has used dry weight as a measure of
growth. However, Sibley et al. (1997b) found that the
grain size of sediments influences the amount of sedi-
ment that C. tentans larvae ingest and retain in their gut.
As a result, in finer-grain sediments, a substantial portion
of the measured dry weight may be comprised of sedi-
ment rather than tissue. While this may not represent a
strong bias in tests with identical grain size distributions
in all treatments, most field assessments are likely to
have varying grain size among sites. This will likely
create differences in dry weight among treatments that
are not reflective of true somatic growth. Forthis reason,
weight of midges should be measured as ash-free dry
weight (AFDW) instead of dry weight. AFDWwill more
directly reflect actual differences in tissue weight by
reducing the influence of sediment in the gut. If test
organisms are to be used for an evaluation of bioaccumu-
lation, it is not advisable to dry the sample before con-
ducting the residue analysis. If conversion from wet weight
to dry weight is necessary, aliquots of organisms can be
weighed to establish wet to dry weight conversion factors.
A consistent procedure should be used to remove the
excess water from the organisms before measuring wet
weight.
15.3.8.3.3 The AFDW of midges should be determined
for the growth endpoint. All living larvae per replicate are
combined and dried to a constant weight (e.g., 60°C for
24 h). Note that the weigh boats should be ashed before
use to eliminate weighing errors due to the pan oxidizing
during ashing. The sample is brought to room tempera-
ture in a desiccator and weighed to the nearest 0.01 mg to
obtain mean weights per surviving organism per replicate.
The dried larvae in the pan are then ashed at 550°C for
2 h. The pan with the ashed larvae is then reweighed and
the tissue mass of the larvae is determined as the differ-
ence between the weight of the dried larvae plus pan and
the weight of the ashed larvae plus pan. For rare in-
stances in which preservation is required, an 8% sugar
formalin solution can be used to preserve samples
(USEPA, 1994a), but the effects of preservation on the
weight and lengths of the midges have not been suffi-
ciently studied. The sugar formalin solution is prepared
by adding 120 g of sucrose to 80 ml of formalin which is
then brought to a volume of 1 L using deionized water.
This stock solution is mixed with an equal volume of
deionized water when used to preserve organisms.
NoTox® (Earth Safe Industries, Belle Mead, NJ) can be
used as a substitute for formalin (Ungeret al., 1993).
15.3.8.4 Monitoring Emergence
15.3.8.4.1 Emergence traps are placed on the reproduc-
tive replicates on Day 20 (emergence traps for the auxil-
iary beakers are added at the corresponding 20-d time
interval for those replicates; Appendix C, Figures C.1 and
C.4). At 23 °C, emergence in control sediments typically
begins on or about Day 23 and continues for about
2 weeks. However, in contaminated sediments, the
emergence period may be extended by several weeks.
15.3.8.4.2 Two categories are recorded for emergence:
complete emergence and partial emergence. Complete
emergence occurs when an organism has shed the pupal
exuviae completely and escapes the surface tension of
the water. If complete emergence has occurred but the
adult has not escaped the surface tension of the water,
the adult will die within 24 h. Therefore, 24 h should
elapse before this death is recorded. Partial emergence
occurs when an adult has only partially shed the pupal
exuviae. These adults will also die, an event which can
be recorded after 24 h. Pupae at the sediment surface or
the air-water interface may emerge successfully during
the 24-h period. However, cannibalism of sediment bound
pupae by larvae may also occur. Data are recorded on
data sheets provided as shown in example data sheet
(Appendix D).
15.3.8.4.3 Between Day 23 and the end of the test,
emergence of males and females, pupal and adult mortal-
ity, and time to death for adults is recorded daily for the
reproductive replicates. On Day 30 (20-d-old organisms),
emergence traps are placed on the auxiliary beakers to
collect the additional males for use with females emerging
from the reproduction replicates (Table 15.2; Appendix C,
Figures C.1 and C.4). Data are recorded on data sheets
provided as shown in the example data sheet (Appendix
D).
15.3.8.5 Collecting Adults for Reproduction
15.3.8.5.1 Adults are collected daily from individual traps
using the aspirator and collector dish (Appendix C,
Figure C.2). With the collector dish nearby, the emer-
gence trap is quickly moved from the beaker onto the
dish. With the syringe plunger fully drawn, the glass
collector tube is inserted through the screened access
hole of the collector dish and the adults gently aspirated
into the syringe barrel. Aspirated adults can easily be
seen through the translucent plastic of the syringe. The
detachable portion of the aspirator unit is then replaced
with a reproduction/oviposit (R/O) chamber. This ex-
change can be facilitated by placing the thumb of the
hand holding the syringe overthe barrel entry port until the
R/O chamber is in place. With the R/O chamber in place,
and the plunger on a solid surface, the barrel of the
syringe is pushed gently downward which forces the
adults to move up into the R/O unit. Adults remaining on
the transfer apparatus may be prodded into the R/O
chamber by gently tapping the syringe. The transfer
process is completed by quickly moving the R/O chamber
to a petri dish containing clean water. At all times during
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the transfer process, it is important to ensure that the
adults are stationary to minimize the possibility of es-
cape.
15.3.8.5.2 At about Day 33 to the end of the test, the
auxiliary males may be needed to support reproduction in
females. Males that emerge from the auxiliary male
replicates are transferred to individual inverted petri dishes
(60 x 15 mm dishes without water and with air holes drilled
in top of the dish; see Appendix C for a listing of equip-
ment.) Each male may be used for mating with females
from corresponding treatments for up to 5 d. Males may
be used for breeding with more than one new emergent
female. Males from a different replicate within the same
sediment treatment may be paired with females of repli-
cates where no males have emerged. Data can be re-
corded on data sheets provided in Appendix D.
15.3.8.6 Monitoring Reproduction
15.3.8.6.1 Each R/O unit is checked daily for dead adults
and egg cases. Dead organisms are removed. In situa-
tions where many adults are contained within an R/O
chamber, it may be necessary to assume that a dead
adult is the oldest male or female in that replicate for the
purpose of recording time to death. To remove dead
adults and egg cases from the R/O chamber, one side of
the chamber is carefully lifted just enough to permit the
insertion of a transfer pipet or tweezers.
15.3.8.6.2 For each emerged female, at least one male,
obtained from the corresponding reproductive replicate,
from another replicate of that treatment, or from the
auxiliary male beakers, is transferred into the R/O unit
using an aspirator. Females generally remain sexually
receptive up to 3 d if they have not already mated. Benoit
et al. (1997) have shown that over 90% of females will
oviposit within 1 d of fertilization; however, a few will
require as long as 72 h to oviposit. A female will lay a
single primary egg case, usually in the early morning
(Sadler, 1935). A second, generally smaller egg case
may be laid; however these second egg cases are prone
to fungus and the viability of embryos is typically poor.
These second egg cases do not need to be counted, or
recorded, and the numbers of eggs are not included in the
egg counts because eggs in second egg cases typically
have lower viability.
15.3.8.7 Counting Eggs, Egg Case Incubation, and
Hatch Determination
15.3.8.7.1 Primary egg cases from the R/O chamber are
transferred to a separate and corresponding petri dish
(60 x 15 mm with about 15 ml of water) to monitor
incubation and hatch. The number of eggs should be
estimated in each egg case by using a "ring method" as
follows: (1) for each egg case, the mean number of eggs
in five rings is determined; (2) these rings should be
selected at about equal distances along the length of the
egg case; (3) the number of eggs/ring multiplied by the
number of number of rings in the egg case will provide an
estimate of the total number of eggs. This can be done in
about 5 min or less for each egg case. Accuracy of
estimating versus a direct count method is very close,
roughly 95% (Benoit et al., 1997). The ring method is best
suited to the "C" shaped egg cases.
15.3.8.7.2 When the integrity of an egg case precludes
estimation by the ring method (egg case is convoluted or
distorted), the eggs should be counted directly. Each egg
case is placed into a 5-cm glass culture tube containing
about 2 ml of 2 N sulfuric acid (H2SO4) and left overnight.
The acid dissolves the gelatinous matrix surrounding the
eggs but does not affect the structural integrity of the
eggs themselves. After digestion, the eggs are collected
with a Pasteur pipet and spread across a microscope
slide for counting under a dissecting microscope. Count-
ing can be simplified by drawing a grid on the underside of
the slide. The direct count method requires a minimum of
10 min to complete and does not permit determination of
hatching success.
15.3.8.7.3 Following estimated egg counts, each egg
case is transferred to a 60- x 15-mm plastic petri dish
containing 15 ml overlying water and incubated at 23°C
until hatching is complete. Although the time required to
initiate hatching at this temperature is about 2 d, the
period of time required to bring about complete hatch may
be as long as 6 d. Therefore, hatching success is
determined after 6 d of incubation. Hatching success is
determined by subtracting the number of unhatched eggs
remaining after the 6 d period from the number of eggs
originally estimated for that egg case. Unhatched eggs
either remain in the gelatinous egg case or are distributed
on the bottom of the petri dish.
15.3.8.7.4 Depending on the objectives of the study,
reproductive output in C. tentans may be expressed as:
(1) number of eggs/female or (2) number of offspring/
female. The former approach estimates reproductive
output (fecundity) in terms of the number of eggs depos-
ited by a female (secondary egg cases are not included)
and does not take into account survival of hatched eggs.
This approach has been shown to adequately discrimi-
nate contaminant (Sibleyetal., 1996) and noncontaminant
(Sibley et al., 1997a) stressors. Since this approach does
not require monitoring egg masses for hatchability, the
time and labor involved in conducting the life-cycle test is
reduced. However, studies that require estimates of
demographic parameters, or include population modeling,
will need to determine the number of viable offspring per
female (Sibley et al., 1997a). This will require determina-
tion of larval hatch (see Section 15.3.8.7.3). Although
larval hatch is listed as a potential endpoint by itself in
this manual (Table 15.4), the sensitivity of this endpoint
has not been fully assessed.
15.3.9 Ending a Test
15.3.9.1 The point at which the life-cycle test is ended
depends upon the sediments being evaluated. In clean
sediments, the test typically requires 40 to 50 d from
initial setup to completion. However, test duration will
increase in the presence of environmental stressors which
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act to reduce growth and delay emergence (Sibley et al.,
1997a). Where a strong gradient of sediment contamina-
tion exists, emergence patterns between treatments will
likely become asynchronous, in which case each treat-
ment needs to be ended separately. For this reason,
emergence is used as a guide to decide when to end a
test.
15.3.9.2 For treatments in which emergence has oc-
curred, the treatment (not the entire test) is ended when
no further emergence is recorded over a period of 7 d (the
7-d criterion). At this time, all beakers of the treatment
are sieved through a #40-mesh screen (425 urn) to re-
cover remaining larvae, pupae, or pupal castes. When no
emergence is recorded in a treatment at any time during
the test, that treatment can be ended once emergence in
the control sediment has ended using the 7-d criterion.
15.4 Interpretation of Results
15.4.1 Data Analysis
15.4.1.1 Endpoints measured in the C. tentans test
include survival, growth, emergence and reproduction.
Section 16 describes general information regarding
statistical analysis of these data, including both point
estimates (i.e., LC50s) and hypothesis testing (i.e.,
ANOVA). The following sections describe species-specific
information that is useful in helping to interpret the results
of long-term sediment toxicity tests with C. tentans.
15.4.2 Age Sensitivity
15.4.2.1 Midges are perceived to be relatively insensitive
organisms in toxicity assessments (Ingersoll, 1995). This
conclusion is based on the practice of measuring survival
of fourth-instar larvae in short-term water-only exposures,
a procedure that may underestimate the sensitivity of
midges to toxicants. The first and second instars of
chironomids are more sensitive to contaminants than the
third or fourth instars. For example, first-instar C. tentans
larvae were 6 to 27 times more sensitive than fourth-instar
larvae to acute copper exposure (Nebeker et al., 1984b;
Gauss etal., 1985; Figure 12.1) and first-instar C. riparius
larvae were 127 times more sensitive than second-instar
larvae to acute cadmium exposure (Williams et al., 1986b;
Figure 12.1). In long-term tests with first-instar larvae,
midges were often as sensitive as daphnids to inorganic
and organic compounds (Ingersoll etal., 1990). Sediment
tests should be started with uniform age and size midges
because of the dramatic differences in sensitivity of
midges by age.
15.4.3 Physical Characteristics of Sediment
15.4.3.1 Grain Size
15.4.3.1.1 Larvae of C. tentans appear to be tolerant of a
wide range of particle size conditions in substrates. Sev-
eral studies have shown that survival is not affected by
particle size in natural sediments, sand substrates, or
formulated sediments in both 10-d and long-term expo-
sures (Ankley et al., 1994; Suedel and Rodgers, 1994;
Sibley et al., 1997b, 1998). Ankley et al. (1994a) found
that growth of C. tentans larvae was weakly correlated
with sediment grain size composition, but not organic
carbon, in 10-d tests using 50 natural sediments from the
Great Lakes. However, Sibley et al. (1997b) found that
the correlation between grain size and larval growth disap-
peared after accounting for inorganic material contained
within larval guts and concluded that growth of C. tentans
was not related to grain size composition in either natural
sediments or sand substrates. Avoiding confounding
influences of gut contents on weight is the impetus for
recommending ash-free dry weight (instead of dry weight)
as the index of growth in the 10-day and long-term
C. tentans tests. Failing to do so could lead to erroneous
conclusions regarding the toxicity of the test sediment
(Sibley et al., 1997b). Procedures for correcting for gut
contents are described in Section 15.3.8.3. Emergence,
reproduction (mean eggs/female), and hatch success
were also not affected by the particle size composition of
substrates in long-term tests with C. tentans (Sibley et
al., 1998).
15.4.3.2 Organic Matter
15.4.3.2.1 Based on 10-d tests, the content of organic
matter in sediments does not appear to affect survival of
C. tentans larvae in natural and formulated sediments, but
maybe important with respect to larval growth. Ankley et
al. (1994a) found no relationship between sediment or-
ganic content and survival or growth in 10-d bioassays
with C. tentans in natural sediments. Suedel and Rodgers
(1994) observed reduced survival in 10-d tests with a
formulated sediment when organic matter was <0.91%;
however, supplemental food was not supplied in this
study, which may influence these results relative to the
10-d test procedures described in this manual. Lacey et
al. (1999) found that survival of C. tentans larvae was
generally not affected in 10-d tests by eitherthe quality or
quantity of synthetic (alpha-cellulose) or naturally derived
(peat, maple leaves) organic material spiked into a formu-
lated sediment, although a slight reduction in survival
below the acceptability criterion (70%) was observed in a
natural sediment diluted with formulated sediment at an
organic matter content of 6%. In terms of larval growth,
Lacey et al. (1999) did not observe any systematic rela-
tionship between the level of organic material (e.g., food
quantity) and larval growth for each carbon source. Al-
though a significant reduction in growth was observed at
the highest concentration (10%) of the leaf treatment in
the food quantity study, significantly higher larval growth
was observed in this treatment when the different carbon
sources were compared at about equal concentrations
(effect of food quality). In the latter study, the following
gradient of larval growth was established in relation to the
source of organic carbon: peat < natural sediment
< alpha-cellulose < leaves. Since all of the treatments
received a supplemental source of food, these data sug-
gest that both the quality and quantity of organic carbon in
natural and formulated sediments may represent an im-
portant confounding factor forthe growth endpoint in tests
with C. tentans (Lacey et al., 1999). However, it is
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important to note that these data are based on 10-d tests;
the applicability of these data to long-term testing has not
been evaluated.
15.4.4 Isolating Organisms at the End of a Test
15.4.4.1 Quantitative recovery of larvae at the end of a
sediment test should not be a problem. The larvae are red
and typically greater than 5 mm long and are readily
retained on the #40-mesh sieve.
15.4.5 Influence of Indigenous Organisms
15.4.5.1 The influence of indigenous organisms on the
response of C. tentans in sediment tests has not been
reported. Survival of a closely related species, C. riparius
was not reduced in the presence of oligochaetes in sedi-
ment samples (Reynoldson etal., 1994). However, growth
of C. riparius was reduced when high numbers of oli-
gochaetes were placed in a sample. Therefore, it is
important to determine the number and biomass of indig-
enous organisms in field-collected sediment in order to
better interpret growth data (Reynoldson et al., 1994;
DeFoe and Ankley, 1998). Furthermore, the presence of
predators may also influence the response of test organ-
isms in sediment (Ingersoll and Nelson, 1990).
15.4.6 Relationship Between Endpoints
15.4.6.1 Relationship Between Growth and
Emergence Endpoints
15.4.6.1.1 An important stage in the life cycle of C. tentans
is the emergence of adults from pupal forms. Emergence
has been used in many studies as an indicator of con-
taminant stress (Wentsel et al., 1978; Pascoe et al.,
1989; Sibley et al., 1996). The use of emergence as an
endpoint in this context is based upon the understanding
that larval growth and emergence are intimately related
such that environmental factors that affect larval develop-
ment may also affect emergence success. Implicit in the
relationship between growth and emergence is the notion
of a weight threshold that needs to be attained by larvae in
order for emergence to take place (Hilsenhoff,1966; Liber
et al., 1996; Sibley et al., 1997a). For example, based on
evaluations conducted in clean control sediment, Liber et
al. (1996) and Sibley et al. (1997a) showed that a mini-
mum tissue mass threshold of approximately 0.6 mg dry
weight or 0.48 mg ash-free dry weight was required before
pupation and emergence could take place (Figure 15.1).
Further, Sibley et al. (1997a) found that maximum emer-
gence (e.g., >60%) in this sediment occurred only after
larvae had attained a tissue mass of about 0.8 mg dry
weight. This value corresponds closely to that suggested
by Ankley et al. (1994a) as an acceptability criterion for
growth in control sediments in 10-d tests with C. tentans.
15.4.6.2 Relationship Between Growth and
Reproduction Endpoints
15.4.6.2.1 Natural or anthropogenic stressors that affect
growth of invertebrates may also affect reproduction,
because of a minimum threshold body mass needed for
reproduction (Rees and Crawley, 1989; Ernsting et al.,
1993; Moore and Dillon, 1993; Sibley et al., 1996,1997a).
Sibley etal. (1996,1997a) reported a significant relation-
ship between growth (dry weight) of larval C. tenfansand
reproductive output (mean number of eggs) of adults in
relation to both food and contaminant (zinc) stressors
(Figure 15.2). The form that this relationship may take
depends upon the range of stress to which the larvae are
exposed and may be linear or sigmoidal. The latter
relationship is typically characterized by an upper maxi-
mum determined by competitive factors (i.e., food and
space availability) and a lower minimum determined pri-
marily by emergence thresholds (See Section 15.4.6.1;
Sibley etal., 1997a).
15.4.6.2.2 Embryo viability (percent hatch of eggs) has
been shown to evaluate the toxicity for waterborne
chemicals (Williams et al.,1986b; Pascoe et al.,1989).
However, percent hatch has not been used extensively as
an endpoint to assess toxicity in contaminated sedi-
ments. Sibley et al. (1996) found that the viability of
embryos was not affected at any of the zinc treatments
for which egg masses were produced; >87% of all eggs
eventually hatched. Additional information regarding the
measurement of embryo viability in round-robin testing is
presented in Section 17.6.
15.4.6.2.3 In contrast to H. azteca (Section 14.4), length
is not commonly utilized as a growth endpoint in C. tentans.
However, length may represent a useful alternative to
weight. For example, recent studies (P.K. Sibley, Univer-
sity of Guelph, Guelph, Ontario, unpublished data) found
a significant relationship (r2=0.99; p <0.001) between ash-
free dry weight and length in larvae of C. tentans reared in
clean control sediment (Figure 15.3). This suggests that
either weight or length could be used to assess growth
in C. tentans. However, the relationship between length
and emergence or reproductive endpoints has not been
evaluated.
15.4.6.3 Relationship Between Growth and
Population Endpoints
15.4.6.3.1 Few studies have attempted to quantitatively
define the relationship between larval growth and popula-
tion-level processes. However, an accurate understand-
ing of the ecological relevance of growth as an endpoint in
sediment toxicity tests can only be achieved in terms of
its effect, if any, on population-level processes. Sibley et
al. (1997a) found a significant relationship between larval
growth and the intrinsic rate of population increase in
C. tentans in relation to a food stressor (Figure 15.4).
When applied in a theoretical population model, it was
further demonstrated that changes in larval growth result-
ing from the stressor gradient were significantly correlated
to the predicted number of offspring recruited to subse-
quent generations.
93
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CD
O
CD
E?
CD
LLJ
-*— •
0
CD
Q_
QU
80
70
60
50
40
30
20
10
0
* *
~ s~ — ^ ^ ^ ^
i
/
1
Y
/
/
/
/
/
/ « Liber et at. (1996)
/ *• Sibley etal. (1997)
/ *
• /i i i i
0
0.5 1.0 1.5 2.0
Larval Dry Weight (mg/individual)
2.5
Figure 15.1 Relationship between weight and emergence of Chironomus tentans.
_
03
E
CD
LL
1/5
D)
D5
LJJ
c
03
CD
900
800
700
600
500
400
300
200
100
0
0 0.5 1 1.5 2
Larval Dry Weight (mg/individual)
Figure 15.2 Relationship between weight and reproduction of Chironomus tentans.
94
2.5
-------
CO
Q
0.01
0.001
0.0001
1
10
30
Length (mm)
Figure 15.3 Relationship between ash-free dry weight (AFDW) and length of Chironomus tentans.
03
V)
CO
O
Q.
O
Q.
a:
.O
c
'l_
•4—»
0.16
0.12
0.08
0.04
Y = 0.048X + 0.018
r2=0.97
0.5
1.5
2.5
AFDW (mg/individual)
Figure 15.4 Relationship between ash-free dry weight (AFDW) and intrinsic rate of natural increase of Chironomus tentans.
95
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75.4.6.4 Relative Endpoint Variability
15.4.6.4.1 Based on coefficient of variation (CV) deter-
mined from a control sediment (West Bearskin), the fol-
lowing variability has been documented for the various
endpoints in the C. tentans life-cycle test (Sibley et al.,
1996; Benoit et al., 1997): Survival (<20%), growth as dry
weight (<15%), emergence (<30%), reproduction as mean
eggs/female (<20%), percent hatch (<10%). Additional
information regarding the variation in these endpoints in
round-robin testing is presented in Section 17.6.
75.4.6.5 Relative Endpoint Sensitivity
15.4.6.5.1 Measurement of sublethal endpoints (e.g.,
growth) can often provide unique information in addition to
measuring survival. A comparison of lethal and sublethal
endpoints relative to toxicity identification is presented in
Table 14.4 for H. azteca. However, few studies have
compared the relative sensitivity of the various endpoints
in the C. tentans life cycle or in 10-d tests. Sibley et al.
(1997a) found that larval C. tentans exposed to a gradient
of food stress did not experience significant effects on
survival, yet did experience a significant reduction in
growth and reproduction. Further, the proportion of larvae
hatching in this study was high (>80%) and not
systematically related to treatment, suggesting that per-
cent hatch may be a relatively insensitive endpoint to
sediment-associated contaminants. This is consistent
with the findings of another study using zinc-spiked sedi-
ments; no effect on embryo viability was observed for
those treatments in which egg masses were produced
(Sibley et al. 1996). Although the responses observed in
the feeding study were not due to a contaminant stressor
per se, the sublethal endpoints were clearly better able to
discriminate the presence of the stressorthan was lethal-
ity. Ankley and DeFoe (1998) studied a variety of con-
taminated sediments and found that the sensitivity of
C. tentans 10-d tests is greatly increased by measure-
ment of growth in addition to survival. Growth of midge in
these 10-d sediment tests was found to be a more sensi-
tive endpoint than survival of Hyalella azteca.
15.4.7 Future Research
15.4.7.1 Additional studies using known concentration
gradients in sediment, should be conducted to better
differentiate the relative sensitivity between lethal and
sublethal endpoints and between sublethal endpoints in
the long-term C. fenfanstest. Additional studies also are
needed to further evaluate the influence of ammonia on
long-term exposures with C. tentans. Section 1.3.8.5
addresses interpretative guidance for evaluating toxicity
associated with ammonia in sediment. Planned water-
only toxicity tests with select chemicals (i.e., cadmium,
ODD, and fluoranthene) should generate data that can be
used to better determine the relative sensitivity of sur-
vival, reproduction, and growth endpoints in tests with C.
tentans. In addition to studies evaluating the relative
sensitivity of endpoints, research is also needed to evalu-
ate the ability of these laboratory endpoints to estimate
responses of benthic organisms exposed in the field to
chemicals in sediments.
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Section 16
Data Recording, Data Analysis and Calculations, and Reporting
16.1 Data Recording
16.1.1 Quality assurance project plans with data quality
objectives and standard operating procedures should be
developed before starting a test. Procedures should be
developed by each laboratory to verify and archive data
(USEPA, 1994e).
16.1.2 A file should be maintained for each sediment test
or group of tests on closely related samples (Section 9).
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 sediment test(s); chemical analysis data on the
sample(s); control data sheets for reference toxicants;
detailed records of the test organisms used in the test(s),
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 instru-
ments; test conditions used; and results of reference-
toxicity tests. Original data sheets should be signed and
dated by the laboratory personnel performing the tests. A
record of the electronic files of data should also be
included in the file.
16.1.3 Example data sheets are included in Appendix D.
16.2 Data Analysis
16.2.1 Statistical methods are used to make inferences
about populations, based on samples from those popula-
tions. In most sediment toxicity and bioaccumulation
tests, test organisms are exposed to chemicals in sedi-
ment to estimate the response of the population of labora-
tory organisms. The organism response to these sedi-
ments is usually compared with the response to a control
or reference sediment, or in some analyses of bioaccu-
mulation test data, with a fixed standard such as a Food
and Drug Administration (FDA) action level. In any toxic-
ity or bioaccumulation test, summary statistics such as
means and standard errors for response variables (e.g.,
survival, chemical concentrations in tissue) should be
provided for each treatment (e.g., pore-water concentra-
tion, sediment).
16.2.1.1 Types of Data.
16.2.1.1.1 Two types of data can be obtained from
sediment toxicity or bioaccumulation tests. The most
common endpoint in toxicity testing is mortality, which is
a dichotomous or categorical type of data. Other endpoints
measured in sublethal evaluations include growth and
reproduction (Sections 14 and 15) or tissue concentra-
tions (e.g., in sediment bioaccumulation tests conducted
with oligochaetes (Section 13) or with polychaetes and
mollusks; USEPA, 1994b). Growth, reproduction, and
bioaccumulation endpoints are representative of continu-
ous data.
76.2.1.2 Sediment Testing Scenarios
16.2.1.2.1 Sediment tests are conducted to determine
whether contaminants in sediment are harmful to or are
bioaccumulated in benthic organisms. Sediment tests are
commonly used in studies designed to (1) evaluate dredged
material, (2) assess site contamination in the environ-
ment (e.g., to rank areas for cleanup), and (3) determine
effects of specific contaminants, or combinations of con-
taminants, through the use of sediment-spiking tech-
niques. Each of these broad study designs has specific
statistical design and analytical considerations, which are
detailed below.
16.2.1.2.2 Dredged Material Evaluation. In these
studies, each site is compared individually with a refer-
ence sediment. The statistical procedures appropriate for
these studies are generally pain/vise comparisons. Addi-
tional information on toxicity testing of dredged material
and analysis of data from dredged material evaluations is
available in USEPA-USACE (1998a).
16.2.1.2.3 Site Assessment of Field Contamination.
Surveys of sediment toxicity or bioaccumulation often are
included in more comprehensive analyses of biological,
chemical, geological, and hydrographic data. Statistical
correlation can be improved and costs may be reduced if
subsamples are taken simultaneously for sediment toxic-
ity or bioaccumulation tests, chemical analyses, and
benthic community structure determinations. There are
several statistical approaches to field assessments, each
with a specific purpose. If the objective is to compare the
response or residue level at all sites individually to a
control sediment, then the pairwise comparison approach
described below is appropriate. If the objective is to
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compare among all sites in the study area, then a multiple
comparison procedure that employs an experiment-wise
error rate is appropriate. If the objective is to compare
among groups of sites, then orthogonal contrasts are a
useful data analysis technique.
16.2.1.2.4 Sediment-spiking Experiments. Sediments
spiked with known concentrations of chemicals can be
used to establish cause-and-effect relationships between
chemicals and biological responses. Results of toxicity
tests with test materials spiked into sediments at different
concentrations may be reported in terms of an LC50,
EC50, IC50, NOEC, orLOEC. Results of bioaccumulation
tests with either field or spiked samples may be reported
in terms of a BSAF (biota sediment accumulation factor;
ASTM, 1999c). The statistical approach outlined above
for spiked-sediment toxicity tests also applies to the
analysis of data from sediment dilution experiments or
water-only reference-toxicity tests.
16.2.2 Experimental Design
16.2.2.1 The guidance outlined below on the analysis of
sediment toxicity and bioaccumulation test data is adapted
from a variety of sources including ASTM (1999c), USEPA
(1991a), USEPA (1994a), USEPA (1994b), and
USEPA-USACE (1998a). The objectives of a sediment
toxicity or bioaccumulation test are to quantify contami-
nant effects on or accumulation in test organisms ex-
posed to natural or spiked sediments or dredged materials
and to determine whether these effects are statistically
different from those occurring in a control or reference
sediment. Each experiment consists of at least two treat-
ments: the control and one or more testtreatment(s). The
test treatments) consists) of the contaminated or poten-
tially contaminated sediment(s). A control sediment is
always required to ensure that no contamination is intro-
duced during the experiment setup and that test organ-
isms are healthy. A control sediment is used to judge the
acceptability of the test (Tables 11.3, 12.3, 13.4, 14.3,
15.3). Some designs also require a reference sediment
that represents an environmental condition or potential
treatment effect of interest. Controls are used to evaluate
the acceptability of the test and might include a control
sediment, a sand substrate (for C. tentans; Section 12.2,
15.2), or water-only exposures (for H. azteca; Section
14.3.7.8). Testing a reference sediment provides a
site-specific basis for evaluating toxicity of the test sedi-
ments. Comparisons of test sediments to multiple refer-
ence or control sediments representative of the physical
characteristics of the test sediment (i.e., grain size, or-
ganic carbon) may be useful in these evaluations
(Section 2.1.2).
16.2.2.2 Experimental Unit
16.2.2.2.1 During toxicity testing, each test chamber to
which a single application of treatment is applied is an
experimental unit. During bioaccumulation testing, how-
ever, the test organism may be the experimental unit if
individual members of the test species are evaluated and
they are large enough to provide sufficient biomass for
chemical analysis. The important concept is that the
treatment (sediment) is applied to each experimental unit
as a discrete unit. Experimental units should be indepen-
dent and should not differ systematically.
16.2.2.3 Replication
16.2.2.3.1 Replication is the assignment of a treatment to
more than one experimental unit. The variation among
replicates is a measure of the within-treatment variation
and provides an estimate of within-treatment error for
assessing the significance of observed differences be-
tween treatments.
16.2.2.4 Minimum Detectable Difference (MOD)
16.2.2.4.1 As the minimum difference between treat-
ments which the test is required or designed to detect
decreases, the number of replicates required to meet a
given significance level and power increases. Because no
consensus currently exists on what constitutes a biologi-
cally acceptable MOD, the appropriate statistical mini-
mum significant difference should be a data quality objec-
tive (DQO) established by the individual user (e.g., pro-
gram considerations) based on their data requirements,
the logistics and economics of test design, and the
ultimate use of the sediment toxicity or bioaccumulation
test results.
16.2.2.5 Minimum Number of Replicates
16.2.2.5.1 Eight replicates are recommended for 10-d
fresh water sediment toxicity testing (Section 11 and 12)
and five replicates are recommended for 10-d marine
testing (USEPA, 1994b). However, four replicates per
treatment are the absolute minimum number of replicates
for a 10-d sediment toxicity test. A minimum of five
replicates per treatment is recommended for bioaccumu-
lation testing (Section 13). It is always prudent to include
as many replicates in the test design as are economically
and logistically possible. USEPA 10-d sediment toxicity
testing methods recommend the use of 10 organisms per
replicate for fresh water testing or 20 organisms per repli-
cate for 10-d marine testing. An increase in the number of
organisms per replicate in all treatments is allowable only
if (1) test performance criteria forthe recommended num-
ber of replicates are achieved and (2) it can be demon-
strated that no change occurs in contaminant availability
due to the increased organism loading. See Tables 14.1
and 15.1 for a description of the number of replicates and
test organisms/replicate recommended for long-term test-
ing of Hyalella azteca or Chironomus tentans.
16.2.2.6 Randomization
16.2.2.6.1 Randomization is the unbiased assignment of
treatments within a test system and to the exposure
chambers ensuring that no treatment is favored and that
observations are independent. It is also important to
(1) randomly select the organisms (but not the number of
organisms) for assignment to the control and test
treatments (e.g., a bias in the results may occur if all of
the largest animals are placed in the same treatment),
98
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(2) randomize the allocation of sediment (e.g., do not take
all the sediment in the top of a jar for the control and the
bottom for spiking), and (3) randomize the location of
exposure units.
16.2.2.7 Pseudoreplication
16.2.2.7.1 The appropriate assignment of treatments to
the replicate exposure chambers is critical to the avoid-
ance of a common error in design and analysis termed
"pseudoreplication" (Hurlbert, 1984). Pseudoreplication oc-
curs when inferential statistics are used to test for treat-
ment effects even though the treatments are not repli-
cated or the replicates are not statistically independent
(Hurlbert, 1984). The simplest form of pseudoreplication
is the treatment of subsamples of the experimental unit
as true replicates. For example, two aquaria are prepared,
one with control sediment and the other with test sedi-
ment, and 10 organisms are placed in each aquarium.
Even if each organism is analyzed individually, the 10
organisms only replicate the biological response and do
not replicate the treatment (i.e., sediment type). In this
case, the experimental unit is the 10 organisms and each
organism is a subsample. A less obvious form of pseudo-
replication is the potential systematic error due to the
physical segregation of exposure chambers by treatment.
For example, if all the control exposure chambers are
placed in one area of a room and all the test exposure
chambers are in another, spatial effects (e.g., different
lighting, temperature) could bias the results for one set of
treatments. Random physical intermixing of the exposure
chambers or randomization of treatment location may be
necessary to avoid this type of pseudoreplication. Pseu-
doreplication can be avoided or reduced by properly iden-
tifying the experimental unit, providing replicate experi-
mental units for each treatment, and applying the treat-
ments to each experimental unit in a mannerthat includes
random physical intermixing (interspersion) and indepen-
dence. However, avoiding pseudoreplication completely
may be difficult or impossible given resource constraints.
16.2.2.8 Optimum Design of Experiments
16.2.2.8.1 An optimum design is one which obtains the
most precise answer for the least effort. It maximizes or
minimizes one of many optimality criteria, which are
formal, mathematical expressions of certain properties of
the model that are fit to the data. Optimum design of
experiments using specific approaches described in
Atkinson and Donev (1992) has not been formally applied
to sediment testing; however, it might be desirable to use
the approaches in experiments. The choice of optimality
criterion depends on the objective of the test, and compos-
ite criteria can be used when a test has more than one goal.
A design is optimum only for a specific model, so it is
necessary to know beforehand which models might be
used (Atkinson and Donev, 1992).
16.2.2.9 Compositing Samples
16.2.2.9.1 Decisions regarding compositing of samples
depend on the objective of the test. Compositing is used
primarily in bioaccumulation experiments when the biom-
ass of an individual organism is insufficient for chemical
analysis. Compositing consists of combining samples
(e.g., organisms, sediment) and chemically analyzing the
mixture ratherthan the individual samples. The chemical
analysis of the mixture provides an estimate of the aver-
age concentration of the individual samples making up
the composite. Compositing also may be used when the
cost of analysis is high. Each organism or sediment
sample added to the composite should be of equal size
(i.e., wet weight) and the composite should be completely
homogenized before taking a sample for chemical analy-
sis. If compositing is performed in this manner, the value
obtained from the analysis of the composite is the same
as the average obtained from analyzing each individual
sample (within any sampling and analytical errors). If true
replicate composites (not subsample composites) are
made, the variance of the replicates will be less than the
variance of the individual samples, providing a more
precise estimate of the mean value. This increases the
power of a test between means of composites over a test
between means of individuals or samples for a given
number of samples analyzed. If compositing reduces the
actual number of replicates, however, the power of the
test will also be reduced. If composites are made of
individuals or samples varying in size, the value of the
composite and the mean of the individual organisms or
sediment samples are no longer equivalent. The variance
of the replicate composites will increase, decreasing the
power of any test between means. In extreme cases, the
variance of the composites can exceed the population
variance (Tetra Tech, 1986). Therefore, it is important to
keep the individuals or sediment samples comprising the
composite equivalent in size. If sample sizes vary, con-
sult the tables in Schaeffer and Janardan (1978) to deter-
mine if replicate composite variances will be higher than
individual sample variances, which would make compos-
iting inappropriate.
16.2.3 Hypothesis Testing and Power
16.2.3.1 The purpose of a toxicity or bioaccumulation
test is to determine if the biological response to a treat-
ment sample differs from the response to a control sample.
Figure 16.1 presents the possible outcomes and deci-
sions that can be reached in a statistical test of such a
hypothesis. The null hypothesis is that no difference
exists among the mean control and treatment responses.
The alternative hypothesis of greatest interest in sedi-
ment tests is that the treatments are toxic, or contain
concentrations of bioaccumulatable compounds, relative
to the control or reference sediment.
16.2.3.2 Statistical tests of hypotheses can be designed
to control for the chances of making incorrect decisions.
In Figure 16.1, alpha (a) represents the probability of
making a Type I statistical error. A Type I statistical error
in this testing situation results from the false conclusion
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Decision
TR =Control
TR > Control
TR =Control
TR > Control
Correct
1 -a
Type I
Error
a
Type II
Error
P
Correct
1-P
(Power)
Treatment response (TR), Alpha (a) represents the probability of
making a Type I statistical error (false positive); beta (P)
represents the probability of making a Type II statistical error
(false negative).
Figure 16.1 Treatment response for a Type I and Type II error.
that the treated sample is toxic or contains chemical
residues not found in the control or reference sample.
Beta (P) represents the probability of making a Type II
statistical error, or the likelihood that one erroneously
concludes there are no differences among the mean
responses in the treatment, control or reference samples.
Traditionally, acceptable values for a have ranged from
0.1 to 0.01 with 0.05 or 5% used most commonly. This
choice should depend upon the consequences of making
a Type I error. Historically, having chosen a, environmen-
tal researchers have ignored p and the associated power
of the test (1-0).
16.2.3.3 Fairweather (1991) presents a review of the need
for, and the practical implications of, conducting power
analyses in environmental monitoring studies. This re-
view also includes a comprehensive bibliography of re-
cent publications on the need for, and use of, power
analyses in environmental study design and data analy-
sis. The consequences of a Type II statistical error in
environmental studies should never be ignored and may,
in fact, be one of the most important criteria to consider in
experimental designs and data analyses that include
statistical hypothesis testing. To paraphrase Fairweather
(1991), "The commitment of time, energy and people to a
false positive (a Type I error) will only continue until the
mistake is discovered. In contrast, the cost of a false
negative (a Type II error) will have both short- and long-term
costs (e.g., ensuing environmental degradation and the
eventual cost of its rectification)."
16.2.3.4 The critical components of the experimental
design associated with the testing of hypotheses outlined
above are (1) the required MOD between the treatment
and control or reference responses, (2) the variance among
treatment and control replicate experimental units, (3) the
number of replicate units for the treatment and control
samples, (4) the number of animals exposed within a
replicate exposure chamber, and (5) the selected prob-
abilities of Type I (a) and Type II (P) errors.
16.2.3.5 Sample size or number of replicates may be
fixed due to cost or space considerations or may be
varied to achieve a priori probabilities of a and p. The
MOD should be established ahead of time based upon
biological and program considerations. The investigator
has little control of the variance among replicate expo-
sure chambers. However, this variance component can
be minimized by selecting test organisms that are as
biologically similar as possible and maintaining test con-
ditions within prescribed quality control (QC) limits.
16.2.3.6 The MOD is expressed as a percentage change
from the mean control response. To test the equality of
the control and treatment responses, a two-sample ttest
with its associated assumptions is the appropriate para-
metric analysis. If the desired MOD, the number of repli-
cates per treatment, the number of organisms per repli-
cate and an estimate of typical among replicate variabil-
ity, such as the coefficient of variation (CV) from a control
sample, are available, it is possible to use a graphical
approach as in Figure 16.2 to determine how likely it is
that a 20% reduction will be detected in the treatment
response relative to the control response. The CV is
defined as 100% x (standard deviation divided by the
mean). In a test design with 8 replicates per treatment
and with an a level of 0.05, high power (i.e., >0.8) to
detect a 20% reduction from the control mean occurs
only if the CVis 15% or less (Figure 16.2). The choice of
these variables also affects the power of the test. If 5
replicates are used per treatment (Figure 16.3), the CV
needs to be 10% or lower to detect a 20% reduction in
response relative to the control mean with a power of 90%.
16.2.3.7 Relaxing the a level of a statistical test in-
creases the power of the test. Figure 16.4 duplicates
Figure 16.2 except that a is 0.10 instead of 0.05. Selec-
tion of the appropriate a level of a test is a function of the
costs associated with making Type I and II statistical
errors. Evaluation of Figure 16.2 illustrates that with aCV
of 15% and an a level of 0.05, there is an 80% probability
(power) of detecting a 20% reduction in the mean treat-
ment response relative to the control mean. However, if
a is set at 0.10 (Figure 16.4) and the CV remains at 15%,
then there is a 90% probability (power) of detecting a 20%
reduction relative to the control mean. The latter example
would be preferable if an environmentally conservative
analysis and interpretation of the data is desirable.
16.2.3.8 Increasing the number of replicates per treat-
ment will increase the power to detect a 20% reduction in
treatment response relative to the control mean
(Figure 16.5). Note, however, that for less than 8 repli-
cates per treatment it is difficult to have high power
(i.e., >0.80) unless the CV is less than 15%. If space or
cost limit the number of replicates to fewer than 8 per
treatment, then it may be necessary to find ways to
reduce the among replicate variability and consequently
the CV. Options that are available to increase the power
of the test include selecting more uniform organisms to
reduce biological variability or increasing the a level of
the test. For CVs in the range of 30% to 40%, even
8 replicates per treatment is inadequate to detect small
reductions (<20%) in response relative to the control
mean.
100
-------
-------
10 20 30 40 50 60 70 80
% Reduction of Control Mean
Figure 16.4 Power of the test vs. percent reduction in treatment response relative to the control mean at various CVs
(8 replicates, alpha = 0.10 [one-tailed]).
1.2 T
0.8- •
O
0.
0.6- •
0.4- .
0.2 • •
CV = 5%
6 8 10 12
No. of Replicates (n)
14
16
Figure 16.5 Effect of CV and number of replicates on the power to detect a 20% decrease in treatment response relative to the
control mean (alpha = 0.05 [one-tailed]).
102
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16.2.3.9 The effect of the choice of a and pon number of
replicates for various CVs, assuming the combined total
probability of Type I and Type II statistical errors is fixed
at 0.25, is illustrated in Figure 16.6. An a of 0.10 therefore
establishes a p of 0.15. In Figure 16.6, if oc = p = 0.125,
the number of replicates required to detect a difference of
20% relative to the control is at a minimum. As a or p
decrease, the number of replicates required to detect the
same 20% difference relative to the control increases.
However, the curves are relatively flat over the range of
0.05 to 0.20, and their shape will change dramatically if
the combined total a + p is changed. Limiting the total of
a + p to 0.10 greatly increases the number of replicates
necessary to detect a preselected percentage reduction
in mean treatment response relative to the control mean.
16.2.4 Comparing Means
16.2.4.1 Figure 16.7 outlines a decision tree for analysis
of survival, growth, or reproduction data subjected to
hypothesis testing. In the tests described herein, samples
or observations referto replicates of treatments. Sample
size n is the number of replicates (i.e., exposure cham-
bers) in an individual treatment, not the number of organ-
isms in an exposure chamber. Overall sample size N is
the combined total number of replicates in all treatments.
The statistical methods discussed in this section are
described in general statistics texts such as Steel and
Torrie (1980), Sokal and Rohlf (1981), Dixon and Massey
(1983), Zar (1984), and Snedecor and Cochran (1989). It
is recommended that users of this manual have at least
one of these texts and associated statistical tables on
hand. A nonparametric statistics text such as Conover
(1 980) might also be helpful.
76.2.4.2 Mean
16.2.4.2.1 The sample mean (x) is the average value, or
Ex/n where
n = number of observations (replicates)
x; = ith observation
Ex, = every x summed = x1+x2 + x3 + ...+xn
16.2.4.3 Standard Deviation
16.2.4.3.1 The sample standard deviation (s) is a mea-
sure of the variation of the data around the mean and is
equivalent to . The sample variance, s2, is given by
the following "machine" or "calculation" formula:
n-l
25
20--
15 --
CO
"a.
03
OL
O
z
10 ••
5 ••
-I—I-
I I I 1 1 I I I I I I 1 I
Alpha (Beta = 0.25 - Alpha)
Figure 16.6 Effect of alpha and beta on the number of replicates at various CVs (assuming combined alpha + beta = 0.25).
103
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Data-Survival. Growth, and Reproduction
I
Test for Normality
Normal <—
N'
Tests for Homogeneity of Variance
Shapiro-Wilk'sTest(N
/
snce
<50)
s.
— ik
Bartlett's
Hartley's
Heterogenous Variances
>Non Normal —
4.
Transformation?
T I
I
Homogenous Variances Noi n ~ 2 )
Yes, n > 2 I
Rankits
/
No
>3 Replicates
4-
ANOVA
I
Equal Replication
t test for
Unequal Variances
No
Bonferroni's
Yes
Comparison-Wise Alpha
Fisher's LSD, Duncan's
Experiment-Wise Alpha
Dunnett's
Yes
Equal Replication
Yes
\ ?
No
Steel's Many-One
Rank Test
Wilcoxon
with Bonferroni's
Endpoint
Figure 16.7 Decision tree for analysis of survival, growth, and reproduction data subjected to hypothesis testing.
16.2.4.4 Standard Error of the Mean
16.2.4.4.1 The standard error of the mean (SE, or
estimates variation among sample means rather than
among individual values. The SE is an estimate of the
standard deviation among means that would be obtained
from several samples of n observations each. Most of the
statistical tests in this manual compare means with other
means (e.g., dredged sediment mean with reference mean)
or with a fixed standard (e.g., FDA action level; ASTM,
1999c). Therefore, the "natural" or "random" variation of
sample means (estimated by SE), rather than the varia-
tion among individual observations (estimated by s), is
required for the tests.
16.2.4.5 Tests of Assumptions
16.2.4.5.1 In general, parametric statistical analyses
such as t tests and analysis of variance are appropriate
only if (1) there are independent, replicate experimental
units for each treatment, (2) the observations within each
treatment follow a normal distribution, and (3) variances
for both treatments are equal or similar. The first assump-
tion is an essential component of experimental design.
The second and third assumptions can be tested using
the data obtained from the experiment. Therefore, before
conducting statistical analyses, tests for normality and
equality of variances should be performed.
16.2.4.5.2 Outliers. Extreme values and systematic
departures from a normal distribution (e.g., a log-normal
distribution) are the most common causes of departures
from normality or equality of variances. An outlier is an
inconsistent or questionable data point that appears un-
representative of the general trend exhibited by the major-
ity of the data. Outliers may be detected by tabulation of
the data, by plotting, or by analysis of residuals. An
explanation should be sought for any questionable data
points. Without an explanation, data points should only be
discarded with extreme caution. If there is no explanation,
the analysis should be performed both with and without
the outlier, and the results of both analyses should be
reported. An appropriate transformation, such as the arc
sine-square root transformation, will normalize many
distributions (USEPA, 1985). Problems with outliers can
usually be solved only by using nonparametric tests, but
careful laboratory practices can reduce the frequency of
outliers.
16.2.4.5.3 Tests for Normality. The most commonly
used test for normality for small sample sizes (N<50) is
the Shapiro-Wilk's test. This test determines if residuals
are normally distributed. Residuals are the differences
between individual observations and the treatment mean.
Residuals, rather than raw observations, are tested be-
cause subtracting the treatment mean removes any dif-
ferences among treatments. This scales the observations
so that the mean of residuals for each treatment and over
104
-------
all treatments is zero. The Shapiro-Wilk's test provides a
test statistic W, which is compared to values of W
expected from a normal distribution. Wwill generally vary
between 0.3 and 1.0, with lower values indicating greater
departure from normality. Because normality is desired,
one looks for a high value of W with an associated
probability greater than the pre-specified a level.
16.2.4.5.3.1 Table 16.1 provides a levels to determine
whether departures from normality are significant. Nor-
mality should be rejected when the probability associated
with W (or other normality test statistic) is less than a for
the appropriate total number of replicates (N) and design.
A balanced design means that all treatments have an
equal number (n) of replicate exposure chambers. A
design is considered unbalanced when the treatment with
the largest number of replicates (nmax) has at least twice
as many replicates as the treatment with the fewest
replicates (nmin). Note that higher a levels are used when
the number of replicates is small, or when the design is
unbalanced, because these are the cases in which depar-
tures from normality have the greatest effects on t tests
and other parametric comparisons. If data fail the test for
normality, even after transformation, nonparametric tests
should be used for additional analyses (See Section
16.2.4.8 and Figure 16.7).
16.2.4.5.3.2 Tables of quantiles of W can be found in
Shapiro and Wilk (1965), Gill (1978), Conover (1980),
USEPA (1989c) and other statistical texts. These refer-
ences also provide methods of calculating W, although
the calculations can be tedious. Forthat reason, commonly
available computer programs or statistical packages are
preferred for the calculation of W.
16.2.4.5.4 Tests for Homogeneity of Variances. There
are a number of tests for equality of variances. Some of
these tests are sensitive to departures from normality,
which is why a test for normality should be performed
first. Bartlett's test or other tests such as Levene's test or
Cochran's test (Winer, 1971; Snedecor and Cochran,
1989) all have similar power for small, equal sample sizes
Table 16.1 Suggested a Levels to Use for Tests of
Assumptions
Test
Normality
Equality of variances
Number of
Observations1
N = 2 to 9
N = 1 0 to 1 9
N = 20 or more
n = 2 to 9
n = 10 or more
a When Design Is
Balanced Unbalanced2
0.10
0.05
0.01
0.10
0.05
0.25
0.10
0.05
0.25
0.10
1 N = total number of observations (replicates) in all treatments
combined; n = number of observations (replicates) in an
individual treatment
(n=5) (Conover etal., 1981). The data must be normally
distributed for Bartlett's test. Many software packages for
t tests and analysis of variance (ANOVA) provide at least
one of the tests.
16.2.4.5.4.1 If no tests for equality of variances are
included in the available statistical software, Hartley's
F can easily be calculated:
Fmax = (larger of sl , s\ ) / (smaller of sl , s\ )
When Fmax is large, the hypothesis of equal variances is
more likely to be rejected. Fmax is a two-tailed test be-
cause it does not matter whicl-Tvariance is expected to be
larger. Some statistical texts provide critical values of
Fmax (Winer, 1971; Gill, 1978; Rohlf and Sokal, 1981).
16.2.4.5.4.2 Levels of a for tests of equality of variances
are provided in Table 16.1. These levels depend upon
number of replicates in a treatment (n) and allotment of
replicates among treatments. Relatively high a's
(i.e., >0.10) are recommended because the power of the
above tests for equality of variances is rather low
(about 0.3) when n is small. Equality of variances is
rejected if the probability associated with the test statistic
is less than the appropriate a.
16.2.4.6 Transformations of the Data
16.2.4.6.1 When the assumptions of normality or homo-
geneity of variance are not met, transformations of the
data may remedy the problem, so that the data can be
analyzed by parametric procedures, rather than by a
nonparametric technique. The first step in these analyses
is to transform the responses, expressed as the propor-
tion surviving, by the arc sine-square root transformation.
The arc sine-square root transformation is commonly
used on proportionality data to stabilize the variance and
satisfy the normality requirement. If the data do not meet
the assumption of normality and there are four or more
replicates pergroup, then the nonparametric test, Wilcoxon
Rank Sum test, can be used to analyze the data. If the
data meet the assumption of normality, Bartlett's test or
Hartley's Ftest 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 ftest, and the degrees of freedom for the test are
adjusted.
16.2.4.6.2 The arc sine-square root transformation con-
sists of determining the angle (in radians) represented by
a sine value. In this transformation, the proportion surviv-
ing is taken as the sine value, the square root of the sine
value is calculated, and the angle (in radians) for the
square root of the sine value is determined. When the
proportion surviving is 0 or 1, a special modification of the
transformation should be used (Bartlett, 1937). An ex-
ample of the arc sine-square root transformation and
modification are provided below.
105
-------
1. Calculate the response proportion (RP) for each repli-
cate within a group, where
RP = (number of surviving organisms)/(number ex-
posed)
2. Transform each RP to arc sine, as follows:
a. For RPs greaterthan zero or less than one:
Angle (in radians) = arc sine J(RP)
b. Modification of the arc sine when RP = 0.
Angle (in radians) = arc sine J—
where n = number of animals/treatment rep.
c. Modification of the arc sine when RP = 1.0.
Angle = 1.5708radians-(radians forRP = 0)
16.2.4.7 Two Sample Comparisons (N=2)
16.2.4.7.1 The true population mean (u) and standard
deviation (a) are known only after sampling the entire
population. In most cases, samples are taken randomly
from the population, and the s calculated from those
samples is only an estimate of o. Student's f-values
account for this uncertainty. The degrees of freedom for
the test, which are defined as the sample size minus one
(n-1), should be used to obtain the correct f-value. Student's
f-values decrease with increasing sample size because
larger samples provide a more precise estimate of u and o.
16.2.4.7.2 When using a ttable, it is crucial to determine
whether the table is based on one-tailed probabilities or
two-tailed probabilities. In formulating a statistical hypoth-
esis, the alternative hypothesis can be one-sided
(one-tailed test) or two-sided (two-tailed test). The null
hypothesis (H0) is always that the two values being ana-
lyzed are equal. A one-sided alternative hypothesis (Ha) is
that there is a specified relationship between the two
values (e.g., one value is greaterthan the other) versus a
two-sided alternative hypothesis (Ha) which is that the two
values are simply different (i.e., either larger or smaller). A
one-tailed test is used when there is an a priori reason to
test for a specific relationship between two means, such
as the alternative hypothesis that the treatment mortality
or tissue residue is greater than the control mortality or
tissue residue. In contrast, the two-tailed test is used
when the direction of the difference is not important or
cannot be assumed before testing.
16.2.4.7.3 Since control organism mortality or tissue
residues and sediment chemical concentrations are pre-
sumed lower than reference or treatment sediment val-
ues, conducting one-tailed tests is recommended in most
cases. For the same number of replicates, one-tailed
tests are more likely to detect statistically significant
differences between treatments (e.g., have a greater
power) than are two-tailed tests. This is a critical consid-
eration when dealing with a small number of replicates
(such as 8/treatment). The other alternative for increasing
statistical power is to increase the number of replicates,
which increases the cost of the test.
16.2.4.7.4 There are cases when a one-tailed test is
inappropriate. When no a priori assumption can be made
as to how the values vary in relationship to one another, a
two-tailed test should be used. An example of an alterna-
tive two-sided hypothesis is that the reference sediment
total organic carbon (TOC) content is different (greater or
lesser) from the control sediment TOC. A two-tailed test
should also be used when comparing tissue residues
among different species exposed to the same sediment
and when comparing bioaccumulation factors (BAFs) or
biota-sediment accumulation factors (BSAFs).
16.2.4.7.5 The f-value for a one-tailed probability can be
found in a two-tailed table by looking up t under the
column for twice the desired one-tailed probability. For
example, the one-tailed f-value for a = 0.05 and df = 20
is 1.725, and is found in a two-tailed table using the
column fora = 0.10.
16.2.4.7.6 The usual statistical test for comparing two
independent samples is the two-sample t test (Snedecor
and Cochran, 1989). The f-statistic for testing the equality
of means xj' and x^ from two independent samples with n1
and n2 replicates and unequal variances is
t-(xr x2) /
nr
where sf and s\ are the sample variances of the two
groups. Although the equation assumes that the vari-
ances of the two groups are unequal, it is equally useful
for situations in which the variances of the two groups are
equal. This statistic is compared with the Student's t
distribution with degrees of freedom (df) given by
Satterthwaite's (1946) approximation:
df =
2
(~2
si' m,
This formula can result in fractional degrees of freedom,
in which case one should round the degree of freedom
down to the nearest integer in orderto use a t table. Using
this approach, the degrees of freedom for this test will be
less than the degrees of freedom for a t test assuming
equal variances. If there are unequal numbers of repli-
cates in the treatments, the t test with Bonferroni's adjust-
ment can be used for data analysis (USEPA, 1994c;
USEPA, 1994d). When variances are equal, an Ftestfor
equality is unnecessary.
16.2.4.8 Nonparametric Tests
16.2.4.8.1 Tests such as the t test, which analyze the
original or transformed data and which rely on the proper-
ties of the normal distribution, are referred to as paramet-
ric tests. Nonparametric tests, which do not require nor-
mally distributed data, analyze the ranks of data and
generally compare medians ratherthan means. The me-
106
-------
dian of a sample is the middle or 50th percentile observa-
tion when the data are ranked from smallest to largest. In
many cases, nonparametric tests can be performed sim-
ply by converting the data to ranks or normalized ranks
(rankits) and conducting the usual parametric test proce-
dures on the ranks or rankits.
16.2.4.8.2 Nonparametric tests are useful because of
their generality, but have less statistical power than corre-
sponding parametric tests when the parametric test as-
sumptions are met. If parametric tests are not appropriate
for comparisons because the normality assumption is not
met, data should be converted to normalized ranks
(rankits). Rankits are simply the z-scores expected for
the rank in a normal distribution. Thus, using rankits
imposes a normal distribution overall the data, although
not necessarily within each treatment. Rankits can be
obtained by ranking the data, then converting the ranks to
rankits using the following formula:
— 7
[(rank - 0.375) /(N + 0.25)]
where z is the normal deviate and N is the total number of
observations. Alternatively, rankits may be obtained from
standard statistical tables such as Rohlf and Sokal (1981).
16.2.4.8.3 If normalized ranks are calculated, the ranks
should be converted to rankits using the formula above. In
comparisons involving only two treatments (N=2), there is
no need to test assumptions on the rankits or ranks;
simply proceed with a one-tailed t test for unequal vari-
ances using the rankits or ranks.
16.2.4.9 Analysis of Variance (N>2)
16.2.4.9.1 Some experiments are set up to compare
more than one treatment with a control, whereas others
may also be interested in comparing the treatments with
one another. The basic design of these experiments is the
same as for experiments evaluating pain/vise compari-
sons. After the applicable comparisons are determined,
the data must be tested for normality to determine whether
parametric statistics are appropriate and whether the
variances of the treatments are equal. If normality of the
data and equal variances are established, then an analysis
of variance (ANOVA) may be performed to address the
hypothesis that all the treatments, including the control,
are equal. If normality or equality of variance are not
established, then transformations of the data might be
appropriate, or nonparametric statistics can be used to
test for equal means. Tests for normality of the data
should be performed on the treatment residuals. A re-
sidual is defined as the observed value minus the treat-
ment mean, that is, rik = oik - (kth treatment mean). Pooling
residuals provides an adequate sample size to test the
data for normality.
16.2.4.9.2 The variances of the treatments should also
be tested for equality. Currently there is no easy way to
test for equality of the treatment means using analysis of
variance if the variances are not equal. In a toxicity test
with several treatments, one treatment may have 100%
mortality in all of its replicates, or the control treatment
may have 100% survival in all of its replicates. These
responses result in 0 variance for a treatment that results
in a rejection of equality of variance in these cases. No
transformation will change this outcome. In this case, the
replicate responses for the treatment with 0 variance
should be removed before testing for equality of vari-
ances. Only those treatments that do not have 0 replicate
variance should be used in the ANOVA to get an estimate
of the within treatment variance. After a variance estimate
is obtained, the means of the treatments with 0 variance
can be tested against the other treatment means using
the appropriate mean comparison. Equality of variances
among the treatments can be evaluated with the Hartley
Fmax test or Bartlett's test. The option of using
nonparametric statistics on the entire set of data is also
an alternative.
16.2.4.9.3 If the data are not normally distributed or the
variances among treatments are not homogeneous, even
after data transformation, nonparametric analyses are
appropriate. If there are four or more replicates per treat-
ment and the number of replicates per treatment is equal,
the data can be analyzed with Steel's Many-One Rank
test. Unequal replication among treatments requires data
analysis with the Wilcoxon Rank Sum test with Bonferroni's
adjustment. Steel's Many-One Rank test is a nonpara-
metric test for comparing treatments with a control. This
test is an alternative to the Dunnett's test, and may be
applied to data when the normality assumption has not
been met. Steel's test requires equal variances across
treatments and the control, but is thought to be fairly
insensitive to deviations from this condition (USEPA,
1991 a). Wilcoxon's Rank Sum test is a nonparametric
test to be used as an alternative to the Steel's test when
the number of replicates are not the same within each
treatment. A Bonferroni's adjustment of the pain/vise error
rate for comparison of each treatment versus the control
is used to set an upper bound of alpha on the overall error
rate. This is in contrast to the Steel's test with a fixed
overall error rate for alpha. Thus, Steel's test is a more
powerful test (USEPA, 1991 a).
16.2.4.9.4 Different mean comparison tests are used
depending on whether an a percent comparison-wise error
rate or an a percent experiment-wise error rate is desired.
The choice of a comparison-wise or experiment-wise
error rate depends on whether a decision is based on a
pairwise comparison (comparison-wise) or from a set
of comparisons (experiment-wise). For example, a
comparison-wise error rate would be used for deciding
which stations along a gradient were acceptable or not
acceptable relative to a control or reference sediment.
Each individual comparison is performed independently at
a smaller a (than that used in an experiment-wise com-
parison), such that the probability of making a Type I error
in the entire series of comparisons is not greater than the
chosen experiment-wise a level of the test. This results in
a more conservative test when comparing any particular
sample to the control or reference. However, if several
samples were taken from the same area and the decision
to accept or reject the area was based upon all comparisons
107
-------
with a reference, then an experiment-wise error rate should
be used. When an experiment-wise error rate is used, the
power to detect real differences between any two means
decreases as a function of the number of treatment
means being compared to the control treatment.
16.2.4.9.5 The recommended procedure for pain/vise
comparisons that have a comparison-wise a error rate
and equal replication is to do an ANOVA followed by a
one-sided Fisher's Least Significant Difference (LSD) test
(Steel and Torrie, 1980). A Duncan's mean comparison
test should give results similar to the LSD. If the treat-
ments do not contain equal numbers of replicates, the
appropriate analysis is the t test with Bonferroni's adjust-
ment. For comparisons that maintain an experiment-wise
a error rate, Dunnett's test is recommended for compari-
sons with the control.
16.2.4.9.6 Dunnett's test has an overall error rate of a,
which accounts for the multiple comparisons with the
control. Dunnett's procedure uses a pooled estimate of
the variance, which is equal to the error value calculated
in an ANOVA.
16.2.4.9.7 To perform the individual comparisons, calcu-
late the t statistic for each treatment and control combina-
tion, as follows:
ti = -
where Y = mean for each treatment
Y,
= mean for the control
Sw = square root of the within mean square
n1 = number of replicates in the control
a = number of replicates for treatment "i"
To quantify the sensitivity of the Dunnett's test, the
minimum significant difference (MSD=MDD) may be cal-
culated with the following formula:
where d = Critical value for the Dunnett's Proce-
dure
Sw = The square root of the within mean square
n = The number of replicates per treatment,
assuming an equal number of replicates
at all treatment concentrations
n1 = Number of replicates in the control
16.2.5 Methods for Calculating LCSOs, ECSOs,
and ICps
16.2.5.1 Figure 16.8 outlines a decision tree for analysis
of point estimate data. USEPA manuals (USEPA, 1991 a;
USEPA, 1994c; USEPA, 1994d) discuss in detail the
mechanics of calculating LC50 (or EC50) or ICp values
using the most current methods. The most commonly
used methods are the Graphical, Probit, trimmed
Spearman-Karberand the Linear Interpolation Methods.
Methods for evaluating point estimate data using logistic
regression are outlined in Snedecorand Cochran (1989).
In general, results from these methods should yield simi-
lar estimates. Each method is outlined below, and recom-
mendations are presented forthe use of each method.
16.2.5.2 Data for at least five test concentrations and the
control should be available to calculate an LC50, although
each method can be used with fewer concentrations.
Survival in the lowest concentration must be at least
50%, and an LC50 should not be calculated unless at
least 50% of the organisms die in at least one of the serial
dilutions. When less than 50% mortality occurs in the
highest test concentration, the LC50 is expressed as
greater than the highest test concentration.
16.2.5.3 Due to the intensive nature of the calculations
forthe estimated LC50 and associated 95% confidence
interval using most of the following methods, it is recom-
mended that the data be analyzed with the aid of com-
puter software. Computer programs to estimate the LC50
or ICp values and associated 95% confidence intervals
using the methods discussed below (except forthe Graphi-
cal Method) were developed by USEPA and can be
obtained by sending a diskette with a written request to
USEPA, National Exposure Research Laboratory, 26 W.
Data Survival Point Estimates
Two or More Partial Mortalities
Yes
No
Significant Chi-Square Test
Yes
One Partial Mortality
No
Yes
Linear Interpolation
Trimmed Spearman-Karber
1
aphi
T
LC50 and 95% Confidence Intervals
Figure 16.8 Decision tree for analysis of point estimate data.
108
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Martin Luther King Drive, Cincinnati, OH 45268 or call
513/569-7076.
16.2.5.4 Graphical Method
16.2.5.4.1 This procedure estimates an LC50 (or EC50)
by linearly interpolating between points of a plot of ob-
served percentage mortality versus the base 10 logarithm
(Iog10) of treatment concentration. The only requirement
for its use is that treatment mortalities bracket 50%.
16.2.5.4.2 For an analysis using the Graphical Method,
the data should first be smoothed and adjusted for mortal-
ity in the control replicates. The procedure for smoothing
and adjusting the data is detailed in the following steps:
Let p0, pr ..., pk denote the observed proportion mortali-
ties for the control and the k treatments. The first step is
to smooth the p: if they do not satisfy p0 - p: - ... - pk. The
smoothing process replaces any adjacent p^'s that do not
conform to p0- p.,- ... - pk with their average. For example,
if p: is less than pM, then
P'-i=P'=(Pl+Pl-i)/2
where p* = the smoothed observed proportion
mortality for concentration ;'.
Adjust the smoothed observed proportion mortality in
each treatment for mortality in the control group using
Abbott's formula (Finney, 1971). The adjustment takes
the form:
where p* =
the smoothed observed proportion
mortality for the control
the smoothed observed proportion
mortality for concentration ;'.
76.2.5.5 The Probit Method
16.2.5.5.1 This method is a parametric statistical proce-
dure for estimating the LC50 (or EC50) and the associated
95% confidence interval (Finney, 1971). The analysis
consists of transforming the observed proportion mortali-
ties with a Probit transformation, and transforming the
treatment concentrations to Iog10. Given the assumption
of normality forthe Iog10 of the tolerances, the relationship
between the transformed variables mentioned above is
about linear. This relationship allows estimation of linear
regression parameters, using an iterative approach. A
Probit is the same as a z-score: for example, the Probit
corresponding to 70% mortality is z70 or = 0.52. The LC50
is calculated from the regression and is the concentration
associated with 50% mortality or z = 0. To obtain a
reasonably precise estimate of the LC50 with the Probit
Method, the observed proportion mortalities must bracket
0.5 and the Iog10 of the tolerance should be normally
distributed. To calculate the LC50 estimate and associ-
ated 95% confidence interval, two or more of the ob-
served proportion mortalities must be between zero and
one. The original percentage of mortalities should be
corrected for control mortality using Abbott's formula
(Section 1 6.2.5.4.1 ; Finney, 1 971 ) before the Probit trans-
formation is applied to the data.
16.2.5.5.2 A goodness-of-fit procedure with the Chi-square
statistic is used to determine whether the data fit the
Probit model. If many data sets are to be compared to
one another, the Probit Method is not recommended,
because it may not be appropriate for many of the data
sets. This method also is only appropriate for percent
mortality data sets and should not be used for estimating
endpoints that are a function of the control response,
such as inhibition of growth or reproduction. Most com-
puter programs that generate Probit estimates also gener-
ate confidence interval estimates for the LC50. These
confidence interval estimates on the LC50 might not be
correct if replicate mortalities are pooled to obtain a mean
treatment response (USEPA-USACE, 1998a). This can
be avoided by entering the Probit-transformed replicate
responses and doing a least-squares regression on the
transformed data.
16.2.5.6 The Trimmed Spearman-Karber Method
16.2.5.6.1 The trimmed Spearman-Karber Method is a
modification of the Spearman-Karber, nonparametric sta-
tistical procedure for estimating the LC50 and the associ-
ated 95% confidence interval (Hamilton etal., 1977). This
procedure estimates the trimmed mean of the distribution
of the Iog10 of the tolerance. If the log tolerance distribu-
tion is symmetric, this estimate of the trimmed mean is
equivalent to an estimate of the median of the log toler-
ance distribution. Use of the trimmed Spearman-Karber
Method is only appropriate for lethality data sets when the
requirements forthe Probit Method are not met (USEPA,
1994c;USEPA, 1994d).
16.2.5.6.2 To calculate the LC50 estimate with the
trimmed Spearman-Karber Method, the smoothed, ad-
justed, observed proportion mortalities must bracket 0.5.
To calculate a confidence interval forthe LC50 estimate,
one or more of the smoothed, adjusted, observed propor-
tion mortalities must be between zero and one.
16.2.5.6.3 Smooth the observed proportion mortalities as
described for the Probit Method. Adjust the smoothed
observed proportion mortality in each concentration for
mortality in the control group using Abbott's formula (see
Probit Method, Section 16.2.5.5). Calculate the amount of
trim to use in the estimation of the LC50 as follows:
where
Trim =
the smoothed, adjusted proportion mor-
tality forthe lowest treatment concentra-
tion, exclusive of the control.
the smoothed, adjusted proportion mor-
tality for the highest treatment concen-
tration.
109
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k = the number of treatment concentrations,
exclusive of the control.
16.2.5.7 Linear Interpolation Method
16.2.5.7.1 The Linear Interpolation Method calculates a
toxicant concentration that causes a given percent reduc-
tion (e.g., 25%, 50%, etc.) in the endpoint of interest and
is reported as an ICp value (1C = Inhibition Concentration;
where p = the percent effect). The procedure was de-
signed forgeneral applicability in the analysis of data from
chronic toxicity tests and for the generation of an endpoint
from a continuous model that allows a traditional quantita-
tive assessment of the precision of the endpoint, such as
confidence limits for the endpoint of a single test or a
mean and coefficient of variation for the endpoints of
multiple tests.
16.2.5.7.2 As described in USEPA (1994c; 1994d), the
Linear Interpolation Method of calculating an ICp as-
sumes that the responses (1) are monotonically
nonincreasing, where the mean response for each higher
concentration is less than or equal to the mean response
for the previous concentration, (2) follow a piecewise
linear response function, and (3) are from a random,
independent, and representative sample of test data. If
the data are not monotonically nonincreasing, they are
adjusted by smoothing (averaging). In cases where the
responses at the low toxicant concentrations are much
higher than in the controls, the smoothing process may
result in a large upward adjustment in the control mean. In
the Linear Interpolation Method, the smoothed response
means are used to obtain the ICp estimate reported for
the test. No assumption is made about the distribution of
the data except that the data within a group being resampled
are independent and identically distributed.
16.2.5.7.3 The Linear Interpolation Method assumes a
linear response from one concentration to the next. Thus,
the 1C is estimated by linear interpolation between two
concentrations whose responses bracket the response of
interest, the (p) percent reduction from the control.
16.2.5.7.4 If the assumption of monotonicity of test
results is met, the observed response means (Y^ should
stay the same or decrease as the toxicant concentration
increases. If the means do not decrease monotonically,
the responses are "smoothed" by averaging (pooling)
adjacent means. Observed means at each concentration
are considered in order of increasing concentration, start-
ing with the control mean (Y.,). If the mean observed
response at the lowest toxicant concentration (Y2) is
equal to or smaller than the control mean (Y1), it is used
as the response. If it is larger than the control mean, it is
averaged with the control, and this average is used for
both the control response (M.,) and the lowest toxicant
concentration response (M2). This mean is then compared
to the mean observedjesponse for the next higher toxi-
cant concentration (Y~3). Again, if the mean observed
response for the next higher toxicant concentration is
smaller than the mean of the control and the lowest
toxicant concentration, it is used as the response. If it is
higher than the mean of the first two, it is averaged with
the mean of the first two, and the resulting mean is used
as the response for the control and two lowest concentra-
tions of toxicant. This process is continued for data from
the remaining toxicant concentrations. Unusual patterns
in the deviations from monotonicity may require an addi-
tional step_pf smoothing. Where Y; decrease monotoni-
cally, the Y~ become M: without smoothing.
16.2.5.7.5 To obtain the ICp estimate, determine the
concentrations C^ and CJ+1 that bracket the response M1
(1 - p/100), where M1 is the smoothed control mean
response and p is the percent reduction in response
relative to the control response. These calculations can
easily be done by hand or with a computer program as
described below. The linear interpolation estimate is cal-
culated as follows:
where C,
M,
= tested concentration whose observed
mean response is greater than
1^(1-p/100).
= tested concentration whose observed
mean response is less than
1^(1-p/100).
= smoothed mean response for the
control.
= smoothed mean response for
concentration J.
MJ + 1 = smoothed mean response
concentration J + 1.
for
p = percent reduction in response relative
to the control response.
ICp = estimated concentration at which there
is a percent reduction from the
smoothed mean control response.
16.2.5.7.6 Standard statistical methods for calculating
confidence intervals are not applicable for the ICp. The
bootstrap method, as proposed by Efron (1982), is used
to obtain the 95% confidence interval forthe true mean. In
the bootstrap method, the test data Y is randomly
resampled with replacement to produce a new set of data
Y * that is statistically equivalent to the original data, but
which produces a new and slightly different estimate of
the ICp (ICp*). This process is repeated at least 80 times
(Marcus and Holtzman, 1988), resulting in multiple "data"
sets, each with an associated ICp* estimate. The distribu-
tion of the ICp* estimates derived from the sets of
resampled data approximates the sampling distribution of
the ICp estimate. The standard error of the ICp is esti-
mated by the standard deviation of the individual ICp*
estimates. Empirical confidence intervals are derived from
the quantiles of the ICp* empirical distribution. For ex-
110
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ample, if the test data are resampled a minimum of 80
times, the empirical 2.5% and the 97.5% confidence
limits are about the second smallest and second largest
ICp* estimates (Marcus and Holtzman, 1988). The width
of the confidence intervals calculated by the bootstrap
method is related to the variability of the data. When
confidence intervals are wide, the reliability of the 1C
estimate is in question. However, narrow intervals do not
necessarily indicate that the estimate is highly reliable,
because of undetected violations of assumptions and the
fact that the confidence limits based on the empirical
quantiles of a bootstrap distribution of 80 samples may be
unstable.
16.2.6 Analysis of Bioaccumulation Data
16.2.6.1 In some cases, body burdens will not approach
steady-state body burdens in a 28-d test (ASTM, 1999c).
Organic compounds exhibiting these kinetics will prob-
ably have a log Kow >5, be metabolically refractory (e.g.,
highly chlorinated PCBs, dioxins), or have low depuration
rates. Additionally, tissue residues of several heavy met-
als may gradually increase over time so that 28 d is
inadequate to approach steady-state. Depending on the
goals of the study and the adaptability of the test species
to long-term testing, it may be necessary to conduct an
exposure longer than 28 d (or a kinetic study) to obtain a
sufficiently accurate estimate of steady-state tissue resi-
dues of these compounds.
76.2.6.2 Biotic Sampling
16.2.6.2.1 In the long-term studies, the exposure should
continue until steady-state body burdens are attained.
ASTM (1999c) recommends a minimum of five sampling
periods (plus t0) when conducting water exposures to
generate bioconcentration factors (BCFs). Sampling in a
geometric progression is also recommended with sam-
pling times reasonably close to S/16, S/8, S/4, S/2, and
S, where S is the time to steady state. This sampling
design assumes a fairly accurate estimate of time to
steady state, which is often not the case with sediment
exposures.
16.2.6.2.2 To document steady state from sediment
exposures, placing a greater number of samples at and
beyond the predicted time to steady state is recom-
mended. With a chemical expected to reach steady state
within 28 to 50 d, samples should be taken at Day 0, 7,
14, 21, 28, 42, 56, and 70. If the time to steady state is
much greaterthan 42 d, then additional sampling periods
at two-week intervals should be added (e.g., Day 84).
Slight deviations from this schedule (e.g., Day 45 ver-
sus Day 42) are not critical, though for comparative
purposes, samples should be taken att28. An estimate of
time to steady state may be obtained from the literature or
estimated from structure-activity relationships, though
these values should be considered the minimum times to
steady state.
16.2.6.2.3 This schedule increases the likelihood of
statistically documenting that steady state has been ob-
tained although it does not document the initial uptake
phase as well. If an accurate estimate of the sediment
uptake rate coefficient (Ks) is required, additional sam-
pling periods are necessary during the initial uptake phase
(e.g., Day 0,2, 4, 7, 10, 14).
76.2.6.3 Abiotic Samples
16.2.6.3.1 The bioavailable fraction of the contaminants
as well as the nutritional quality of the sediment are more
prone to depletion in extended tests than during the 28-d
exposures. To statistically document whether such deple-
tions have occurred, replicate sediment samples should
be collected for physical and chemical analysis from each
sediment type at the beginning and the end of the expo-
sure. Archiving sediment samples from every biological
sampling period also is recommended.
76.2.6.4 Short-term Uptake Tests
16.2.6.4.1 Compounds may attain steady state in the
oligochaete, Lumbriculus variegatus, in less than 28 d
(Kukkonen and Landrum, 1993). However, before a shorter
test is used, it must be ascertained that the analytes of
interest do indeed achieve steady state in L. variegatus in
<28 d. Biotic and abiotic samples should be taken at
Day 0 and 10 following the same procedure used for the
28-d tests. If time-series biotic samples are desired,
sample on Day 0,1,3, 5, 7, and 10.
76.2.6.5 Estimating Steady State
16.2.6.5.1 In tests where steady state cannot be docu-
mented, it may be possible to estimate steady-state
concentrations. Several methods have been published
that can be used to predict steady-state chemical con-
centrations from uptake and depuration kinetics (Spacie
and Hamelink, 1982; Davies and Dobbs, 1984). All of
these methods were derived from fish exposures and
most use a linear uptake, first-order depuration model that
can be modified for uptake of chemicals from sediment.
To avoid confusing uptake from water versus sediment,
Ks, the sediment uptake rate coefficient, is used instead
of K1. The Ks coefficient has also been referred to as the
uptake clearance rate (Landrum et al., 1989). Following
the recommendation of Stehly et al. (1990), the gram
sediment and gram tissue units are retained in the
formulation:
Ct(t) = KsxCs/K2x(1-e-K2xt)
where Ct = chemical concentration in tissue at
time t
Cs = chemical concentration in sediment
Ks = uptake rate coefficient in tissue (g sed
g-1 day1)
K2 = depuration constant (day1)
t = time (days)
111
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As time approaches infinity, the maximum or equilibrium
chemical concentration within the organism (Ctmax) be-
comes
Ctmax= CsxKs/K2
Correspondingly, the bioaccumulation factor (BAF) for a
compound may be estimated from
BAF= Ks/K2
16.2.6.5.2 This model assumes that the sediment con-
centration and the kinetic coefficients are invariant.
Depletion of the sediment concentrations in the vicinity of
the organism would invalidate the model. Further, the rate
coefficients are conditional on the environment and health
of the test organisms. Thus, changes in environmental
conditions such as temperature or changes in physiology
such as reproduction will also invalidate the model. De-
spite these potential limitations, the model can provide
estimates of steady-state tissue residues.
16.2.6.5.3 The kinetic approach requires an estimate of
Ks and K2, which are determined from the changes in
tissue residues during the uptake phase and depuration
phase, respectively. The uptake experiment should be
short enough that an estimate of Ks is made during the
linear portion of the uptake phase to avoid an unrealisti-
cally low uptake rate due to depuration. The depuration
phase should be of sufficient duration to smooth out any
loss from a rapidly depurated compartment such as loss
from the voiding of feces. Unless there is reason to
suspect that the route of exposure will affect the depura-
tion rate, it is acceptable to use a K2 derived from a water
exposure. For further discussion of this method for
bioconcentration studies in fish, see Davies and Dobbs
(1984), Spacieand Hamelink(1982), and ASTM (1999b).
For application of this procedure for sediment, see ASTM
(1999c). Recent studies of the accumulation of
sediment-associated chemicals by benthos suggest that
the kinetics for freshly dosed sediments may require a
more complex formulation to estimate the uptake clear-
ance constant than that presented above (Landrum, 1989).
16.2.6.5.4 This model predicts that equilibrium would be
reached only as time becomes infinite. Therefore, for
practical reasons, apparent steady state is defined here
as 95% of the equilibrium tissue residue. The time to
reach steady state can be estimated by
S = ln[1 / (1.00-0.95)]/K2 = 3.0/K2
where S = time to apparent steady state (days)
Thus, the key information is the depuration rate of the
compound of interest in the test species or phylogeneti-
cally related species. Unfortunately, little of this data has
been generated for benthic invertebrates. When no depu-
ration rates are available, the depuration rate constant for
organic compounds can then be estimated from the rela-
tionship between Kow and K2 for fish species (Spacie and
Hamelink, 1982):
K2 = antilog[1.47-0.414 xlog(Kow)]
The relationship between S and K2 and between K2 and
Kow is summarized in Table 16.2. Estimated time (days)
to reach 95% of chemical steady-state tissue residue (S)
and depuration rate constants (K2) are calculated from
octanol-water partition coefficients using a linear uptake,
first-order depuration model (Spacie and Hamelink, 1982).
The K2 values are the amount depurated (decimal fraction
of tissue residue lost per day). Table 16.2 may be used to
make a rough estimate of the exposure time to reach
steady-state tissue residues if a depuration rate constant
for the compound of interest from a phylogenetically
similar species is available. If no depuration rate is avail-
able, then the table may be used for estimating the S of
organic compounds from the Kow value. However, as
these data were developed from fish bioconcentration
data, its applicability to the kinetics of uptake from
sediment-associated chemicals is unknown. The portion
of organics readily available for uptake may be small in
comparison to the total sediment organic concentration
(Landrum, 1989). Therefore S values generated by this
model should be considered as minimum time periods.
16.2.6.5.5 Using a linear uptake, first-order depuration
model to estimate exposure time to reach steady-state
body burden for metals is problematical for a number of
reasons. The kinetics of uptake may be dependent upon a
small fraction of the total sediment metal load that is
bioavailable (Luoma and Bryan, 1982). Depuration rates
may be more difficult to determine, as metals bound to
proteins may have very low exchange rates (Bryan, 1976).
High exposure concentrations of some metals can lead to
the induction of metal binding proteins, like metallothionein,
which detoxify metals. These metal-protein complexes
within the organism have extremely low exchange rates
with the environment (Bryan, 1976). Thus, the induction of
metal binding proteins may result in decreased depuration
rate constants in organisms exposed to the most polluted
sediments. Additionally, structure-activity relationships
that exist for organic chemicals (e.g., relationship be-
tween Kow and BCFs) are not well developed for metals.
Table 16.2
Log Kow
Estimated Time to Obtain 95 Percent of Steady-
state Tissue Residue
K2
S (days)
1
2
3
4
5
6
7
8
9
0.114
0.44
0.17
0.0065
0.0025
0.00097
0.00037
0.00014
0.00006
0.2
0.5
1.4
3.5
9.2
24
61
160
410
112
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16.3 Data Interpretation
16.3.1 Sediments spiked with known concentrations of
chemicals can be used to establish cause and effect
relationships between chemicals and biological responses.
Results of toxicity tests with test materials spiked into
sediments at different concentrations may be reported in
terms of an LC50 (median lethal concentration), an EC50
(median effect concentration), an IC50 (inhibition concen-
tration), or as a NOEC (no observed effect concentration)
or LOEC (lowest observed effect concentration; Section
3). Consistent spiking procedures should be followed in
orderto make interlaboratory comparisons (Section 8.3).
16.3.2 Evaluating effect concentrations for chemicals in
sediment requires knowledge of factors controlling the
bioavailability. Similar concentrations of a chemical in
units of mass of chemical per mass of sediment dry
weight often exhibit a range in toxicity in different sedi-
ments (Di Toro et al., 1991; USEPA, 1992c). Effect
concentrations of chemicals in sediment have been corre-
lated to interstitial water concentrations, and effect con-
centrations in interstitial water are often similar to effect
concentrations in water-only exposures. The bioavailabil-
ity of nonionic organic compounds are often inversely
correlated with the organic carbon concentration of the
sediment. Whatever the route of exposure, the correla-
tions of effect concentrations to interstitial water concen-
trations indicate that predicted or measured concentra-
tions in interstitial water can be useful for quantifying the
exposure concentration to an organism. Therefore, infor-
mation on partitioning of chemicals between solid and
liquid phases of sediment can be useful for establishing
effect concentrations.
16.3.3 Toxic units can be used to help interpret the
response of organisms to multiple chemicals in sediment.
A toxic unit is the concentration of a chemical divided by
an effect concentration. For example, a toxic unit of
exposure can be calculated by dividing the measured
concentration of a chemical in pore water by the water-only
LC50 forthe same chemical (Ankley et al., 1991 a). Toxic-
ity expressed as toxic units may be summed and this
may provide information on the toxicity of chemical mix-
tures (Ankley etal., 1991 a).
16.3.4 Field surveys can be designed to provide either a
qualitative reconnaissance of the distribution of sediment
contamination or a quantitative statistical comparison of
contamination among sites (Burton and Ingersoll, 1994).
Surveys of sediment toxicity are usually part of more
comprehensive analyses of biological, chemical, geologi-
cal, and hydrographicdata. Statistical correlation can be
improved and costs reduced if subsamples are taken
simultaneously for sediment toxicity or bioaccumulation
tests, chemical analyses, and benthic community
structure.
16.3.5 Descriptive methods, such as toxicity tests with
field-collected sediment, should not be used alone to
evaluate sediment contamination. An integration of sev-
eral methods using the weight of evidence is needed to
assess the effects of contaminants associated with sedi-
ment (Long and Morgan, 1990; Ingersoll et al., 1996,
1997; MacDonald etal., 1996). Hazard evaluations inte-
grating data from laboratory exposures, chemical analy-
ses, and benthic community assessments provide strong
complementary evidence of the degree of pollution-induced
degradation in aquatic communities (Chapman et al.,
1992, 1997; Burton, 1991; Canfield et al., 1994, 1996,
1998).
16.3.6 Toxicity Identification Evaluation (TIE) procedures
can be used to help provide insights as to specific con-
taminants responsible for toxicity in sediment (USEPA,
1991b; Ankley and Thomas, 1992). For example, the
toxicity of contaminants such as metals, ammonia, hy-
drogen sulfide, and nonionic organic compounds can be
identified using TIE procedures.
16.3.7 Interpretation of Comparisons of Tissue
Residues
16.3.7.1 If the mean control tissue residues at Day 28 are
not significantly greaterthan the Day 0 tissue residues, it
can be concluded that there is no significant contamina-
tion from the exposure system or from the control sedi-
ment. If there is significant uptake, the exposure system
or control sediment should be reevaluated as to suitabil-
ity. Even if there is a significant uptake in the controls, it
is still possible to compare the controls and treatments as
long as the contaminant concentrations in the test tissue
residues are substantially higher. However, if control val-
ues are high, the data should be discarded and the
experiment conducted again after determining the source
of contamination.
16.3.7.2 Comparisons of the 28-d control (or reference)
tissue residues and 28-d treatment tissue residues deter-
mines whether there was statistically significant bioaccu-
mulation due to exposure to test sediments. Comparisons
between control and reference tissue residues at Day 28
determine whether there was a statistically significant
bioaccumulation due to exposure to the reference sedi-
ment. If no significant difference is detected when treatment
tissue residues are compared to a set criterion value
(e.g., FDA action level) with a one-tailed test, the residues
must be considered equivalent to the value even though
numerically the mean treatment tissue residue may be
smaller.
76.3.7.3 BAFsandBSAFs
16.3.7.3.1 Statistical comparisons between ratios such
as BAFs or BSAFs are difficult due to computation of
error terms. Since all variables used to compute BAFs
and BSAFs have errors associated with them, it is neces-
sary to estimate the variance as a function of these
errors. This can be accomplished using approximation
techniques such as the propagation of error (Beers, 1957)
or a Taylor series expansion method (Mood et al., 1974).
BAFs and BSAFs can then be compared using these
estimates of the variance. ASTM (1999c) provides ex-
amples of this approach.
113
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16.3.7.4 Comparing Tissue Residues of Different
Compounds
16.3.7.4.1 In some cases, it is of interest to compare the
tissue residues of different compounds. For example,
Rubinstein et al. (1987) compared the uptake of thirteen
different PCB congeners to test for differences in bioavail-
ability. Because the values for the different compounds
are derived from the same tissue samples, they are not
independent and tend to be correlated, so standard ttests
and ANOVAs are inappropriate. A repeated measures
technique (repeated testing of the same experimental
unit) should be used where the experimental unit (individual)
is considered as a random factor and the different com-
pounds as a second factor. See Rubinstein et al. (1987)
and Lake et al. (1990) for an example of the application of
repeated measures to bioaccumulation data.
16.4 Reporting
16.4.1 The record of the results of an acceptable sedi-
ment test should include the following information either
directly or by referencing available documents:
16.4.1.1 Name of test and investigator(s), name and
location of laboratory, and dates of start and end of test.
16.4.1.2 Source of control or test sediment, and method
for collection, handling, shipping, storage and disposal of
sediment.
16.4.1.3 Source of test material, lot number if applicable,
composition (identities and concentrations of major
ingredients and impurities if known), known chemical and
physical properties, and the identity and concentration^)
of any solvent used.
16.4.1.4 Source and characteristics of overlying water,
description of any pretreatment, and results of any dem-
onstration of the ability of an organism to survive or grow
in the water.
16.4.1.5 Source, history, and age of test organisms;
source, history, and age of brood stock, culture procedures;
and source and date of collection of the test organisms,
scientific name, name of person who identified the organ-
isms and the taxonomic key used, age or life stage,
means and ranges of weight or length, observed diseases
or unusual appearance, treatments used, and holding
procedures.
16.4.1.6 Source and composition of food; concentrations
of test material and other contaminants; procedure used
to prepare food; and feeding methods, frequency and
ration.
16.4.1.7 Description of the experimental design and test
chambers, the depth and volume of sediment and overly-
ing water in the chambers, lighting, number of test cham-
bers and number of test organisms/treatment, date and
time test starts and ends, temperature measurements,
dissolved oxygen concentration (ug/L) and any aeration
used before starting a test and during the conduct of a
test.
16.4.1.8 Methods used for physical and chemical charac-
terization of sediment.
16.4.1.9 Definition(s) of the effects used to calculate
LC50 or ECSOs, biological endpoints for tests, and a
summary of general observations of other effects.
16.4.1.10 A table of the biological data for each test
chamber for each treatment, including the controls), in
sufficient detail to allow independent statistical analysis.
16.4.1.11 Methods used for statistical analyses of data.
16.4.1.12 Summary of general observations on other
effects or symptoms.
16.4.1.13 Anything unusual about the test, any deviation
from these procedures, and any other relevant information.
16.4.2 Published reports should contain enough informa-
tion to clearly identify the methodology used and the
quality of the results.
114
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Section 17
Precision and Accuracy
17.1 Determining Precision and Accuracy
17.1.1 Precision is a term that describes the degree to
which data generated from replicate measurements differ
and reflects the closeness of agreement between ran-
domly selected test results. Accuracy is the difference
between the value of the measured data and the true
value and is the closeness of agreement between an
observed value and an accepted reference value. Quanti-
tative determination of precision and accuracy in sedi-
ment testing of aquatic organisms is difficult or may be
impossible in some cases, as compared to analytical
(chemical) determinations. This is due, in part, to the
many unknown variables that affect organism response.
Determining the accuracy of a sediment test using field
samples is not possible since the true values are not
known. Since there is no acceptable reference material
suitable for determining the accuracy of sediment tests,
the accuracy of the test methods has not been deter-
mined (Section 17.2).
17.1.2 Sediment tests exhibit variability due to several
factors (Section 9). Test variability can be described in
terms of two types of precision, either single laboratory
(intralaboratory or repeatability; Section 17.5.1) precision
or multi-laboratory (interlaboratory or reproducibility; Sec-
tion 17.5.2, 17.5.3 and 17.6) precision. Intralaboratory
precision reflects the ability of trained laboratory person-
nel to obtain consistent results repeatedly when perform-
ing the same test on the same organism using the same
toxicant. Interlaboratory precision (also referred to as
round-robin or ring tests) is a measure of the reproducibil-
ity of a method when tests are conducted by a number of
laboratories using that method and the same organism
and samples. Generally, intralaboratory results are less
variable than interlaboratory results (USEPA, 1991 a;
USEPA, 1991c; USEPA, 1994b; USEPA, 1994c; Hall et
al., 1989; Grothe and Kimerle, 1985).
17.1.3 A measure of precision can be calculated using
the mean and relative standard deviation (percent coeffi-
cient of variation, or CV% = standard deviation/mean x
100) of the calculated endpoints from the replicated end-
points of a test. However, precision reported as the CV
should not be the only approach used for evaluating
precision of tests and should not be used for the NOEC
levels derived from statistical analyses of hypothesis
testing. The CVs can be very high when testing extremely
toxic samples. For example, if there are multiple replicates
with no survival and one with low survival, the CV might
exceed 100%, yet the range of response is actually quite
consistent. Therefore, additional estimates of precision
should be used, such as range of responses, and mini-
mum detectable differences (MOD) compared to control
survival or growth. Several factors can affect the preci-
sion of the test, including test organism age, condition
and sensitivity; handling and feeding of the test organ-
isms; overlying water quality; and the experience of the
investigators in conducting tests. For these reasons, it is
recommended that trained laboratory personnel conduct
the tests in accordance with the procedures outlined in
Section 9. Quality assurance practices should include
the following: (1) single laboratory precision determina-
tions that are used to evaluate the ability of the laboratory
personnel to obtain precise results using reference toxi-
cants for each of the test organisms and (2) preparation of
control charts (Section 17.4) for each reference toxicant
and test organism. The single laboratory precision determi-
nations should be made before conducting a sediment test
and should be periodically performed as long as whole-
sediment tests are being conducted at the laboratory.
MA A Intralaboratory precision data are routinely calcu-
lated for test organisms using water-only 96-h exposures
to a reference toxicant, such as potassium chloride (KCI).
Intralaboratory precision data should be tracked using a
control chart. Each laboratory's reference-toxicity data
will reflect conditions unique to that facility, including
dilution water, culturing, and other variables (Section 9).
However, each laboratory's reference-toxicity CVs should
reflect good repeatability.
17.1.5 Interlaboratory precision (round-robin) tests have
been completed with both Hyalella azteca and Chirono-
mus tentans using 4-d water-only tests and 10-d whole-
sediment tests described in Section 11.2 and 12.2
(Section 17.5). Section 17.6 describes results of round-
robin evaluations with long-term sediment toxicity tests
described in Sections 14 and 15 for H. azteca and
C. tentans.
17.2 Accuracy
17.2.1 The relative accuracy of toxicity tests cannot be
determined since there is no acceptable reference mate-
rial. The relative accuracy of the reference-toxicity tests
can only be evaluated by comparing test responses to
control charts.
115
-------
17.3 Replication and Test Sensitivity
17.3.1 The sensitivity of sediment tests will depend in
part on the number of replicates per concentration, the
probability levels (alpha and beta) selected, and the type
of statistical analysis. For a specific level of variability,
the sensitivity of the test will increase as the number of
replicates is increased. The minimum recommended num-
ber of replicates varies with the objectives of the test and
the statistical method used for analysis of the data
(Section 16).
17.4 Demonstrating Acceptable
Laboratory Performance
17.4.1 Intralaboratory precision, expressed as a coeffi-
cient of variation (CV), can be determined by performing
five or more tests with different batches of test organ-
isms, using the same reference toxicant, at the same
concentrations, with the same test conditions (e.g., the
same test duration, type of water, age of test organisms,
feeding), and same data analysis methods. A reference-
toxicity concentration series (dilution factor of 0.5 or
higher) should be selected that will provide partial mortali-
ties at two or more concentrations of the test chemical
(Section 9.14, Table 9.1, 9.2). See Section 9.16 for
additional detail on reference-toxicity testing.
17.4.2 It is desirable to determine the sensitivity of test
organisms obtained from an outside source. The supplier
should provide data with the shipment describing the
history of the sensitivity of organisms from the same
source culture.
17.4.3 Before conducting tests with potentially contami-
nated sediment, it is strongly recommended that the
laboratory conduct the tests with control sediment(s)
alone. Results of these preliminary studies should be
used to determine if use of the control sediment and other
test conditions (i.e., water quality) result in acceptable
performance in the tests as outlined in Tables 11.1,12.1,
13.1,14.1,and15.1.
17.4.4 A control chart should be prepared for each
combination of reference toxicant and test organism.
Each control chart should include the most current data.
Endpoints from five tests are adequate for establishing
the control charts. In this technique, a running plot is
maintained for the values (X:) from successive tests with
a given reference toxicant (Figure 17.1), and the end-
points (LC50, NOEC, ICp) are examined to determine if
they are within prescribed limits. Control charts as de-
scribed in USEPA (1991 a) and USEPA (1993b) are used
to evaluate the cumulative trend of results from a series
of samples. The mean and upper and lower control limits
(±2 SD) are recalculated with each successive test result.
After two years of data collection, or a minimum of 20
data points, the control (cusum) chart should be main-
tained using only the 20 most recent data points.
17.4.5 The outliers, which are values falling outside the
upper and lower control limits, and trends of increasing or
decreasing sensitivity, are readily identified using control
charts. With an alpha of 0.05, one in 20 tests would be
expected to fall outside of the control limits by chance
alone. During a 30-d period, if two reference-toxicity tests
out of a total of the previous 20 fall outside the control
limits, the sediment toxicity tests conducted during the
time in which the second reference-toxicity test failed are
suspect and should be considered as provisional and
subject to careful review.
17.4.5.1 A sediment test may be acceptable if specified
conditions of a reference-toxicity test fall outside the
expected ranges (Section 9). Specifically, a sediment
test should not necessarily be judged unacceptable if the
LC50 for a given reference-toxicity test falls outside the
expected range or if mortality in the control of the reference-
toxicity test exceeds 10% (Tables 9.1 and 9.2). All the
performance criteria outlined in Tables 11.3, 12.3, 13.4,
UPPER CONTROL LIMIT
O
LU
O
CENTRAL TENDENCY
LOWER CONTROL LIMIT
J
10
15
20
O
o
O"
O"
UPPER CONTROL LIMIT (X + 2 S)
CENTRAL TENDENCY
B
LOWER CONTROL LIMIT (X - 2 S)
I
I ^
0 5 10 15 20
TOXICITY TEST WITH REFERENCE TOXICANTS
where
X1
n
x
S
Figure 17.1
n-l
= Successive toxicity values of toxicity tests.
= Number of tests.
= Mean toxicity value.
= Standard deviation.
Control (cusum) charts: (A) hypothesis testing
results; and (B) point estimates (LC, EC, or 1C).
116
-------
14.3, and 15.3 must be considered when determining the
acceptability of a sediment test. The acceptability of the
sediment test would depend on the experience and judg-
ment of the investigator and the regulatory authority.
17.4.6 If the value from a given test with the reference
toxicant falls more than two standard deviations (SD)
outside the expected range, the sensitivity of the organ-
isms and the overall credibility of the test system may be
suspect (USEPA, 1991 a). In this case, the test procedure
should be examined for defects and should be repeated
with a different batch of test organisms.
17.4.7 Performance should improve with experience, and
the control limits for point estimates should gradually
narrow. However, control limits of ±2 SD, by definition,
will be exceeded 5% of the time, regardless of how well a
laboratory performs. Highly proficient laboratories that
develop a very narrow control limit may be unfairly penal-
ized if a test that falls just outside the control limits is
rejected cte facto. Forthis reason, the width of the control
limits should be considered in determining whether or not
an outlier is to be rejected. This determination may be
made by the regulatory authority evaluating the data.
17.4.8 The recommended reference-toxicity test con-
sists of a control and five or more concentrations in which
the endpoint is an estimate of the toxicant concentration
that is lethal to 50% of the test organisms in the time
period prescribed by the test. The LC50 is determined by
an appropriate procedure, such as the trimmed
Spearman-Karber Method, Probit Method, Graphical
Method, orthe Linear Interpolation Method (Section 16).
17.4.9 The point estimation analysis methods recom-
mended in this manual have been chosen primarily be-
cause they are well-tested, well-documented, and are
applicable to most types of test data. Many other meth-
ods were considered in the selection process, and it is
recognized that the methods selected are not the only
possible methods of analysis of toxicity data.
17.5 Precision of Sediment Toxicity Test
Methods: Evaluation of 10-d
Sediment Tests and Reference-
toxicity Tests
17.5.1 Intralaboratory Performance
17.5.1.1 Intralaboratory performance of the Hyalella azteca
and Chironomustentans 10-d tests (as described in Tables
11.1 and 12.1) was evaluated at the USEPA Office of
Research and Development Laboratory (Duluth, MN) us-
ing one control sediment sample in June 1993. In this
study, five individuals simultaneously conducted the 10-d
whole-sediment toxicity tests as described in Tables 11.1
and 12.1 with the exception of the feeding rate of 1.0 mL
ratherthan 1.5 mL for C. tentans. The results of the study
are presented in Table 17.1. The mean survival for
H. azteca was 90.4% with a CV of 7.2% and the mean
survival for C. tentans was 93.0% with a CV of 5.7%. All
of the individuals met the survival performance criteria of
80% for H. azteca (Table 11.3) or 70% for C. tentans
(Table 12.3).
17.5.2 Interlaboratory Precision: 1993
Evaluation of the 10-d Sediment Tests
and the Reference-toxicity Tests
17.5.2.1 Interlaboratory precision using reference-toxicity
tests or 10-d whole-sediment toxicity tests using the
methods described in this manual (Tables 9.1, 9.2,11.1,
and 12.1) were conducted by federal government labora-
tories, contract laboratories, and academic laboratories
that had demonstrated experience in sediment toxicity
testing for a first time in 1993 (Section 17.5.2.2 and
Burton et al., 1996b) and a second time in 1996/1997 (the
"1996/1997 study"; Section 17.5.3). In the 1993 study the
only exception to the methods outlined in Table 9.1 and
9.2 was that 80% ratherthan the current recommendation
of 90% survival was used to judge the acceptability of the
reference-toxicity tests. The 1993 round-robin study
was conducted in two phases for each test organism.
The experimental design for the 1993 round-robin study
required each laboratory to conduct 96-h water-only
reference-toxicity tests in Phase 1 and 10-d whole-
sediment tests in Phase 2 with Hyalella azteca or
Chironomus tentans over a period of six months. Crite-
ria for selection of participants in the 1993 round-robin
study were that the laboratories: (1) had existing cultures
of the test organisms, (2) had experience conducting
tests with the organisms, and (3) would participate volun-
tarily. The test methods for the reference-toxicity tests
and the whole-sediment toxicity tests were similar among
laboratories. Standard operating procedures detailing the
test methods were provided to all participants. Culture
methods were not specified and were not identical across
laboratories.
Table 17.1
Intralaboratory Precision for Survival of Hyalella
azteca and Chironomus tentans in 10-d Whole-
sediment Toxicity Tests, June 19931
Percent Survival
Individual
H. azteca
C. tentans
A
B
C
D
E
N
Mean
CV
85
93
90
84
100
5
90.4
7.2%
85
93
93
94
100
5
93.0
5.7%
Test sample was from a control sediment (T.J. Norberg-King,
USEPA, Duluth, MN, personal communication). The test was
conducted at the same time by five individuals at the USEPA Office
of Research and Development Laboratory (Duluth, MN). The source
of overlying water was from Lake Superior.
117
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Table 17.2 Participants in 1993 Round-robin Studies1
Chironomus tentans
Hyalella azteca
Laboratory
A
B
C
D
E
F
G
H
I
J
K
L
N
96-h
KCI
Test
Dec 92
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
3
4
10
96-h
KCI
Test
May 93
N
Y
N
Y
Y
Y
Y
N
Y
Y
3
4
7
10-d
Sediment
Test
May 93
N
Y
Y
Y
Y
Y
Y
N
Y
Y
3
4
8
96-h
KCI
Test
Oct 92
Y
Y
Y
N
Y
Y
Y
Y
-2
Y
Y
Y
10
10-d
Sediment
Test
Mar 93
N
Y
Y
N
Y
Y
Y
N
Y
Y
Y
Y
9
1 Y = Laboratory participated in testing sediment samples.
2 Test in January 1993.
3 Participated using C. riparius only.
4 Did not intend to participate with C. tentans.
17.5.2.2 In the second series of round-robin tests con-
ducted in 1996/1997, 10-d and long-term toxicity testing
methods were evaluated with Hyalella azteca and
Chironomus tentans. Results from these interlaboratory
comparisons conducted in 1996/1997 are presented in
detail in Sections 17.5.3 and 17.6. The second series of
interlaboratory comparisons conducted in 1996/1997 did
not restrict testing to laboratories with experience. As in
1993, the participants in the 1996/1997 round-robin study
included government, contract, and academic laborato-
ries. In the 1996/1997 study, no water-only reference-
toxicity tests were conducted.
17.5.2.3 Ten laboratories participated in the H. azteca
reference-toxicity test in the 1993 study (Table 17.2). The
results from the tests with KCI are summarized in Table
17.3. The test performance criteria of >80% control sur-
vival was met by 90% of the laboratories resulting in a
mean control survival of 98.8% (CV = 2.1%). The mean
LC50 was 305 mg/L (CV = 14.2%) and the LC50s ranged
from 232 to 372 mg/L KCI.
17.5.2.4 In the 10-d whole-sediment tests with H. azteca,
nine laboratories tested the three sediments described
above and five laboratories tested a fourth sediment from
a heavily contaminated site in the 1993 study (Table
17.4). All laboratories completed the tests; however, Labo-
ratory C had 75% survival, which was below the accept-
Table 17.3 Interlaboratory Precision for Hyalella azteca 96-h
LCSOs from Water-only Static Acute Toxicity
Tests Using a Reference Toxicant (KCI)
(October 1992)
KCI Percent
LC50 Confidence Intervals Control
Laboratory (mg/L) Lower Upper Survival
A
B
C
D
E
F
G
H
I
J
L
N
Mean 1
CV 1
N
Mean 2
CV2
372
321
232
1
325
276
297
336
1422
337
250
10
289.03
23.0%3
9
305.04
14.2%4
352
294
205
1
282
240
267
317
101
286
222
395
350
262
1
374
316
331
356
200
398
282
100
98
100
1
100
98
73
100
93
100
100
10
96.2%
8.3%
9
98.8
2.1%
1 Laboratory did not participate in H. azteca test in October.
2 Results are from a retest in January using three concentrations only;
results excluded from analysis.
3 Mean 1 and CV 1 include all data points
4 Mean 2 and CV 2 exclude data points for all sediment samples from
laboratories that did not meet minimum control survival of >80%.
able test criteria for survival (Table 1 1 .3). Forthese tests,
the CV was calculated using the mean percent survival
for the eight laboratories that met the performance criteria
for the test. The CV for survival in the control sediment
(RR 3) was 5.8% with a mean survival of 94.5% and
survival ranging from 86% to 100%. For sediments RR 2
and RR 4, the mean survival was 3.3% and 4.3%, respec-
tively (Table 17.4). For RR 2, survival ranged from 0% to
24% (CV = 253%) and for RR 4, the survival ranged from
0% to 11% (CV = 114%). Survival in the moderately
contaminated sediment (RR 1) was 54.2% with survival
ranging from 23% to 76% (CV = 38.9%). When the RR 1
data for each laboratory were compared to the control for
that laboratory, the range for the minimum detectable
difference (MOD) between the test sediments and the
control sediment ranged from 5 to 24% with a mean of
17.5.2.5 The Phase 1 C. tentans reference-toxicity test
was conducted with KCI on two occasions in the 1993
study (Tables 1 7.5 and 1 7.6). Both tests were conducted
in 20 mL of test solution in 30-mL beakers using 10
replicates per treatment with 1 organism per beaker.
Animals were fed 0.25 mLof a 4 g/L solution ofTetrafin®
on Day 0 and Day 2 (Table 9.1). For the first reference-
toxicity test comparison, 10 laboratories participated, and
118
-------
Table 17.4 Interlaboratory Precision for Survival of Hya lei la azteca in 10-d Whole-sediment Toxicity Tests Using Four
Sediments (March 1993)
Laboratory
RR 1
Mean Percent Survival (SD) in Sediment Samples
RR 2
RR 3 (Control)
RR 4
A
B
C
D
E
F
G
H
I
J
K
L
N
Mean 13
CV1
N
Mean 24
CV2
1
76.2
57.S22
1
46.2
72.5
50.0
1
73.7
65.0
22.5
27.5
9
54.6
36.2%
8
54.2
38.9%
(20.7)
(14.9)
(17.7)
(12.8)
(28.3)
(32.0)
(9.3)
(18.3)
(16.7)
1
2.5
1.22
1
0
23.7
0
1
0
0
0
0
9
3.0
256%
8
3.3
253%
(7.1)
(0)
(0)
(18.5)
(0)
(0)
(0)
(0)
(0)
1
97.5
75.02
1
97.5
98.7
100
1
86.2
96.2
95.0
86.2
9
93.0
9.0%
8
94.5
5.8%
(4.6)
(17.7)
(7.1)
(3.5)
(0)
(10.6)
(5.2)
(5.3)
(18.5)
1
11.2
1.22
1
—
0
3.3
1
—
2.5
—
—
5
3.6
121%
4
4.3
114%
(13.6)
(0)
(0)
(5.2)
(7.1)
1 Laboratory did not participate in H. azteca test in March.
2 Survival in control sediment (RR 3) below minimum acceptable level.
3 Mean 1 and CV 1 include all data points.
4 Mean 2 and CV 2 exclude data points for all sediment samples from laboratories that did not meet minimum control survival of ^80%.
eight laboratories met the survival criteria of the round
robin, which was >80% survival (Table 17.5). The mean
LC50 for the eight laboratories that met the survival
criterion was 4.25 g/L (CV of 51.8%). The LC50s ranged
from 1.25 to 6.83 g/L. Length and instar were determined
for a subset of organisms at the start of the tests for some
of the laboratories. When length was correlated with the
LC50, the larger animals were less sensitive than the
smaller animals. The effect level was significantly corre-
lated (r2 = 0.78) with the organism size, which ranged from
1.56 mm to 10.87 mm (ages of animals ranged from 7-to
13-d post-deposition). The majority of these animals were
the third instar, with the smallest animals in their first
instar and the largest animals a mix of third and fourth
instar (Table 17.5) as determined by head capsule width.
17.5.2.6 For the second Phase 1 KCI reference-toxicity
tests with C. tentans, seven laboratories participated in
the 1993 study (Table 17.6). The test conditions were
identical to those in the previous reference-toxicity test
except that a minimum size was specified rather than
using initial age of the animals. Each laboratory was
instructed to start the test when larvae were at least 0.4 to
0.6 mm long. Therefore, a more consistent size of test
organisms was used in this test. Six out of the seven
laboratories met the >80% control survival criterion with a
mean LC50 of 5.37 g/L (CV= 19.6%). The LC50s ranged
from 3.61 to 6.65 g/L.
17.5.2.7 Eight laboratories participated in the 10-d whole-
sediment testing with C. tentans. The same three sedi-
ments used in the H. azteca whole-sediment test were
used for this test in the 1993 study (Table 17.7). All test
conditions were those as described in Table 12.1 with the
exception of the feeding rate of 1.0 mL ratherthan 1.5 mL
for C. tentans. Three laboratories did not meet the control
criteria for acceptable tests of >70% survival in the con-
trol (RR 3) sediment (Table 12.3). Forthe five laboratories
that successfully completed the tests, the mean survival
in the control sediment (RR 3) was 92.0% (CV of 8.3%)
and survival ranged from 81.2% to 98.8%. Forthe RR 2
sediment sample, the mean survival among the five
laboratories was 3.0% (CV = 181%) and for the RR 1
sediment sample, the mean survival was 86.8%
(CV = 13.5%). A significant effect on survival was not
evident for the RR 1 sample, but growth was affected
(Table 17.8). When the RR1 data for each laboratory we re
compared to the control for that laboratory, the MOD for
survival among laboratories ranged from 2.3 to 12.1%
with a mean of 8% (SD = 4).
17.5.2.8 For C. tentans, growth in 10-d tests is a sensi-
tive indicator of sediment toxicity (Ankley et al., 1993)
and growth was also measured in the round-robin com-
parison in the 1993 study (Table 17.8). Using the data
from five laboratories with acceptable control survival in
the control sediment (RR 3), the mean weight of C.
tenfansforthe control sediment (RR 3) was 1.254 mg (CV
119
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Table 17.5 Interlaboratory Precision for Chironomus tentans 96-h LCSOs from Water-only Static Acute Toxicity Tests Using a
Reference Toxicant (KCI) (December 1992)
Labora-
tory
A
B
C
D
E
F
G
H
I
J
N
Mean 15
CV1
N
Mean 26
CV2
KCI
LC50
(g/L)
6.19
6.83
5.00
3.17
2.002
1.25
6.28
2.89
6.66
1.77
10
4.20
52.7%
8
4.25
51 .8%
Confidence
Lower
5.37
6.38
4.16
2.29
2
3
5.26
2.39
6.01
0.59
Interval
Upper
7.13
7.31
6.01
4.40
—
—
7.50
3.50
7.24
5.26
Control
Survival
(%)
751
100
100
100
80
80
95
95
100
651
10
89.0
14.5%
8
93.8
9.3%
1 Control survival below minimum acceptable level.
2 Unable to calculate LC50 with trimmed Spearman Karber; no confidence interval
3 Confidence intervals cannot be calculated as no partial mortalities occurred.
4 No animals were measured.
5 Mean 1
6 Mean 2
and CV 1 include all data points.
and CV 2 exclude data points for all samples from
Mean
Length
(mm)
10.87
10.43
5.78
5.86
6.07
1.56
7.84
6.07
4
4.42
8
6.6
46.6%
7
6.2
39.5%
could be
Instar
at
Start
of Test
3,4
3
3
3
3
1
3
3
4
2,3
calculated.
laboratories that did not meet minimum control survival
Age at
Start
of Test
(day)
1
13
11
11
11
12
11
7
10
7
10
10.3
17.9%
8
10.75
15.2%
of>80%.
Table 17.6 Interlaboratory Precision for Chironomus tentans 96-h LCSOs from Water-only Static Acute Toxicity Tests Using a
Reference Toxicant (KCI) (May 1993)
Labora-
tory
A
B
C
D
E
F
G
H
I
J
N
Mean 14
CV1
N
Mean 2 5
CV2
KCI
LC50
(9/L)
1
6.65
1
5.30
5.11
3.61
5.36
1
5.30
6.20
7
5.36
17.9%
6
5.37
19.6%
Lower
—
2
4.33
4.18
2.95
4.43
—
4.33
4.80
Confidence Interval
Upper
—
—
—
6.50
6.24
4.42
6.49
—
6.52
7.89
1 Did not participate in reference-toxicity test in April.
2 Confidence intervals cannot be calculated as no partial mortalities occurred.
3 Control survival below minimum acceptable level.
4 Mean 1 and CV 1 include all data points.
5 Mean 2 and CV 2 exclude data points for all samples from laboratories that did not
Control
Survival
(%)
—
90
—
553
100
90
93
—
95
100
7
89
17.5%
6
94.7
4.8%
meet minimum control survival
Age at
Start
of Test
(day)
—
12
—
10
11
10
12
10-11
13
7
11.1
9.46%
6
11.2
9.13%
of>70%.
120
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Table 17.7 Interlaboratory Precision for Survival of Chironomus tentans in 10-d Whole-sediment Toxicity Tests Using Three
Sediments (May 1993)
Mean Percent Survival (SD) in Sediment Samples
Laboratory
A
B
C
D
E
F
G
H
I
J
N
Mean 13
CV1
N
Mean 24
CV2
1 Did not participate
2 Survival in control
3 Mean 1 anrl PA/ 1
RR 1
1
67.5
15.02
60.02
85.0
87.52
90.0
1
97.5
93.8
8
74.5
36.7%
5
86.8
135%
in C. tentans
sediment (RR
inrh irlp all Hat:
(14.9)
(12.0)
(20.0)
(11.9)
(12.5)
(13.1)
(4.6)
(11.8)
test in May.
3) below minimum acceptable
=1 nnintQ
1
2.5
O2
O2
0
O2
12.5
1
0
0
8
1.88
233%
5
3.0
181%
level.
RR 2
(7.1)
(0)
(0)
(0)
(0)
(3.5)
(0)
(0)
RR 3
1
98.8
62.52
66.32
93.8
43.82
87.5
1
98.8
81.2
8
79.1
25.1%
5
92.0
8.3%
(Control)
(3.5)
(26.0)
(27.7)
(9.2)
(30.2)
(10.3)
(3.5)
(8.3)
Mean 2 and CV 2 exclude data points for all sediment samples from laboratories that did not meet minimum control survival of >70%.
Table 17.8 Interlaboratory Precision for Growth of Chironomus tentans in 10-d Whole-sediment Toxicity Tests Using Three
Sediments (May 1993)
Growth—Dry Weight in mg (SD) in Sediment Samples
Laboratory
A
B
C
D
E
F
G
H
I
J
N
Mean 13
CV1
N
Mean 24
CV2
RR 1
1
0.370
0.8832
0.21 52
0.657
0.21 02
0.718
1
0.639
0.347
8
0.505
49.9%
5
0.546
31 .9%
(0.090)
(0.890)
(0.052)
(0.198)
(0.120)
(0.114)
(0.149)
(0.050)
1
0
O2
O2
0
O2
0
1
0
0
8
—
—
5
—
—
RR2
(0)
(0)
(0)
(0)
(0)
(0)
(0)
(0)
RR 3
1
1.300
0.5042
1 .0702
0.778
0.61 02
1.710
1
1.300
1.180
8
1.056
38.3%
5
1.254
26.6%
(Control)
(0.060)
(0.212)
(0.107)
(0.169)
(0.390)
(0.250)
(0.006)
(0.123)
1 Did not participate in testing in May.
2 Survival in control sediment (RR 3) below minimum acceptable level.
3 Mean 1 and CV 1 include all data points.
Mean 2 and CV 2 exclude data points for all sediment samples from laboratories that did not meet minimum control survival of >70%.
121
-------
= 26.6%). The C. tentans in the moderately contaminated
sediment (RR 1) had a mean weight of 0.546 mg (CV =
31.9%). No growth measurements were obtained for C.
tentans in sediment RR 2 because of the high mortality.
The mean minimum detectable difference for growth among
laboratories meeting the survival performance criteria was
11% (SD = 5) and the MOD ranged from 4.8 to 23.6%
when the RR 1 data were compared to the RR 3 data.
17.5.3 Intel-laboratory Precision: 1996/1997
Evaluation of 10-d Sediment Tests
17.5.3.1 The 1996/1997 Precision Evaluation: 10-d
Whole-sediment Toxicity Testing. The results of the 10-
d toxicity interlaboratory comparisons conducted in 19967
1997 are presented in Tables 17.9 to 17.12. A total of 18
laboratories participated in the 1996/1997 study; however,
not all samples were tested by all laboratories.
Laboratories performed the tests during a specified time
period and followed methods outlined in Tables 11.1 and
12.1. Field samples were pretested to identify moderately
toxic samples. Samples were prepared and subsampled
at one time to increase consistency among the
subsamples. Samples were shipped to the testing
laboratories by express mail. Laboratories used their own
water supplies and were asked to use moderately hard
water (hardness about 100mg/LasCaCO3). The following
samples were evaluated in the 10-d toxicity tests: a field
control sediment from West Bearskin Lake, MN (WB), a
formulated sediment (FS, formulated with alpha-cellulose;
Kemble et al., 1999), two contaminated sediments (Little
Scioto River, OH (LS); Defoe Creek site, Keweenaw, Ml
(DC)), and FS spiked with three concentrations of
cadmium (0.3,1.0, and 3.0 mg/kg Cd). The LS sample was
primarily contaminated with polycyclic aromatic
hydrocarbons and the DC sample was primarily
contaminated with copper. Some laboratories did not
conduct tests on all samples due to logistical constraints.
In addition, ash-free dry weight (AFDW) was not measured
by laboratories which did not have access to a muffle
furnace.
17.5.3.2 The 1996/1997Precision Evaluation-Hyalella
azteca. Eighteen laboratories participated in the 19967
1997 H. azteca 10-d comparison (Table 17.9). A total of
82% of the laboratories had acceptable survival (>80%)
and for these tests the average survival (and CV) was 92%
(CV=5%) in the WB control sediment and 89% (CV=12%)
in the formulated sediment (FS). The two contaminated
field sediments (DC, LS) were moderately toxic, with the
mean survival of 45% (CV=38%) in DC sediment and 57%
(CV=49%) in LS sediment. The mean MDDs of the two
contaminated samples for all laboratories relative to the
WB control sediment were low (14% for both the DC and the
LS sediments). The range of MDDs relative to the WB
control sediment among all laboratories was 8 to 23% for
the DC sediment and 2 to 22% fortheLS sediment. A dose
response effect was observed with the Cd-spiked
formulated sediments. Moderate toxicity was observed in
the 1 mg/kg Cd sample with a mean survival of 49%
(CV=40%). The mean MOD and range for the 1 mg/kg Cd
sample for all laboratories was 16% (5.7 to 26%). It is
apparent from the MDDs that some laboratories had low
variability while others had only moderate levels of
variability.
17.5.3.3 The 1996/1997 Precision Evaluation -
Chironomus tentans. Eighteen laboratories participated
in the 1996/1997 C. tentans 10-d survival and growth
comparison (Table 17.10) with the same samples used in
the toxicity test as described above. A total of 15
laboratories (89%) had acceptable survival (>70%), and for
these tests, the mean survival was 89% (CV=9.4%) in the
WB control sediment and 88% (CV=10.2%) in the
formulated sediment (FS). The two contaminated field
sediments were only slightly toxic to the midge (mean
survival of 80% (CV=16%) for the DC sediment and 71%
(CV=33%) for LS sediment). The mean MDDs relative to
the WB control sediment, across all laboratories for the two
contaminated samples were low (12% for the DC sediment
and 11 % for LC sediment). The range of MDDs relative to
the WB control sediment among laboratories were 6.1 to
22% forthe DC sediment and 5.1 to 18% for LS sediment.
No toxicity was observed for survival in the cadmium tests.
The mean survival of midge in the 1 mg/kg Cd treatment
was 92% (CV=5.6%). The mean MOD and range for the 1
mg/kg Cd sample was 12% (6.9 to 30%). It is apparent from
the MDDs that some laboratories had low variability while
others had slightly lower variability.
17.5.3.4 Growth of C. tentans was evaluated by up to 16
laboratories in 1996/1997, depending on the sample and
whether or not they had capabilities to determine AFDW.
For dry weight analyses, 12 of 15 laboratories had
acceptable dry weight (>0.6 mg/individual) and survival
>70% in the WB control sediment, while 12 of 15 of the
laboratories had acceptable dry weight and survival in the
formulated sediment (FS; Table 17.11). For AFDW, 7 of 11
laboratories had acceptable weight (>0.48 mg/individual)
and survival >70% in WB control sediment (field control)
(WB) and 7 of 11 laboratories reported acceptable weight in
the formulated sediment (FS; Table 17.12). For the
midges, the mean dry weight was 1.39 mg/organism
(CV=33%) in the WB control sediment and 1.50 mg/
organism (CV=31%) in the formulated sediment (FS) for
laboratories that met the control survival in WB control
sediment. For AFDW, mean AFDW was 0.92 mg/organism
(CV=30%) in the WB control sediment and 1.161 mg/
organism (CV=33%) in the formulated sediment (FS).
Exposure to the contaminated DC sediment reduced the
weight of the midge (mean weight of 0.49 mg/organism
(CV=60%) as dry weight, while the mean weight of 0.24 mg/
organism (CV=45%) was determined for the AFDW), yet
exposure to LS sediment did not reduce weight of midges
(1.45 mg dry weight (CV=45%); 0.86 mg AFDW
(CV=27%)). The mean MDDs relative to WB control
sediment, across all laboratories for the two contaminated
samples, were low (0.17 mg/organism dry weight for the DC
sediment and 0.28 mg dry weight for LS sediment). The
range of MDDs among laboratories for dry weight was 0.04
to 0.53 mg/organism for DC sediment and 0.09 to 1.04 mg/
organism for LS sediment. The AFDW data exhibited a
similar pattern. Mean MOD as AFDW was 0.12 mg for the
DC sediment and 0.16 mg forthe LS sediment. The range
122
-------
Table 17.9 Intel-laboratory Precision for Survival (%) of Hyalella azteca in 10-d Whole-sediment Toxicity Tests (1996/1997)
to
Laboratory
A
B
C
E
F
G
H
I
K
M
N
0
P
Q
S
U
V
X
N-1d
Mean-1
SD-1
CV-1
N-2"
Mean- 2
SD-2
CV-2
71a
75a
NT
85
94
83
95
95
95
86
91
91
88
91
68"
94
95
99
WB
(23.0)
(24.5)
NT
(15.1)
(5.2)
(15.8)
(7.6)
(5-4)
(7.6)
(17.7)
(6.4)
(8.4)
(7.1)
(8.4)
(17.5)
(7.4)
(10.0)
(3.5)
17
88
9.1
10.3
14
92
4.6
5.0
Mean Percent Survival (SD) in Sediment Samples and
Sediment
DC LS FS
Oa
49a
NT
31
31
38
61
33
79
23
48
50
56
20
34a
60
35
59
(27.5)
(19.6)
(18.1)
(15.8)
(19.6)
(13.8)
(9.9)
(21.9)
(10.4)
(14.1)
(27.2)
(16.0)
(24.5)
(30.2)
(20.8)
(12.5)
17
42
18.9
45.6
14
45
17.1
38.3
NTb
84a
NT
71
19
28
64
85
94
50
29
74
60
84
80a
63
75
0
(30.7)
(34.4)
(16.4)
(12.8)
(20.7)
(9.3)
(7.4)
(22.7)
(23.6)
(10.6)
(27.3)
(22.0)
(23.9)
(21.2)
(20.8)
16
60
27.4
45.7
14
57
27.9
49.1
40a
90a
95°
83
60
90
99
99
100
85
85
95
85
96
70a
95
93
85
(37.8)
(7.6)
(5.8)
(14.9)
(20.0)
(9.3)
(3.5)
(3.5)
(0)
(16.9)
(14.1)
(5.4)
(10.7)
(52)
(25.1)
(5.4)
(15.0)
(15.1)
17
85
15.7
18.4
14
89
10.4
11.6
Cd-spiked Control Sediment
Cadmium -FS Spikes (mg/kg)
0.3-Cd 1-Cd
NT
NT
90°
68
40
NT
NT
83
98
80
100
78
83
98
NT
88
93
NT
(14.1)
(9.6)
(8.2)
(20.6)
(5-0)
(14.1)
(0)
(22.2)
(16.2)
(5.0)
(12.6)
(5.0)
11
83
17.2
20.9
11
83
17.2
20.9
NT
NT
73C
83
28
NT
NT
28
60
65
70
55
48
23
NT
38
40
NT
11
49
19.4
39.7
11
49
19.4
39.7
(9.6)
(9.6)
(5.0)
(17.1)
(8.2)
(19.2)
(8.2)
(26.5)
(16.2)
(28.7)
(15.0)
(14.1)
3-Cd
NT
NT
0°
3
3
NT
NT
0
0
0
3
0
0
0
NT
0
0
NT
11
1
1.4
171.3
11
1
1.4
171.3
(5.0)
(5.0)
(5.0)
Control survival below acceptable level of 80% in WB sediment.
NT = not tested.
Not included in any mean as WB control sediment was not tested.
N-1, Mean-1, SD-1 and CV-1 include all data (except Laboratory C) whether control met acceptable limits or not in WB sediment.
N-2, Mean-2, SD-2 and CV-2 include only data for sediment samples from laboratories that met the control performance acceptability criteria in WB sediment.
-------
Table 17.10 Interlaboratory Precision for Survival (%) of Chimnomus tentans in 10-d Whole-sediment Toxicity Tests (1996/1997)
Laboratory^
A
B
C
E
F
G
H
1
J
K
L
N
O
P
Q
R
S
X
N-1d
Mean-1
SD-1
CV-1
N-2a
Meari-2
SD-2
CV-2
81
100
NT
94
99
85
96
90
38C
96
84
83
51°
78
91
82
75
100
WB
(13.6)
(0)
(7.4)
(3.5)
(10.7)
(7.4)
(7.6)
(25.5)
(5.2)
(13.0)
(12.8)
(21.0)
(10.4)
(8.4)
(3.4)
(14.1)
(0)
17
84
16.9
20.0
15
89
8.3
9.4
Mean Percent
Sediment
DC
79
89
NT
93
84
76
93
83
25°
84
70
46
61 c
70
93
71
75
89
(6.4)
(9.1)
(11.7)
(10.6)
(20.7)
(7.1)
(13.9)
(20.7)
(10.6)
(13.1)
(32.9)
(18.1)
(17.7)
(8.9)
(15.4)
(27.8)
(12.5)
17
75
18.0
23.9
15
80
12.5
15.7
NTa
93
NT
84
84
19
94
74
83°
NT
86
86
91°
41
94
56
60
51
Survival (SD)
LS
(8.9)
(13.0)
(7.4)
(27,5)
(7.4)
(10,6)
(13.9)
(11.9)
(11.9)
(8.4)
(24,2)
(11-9)
(13.2)
(15.1)
(21,7)
15
68
27.9
41.0
13
71
23.6
33.3
in Sediment Samples a
FS
88
90
98b
96
88
74
100
86
48C
98
86
91
51C
88
99
77
71
98
(10.35)
(7.56)
(5.00)
(5.18)
(8.86)
(24.46)
(0)
(14.08)
(35.76)
(4.63)
(13.02)
(17.27)
(14.58)
(13.89)
(3.54)
(5.89)
(18.08)
(7.07)
17
84
15.5
18.5
15
89
9.1
10.2
id Cd-Spiked Control Sediment
Cadmium -FS Spikes (mg/kg)
0.3-Cd 1-Cd
NT
NT
98b
83
95
NT
NT
85
23C
NT
NT
88
85C
93
98
81
NT
NT
(5.0)
(17.1)
(5.8)
(12.9)
(22.2)
(12.6)
(5.8)
(9.6)
(5.0)
(8.0)
9
81
22.6
27.8
7
89
6.5
7.3
NT
NT
95"
85
93
NT
NT
93
63°
95
NT
95
90C
93
98
83
NT
NT
(5.8)
(5.8)
(9.6)
(9.6)
(28.7)
(10.0)
(5.8)
(8.2)
(9.6)
(5.0)
(11.8)
10
89
10.2
11.4
8
92
5.2
5.6
NT
NT
85fc
73
98
NT
NT
83
40C
NT
NT
70
95°
73
98
72
NT
NT
3-Cd
(19.1)
(9.6)
(5.0)
(15.0)
(24.5)
(8.2)
(5.8)
(9.6)
(5.0)
(29.4)
9
78
18.4
23.6
7
81
12.3
15.2
NT = not tested.
Not included in any mean as WB control sediment was not tested.
Control survival below acceptable level of 70% in WB sediment.
N-1, Mean-1, SD-1 and CV-1 include all data (except Laboratory C) whether control met acceptable limits or not in WB sediment.
N-2, Mean-2, SD-2 and CV-2 include only data for sediment samples from laboratories that met the control performance acceptability criteria in WB sediment.
-------
Table 17.11 Interiaboratory Precision for Growth (mg/Individual dry weight) of Chironomus tentansm 10-d Whole-sediment Toxicity Tests (1996/1997)
Laboratory
A
B
C
E
F
H
I
d
K
N
O
P
Q
R
S
X
N-1e
Mean-1
SD-1
CV-1
N-21
Mean-2
SD-2
CV-2
0.94
1.02
NT
2,47
1,69
0.92
1,55
0.90=
1.48
0.22"
0,99'
1,36
1,01
1.31
1 .73
0.97
WB
(0.15)
(0.06)
(0.30)
(0.17)
(0.12)
(0.27)
(0.83)
(0,12)
(0.11)
(0.17)
(0.18)
(0.29)
(0.29)
(0,29)
(0.10)
15
1.24
0.51
41.6
12
1.39
0.45
33.2
Mean Growth as Dry Weight
Sediment
DC LS
0.38
0.24
NT
1.05
0.41
0.24
0.37
Q.15C
0.20
0.06d
0.07°
1.01
0.21
0.58
0.48
0.68
(0.09)
(0.03)
(0.21)
(0.13)
(0.05)
(0.17)
(0.06)
(0.03)
(0.02)
(0.03)
(0.21)
(0.09)
(0.28)
(0.21)
(0.14)
15
0.41
.31
75.3
12
0.49
0.29
60.2
NTa
0.90
NT
2.69
1.62
0.93
1.80
0.91°
NT
0.30"
0.81°
0.87
1.31
1.06
2.36
0.95
(0.34)
(0.42)
(0.29)
(0.06)
(0.40)
(0.69)
(0.06)
(0.07)
(0.31)
(0.27)
(0.36)
(0.35)
(0.36)
13
1,27
0.67
53.1
10
1.45
0.65
45.1
(SD) in Sediment Sanif
FS
1.22
1.37
0.86"
2.29
2.43
1.29
1.74
0.36°
1.68
0.32e
1.37C
0.99
1.08
1.51
1.26
1.09
(0.27)
(0.12)
(0.12)
(0.51)
(0.40)
(0.21)
(0.49)
(0.23)
(0.18)
(0.10)
(0.29)
(0.29)
(0.17)
(0.34)
(0.80)
(0.22)
15
1.33
0.58
43.3
12
1,50
0.47
31.1
>Ies and Cd-Spiked Control Sediment
Cadmium -FS Spikes
0.3-Cd 1-Cd
NT
NT
0.83b
3.44
2.48
NT
2.58
1.02s
NT
0.35"
0.67=
1.63
1.06
1.25
NT
NT
(0.14)
(0.29)
(0.26)
(0.25)
(0.87)
(0.17)
(0.09)
(0.68)
(0.15)
(0.38)
9
1.61
1.02
63.3
6
2.1
0.92
44.2
NT
NT
0.83"
2.42
2.50
NT
2.05
0.42"
1.29
0.27 "
0,55C
1.54
1.16
1.37
(0.14)
(0.41)
(0.29)
(0.57)
(0,25)
(0.05)
(0.04)
(0.06)
(0.18)
(0.18)
(0.28)
NT
NT
10
1.36
0.80
58.7
7
1.76
0,56
31.5
(mg/kg)
NT
NT
0.20 b
2.90
1.02
NT
2.05
0.18"
NT
0.12"
0.15°
1,11
1.16
0.70
NT
NT
3-Cd
(0.09)
(0.58)
(0.43)
(0.50)
(0.05)
(0,02)
(0.02)
(0.03)
(0.10)
(0.24)
9
1.04
0.93
69.6
6
1.49
0.83
55.3
NT = not tested.
Not Included In any mean as WB control sediment was not tested,
Control survival below acceptable level of 70% in WB sediment.
Control weight below acceptable level of 0.60 mg/organism in WB sediment.
N-1, Mean-1, SD-1 and CV-1 include all data (except Laboratory C) whether control met acceptable limits or not in WB control sediment
N-2, Mean-2, SD-2 and CV-2 include only data for sediment samples from laboratories that met the control performance acceptability criteria in WB sediment.
-------
Table 17.12 Interlaboratory Precision for Growth (mg/lndividual as ash-free dry weight) of Chironomus tentans in 10-d Whole-sediment Toxicity Tests (1996/1997)
Mean Growth as Ash-free Dry Weight (SD) in Sediment Samples and Cd-spiked Control Sediment
Sediment
Cadmium -FS Spikes (mg/kg)
Laboratory
B
C
E
F
I
K
L
0
P
Q
R
X
N-1e
Mean-1
SD-1
CV-1
N-2(
Mean-2
SD-2
CV-2
WB
0.79
NT
0.25C
0.50
1,35
1,06
1.07
0,30°'d
0.36"
0.76
0.88
0.15d
11
0.677
0.39
58.1
7
0.916
0.27
29.8
(0.03)
(0.09)
(0.11)
(0.26)
(0.09)
(0.28)
(0.05)
(0.33)
(0.24)
(0.27)
(0.04)
0.18
NT
DC
(0.03)
0.1 Oc (0.03)
0.13
0.32
0.17
0.34
(0.12)
(0.13)
(0.02)
(0.09)
0.01 ='" (0.01)
0.29d
0.15
0.40
0.20d
(0.03)
(0.08)
(0.16)
(0.09)
11
0.208
0.12
56.1
7
0.241
0.11
45.0
LS
0.69
NT
0.2°
0.73
1.16
NT
1.13
0.26c'rf
0.18d
0.78
0.64
0.49d
10
0.630
0.35
54.9
6
0.855
0.23
26.8
(0.07)
(0.07)
(0.16)
(0.27)
(0.23)
(0.06)
(0.10)
(0.16)
(0.17)
(0.21)
FS
1.04 (0.09)
0.20b (0.05)
0.24C (0.06)
1.14 (0.39)
1.99 (1.50)
1.12 (0.09)
1.11 (0.18)
0.60cd (0.15)
0.1 5d (0.05)
0.79 (0.12)
0.94 (0.20)
0.30d (0.18)
11
0.856
0.53
61.8
7
1.161
0.39
33.2
0.3-Cd
NTa
0.1 9b (0.03)
0.48° (0.12)
0.94 (0.10)
2.01 (0.19)
NT
NT
0.22Cid (0.03)
0.46d (0.41)
0.74 (0.12)
0.74 (0.21)
NT
7
0.799
0.58
73.1
4
1.108
0.61
55.0
1-Cd
NT
0.23b (0.12)
0.27C (0.08)
1.00 (0.31)
1.56 (0.35)
0.91 (0.03)
NT
0.16c'd (0.03)
0.29" (0.07)
0.78 (0.22)
0.86 (0.22)
NT
8
0.729
0.47
64.6
5
1.022
0.31
30.4
NT
3-Cd
0.03" (0.03)
0.38° (0.18)
0.45
1.55
NT
NT
(0.24)
(0.41)
0.03c'd (0.01)
0.21
0.78
0.46
NT
" (0.05)
(0.04)
(0.17)
7
0.551
0.50
90.2
4
0.810
0.52
63.8
NT = not tested.
Not included in any mean as WB control sediment was not tested.
Control weight below acceptable weight criteria of 0.48 mg/organism in WB sediment.
Control survival below acceptable level of 70% in WB sediment.
N-1, Mean-1, SD-1 and CV-1 include all data (except Laboratory C) whether control met acceptable limits or not in WB sediment.
N-2, Mean-2, SD-2 and CV-2 include only data for sediment samples from laboratories that met the control performance acceptability criteria in WB
sediment.
-------
of MDDs for AFDW across laboratories was 0.03 to 0.22 mg
for the DC sediment and 0.04 to 0.25 mg for LS sediment.
No toxicity relative to weight was observed in the cadmium
tests. The mean dry weight of midge in the 1 mg/kg Cd
treatment was 1.76 mg/organism (CV=32%). The mean
MOD and range for the 1 mg/kg Cd sample was 0.28 mg/
organism (0.09 to 0.57). The AFD W for the 1 mg/kgsample
was 1.022 mg/organism (CV=30%) with MDDs of 0.19 mg
(0.04 to 0.36).
17.5.4 These round-robin tests conducted in 1993
(Section 17.5.2) and in 1996/1997 (Section 17.5.3)
exhibited similar or better precision compared to many
chemical analyses and effluent toxicity test methods
(USEPA, 1991 a; USEPA, 1991c). The success rate for
test initiation and completion of the USEPA's round-robin
evaluations is a good indication that a well equipped and
trained staff will be able to successfully conduct these
tests. This is an important consideration for any test
performed routinely in any regulatory program.
17.6 Precision of Sediment Toxicity Test
Methods: Evaluation of Long-term
Sediment Tests
17.6.1 Interlaboratory precision evaluations of the long-
term H. azteca and C. tentans tests, using the methods
described in Sections 14 and 15, were conducted byfederal
government, contract, and academic laboratories that had
demonstrated experience in sediment toxicity testing,
although only two of the laboratories had prior experience
with the long-term test methods described in this manual.
This round-robin study was conducted in two phases: a
Preliminary Round-robin (PRR) and a Definitive Round-
robin (DRR). The objective of the PRR was to provide
participating laboratories with an opportunity to become
acquainted with the techniques necessary to conduct the
two tests and to solicit commentary and recommendations
regarding potential improvements for the definitive
evaluation. Criteria for selection of participants in both
phases were that the laboratories: (1) had existing cultures
of the test organisms, (2) had experience conducting 10-d
tests with the organisms, and (3) would participate
voluntarily. Methods for conducting toxicity tests were
similar among laboratories, and each laboratory was
supplied with detailed operating procedures outlining these
methods. Methods for culturing were not specified and
were not identical across laboratories (as long as each
laboratory started with the appropriate age test organisms).
The PRR (phase 1) included the WB control sediment
(West Bearskin, MN; WB) and the formulated sediment
(FS) in which alpha-cellulose represented the primary
carbon source (Kembleetal., 1999; Table 17.13). The DRR
(phase 2) also included a copper-contaminated sediment
from Cole Creek, Keweenaw, Ml (CC), and a PAH-
contaminated sediment from the Little Scioto River, OH
(LS). In addition to the WB control sediment and the FS
sediment described above, an additional sediment, in
which peat (PE) represented the primary carbon source,
was also tested (Table 17.13).
17.6.2 Twelve laboratories participated in the PRR with H.
azteca. In these tests, 100% of laboratories passed the
acceptability criterion for survival (>80%) in the WB control
sediment at 28 d (Table 17.14) with survival ranges of 83
to 98% at 28 d, 71 to 93% at 35 d and 63 to 92% at 42 d.
In the formulated sediment (FS), 80% of the laboratories
met the survival criterion at 28 d (range: 47 to 98%).
Survival ranges in FS sediment at 35 d were 48 to 98% and
at 42 d the survival ranges were 48 to 98%. For growth
measured as length in the WB sediment, 92% of the
laboratories reported the mean length of the organisms to
be >3.2 mm at 28 d (range: 3.07 to 5.64 mm). For the FS
sediment, 100% of the laboratories reported length >3.2
mm with lengths ranging from 3.54 to 5.44 mm. For growth
measured as dry weight, >66% of the laboratories met the
minimum weight criterion (>0.15 mg/organism) in WB
(range: 0.10 to 1.16 mg/individual). In the FS samples,
100% of the laboratories met this growth criterion, with
weight ranges from 0.15 to 0.90 mg/individual. The criterion
for reproductive output for H. azteca (>2 young/female) was
met by 78% of laboratories in the WB (range: 0 to 27 young/
female). In the FS samples, 89% of the laboratories met
the reproductive requirement with ranges of 0.62 to 22
young/female.
17.6.3 Ten laboratories participated in the PRR with C.
tentans. In these tests, 90% of laboratories passed the
acceptability criterion for survival at 20 d (>70%) in WB
(range: 67 to 96%; Table 17.14), and in the FS sediment,
60% of the laboratories met the acceptability criterion
(range: 42 to 83%). For growth measured as dry weight,
100% of laboratories passed the criterion (>0.6 mg/
individual) in WB (range: 1.45 to 3.78 mg/individual). For
the FS samples, 86% of the laboratories passed the
criterion (range: 0.50 to 3.40 mg/individual). For growth as
AFDW, 100% of the laboratories passed the criterion of
>0.48 mg in the WB (range: 0.86 to 3.22 mg/individual)
(Table 17.14). In the FS sediment, 88% of the laboratories
met the growth criterion (as dry weight) with ranges of
weights from 0.42 to 2.72 mg/individual. The criterion for
emergence (>50%) was met by 70% of the laboratories in
WB sediment. In the FS, 50% of the laboratories met the
emergence criterion. The criterion for reproductive output
in C. tentans (>800 eggs/female) was exceeded by 90% of
laboratories in WB control sediment (range: 504 to 1240
eggs/female). In FS, 86% of laboratories met this criterion
in the FS (range: 0 to 1244 eggs/female). The suggested
criterion for percent hatch (>80%) was met by 88% of
laboratories in WB (range: 0 to 98%), and in FS, 67% of
laboratories (range: 0 to 98.7%).
17.6.4 In both the H. azteca and C. tentans tests, the
results of the PRR demonstrated that the majority of
laboratories met the acceptability criteria for those
endpoints for which criteria had been established (e.g.,
survival and growth). The highest proportion of failures in
the midge test occurred with post-pupation endpoints
(emergence, percent hatch) and may reflect the fact that
the criteria developed for these endpoints are based on
evaluations conducted at a single laboratory (Sibley et al.,
1996; Sibley etal., 1997b; Benoitetal., 1997). In the PRR,
some laboratories experienced unacceptably low oxygen
127
-------
Table 17.13 Physical Characteristics of the Sediments Used in the Preliminary and Definitive Round-robin Evaluations of Long-
term Methods for Sediment Toxicity Testing (Section 17.6).
Sediment
FS° (a high sand/low TOG)
WB
PE
Total
Organic
Carbon (%)
2.2
3.3
10
Particle Size (%)
Water
Content
31
31
NDa
Sand
74
74
ND
Clay
16
16
ND
Silt
11
10
ND
Sediment Type
Sandy Loam
Sandy Loam
Clay
ND = not determined
Table 17.14 Percentage of Laboratories Meeting Performance Levels for the Following Endpoints in the WB Control Sediment
Evaluated in the Long-term Round-robin Tests.
Performance Level
28-d survival > 80%
28-d growth > 3,2 mm length
28-d growth >0.15 mg dry weight
28- to 42-d reproduction (> 2 young/female)
20-d survival >70%
20-d growth >0.6 mg (dry weight)
20-d growth >0.48 mg (ash-free dry weight)
Emergence >50%
Number of eggs/egg case > 800
Percentage hatch >80%
Preliminary Round
Hvalella azteca
100
92
66
78
Chironomus tentans
90
100
100
70
90
88
Definitive Round
88
71
88
71
63
63
67
50
63
57
128
-------
Table 17.15 Intel-laboratory Comparison of Day 28 Percent Survival (Mean ± SD) of H. azteca in a Long-term Sediment
Exposure Using Five Sediments (WB = West Bearskin, CC = Cole Creek, LS = Little Scioto River, FS =
Formulated Sediment (using alpha-cellulose as organic carbon source), and PE = Formulated Sediment (using
peat moss as organic carbon source)).
Sediment
Laboratory
E
F
H
K
L
N
Q
U
X
N-1d
Mean-1
SD-1
CV (%)-1
N-2e
Mean-2
SD-2
CV (%)- 2
100
62b
93
95
83
89
98
100
NT
WB
(0)
(33.0)
(9.6)
(10.0)
(12.2)
(16.8)
(5.0)
(0)
8
90
12.8
14.3
7
94
6.4
6.8
97
84b
85
98
88
92
93
100
NT
CC
8
92
6.0
6.5
7
93
5.5
5.9
(4.9)
(21.1)
(5.8)
(4.5)
(8.7)
(8.4)
(9.6)
(0)
94
90C
98
96
84
91
80
98
NT
LS
8
91
6.4
7.0
7
91
6.9
7.5
(6.7)
(20.9)
(5.0)
(6.7)
(12.4)
(6.7)
(27.1)
(5.0)
94
38h
NT
NT
78
NT
90
NT
83°
FS
(7.9)
(35.2)
(14.2)
(14.4)
(10.7)
4
75
25.6
34.6
3
87
8.7
9.9
NTa
93"
68
54
NT
88
NT
93C
PE
4
75
17.8
23.6
3
70
16.8
24.1
(23.0)
(37.8)
NT
(40.6)
(9.6)
(7.5)
NT = not tested
Control survival below acceptable level of 80% in WB sediment.
Not included in any mean as WB control sediment was not tested.
N-1, Mean-1, SD-1 and CV-1 include all data (except Laboratory X) whether control met acceptable limits or
not in WB sediment.
N-2, Mean-2, SD-2 and CV-2 include only data for sediment samples from laboratories that met the 28-d
control performance acceptability criteria in WB sediment.
levels during evaluation of the C. tentans test which was
attributed to high feeding rates. To address this issue, the
feeding rate for the DRR of the C. tentans test was reduced
from 1.5 to 1.0 mL/d of Tetrafin.
17.6.5 In total, eight laboratories participated in the DRR
with H. azteca; however all laboratories did not test all
sediments. Mean survival for those laboratories that met
the control survival test acceptability criteria at 28 d in the
WB control sediment was 94% (CV=6.8%). In FS, the
mean survival was 87% (CV=9.9%), and in the PE it was
70% (CV=24%; Table 17.15). Mean survival at 35 d with
laboratories that met the >80% control survival criterion at
28 d was as follows: WB had 92% survival (CV=7.2%), FS
had 88% survival (CV=15.1%)and PE had survival of 63%
(CV=34.0%; Table 17.16). Mean survival at 42 d with
laboratories that met the >80% 28-d control survival
criterion was as follows: WB had 92% survival (CV=7.4%),
FS had 84% survival (CV=14.1%) and PE had 60%
survival (CV=38.2% with 3 laboratories; Table 17.16). At
28 d, 88% of the laboratories met the control survival
criteria in the WB control sediment (Table 17.14). When
acceptable 28-d control survival was reported in WB
sediment, 71% of the laboratories met the length criterion
(>3.2 mm) for H. azteca (Table 17.14). Forthose laboratories
that met the 28-d survival criterion and the growth criterion,
the mean growth (measured as length) of H. azteca at 28
d was 4.17 mm (CV=12.4%) in WB, 3.51 mm (CV=22.6%)
in the FS and 3.24 mm (CV=36.6%) in the PE (Table 17.18).
For growth measured as dry weight for the WB control
sediment, 88% of the laboratories met the weight criterion
of >0.15 mg/individual when acceptable 28-d control
survival was reported (Table 17.19) The mean growth of H.
azteca (mg/individual dry weight) in each sample where 28-
d control survival and growth was met was: 0.25 mg
(CV=27.8%) in WB, 0.30 mg (CV=68.6%) in FS, and 0.18
mg (CV=34.0%; Table 17.19) in PE. For the WB control
sediment, 71% of the laboratories met the reproduction
criteria (>2 young/female) when acceptable 28-d control
survival was reported (Table 17.14). The mean
reproduction from 28 to 42 d for laboratories that met both
the reproduction criteria and 28-d survival criteria was 3.13
young/female (CV=48.9%) for WB. For the FS, only one
laboratory that had acceptable survival in WB control
sediment at 28 d also had acceptable reproduction at 42 d,
with a mean of 2.3 young/female. Forthe PE sediment, the
only laboratory that had acceptable survival did not have
acceptable young production, as only 0.08 young/female
were obtained (Table 17.20).
129
-------
Table 17.16 Interiaboratory Comparison of Day 35 Percent Survival (Mean ± SD) of H. azteca in a Long-term Sediment
Exposure Using Five Sediments (WB = West Bearskin, CC = Cole Creek, LS = Little Scioto River, FS =
Formulated Sediment (using alpha-cellulose as organic carbon source), and PE = Formulated Sediment (using
peat moss as organic carbon source)).
Laboratory
E
F
H
K
L
N
Q
U
X
N-1°
Mean-1
SD-1
P\/ (°/ M
L< v 1 /o ) 1
N-2e
Mean-2
SD-2
CV (%) -2
98
70b
95
91
78
93
94
95
NT
WB
(7.1)
(31.6)
(7.6)
(18.1)
(10.4)
(7.1)
(9.2)
(7.6)
8
89
9.8
11.0
7
92
6,6
7.2
96
73b
96
96
83
88
86
98
NT
CC
(5.2)
(31.5)
(7.4)
(5.2)
(8.9)
(11.7)
(27.7)
(4.6)
8
89
8.9
10.0
7
92
6.2
6.7
Sediment
LS
96 (5.2)
86b (27.2)
95 (5.4)
90 (10.7)
84 (13.0)
83 (7.6)
88 (13.9)
86 (10.6)
NT
8
88
4.9
5.6
7
89
5.2
5.9
FS
98 (4.6)
33b (38.5)
NT
NT
73 (17.5)
NT
93 (8.9)
NT
74C (16.9)
4
74
21.0
30.4
3
88
13.2
15.1
NT8
86b
46
NT
57
NT
88
NT
95C
PE
(35.0)
(26.7)
(37.4)
(12.8)
(5.4)
4
69
30.0
40.1
3
63
21.5
34.0
NT = not tested
Control survival below acceptable level of 80% in WB sediment at 28 d.
Not included in any mean as WB control sediment was not tested.
N-1, Mean-1, SD1 and CV (%)-1 include all data (except Laboratory X) whether control met acceptable limits
or not in WB sediment.
N-2, Mean-2, SD-2 and CV-2 include only data for sediment samples from laboratories that met the 28-d
control survival performance acceptability criteria in WB sediment.
17.6.6 Overall, nine laboratories participated in the DRR
with C. tentansbut not all laboratories tested all sediments.
Mean survival (with CV in parentheses) for those
laboratories that met the control criterion of >80% survival
at 20 d was 85% (CV=5%) for WB sediment. In addition,
mean survival at 28 d, in the FS was 86% (CV=14.4%) and,
in the PE sediment was 75% (CV=13.9%) (Table 17.21).
In total, 63% of the laboratories met the acceptability
criterion for survival (>70%) for the WB control sediment
in the C. tentans test (Table 17.14). For laboratories
reporting dry weights, the mean growth of C. tentans at 20
d (criterion of >0.60 mg/individual dry weight and >70%
survival) was 1.45 mg (CV=58.6%) for WB sediment. In
addition, mean growth (as dry weight) was 1.63 mg/
individual (CV=20.9%) for the FS and 1.43 mg/individual
(CV=47.9%) for the PE sediment (Table 17.22). For
laboratories reporting weights as AFDW, the mean growth
of C. tentansal 20 d (criterion of >0.48 mg/individual AFDW
and >70% survival) was 0.81 mg (CV=53.3%) for WB, 1.05
mg/individual (CV=18.1%) for FS, and 0.64 mg/individual
(CV=12.7%)forPE (Table 17.23). For growth as dry weight
in the WB control sediment, 63% of the laboratories met the
acceptability criterion for survival and growth (as dry
weight) in the C. fenfanstest, while for AFDW, 67% of the
laboratories met the test acceptability criterion of >0.48
mg/AFDW per individual (Table 17.14). Mean percent
emergence for those laboratories that met the emergence
criterion of >50% reported emergence in WB control
sediment as 69.8% (CV=29.5%). In addition, mean
emergence was 50.5% in FS (CV=68.6%) and 55.8% in PE
(CV=30.3%) sediment (Table 17.24). In total 50% of the
laboratories met the acceptability criterion for both 20-d
survival and emergence in the WB control sediment (Table
17.14). The success rate for the number of eggs /case and
the control survival criterion was 63% in WB. Mean number
of eggs/female was 1118 eggs/case (CV=15.0%) in WB.
The FS and PE sediments had 1024 eggs/case
(CV=30.4%) and 867 eggs/case (CV=29.3%), respectively
(Table 17.25). The mean percent hatch for laboratories with
acceptable control survival and acceptable number of
eggs/case was 90% (CV=10.8%) for WB control sediment
(Tablel 7.26), and 57% of the laboratories that tested these
130
-------
Table 17.17 Interlaboratory Comparison of Day 42 Percent Survival (Mean ± SD) of H. azteca in a Long-term Sediment
Exposure Using Five Sediments (WB = West Bearskin, CC = Cole Creek, LS = Little Scioto River, FS =
Formulated Sediment (using alpha-cellulose as organic carbon source), and PE = Formulated Sediment (using
peat moss as organic carbon source)).
Sediment
Laboratory
E
F
H
K
L
N
Q
U
X
N-1d
Mean-1
SD-1
CV (%)-1
N-2e
Mean-2
SD-2
CV (%)2
95
61b
90
91
75
89
93
93
NT
WB
(7.6)
(31.8)
(9.3)
(18.1)
(10.7)
(8.4)
(11.7)
(8.9)
8
86
11.7
13.6
7
92
6.6
7.4
93
68b
90
96
83
81
81
95
NT
CC
(7.1)
(33.7)
(9.3)
(5.2)
(8.9)
(17.3)
(30.9)
(5.4)
8
86
9.6
11.2
7
88
6.6
7.5
95
85b
93
88
84
79
88
86
NT
LS
(5.4)
(26.7)
(8.9)
(12.8)
(13.0)
(10.7)
(13.9)
(10.6)
8
87
5.1
5.8
7
87
5.4
6.2
93
30b
NT
NT
70
NT
89
NT
43C
FS
(8.9)
(37.6)
(16.0)
(13.6)
(23.2)
4
70
28.6
40.7
3
84
12.1
14.1
NTa
83"
40
NT
55
NT
85
NT
84°
PE
(33.7)
(26.2)
(36.7)
(16.0)
(9.2)
4
65
21.8
33.3
3
60
22.9
38.2
NT = not tested
Control survival below acceptable level of 80% in WB sediment at 28 d.
Not included in any mean as WB control sediment was not tested.
N-1, Mean-1, SD1 and CV (%)-1 include all data (except Laboratory X) whether control met acceptable limits
or not in WB sediment.
N-2, Mean-2, SD-2 and CV (%)-2 include only data for sediment samples from laboratories that met the
28-d control survival performance acceptability criteria in WB sediment.
sediments met the test acceptability criteria for
hatchability.
17.6.7 In total, the proportion of laboratories that met the
various endpoint criteria in WB control sediment in the DRR
was higher for H. azteca than it was for C. tentans. The
most likely reason for the lower success with C. tentansm
the DRR was the reduction in feeding rate (from 1.5 to 1.0
ml of Tetrafin/beaker/d) relative to the PRR. In the PRR
with C. tentans, the proportion of laboratories meeting the
various endpoint criteria was generally higher (see Table
17.14), particularly for post-pupation endpoints (emergence,
reproduction, and percent hatch). Therefore, this manual
recommends that the higher feeding rate of 1.5 ml/beaker/
d be used in long-term tests with C. tentans (Section 15).
17.6.8 In the DRR, mean survival (CV in parentheses) of
H. azteca in the LS sediment (contaminated with PAHs;
using only values where the 28-d control survival criterion
was met) was 91% (CV=7.5%) at 28 d, was 89%
(CV=5.9%) at 35 d and 87% (CV=6.2%) at 42 d (Tables
17.15 to 17.17). Mean survival of C. tentansa\20d in the
LS sediment was 40% (CV=82.6%; Table 17.21). The
growth of H. azteca in LS sediment resulted in a mean
length of 4.37 mm (CV=10.1%; Table 17.18) and a mean
dry weight of 0.31 mg/individual (CV=38.2%; Table 17.19).
Mean growth of C. tentans in LS was 1.72 mg/individual
(CV=66.2%) as dry weight (Table 17.22) and 2.31 mg/
individual (CV=59.1%) as AFDW (Table 17.23). For both
species, all growth endpoints were highest for LS relative
to the other sediments evaluated, except for H. azteca dry
weight which had a comparable mean as the other four
sediments. The mean proportion of C. tentans larvae
emerging from LS was 35.7% (CV=71.2%; Table 17.24).
This value was roughly half of the emergence from the
control sediments. Mean reproductive output of H. azteca
in LS sediment, for those laboratories with acceptable
control survival, was 3.08 young/female (CV of 41.0%;
Table 17.20). The mean reproductive output of C. tentans
in the LS sediment for laboratories that met the control
survival criteria was 980 eggs/female (CV=20.1%; Table
17.25), which was similar to the WB, FS, and PE
sediments. Mean percent hatch of C. tentans eggs was
94% (CV=6.5%) for the laboratories that met at least 70%
control survival (Table 17.26).
131
-------
Table 17.18. Interlaboratory Comparison of Day 28 Length (Mean mm/Individual ± SD) of H. azteca in a Long-term Sediment
Exposure Using Five Sediments (WB = West Bearskin, CC = Cole Creek, LS = Little Scioto River, FS = Formulated
Sediment (using alpha-cellulose as organic carbon source), and PE = Formulated Sediment (using peat moss as
organic carbon source)].
Sediment
Laboratory
E
F
H
K
L
N
Q
U
X
N-1e
Mean-1
SD-1
CV (%)-1
N-2"
Mean-2
SD-2
CV (%)-2
4.15
3.02'
3.77
4.18
5.02
NR
3.11°
3.74
NT
WB
(0.23)
(0.28)
(0.32)
(0.12)
(0.11)
(0.10)
(0.08)
7
3.86
0.68
17.8
5
4.17
0.52
12.4
4.00
4.66
2.72
4.39
4.97
NR
3.17C
3.99
NT
CC
D,C
7
3.99
0.76
20.1
5
4.01
0.83
20.6
(0.11)
(0.17)
(0.14)
(0.29)
(0.27)
(0.18)
(0.17)
4.29
5.23 E
3.77
4.95
4.62
NR
4.29°
4.21
LS
(0.16)
'•= (0.41)
(0.17)
(0.22)
(0.40)
(0.45)
(0.13)
NT
7
4.40
0.50
10.9
5
4.37
0.44
10.1
2.96
3.70"
NT
NT
4.07
NR
4.51C
4.21
3.25d
FS
(0.03)
(0.30)
(0.39)
(0.46)
NA
(0.20)
5
3.81
0.66
17.3
2
3.51
0.79
22.6
NTa
5.03b
2.40
NT
4.08
NR
3.27°
NT
3.35d
PE
(0.06)
(0.41)
(0.64)
(0.03)
(0.21)
4
3.69
0.99
30.4
2
3.24
1.19
36.6
NT = not tested; NR = not reported; NA = not applicable.
Control survival below acceptable level of 80% in WB sediment at 28 d.
Length below acceptable level of 3.2 mm in length in WB control sediment.
Not included in any mean as WB control sediment was not tested.
N-1, Mean-1, SD1 and CV (%)-1 include all data (except Laboratory X) whether control met acceptable limits or not
in WB sediment.
N-2, Mean-2, SD-2 and CV-2 include only data for sediment samples from laboratories that met the 28-d control
survival performance acceptability criteria in WB sediment.
17.6.9 Across all laboratories that met the 28-d survival
criterion of >80% for H. azteca, the mean survival in the
contaminated CC sediment sample was 93% (CV=5.9%)
at 28 d, 92% (CV=7.2%) at 35 d, and 88% at 42 d
(CV=7.5%; Tables 17.15 to 17.17). Mean survival of C.
tentans at 20 d for laboratories that met the 20-d control
survival criteria was 75% (CV=30.9%; Table 17.21). In CC
sediment, the mean growth of H. azteca was 4.01 mm
(CV=20.6%) as length (Table 17.18) and 0.24 mg/individual
(CV=75.2%) as dry weight (Table 17.19). Mean growth of
C. tentans in CC sediment was 0.68 mg/individual
(CV=66.0%) as dry weight (Table 17.22) and 0.37 mg/
individual (CV=49.6%) as AFDW (Table 17.23). The
growth was reduced about 50% in the CC sediment in
comparison to the WB, FS, and PE sediments for C.
tentans only. The mean proportion of C. tentans larvae to
emerge from CC sediment was 38% (CV=60.5%; Table
17.24). Similar to the LS sediment sample, this emergence
was reduced to about half of that observed in the control
sediments. Mean reproductive output of H. azteca in CC
sediment, for those laboratories with acceptable 28-d
control survival, was 1.64 young/female (CV=103.3%) in
contrast to the mean for WB of 3.13 young/female
(CV=48.9%; Table 17.20). The mean reproductive output
of C. tentans eggs in the CC sediment for laboratories that
met the 20-d control survival criteria was 621 eggs/female
(CV=52.4%) (Table 17.25) which was the lowest egg
production for all sediments, which averaged between 404-
1194 eggs/female. The mean percent hatch of C. tentans
eggs was 69% (CV=49.5%) for the laboratories that met at
least 70% control survival (Table 17.26); all other
sediments had percent hatches for survival averaging 90 to
94%.
17.6.10 For the chronic H. azteca test, the mean MOD for
survival relative to the WB control sediment for the CC
sediment across all laboratories was only 7.7% (2.4 to
19.5%) at 28 d and 12.8% (6.4 to 281.7%) at day 42. The
MDDs for survival of amphipods were also small in the LS
sediment: 10.8% (3.3 to 26%) at 28 d and 11.5% (5.7 to
26%) at 42 d. The mean MDDs relative to WB control
sediment were also low for the 28-d amphipod weights as
the mean MOD for the CC sediment relative to WB control
sediment was 0.06 mg (0.04 to 0.14 mg) and the mean MOD
132
-------
Table 17.19. Interlaboratory Comparison of Day 28 Dry Weight (Mean mg/lndividual ± 3D) of H. azteca in a Long-term Sediment
Exposure Using Five Sediments (WB = West Bearskin, CC = Cole Creek, LS = Little Scioto River, FS = Formulated
Sediment (using alpha-cellulose as organic carbon source), and PE = Formulated Sediment (using peat moss as
organic carbon source)).
Sediment
Laboratory
E
F
H
K
L
N
Q
U
X
N-1e
Mean-1
SD-1
CV (%)-1
N-2f
Mean-2
SD-2
CV (%)-2
0.29
0.01bc
0.25
0.31
0.36
0.23
0.16
0.19
NT
WB
(0.04)
(0.01)
(0.06)
(0.04)
(0.04)
(0.10)
(0.04)
(0.02)
8
0.22
0.11
48.8
7
0.25
0.07
27.8
0.23
0.49b
0.10
0.56
0.41
0.09
0.09
0.21
NT
CC
(0.02)
(0.04)
(0)
(0.05)
(0.07)
(0.03)
(0.01)
(0.03)
8
0.27
0.19
69.6
7
0.24
0.18
75.2
LS
0.34
0.78b'c
0.20
0.58
0.32
0.25
0.31
0.27
NT
8
0.38
0.20
52.1
7
0.31
0.12
38.2
(0.07)
(0.18)
(0)
(0.09)
(0,12)
(0.09)
(0,09)
(0.04)
0.12
0.11
NT
NT
0.40
NT
0.39
NT
0.22
FS
(0.02)
bc (0.15)
(0.10)
(0.06)
d (0.17)
4
0.23
0.16
71.2
3
0.30
0.21
68.6
NT"
0.73b"
0,15
NT
0.24
NT
0.13
NT
0.42d
PE
4
0.31
0.28
90.0
3
0.18
0.06
34.0
(0.10)
(0.06)
(0.05)
(0.01)
(0.37)
NT = not tested.
Control survival below acceptable level of 80% in WB sediment at 28 d.
Weight below test acceptable criteria of 0.15 mg/organism in WB control sediment.
Not included in any mean as WB control sediment was not tested.
N-1, Mean-1, SD1 and CV (%)-1 include all data (except Laboratory X) whether control met acceptable limits or
not in WB sediment.
N-2, Mean-2, SD-2 and CV-2 include only data for sediment samples from laboratories that met the 28-d control
survival performance acceptability criteria in WB sediment.
for length was 0.26 mm (0.18 to 0.33 mm). The mean MOD
for LS sediment for amphipod growth as weight was 0.10
mg (0.05 to 0.16 mg) and length of 0.33 mm (0.14 to 0.44
mm). The mean MOD for the mean number of young per
female was 1.92 (0.09 to 2.4) in CC sediment and 2.06 (0.57
to 3.1) in LS sediment relative to WB control sediment.
17.6.11 The summary of the MDDs relative to the WB
control sediment for CC and LS samples and the chronic C.
fenfanstest is discussed by endpoint. For percent survival
at 20 d, the mean MDDs relative to WB control sediment for
CC and LS sediments were 14.4% (range of 5.9 to 19.1 %)
and 15.6% (5.8 to 25.3%), respectively. For 20 d dry
weights, the mean MDDs were 24.9% (CC) and 64.2% (LS)
with ranges of 15.6 to 30.4% and 25.1 to 126.9%,
respectively. The mean MOD and range for the AFDW
relative to the WB control sediment was 29.9% (22.9 to
44.6%) forthe CC sediment and 68.7% (22.9 to 125.0%) for
LS sediment. For emergence the mean MOD for the CC
sediment was 19.4% (10.5 to 25.0%) and the mean LS
MOD was 17.9 (8.2 to 23.0%). The number of eggs
produced had a mean MOD relative to the WB control
sediment of 19.4% (11.0 to 29.3%) forthe CC sediment and
24.4% (11.9 to 37.4%) for LS sediment, while hatch had a
mean MOD of 42.2% (7.4 to 77.3%) for the CC sediment
and 30.5% for LS sediment (9.3 to 53.7%).
17.6.12 These chronic round-robin tests exhibited similar
or better precision compared to many chemical analyses
and effluent toxicity test methods (USEPA, 1991 a;
USEPA, 1991c). The success rate for test initiation and
completion of the USEPA's round-robin evaluations is a
good indication that a well equipped and trained staff will be
able to successfully conduct these tests. These are very
important considerations for any test performed routinely in
any regulatory program.
133
-------
Table 17.20 Interlaboratory Comparison of Reproduction (Mean Number of Young/Female ± SD) of H, azfeca in a Long-term
Sediment Exposure Using Five Sediments (WB = West Bearskin, CC = Cole Creek, US = Little Scioto River, FS =
Formulated Sediment (using alpha-cellulose as organic carbon source), and PE = Formulated Sediment (using peat
moss as organic carbon source)).
Sediment
Laboratory
E
F
H
K
L
N
Q
U
X
N-f
Mean-1
SD-1
CV (%)-1
N-21
Mean-2
SD-2
CV (%)-2
5.7
4.0b
2.3
3.3
NAa
2.0
0.09°
2.4
NT
WB
(3.1)
(4.7)
(2.6)
(1.9)
(1.5)
(0.1)
(1.5)
7
2.8
1.8
62.6
5
3.13
1.53
48,9
CC
4.2
7.5'
0.3
1.2
NA
0.2
0.04C
2.4
NT
7
2.2
2.7
121.6
5
1.54
1.69
103.3
(2.2)
(7.6)
(0.2)
(1.4)
(0.7)
(0.04)
(1.7)
4.2
19.4b
1.2
4.1
NA
2.2
0.6C
3.5
NT
US
(1.6)
(4.4)
(1.3)
(4.5)
(1.3)
(0.9)
7
5.0
6.5
128.2
5
3.08
1.27
41.0
FS
2.3 (2.9)
5.4b (2.1)
NT
NT
NA
NT
0.2C (0.2)
NT
0.12d (0.73)
3
2.6
2.7
100.5
1
2.3
--
--
PE
NTa
16.5b
0.08
NT
NA
NT
0.3°
NT
0.5d
3
5.9
9.2
157.3
H
0.08
--
--
(9.4)
(1.8)
(0.4)
(0.7)
NT = not tested; NA = not applicable; young count not reported per female.
Survival below test acceptable criteria in WB control sediment at 28 d.
Reproduction below test acceptable criteria in WB control sediment of 2 young/female.
Not included in any mean as WB control sediment was not tested.
N-1, Mean-1, SD1 andCV (%)-1 include all data (except Laboratory X) whether control met acceptable limits or
not in WB sediment.
N-2, Mean-2, SD-2 and CV-2 include only data for sediment samples from laboratories that met the 28-d control
survival performance acceptability criteria in WB sediment.
134
-------
Table 17.21 Intel-laboratory Comparison of Day 20 Percent Survival (Mean ± 3D) of C. tentans in a Long-term Sediment Exposure
Using Five Sediments (WB = West Bearskin, CC = Cole Creek, US = Little Scioto River, FS = Formulated Sediment
(using alpha-cellulose as organic carbon source), and PE = Formulated Sediment (using peat moss as organic carbon
source)),
Sediment
Laboratory
E
F
H
1
K
N
Q
V
X
N-1d
Mean-1
SD-1
CV (%)-1
N-2e
Mean-2
SD-2
CV (%)-2
94
79
44b
54b
79
48b
77
98
NT
WB
8
72
20.6
28.7
5
85
9.8
11.5
(8)
(16)
(4)
(8)
(14)
(14)
(8)
(4)
98
40
69b
44b
74
50b
69
94
NT
CC
8
67
21.7
32.3
5
75
23.2
30.9
(4)
(4)
(21)
(14)
(7)
(18)
(10)
(8)
19
17
42b
15b
58
60b
16
90
NT
LS
8
40
28.0
70.6
5
40
33.1
82.6
(13)
(7)
(23)
(12)
(15)
(21)
(4)
(4)
94
81
40"
NT
NT
NT
71
98
75C
FS
(8)
(8)
(10)
(11)
(4)
(30)
5
77
23.2
30.2
4
86
12.4
14.4
NTa
65
NT
56b
NT
NT
75
85
63C
PE
(10)
(10)
(18)
(14)
(5)
4
71
12.9
18.3
3
75
10.5
13.9
Ni = not tested.
Survival below test acceptable criteria of 70% in WB control sediment at 20 d.
Not included in any mean as WB control sediment was not tested.
N-1, Mean-1, SD1 and CV (%)-1 include all data (except Laboratory X) whether control met acceptable limits or
notinWB sediment.
N-2, Mean-2, SD-2 and CV-2 include only data for sediment samples from laboratories that met the 28-d control
survival performance acceptability criteria in WB sediment.
135
-------
Table 17.22 Interlaboratory Comparison of Dry Weight (Mean mg/lndividual ± SD) of C. tentans in a Long-term Sediment Exposure
Using Five Sediments (WB = West Bearskin, CC = Cole Creek, LS = Little Scioto River, FS = Formulated Sediment
(using alpha-cellulose as organic carbon source), and PE = Formulated Sediment (using peat moss as organic carbon
source}}.
Sediment
Laboratory
E
F
H
1
K
N
Q
V
X
N-1"
Mean-1
SD-1
CV (%)-1
N-2e
Mean-2
SD-2
CV (%)-2
1.16
0,94
2,18b
1,96"
1.45
1 .33"
0.79
2.90
NT
WB
(0.09)
(0.28)
(0.13)
(0.49)
(0.32)
(0.91)
(0.25)
(0.73)
8
1.59
0.71
44.7
5
1.45
0.85
58.6
0.71
0.33
0,88"
2.00b
0.71
0.99b
0.26
1.39
NT
CC
(0.17)
(0.07)
(0.22)
(0.84)
(0.16)
(0.63)
(0.04)
(0.34)
8
0.91
0.57
62.6
5
0.68
0.45
66.0
0.83
3.49
2,85"
2.31b
2.05
1.39b
1.57
0.66
NT
LS
(0.32)
(1.23)
(0.58)
(1.17)
(0.29)
(0.66)
(0.60)
(0.24)
8
1.89
0.98
51.6
5
1.72
1.14
66.2
1.85
1.84
2.43b
NT
NT
NT
1.13
1.71
1.41':
FS
5
1.79
0.46
25.8
4
1.63
0.34
20.9
(0.76)
(0.30)
(0.30)
(0.24)
(0.52)
(0.26)
NT8
1.15
NT
2.65
NT
NT
0.93
2.21
1.83':
PE
(0.19)
(1.49)
(0.45)
(0.38)
(0.23)
4
1.74
0.83
47.7
3
1.43
0.68
47.9
* NT = not tested.
b Survival below test acceptable criteria of 70% in WB control sediment at 20 d.
0 Not included in any mean as WB control sediment was not tested.
'' N-1, Mean-1, SD1 and CV (%)-1 include all data (except Laboratory X) whether control met acceptable limits or
not in WB sediment.
9 N-2, Mean-2, SD-2 and CV-2 include only data for sediment samples from laboratories that met the 28-d control
survival performance acceptability criteria in WB sediment.
Note: All dry weight measurements for WB sediment were above the acceptable levsl of 0.6 mg/organism as dry weight.
136
-------
Table 17.23 Intel-laboratory Comparison of Ash-free Dry Weight (Mean mg/lndividual ± SD) of C. tentans in a Long-term Sediment
Exposure Using Five Sediments (WB = West Bearskin, CC = Cole Creek, LS = Little Scioto River, FS = Formulated Sediment
{using alpha-cellulose as organic carbon source), and PE = Formulated Sediment (using peat moss as organic carbon
source)).
Sediment
Laboratory
E
F
H
I
K
N
Q
V
X
N-1d
Mean-1
SD-1
CV (%)-1
N-2"
Mean-2
SD-2
CV (%)-2
0,87
0.65
1.74b
NMa
1.16
0.78"
0.57
NM
NT
WB
(0.12)
(0.18)
(0.13)
(0.28)
(0.31)
(0.27)
6
0.96
0.43
45.0
4
0.81
0.43
53.3
0.54
0.22
0.69b
NM
0.51
0.99b
0.20
NM
NT
CC
6
0.53
0.30
56.7
4
0.37
0.18
49.6
(0.17)
(0.03)
(0.19)
(0.09)
(0.48)
(0.03)
4.22
2.38
1.93"
NM
1.44
0.71b
1.20
NM
NT
LS
(1.80)
(0.84)
(0.43)
(0.29)
(0.47)
(0.50)
6
1.98
1.24
62.6
4
2.31
1.36
59.1
1.13
1.18
1.89"
NM
NT
NT
0.83
NM
0.30C
FS
(0.31)
(0.20)
(0.40)
(0.15)
(0.04)
4
1.26
0.58
35.7
3
1.05
0.19
18.1
NT'
0.69
NT
NM
NT
NT
0.58
NM
0.53C
PE
2
0.64
0.08
12.2
2
0.64
0.08
12.7
(0.19)
(0.26)
(0.11)
'" NT = not tested; NM = not measured.
b Survival below test acceptable criteria of 70% in WB control sediment at 20 d.
u Not included in any mean as WB control sediment was not tested.
d N-1, Mean-1, SD1 and CV (%)-1 include all data (except Laboratory X) whether control met acceptable limits or
not in WB sediment.
N-2, Mean-2, SD-2 and CV-2 include only data for sediment samples from laboratories that met the 28-d control
survival performance acceptability criteria in WB sediment,
Note: All dry weight measurements for WB sediment above acceptable level of 0.48 mg/organism as AFDW.
137
-------
Table 17.24 Interlaboratory Comparison of Percent Emergence {Mean ± SD) of C. tentans in a Long-term Sediment Exposure Using Five
Sediments (WB = West Bearskin, CC = Cole Creek, LS = Little Scioto River, FS = Formulated Sediment (using alpha-cellulose
as organic carbon source), and PE = Formulated Sediment (using peat moss as organic carbon source)).
Sediment
Laboratory
E
F
H
I
K
N
Q
V
X
N-1e
Mean-1
SD-1
CV (%)-1
N-2f
Mean-2
SD-2
CV (%)-2
65.6
20.8b
28.2b'°
1 1 .8°'°
57.3
30.2b'°
56.3
100
NT
WB
8
46.3
29.1
62.8
4
69.8
20.6
29.5
(14.4)
(7.7)
(8.9)
(12.0)
(18.6)
(17.8)
(13.9)
(0)
CC
41.7
5.2b
28.2bc
22gb.=
24.0
1 1 .5b'=
16.7
67.7
NT
8
27.2
19.7
72.4
4
37.5
22.7
60.5
(19.9)
(8.8)
(13.3)
(19.2)
(13.7)
(6.2)
(10.0)
(16.9)
18.8
12.5b
46.9"
5.6b
49.0
32.3"
10.4
64.6
NT
LS
(18.8)
(16.6)
(15.4)
(4.1)
(10.4)
(10.4)
(8.6)
(13.2)
8
30.0
21.6
71.9
4
35.7
25.4
71.2
75
29.2b
26 Ob
NT
NT
NT
26.0
NT
46.5°
FS
(21.8)
(14.1)
(14.4)
(14.3)
(20.2)
4
39.1
24.0
61.5
2
50.5
34.6
68.6
PE
NT'
31.2b
NT
8.3b'°
NT
NT
43.8
67.7
50 Jd
4
37.8
2.4.8
65.7
2
55.8
16.9
30.3
(15.3)
(10.7)
(20.8)
(9,4)
(24.2)
NT = not tested.
Emergence below test acceptable criteria of 50% in WB control sediment.
Survival below test acceptable criteria of 70% in WB control sediment at 20 d.
Not included in any mean as WB control sediment was not tested.
N-1, Mean-1, SD1 and CV (%)-1 include ad data (except Laboratory X) whether control met acceptable limits or
not in WB sediment.
N-2, Mean-2, SD-2 and CV-2 include only data for sediment samples from laboratories that met the 28-d control
survival performance acceptability criteria in WB sediment.
138
-------
Table 17.25 Intel-laboratory Comparison of the Number of Eggs/Female (Mean ± SD) in a Long-term Sediment Exposure Using Five
Sediments (WB = West Bearskin, CC = Cole Creek, LS = Little Scioto River, FS = Formulated Sediment (using alpha-cellulose
as organic carbon source), and PE = Formulated Sediment (using peat moss as organic carbon source)).
Sediment
Laboratory
E
F
H
1
K
N
Q
V
X
N-1d
Mean-1
SD-1
CV (%)-1
N-23
Mean-2
SD-2
CV (%)-2
1258
998
1397b
1261b
1023
1047b
978
1333
NT
WB
8
1162
168
14.4
5
1118
168
15.0
(429)
(243)
(408)
(225)
(177)
(410)
(168)
(227)
CC
523
444
91 9b
538b
538
484"
404
1194
NT
8
631
277
43.9
5
621
325
52.4
(124)
NA
(306)
(117)
(117)
(345)
(204)
(63)
LS
1025
722
1069b
NT
835
728b
1190
1127
NT
7
951
193
20.1
5
980
197
20.1
(366)
(711)
(580)
(86)
(479)
(126)
(191)
1260
671
995"
NT
NT
NT
1141
NT
828'n
FS
(178)
(133)
(615)
(391)
(286)
5
1017
255
25.1
3
1024
311
30.4
PE
NT"
721
NT
988C
NT
NT
720
1160
827C
5
897
21S
24.1
4
867
254
29.3
(200)
(290)
(105)
(120)
(214)
a NT = not tested; NA = not applicable.
b Survival below test acceptable criteria of 70% in WB control sediment at 20 d.
" Not included in any mean as WB control sediment was not tested.
d N-1, Mean-1, SD1 and CV (%)-1 include all data (except Laboratory X) whether control met acceptable limits or
not in WB sediment,
8 N-2, Mean-2, SD-2 and CV-2 include only data for sediment samples from laboratories that met the 28-d control
survival performance acceptability criteria in WB sediment.
Note: The number of eggs acceptable criteria (iSOO eggs) was above acceptable level for all laboratories in WB sediment.
139
-------
Table 17.26 Intel-laboratory Comparison of Percent Hatch (Mean ± SD) of C. tentans in a Long-term Sediment Exposure Using Five
Sediments (WB = West Bearskin, CC = Cole Creek, LS = Little Scioto River, FS = Formulated Sediment (using alpha-
cellulose as organic carbon source), and PE = Formulated Sediment (using peat moss as organic carbon source)).
Sediment
Laboratory
E
F
H
1
K
N
Q
V
X
N-r
Mean-1
SD-1
CV (%)-l
N-2'
Mean-2
SD-2
CV (%)-2
80
99
93b
NM'
62°
683C
80
91
NT
WB
7
82
13.5
16.6
4
90
9.7
10,8
(17.0)
(0.2)
(3.5)
(23,5)
(35.8)
(35.2)
(8.4)
37
97
80 b
NM
78C
47b.c
31
81
NT
CC
7
64
25.6
39.8
4
69
34.3
49.5
(33.0)
NA
(24.6)
(38.5)
(47.3)
(53.3)
(33.0)
51
99
71 b
NM
74=
54b,c
95
87
NT
LS
7
76
18.9
24.9
4
94
6.1
6.5
(39.0)
NA
(36.5)
(14.0)
(40.8)
(3.2)
(10.8)
77
97
74 b
NM
NT
NT
89
NT
60d
FS
(16.1)
(2,3)
(49.2)
(19.4)
(44.0)
4
84
10.7
12.7
3
93
5.5
5.9
NT
99
NT
NM
NT
NT
88
96
80d
PE
3
94
6.0
6.4
3
94
6.0
6.4
(0.4)
(18.3)
(1.7)
(27.1)
NT = not tested; NM = not measured; NA = not applicable.
Survival below test acceptable criteria of 70% in WB control sediment at 20 d.
Hatch below test acceptable criteria of 80% in WB control sediment.
Not included in any mean as WB control sediment was not tested.
N-1, Mean-1, SD1 and CV (%)-1 include all data (except Laboratory X) whether control met acceptable limits or
not in WB sediment.
N-2, Mean-2, SD-2 and CV-2 include only data for sediment samples from laboratories that met the 28-d control
survival performance acceptability criteria in WB sediment.
140
-------
Section 18
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155
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156
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Appendix A
Exposure Systems
A.1 Renewal of overlying water is recommended during
sediment tests (Section 11.3,12.3,13.3,14.3,15.3). The
overlying water can be replaced manually (e.g., siphon-
ing) or automatically. Automatic systems require more
equipment and initially take more time to build, but manual
addition of water takes more time during a test. In
addition, automated systems generally result in less sus-
pension of sediment compared to manual renewal of
water.
A.2 At any particular time during the test, flow rates
through any two test chambers should not differ by more
than 10%. Mount and Brungs (1967) diluters have been
modified for sediment testing, and other diluter systems
have also been used (Maki, 1977; Ingersoll and Nelson,
1990; Benoitetal., 1993; Zumwalt et al., 1994; Brunson
et al., 1998; Wall et al., 1998; Leppanen and Maier, 1998).
The water-delivery system should be calibrated before a
test is started to verify that the system is functioning
properly. Renewal of overlying water is started on Day -1
before the addition of test organisms or food on Day 0.
Water-delivery systems are described by Benoit et al.
(1993) in Section A.3 and by Zumwalt et al. (1994) in
Section A.4. A 60-mL syringe with a mesh screen over
the end can be used to manually remove and replace
overlying water (J. Lazorchak, USEPA, Cincinnati, OH,
personal communication).
A.3 Benoit et al. (1993) describe a sediment testing
intermittent-renewal (STIR) system (stationary or por-
table) for invertebrate toxicity testing with sediment. The
STIR system has been used to conduct both short-term
and long-term sediment toxicity tests with amphipods and
midges (Sections 11, 12, 14, 15). Either stationary or
portable systems enable the maintenance of acceptable
water quality (e.g., dissolved oxygen) by automatically
renewing overlying water in sediment tests at rates rang-
ing from 1 to 21 volume renewals/d. The STIR system not
only reduces the labor associated with renewal of overly-
ing water but also affords a gentle exchange of water that
results in virtually no sediment suspension. Both
gravity-operated systems can be installed in a compact
vented enclosure. The STIR system has been used for
conducting 10-d whole-sediment tests with Chironomus
tentans, Hyalella azteca and Lumbriculus variegatus.
A.3.1 STIR systems described in Benoit et al. (1982) can
be modified to conduct sediment tests and at the same
time maintain their original capacity to deliver varying
concentrations of toxicants for water-only toxicity tests. A
STIR system (stationary or portable) solely for sediment
toxicity tests was designed, which offers a simple, inex-
pensive approach for the automated renewal of variable
amounts of overlying water (Figures A.1 and A.2). This
system is described below. The system can be built as a
two-unit system (Section A.3.2) or with more exposure
treatments (Section A.3.4). All exposure systems consist
of exposure holding tanks, head tanks, head tank support
stands, and a water bath (Section A.3.2 and A.3.3). The
automated delivery system includes design descriptions
for a support stand, water renewal supply, and water-
delivery apparatus (Section A.3.4).
A.3.2 Two-unit Portable STIR System
Construction (Figures A.1 and A.2)
A.3.2.1 Exposure Holding Tanks (2) (Figure A.3).
1. Outer diameter: 15.8 cm wide x 29.3 cm long x 11.7
cm high
2. Cutting dimensions: (double-strength glass, 3 mm)
2 Bottoms: 15.8 cm x 29.3 cm
4 Sides: 11.4 cm x 28.7 cm
4 Ends: 11.4 cm x 15.8 cm
3. Hole: 1.6 cm centered between sides and 7.2 cm
from bottom edge of 11.4 cm high end piece.
4. Standpipe Height: 10.3 cm above inside of tank bot-
tom.
A.3.2.2 Head Tanks (2) (4-L capacity; Figure A.3)
1. Outer diameter: 15.8 cm wide x 24 cm long x 14.5 cm
high
2. Cutting dimensions: (acrylic plastic, 6 mm)
2 Bottoms: 15.8 cm x 24 cm
4 Sides: 13.9 cm x 22.8 cm
4 Ends: 13.9 cm x 15.8 cm
3. Acrylic plastic sheets should be cut with a smooth
cutting fine toothed table saw blade. Dimension cut
pieces can most easily be glued together with
Weld-On® #16 clear-thickened cement for acrylic
157
-------
Figure A.1 Portable table top STIR system described in Benoit et al. (1993).
158
-------
Calibrated Volume Sight Tube
(1.3cm Clear Tube)
t
Head Box
•(30.5x30.5x38cm High)
Adjustable Float on
Threaded Rod
-Toilet Tank Valve
:—Water Inlet
Timer Controled Solenoid Valves
Water Distribution ^
Manifold with Open Ends
(1.3cm plastic pipe)
Water Bath
Head Tank
4L
Calibrated
Row Tube
i
Y
Circulator
Pump
Junction
Box
< = = = = = = = -. CH!F= = =Water Bath Row-
/
Thermostat
Optional
Automated
Water
Delivery
_Apparatus
Pipe to Hose Adaptor
Holding
Tank
ft
Self
Starting
Siphon
Outlet
n
PI
Optional 1,2
or 3 Unit
"Add on"
Water Bath
0)
•p
CO
Q.
o
All tanks and water bath drain to common 19L jug with air
vent and optional hose from jug to floor drain.
Figure A.2 Portable table top STIR system with several additional options as described in Benoit et al. (1993).
plastic (Industrial Polychemical Service, P.O. Box
471, Gardena.CA, 90247).
4. Hole: 1.6 cm centered between sides and 2 cm from
front edge of 24-cm-long bottom piece. Holes can
most easily be drilled in acrylic plastic by using a
wood spade bit and drill press.
5. Flow Tubes: 10-mL pipet tip initially cut off at the 6-
ml_ mark and inserted flush with top of #0 stopper.
Top of stopper should be inserted nearly flush with
head tank bottom. With 2 L of water in head tank,
calibrate flow tube to deliver 32 mL/min.
A.3.2.3 Head Tank Support Stand (1) (Figure A.3)
1. Outer diameter: 16.7 cm wide x 33.7 cm long x 17.8
cm high
2. Cutting dimensions: (acrylic plastic, 6 mm)
1 Bottom: 16.7 cm x 33.7 cm
2 Sides: 17.2 cm x 32.5 cm
2 Ends: 17.2 cm x 16.7 cm
3. Size is such that both head tanks fit into support
stand for storage and transport.
A.3.2.4 Water Bath (1) (Figure A.3)
1. Outer diameter: 33 cm wide x 40.6 cm long x 7.4 cm
high
2. Cutting dimensions: (acrylic plastic, 6 mm)
1 Bottom: 33 cm x 55.9 cm
2 Ends: 33 cm x 6.8 cm
2 Sides: 39.4 cm x 6.8 cm
159
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Width (end)
Exposure Holding Tank
Width (end)
Head Tank
2.5cm
Water pump inlet
Water pump outlet
2.5cm
Overflow
1.6cm
Length (side)
Basic Water Bath
Water pump inlet
Water pump outlet
Thermostat O^—°verflowdrain
Length (side)
Basic Water Bath with Optional Holes for Water Bath
Width (end)
Add-on Water Bath for One Additional Unit
Figure A.3 Tanks for the STIR system in Benoit et al. (1993).
160
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3. Holes:
a. Overflow drain; 1.6 cm centered 2.9 cm from
bottom edge of 39.4-cm-long side piece and 17.8
cm from right edge.
b. Thermostat; 3.2 cm centered 2.5 cm from bottom
edge of 39.4-cm-long side piece and 3.2 cm from
left edge.
c. Water pump outlet; 2.5 cm centered 2.5 cm from
bottom edge of 33-cm-long end piece and 8.3 cm
from back edge.
d. Water pump inlet; 2.5 cm centered 2.5 cm from
bottom edge of 33-cm-long end piece and 2.0 cm
from back edge.
4. A small 90° elbow made of glass or plastic is at-
tached to the water pump inlet tube and turned down-
ward so the circulator pump will not pick up air at the
water surface.
5. The bottom piece for the water bath includes 15.3-cm
extension for motor mount and the thermostat electri-
cal junction box.
6. Motor Mount: 5.1 cm wide x 11.4 cm long x 3.8 cm
thick mount made from 6 pieces of 6-mm acrylic
plastic. Four of these pieces are glued together. The
other two pieces are glued together, motor attached
to the edge with two screws and the two pieces (with
motor attached) are then screwed to the top of the
four pieces. The entire unit is then glued to water bath
extension after 6-mm PVC piping is attached and
secured with stoppers to the inlet and outlet water
bath holes.
7. Thermostat Conduit Junction Box: (1.3-cm small left
back (SLB)) is attached to the water bath extension
by screwing a 1.3-cm PVC plug into junction box and
securing this plug with a screw, countersunk up through
the bottom and into the PVC plug.
A.3.2.5 Latex Rubber Mold
A.3.2.5.1 If you plan to construct a substantial number of
exposure test beakers, as described in Benoit et al.
(1993), then it would be to your advantage to make a latex
rubber mold to give support to the underside of the glass
when drilling holes. It significantly reduces the number of
broken beakers. Liquid latex, with hardener that can be
purchased from the local hardware store is commonly
used to coat the handles of tools. The rubber mold is
constructed as follows:
1. Mix latex with hardener as per instructions.
2. Fill one exposure test beaker with the mixture.
3. Suspend one 5-cm eye bolt (5-mm diameter) with nut
on end so that the eye is protruding just above the top
of the mixture.
4. Allow the latex plenty of time to "set up."
5. With proper eye protection and wearing heavy gloves,
gently break the beaker with a small hammer and
remove all of the glass from the mold.
6. Using a long drill bit for wood, drill an air vent hole
through the mold from top through bottom.
7. When using the mold, wet the mold and the beaker
with water before inserting. Place the beaker, with
pre-marked location of holes, on its side in a 3.5-L
stainless steel pan filled with coolant water so that
the beaker is just below the surface. The beaker is
then held in position with one hand while the other
hand operates the drill press. Operator should wear
proper eye protection.
8. Afterthe two holes are drilled, the mold can be easily
removed, with some effort, by inserting the eye bolt
into the handle of a securely attached "C" clamp and
physically pulling the beaker from the mold.
A.3.3 Suggested Options for More Exposure
Treatments (examples given are for a
three-unit treatment system)
A.3.3.1 Exposure Holding Tanks and Head Tanks
A.3.3.1.1 Same dimensions as for two-unit system ex-
cept that three (3) of each should be made.
A.3.3.2 Head Tank Support Stand (1) (Figure A.3)
1. Outer diameter: 16.7 cm wide x 49.5 cm long x
17.8cm high
2. Cutting dimensions: (acrylic plastic, 6 mm)
1 Bottom:
2 Sides:
2 Ends:
16.7 cm x 49.5 cm
17.2 cm x 48.3 cm
17.2 cm x 16.7 cm
3. Size is such that the three head tanks will fit into the
support stand for storage and transport.
A.3.3.3 Water Bath (1) (Figure A.3)
1. Outer diameter: 33 cm wide x 56.4 cm long x 7.4 cm
high
2. Cutting dimensions: (acrylic plastic, 6 mm)
1 Bottom:
2 Ends:
2 Sides:
33 cm x71.7 cm
33 cmx6.8 cm
55 cm x6.8 cm
3. Holes: All hole sizes and locations are the same as
for the two-unit system except that overflow drain is
located 25.7 cm from right edge of 55-cm side. Also,
two optional 1.6-cm holes centered 2.5 cm from
bottom edge of 33-cm-long end piece and 1.8 cm
161
-------
from corner edges are shown in the drawing for future
additions of "add-on" water baths.
4. Motor mount and junction box installations are the
same as for two-unit system.
A.3.3.4 "Add-on" Water Bath (example given is for
one additional unit treatment system;
Figure A.3)
1. Outer diameter: 18.5 cm wide x 33 cm long x 8 cm
high
2. Cutting dimensions: (acrylic plastic, 6 mm)
1 Bottom: 18.5 cm x 33 cm
2 Ends: 17.3 cm x 7.4 cm
2 Sides: 33 cm x 7.4 cm
3. Holes: Inlet and outlet holes (1.6 cm) are centered
2.5 cm from bottom edge of 33-cm long side piece
and 1.8 cm from corner edges.
4. The above holes will match the previously drilled
holes in the main water bath. The "add-on" water bath
is connected using #2 stoppers and 6.4-cm lengths of
clear plastic tubing (1.3-cm diameter). The circulator
pump outlet tubing (Tygon®) in the main water bath is
extended through the inlet connection as shown in
Figure A.2. Circulating water is then forced into the
"add-on" bath and flows back to the main water bath
by gravity.
5. Note that the walls of the "add-on" bath are 6 mm
higher than the main water bath to accommodate the
small head of water that builds up.
6. "Add-on" water baths tend to run a little warmer (0.2°C)
than main water bath test temperatures.
A.3.4 Optional Automated Water-delivery
Apparatus for Table Top STIR Systems
(examples given are for a three-unit
treatment system)
A.3.4.1 Support Stand
A.3.4.1.1 A stand to support the automated water-deliv-
ery apparatus, shown in Figure A.2, can be made from
bolted slotted angle iron bolted with corner braces. A
convenient size to construct is 30 cm wide x 85 cm long x
43 cm high. The head box in Figure A.2 sits on top of the
stand, and the water distribution manifold as shown in
Figure A.2 is placed directly under the top of the stand
with two 1.3-cm conduit hangers. A small portion of each
angle iron cross piece is cut away to allow the pipe to be
clamped into the conduit hanger. This also keeps the
manifold up high enough for sufficient clearance between
the head tanks and the 6-mm pipe to hose adapters as
shown in Figure A.2.
A.3.4.2 Water Renewal Supply
A.3.4.2.1 If tests will be conducted in the local water
supply, then the head box water inlet shown in Figure A.2
is simply plumbed into the supply line. However, if the
tests are conducted with transported water or with recon-
stituted water, the head box water inlet can be connected
to a Nalgene® drum with flexible Tygon® tubing. With a
four-volume test beaker water renewal flow rate per day,
both 114-L and 208-L Nalgene® drums will hold a 5-d
supply for a 3-unit treatment system and a 5-unit treat-
ment system, respectively. If the water supply drum is
located below the head box, then an open air water pump
such as a March® model MDXT pump (RFC Equipment
Corp., Minneapolis, MN 55440) can be used between the
drum and head box.
A.3.4.3 Operation of Water-delivery Apparatus
A.3.4.3.1 The head box water inlet solenoid valve
(Figure A.2) and the open air water pump (if needed) are
connected to the same timer control switch. The head box
water outlet solenoid valve is connected to another sepa-
rate timer control switch. With fourtest beaker renewals/d
and a 3-unit treatment system, the head box toilet float
valve is pre-adjusted to allow the head box to fill to the
12-L mark on the sight tube (Figure A.2).
A.3.4.3.2 With head box filled, the renewal cycle begins
when the first timer opens the head box outlet solenoid
valve. The distribution manifold is quickly flooded and the
12 Lof renewal water divided equally to each of the three
4-L head tanks. Since the timers have a minimum setting
of one hour on-off periods, the first timer is set to shut off
the head box outlet solenoid valve one hour after it opens.
A.3.4.3.3 About 30 min later, the second timer is set to
open the head box water inlet solenoid valve (and pump if
needed). As head box water volume reaches the 12-L
mark, the pre-adjusted toilet tank valve stops the water
flow. One hour after they come on, the second timer will
shut off the solenoid valve inlet and water pump.
A.3.4.3.4 The automated system is then ready for the
next renewal cycle that is set to begin 12 h after the first
cycle. Head box volume dimensions are such that up to
five-unit treatment systems can be tested simultaneously
as shown in Figure A.2.
A.3.5 A criticism of the system described by Benoit et al.
(1993) is that the (up to) 8 beakers placed in each holding
tank are not true replicates because of the potential for
exchange of water overlying the sediments among the
beakers. However, this concern is largely semantic with
regard to actual test results. The rationale for this position
is described below. The data described below are unpub-
lished data from USEPA Duluth (G.T. Ankley, USEPA,
Duluth, MN, personal communication).
A.3.5.1 Beakers within a test tank should contain an
aliquot of the same homogenized sediment and the same
test species. The replication is intended to reflect variability
162
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in the biology (e.g., health) of the organism, as well as
placement and recovery of the animals from the test
sediments (i.e., operator variability). To treat even com-
pletely separate tanks containing homogenized sediment
from the same source as true replicates (of the sediment
"treatment") is inaccurate and is pseudoreplication. Hence,
because the same sediment is tested in each beaker in a
particular tank, and because the replication is focused on
defining variability in the biology of the organism (and the
operator), this is essentially a nonissue from a theoretical
standpoint.
A.3.5.2 From a practical standpoint, it is important to
determine the potential influence of one beaker on another
over the course of a test. To determine this, a study was
designed (which is not advocated) in which treatments
were mixed within a tank. In the first experiment, four
beakers of highly metal-contaminated sediment from the
Keweenaw Waterway, Ml, were placed in the same tank
as four beakers containing clean sediment from West
Bearskin Lake, MN. This was done in two tanks; in one
tank, 10 amphipods (Hyalella azteca) were added to each
beaker, while in the other tank, 10 midges (Chironomus
tentans) were placed in each beaker. Controls for the
experiment consisted of the West Bearskin sediments
assayed in separate "clean" tanks. The four contaminated
beakers were placed "upstream" of the four clean beakers
to attempt to maximize possible exchange of contami-
nant. At the end of the test, organism survival (and growth
for C. tentans) was measured in two of the beakers from
each site and sediment Cu concentrations were deter-
mined in the other two beakers from each site. The
Keweenaw sediments contained concentrations of Cu in
excess of 9,000 ug/g (dry wt), and were toxic to both test
species (Table A.1). Conversely, survival of both
C. tentans and H. azteca was high in the West Bearskin
sediments from the Keweenaw tank, and was similar to
survival in West Bearskin sediments held in separate
tanks. Most important, there was no apparent increase in
Cu concentrations in the West Bearskin sediments held in
the Keweenaw tank (Table A.1).
Table A.1 Sediment Copper Concentrations and Organism
Survival and Growth at the End of a 10-d Test with
West Bearskin Sediment in an Individual Tank
Versus 10-d Cu Concentrations and Organism
Survival and Growth in West Bearskin Sediment
Tested in the Same Tank as Keweenaw Waterway
Sediment1
Sediment Tank Species
Survival Dry wt Cu
(%) (mg/organism) (ug/g)
WB2
WB
KW4
WB
WB
KW
1
2
2
3
4
4
Amphipod
Amphipod
Amphipod
Midge
Midge
Midge
90
100
20
95
100
5
ND3
ND
ND
1.34
1.33
ND
22.4
13.8
9397.0
12.3
15.6
9167.0
All values are the mean of duplicate observations (G.T. Ankley,
USEPA, Duluth, MN, unpublished data)
West Bearskin
Not determined
Keweenaw Waterway
A.3.5.3 A similar design was used to determine transfer
of contaminants among beakers containing sediments
spiked with the organochlorine pesticide dieldrin. In this
experiment, sediment from Airport Pond, MN, was spiked
with dieldrin and placed in the same tank as clean unspiked
Airport Pond sediments. Two different concentrations
were assayed as follows: (1) in the midge test, sediment
concentrations were about 150 ug dieldrin/g (dry weight)
and (2) in the amphipod test, sediments contained in
excess of 450 ug dieldrin/g sediment. The control for the
experiment again consisted of clean Airport Pond sedi-
ment held in a separate tank. The spiked sediments were
toxic to both test species, and survival of organisms held
in the clean Airport Pond sediments was similar in the two
different tanks. However, there was an effect on the
growth of C. tentans from the clean Airport Pond sedi-
ment assayed in the tank containing the spiked sediment.
This corresponded to the presence of measurable dieldrin
concentrations in unspiked Airport Pond sediments in the
tank with the mixed treatments (Table A.2). The concentra-
tions of dieldrin in the unspiked sediment, although de-
tectable, were on the order of 5,000-fold lower than the
spiked sediments, indicating relatively minimal transfer of
pesticide.
A.3.5.4 Using a similar design, an investigation was
made to evaluate if extremely low dissolved oxygen (DO)
concentrations, due to sediment oxygen demand, in four
beakers in a test system would result in a decrease in DO
in other beakers in the tank. In this experiment, trout chow
was added to each of four beakers containing clean
Pequaywan Lake sediment, and placed in a test tank with
four beakers containing Pequaywan Lake sediment with-
out exogenous organic carbon. Again, the control con-
sisted of Pequaywan Lake sediment held in a separate
tank under otherwise identical test conditions. Assays
were conducted, without organisms, for 10 d. At this time,
DO concentrations were very low in the beakers contain-
ing trout chow-amended sediment (ca., 1 mg/L, n = 4).
However, overlying water DO concentrations in the
Table A.2 Sediment Dieldrin Concentrations and Organism
Survival and Growth at the End of a 10-d Test with
Airport Pond Sediment in an Individual Tank
Versus 10-d Dieldrin Concentrations and Organism
Survival and Growth in Airport Pond Sediment
Tested in the Same Tank as Dieldren-spiked Airport
Pond Sediment1
Sediment Tank Species
Survival Dry wt Dieldrin
(%) (mg/organism) (ug/g)
AP2
AP
DAP4
AP
AP
DAP
1
2
2
3
4
4
Amphipod
Amphipod
Amphipod
Midge
Midge
Midge
75
80
20
85
85
0
ND3
ND
ND
1.71
0.13
ND
<0.01
0.07
446.4
<0.01
0.04
151.9
All values are the mean of duplicate observations (G.T. Ankley,
USEPA, Duluth, MN, unpublished data)
Airport Pond
Not determined
Dieldren-spiked Airport Pond
163
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"untreated" vs. the "treated" beakers in a separate tank
were similar, i.e., 6.8 vs. 6.9 mg/L, respectively. This
indicates that from a practical standpoint, even under
extreme conditions of mixed treatments (which again, is
not recommended), interaction between beakers within a
tank is minimal.
A.3.5.5 One final observation germane to this issue is
worth noting. If indeed beakers of homogenized sediment
within a test tank do not serve as suitable replicates, this
should be manifested by a lack of variability among
beakers with regard to biological assay results. This has
not proven to be the case. For example, in a recent
amphipod test with a homogenized sediment from the
Keweenaw Waterway in which all eight replicates were
held in the same tank, mean survival for the test was
76%; however, survival in the various beakers ranged
from 30 to 100%, with a standard deviation of 21%.
Clearly, if the test system were biased so as to reduce
variability (i.e., result in unsuitable replicates due to com-
mon overlying water), this type of result would not be
expected.
A.3.5.6 In summary, in both a theoretical and practical
sense, use of the system described by Benoit et al.
(1993) results in valid replicates that enable the evalua-
tion of variability due to factors related to differences in
organism biology and operator effects. To achieve this, it
is important that treatments not be mixed within a tank;
rather, the replicates should be generated from the same
sediment sample. Given this, and the fact that it is
difficult to document interaction between beakers using
even unrealistic (and unrecommended) designs, leads to
the conclusion that variability of replicates from the test
system can be validly used for hypothesis testing.
A.4 Zumwalt et al. (1994) also describe a water-delivery
system that can accurately deliver small volumes of
water (50 ml/cycle) to eight 300-mL beakers to conduct
sediment tests. The system was designed to be compa-
rable with the system described by Benoit et al. (1993).
This water-delivery system has been used in a variety of
applications (i.e., Kemble et al., 1998a,b; Ingersoll et al.,
1998).
A.4.1 Eight 35-mL polypropylene syringes equipped with
18-gauge needles are suspended from a splitting chamber
(Figure A.4). The system is suspended above eight bea-
kers and about 1 L of water/cycle is delivered manually or
automatically to the splitting chamber. Each syringe fills
and empties 50 ml into each beaker and the 600 ml of
excess water empties out an overflow in the splitting
chamber (Section A.4.3.1). The volume of water delivered
per day can be adjusted by changing either the cycling
rate or the size of the syringes. The system has been
used to renew overlying water in whole-sediment toxicity
tests with H. azteca and C. tentans. Variation in delivery
of water among 24 beakers was less than 5%. The
system is inexpensive (<$100), easy to build (<8 h), and
easy to calibrate (<15 min).
A.4.2 Water-Splitting Chamber
A.4.2.1 The glass water-splitting chamber is 14.5 cm
wide, 30 cm long, and 6.5 cm high (inner diameter). Eight
3.8-cm holes and one 2.5-cm hole are drilled in a 15.5 cm
x30.5 cm glass bottom before assembly (Figure A.4 and
Table A.3). The glass bottom is made from 4.8- (3/16 inch)
or6.4-mm (1/4 inch) plate glass. An easy way to position
the 3.8-cm holes is to place the eight 300-mL beakers
(2 wide x 4 long) under the bottom plate and mark the
center of each beaker. The 2.5-cm hole for overflow is
centered at one end of the bottom plate between the last
two holes and endplate (Figure A.4). After drilling the
holes in the bottom plate, the side (6.5 x 30.5 cm) and end
(6.5 x 14.5 cm) plates are cut from 3.2-mm (1/8 inch)
double-strength glass and the splitting box is assembled
using silicone adhesive. Sharp glass edges should be
sanded smooth using a whetstone or a piece of
carborundum wheel. After the splitting chamber has dried
for 24 h, four 12-mm (outer diameter) stainless-steel
tubes (7 cm long) are glued to each corner of the splitting
chamber (the surface of the steel tubes is scored with
rough emery paper to allow better adhesion of the sili-
cone). These tubes are used as sleeves for attaching the
legs to the splitting chamber. The legs of the splitting
chamber are threaded stainless-steel rods (9.5 mm [3/8
inch] diameter, 36 cm long). The location of the tubes
depends on the way that the beakers are to be accessed
in the waterbath. If the tubes are placed on the side of the
splitting chamber, a 3.2-mm-thickx2-cm-widex7-cm-long
spacer is required so beakers and the optional waterbath
can be slid out the ends (Figure A.4). If the sleeves and
legs are attached to the ends of the splitting chamber, the
beakers and waterbath can be removed from the side.
The legs are inserted into the 12-mm tubes and secured
using nylon nuts orwingnuts. The distance between the
tips of the needles to the surface of the water in the
300-mL beakers is about 2 cm. Four 1-L beakers could
also be placed underthe splitting chamber.
A.4.2.2 A #7 silicone stopper drilled with a 21 -mm (outer
diameter) core borer is used to hold each 35-mL polypro-
pylene syringe (45 mL total capacity) in place. Glass
syringes could be used if adsorption of contaminants on
the surface of the syringe is of concern. A dilute soap
solution can be used to help slide the syringe into the
#7 stopper (until the end of the syringe is flush with the top
of stopper). Stoppers and syringes are inserted into 3.8-cm
holes and are visually leveled. A #5 silicone stopper
drilled with an 8-mm (outer diameter) core borer is placed
in the 2.5 cm overflow hole. An 8-mm (outer diameter)
glass tube (7.5 cm long) is inserted into the stopper. Only
3 mm of the overflow tube should be left exposed above
the stopper. This overflow drain is placed about 3 mm
lower than the top of the syringes. A short piece of
6.4-mm (1/4 inch; inner diameter) tubing can be placed on
the lower end of drain to collect excess water from the
overflow.
A.4.2.3 The splitting chamber is leveled by placing a
level on top of the chamber and adjusting the nylon nuts.
Eighteen-gauge needles are attached to the syringes.
164
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Figure A.4 Water splitting chamber described in Zumwalt et al. (1994).
165
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Table A.3 Materials Needed for Constructing a Zumwalt et
al. (1994) Delivery System
Equipment
Drill press
Glass drill bits (2.54 cm [1 inch] and 3.8 cm [1.5 inch])
Cork boring set
Table-top saw equipped with a carborundum wheel
Small level (about 30 cm long)
Supplies
300-mL beakers (lipless, tall form; e.g., Pyrex Model 1040)
Stainless-steel screen (50- x 50-mesh)
9.5-mm (3/8 inch x 16) stainless-steel threaded rod
9.5-mm (3/8 inch x 16) nylon wingnuts
9.5-mm (3/8 inch x 16) nylon nuts
35-mL Mono-ject syringes (Sherwood Medical, St. Louis, MO)
18-gauge Mono-ject stainless-steel hypodermic needles
Silicone stoppers (#0, 5, and 7)
Plate glass (6.4 mm [1 /4 inch], 4.8 mm [3/16 inch], 3.2 mm [1 /8 inch])
Glass tubing (8-mm outer diameter)
Stainless-steel tubing (12-mm outer diameter)
Silicone adhesive (without fungicide)
5-way stainless-steel gang valves and
Pasteur pipets (14.5cm [5.75 inch])
About 6 mm of the needle should remain after the sharp
tip has been cut off using a carborundum wheel. Jagged
edges left in the bore of the needle can be smoothed
using a small sewing needle or stainless-steel wire.
A.4.2.4 When about 1 L of water is delivered to the
splitting chamber, the top of each syringe should be
quickly covered with water. The overflow tube will quickly
drain excess water to a level just below the tops of the
syringes. The syringes should empty completely in about
4 min. If water remains in a syringe, the needle should be
checked to ensure that it is clean and does not have any
jagged edges.
A.4.3 Calibration and Delivery of Water to the
Splitting Chamber
A.4.3.1 Flow adjustments can be made by sliding either
the stoppers or syringes up or down to deliver more or
less water. A splitting chamber with eight syringes can be
calibrated in less than 15 min. Delivery of water to the
splitting chamber can be as simple as manually adding
about 1 L of water/cycle. Water can be added automati-
cally to the splitting chamber using a single cell or a
Mount and Brungs (1967) diluter that delivers about 1 L/
cycle on a time delay. About 50 ml will be delivered to
each of the 8 beakers/cycle and 600 ml will flow out the
overflow. A minimum of about 1 L/cycle should be dumped
into the splitting chamber to ensure each syringe fills to
the top. If the quantity of water is limited at a laboratory,
the excess water that drains through the overflow can be
collected and recycled.
A.4.4 Waterbath and Exposure Beakers
A.4.4.1 The optional waterbath surrounding the beakers
is made from 3.2-mm (1/8-inch) double-strength glass and
is 15.8cmwidex29.5cm long x11.7 cm high (Figure A.4
[Figure A.3 in the Benoit et al., 1993 system]). Before the
pieces are assembled, a 1.4-cm hole is drilled in one of
the end pieces. The hole is 7.2 cm from the bottom and
centered between each side of the end piece. A glass
tube inserted through a #0 silicone stopper can be used to
drain water from the waterbath. A notch is made in each
300-mL beaker by making two cuts with a carborundum
wheel 1.9 cm apart to the 275 ml level. The beaker is
etched across the bottom of the cuts, gently tapped to
remove the cut section, and the notch is covered with 50-
x 50-mesh stainless-steel screen using silicone adhe-
sive. The waterbath illustrated in Figure A.4 is optional if
the splitting chambers and beakers are placed in a larger
waterbath to collect waste water. This smaller waterbath
could be used to collect waste water and a surrounding
larger waterbath could be used for temperature control.
A.4.5 Operation and Maintenance
A.4.5.1 Maintenance of the system is minimal. The
syringes should be checked daily to make sure that all of
the water is emptying with each cycle. As long as the
syringe empties completely, the rate of flow out of the
syringes is not important because a set volume of water
is delivered from each syringe. If the syringe does not
empty completely with each cycle, the needle tip should
be replaced or cleaned with a thin wire or sewing needle. If
the screens on the beakers need to be cleaned, a tooth-
brush can be used to brush the outside of screens.
A.4.5.2 Overlying water can be aerated by suspending
Pasteur pipets (e.g., Pyrex disposable 14.5-cm [5.75 inch]
length) about 3 cm above the sediment surface in the
beakers. Five-way stainless-steel gang valves are sus-
pended from the splitting chamber using stainless-steel
hooks. Latextubing (3.2-mm [1/8 inch] inner diameter) is
used to connect valves and pipets. Flow rate of air should
be maintained at about 2 to 3 bubbles/s and the pipets
can be placed on the outside of the beakers when samples
of overlying water are taken during a test.
A.4.5.3 The splitting chambers were used to deliverwater
in a toxicity test with the midge Chironomus tentans
exposed to metal-contaminated sediments (Zumwalt et
al., 1994). Ten third-instar midges were exposed in 300-mL
beakers containing 100 ml of sediment and 175 ml of
overlying water at 23°C. Midges in each beaker received a
daily suspension of 4 mg Tetrafin® flake food and sur-
vival and growth were measured after 10d. Splitting
chambers delivered 50 ml/cycle of overlying water to
each of the eight replicate beakers/sediment sample. One
liter of water was delivered with a single-cell diluter to
each splitting chamber 4 times/d. This cycle rate resulted
in 1.1 volume additions of overlying water/d to each
beaker ([4 cycles/d x 50-mL volume/cycle]/175 ml of
overlying water). The variation in delivery of water be-
tween 24 beakers was less than 5%.
A.4.5.4 Hardness, alkalinity, and conductivity in water
overlying the sediments averaged about 20% higher than
inflowing water. These water-quality characteristics tended
to be more similar to inflowing water at the end of the
166
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exposure compared with the beginning of the exposure.
The average pH was about 0.3 units lower than inflowing
water. Ammonia in overlying water ranged from 0.20 to
0.83 mg/L. The dissolved oxygen content was about 1 mg/L
lower than inflowing water at the beginning of the expo-
sure and was about 2 to 3 mg/L lowerthan inflowing water
by the end of the exposure. Survival and growth of midges
were reduced with exposure to metal-contaminated sedi-
ments. Water delivered at a similar rate to a second set of
beakers using a system described by Benoit et al. (1993)
resulted in similar overlying water quality and similartoxic
effects on midges.
A.4.5.5 The system has been used to deliver 33 %o salt
water to exposure chambers for 10d. Precipitation of
salts on the tips of the needles reduced flow from the
syringes. Use of a larger bore needle (16-gauge) reduced
clogging problems; however, daily brushing of the needle
tips is required. Use of larger bore needles with 300-mL
beakers containing 100 ml of sediment and 175 ml of
overlying water results in some suspension of sediment in
the overlying water. This suspension of sediment can be
eliminated if the stream of water from the larger bore
needle falls on a baffle (e.g., a piece of glass) at the
surface of the water in the beaker.
A.5 Brunson et al. (1998) describe a water-delivery
system for use with larger exposure chambers in the
Lumbriculus variegatus sediment exposures (Section 13).
Exposures of oligochaetes by Brunson et al. (1998) were
conducted for 28 d in 4-L glass beakers containing 1 L of
sediment and 3 L of overlying water. Four replicate
chambers were tested for each sediment sample evalu-
ated. Each beaker was calibrated to 4 L using a glass
standpipe that exited through the beaker wall and was
held in place with a silicon stopper. Beakers received
2 volume additions (6 L) of overlying water per day. Water
was delivered using a modified Mount and Brungs diluter
system that was designed to deliver 1 L/cycle (Ingersoll and
Nelson, 1990). An in-line flow splitter was attached to each
delivery line to split the water flow evenly to each of four
beakers. These splitters were constructed of 1/4 inch PVC
pipe with four silicone stoppers and 14-gauge stainless-
steel hypodermic needles with the points and connector
ends cut off the needles (Figure A.5). Glass stands were
used to support the splitters, keeping them level to maintain
a constant volume delivery to each beaker (+ 5%).
Figure A.5. Diagram of in-line flow splitter used to deliver overlying water in the sediment exposures of Lumbriculus
variegatus (Brunson et al., 1998).
167
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168
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Appendix B
Food Preparation
B.1 Yeast, Cerophyl®, and Trout Chow
(YCT) for Feeding the Cultures and
Hyalella azteca
B.1.1 Food should be stored at 4°C and used within two
weeks from preparation; however, once prepared, YCT
can be frozen until use.
B.1.2 Digested trout chow is prepared as follows:
1. Preparation of trout chow requires one week.
Use 1/8 inch pellets prepared according to cur-
rent U.S. Fish and Wildlife Service specifica-
tions. Suppliers of trout chow include Zeigler
Bros., Inc., P.O. Box 95, Gardners, PA, 17324
(717/780-9009); Glencoe Mills, 1011 Elliott,
Glencoe, MN, 55336 (320/864-3181); and Murray
Elevators, 118 West 4800 South, Murray, UT
84107(800/521-9092).
2. Add 5.0 g of trout chow pellets to 1 L of deionized
water. Mix well in a blender and pour into a 2-L
separatory funnel or similar container. Digest be-
fore use by aerating continuously from the bot-
tom of the vessel for one week at ambient labora-
tory temperature. Water lost due to evaporation is
replaced during digestion. Because of the offen-
sive odor usually produced during digestion, the
vessel should be placed in a ventilated area.
3. At the end of the digestion period, allow material
to settle for a minimum of 1 h. Filterthe superna-
tant through a fine mesh screen (e.g., Nitex®
110 mesh). Combine with equal volumes of the
supernatant from Cerophyl® and yeast prepara-
tion (below). The supernatant can be used fresh,
or it can be frozen until use. Discard the remain-
ing particulate material.
B.1.3 Yeast is prepared as follows:
1. Add 5.0 g of dry yeast, such as Fleishmann's®
Yeast, Lake State Kosher Certified Yeast, or
equivalent, to 1 L of deionized 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 super-
natant from the trout chow (above) and Cerophyl®
preparations (below). Discard excess material.
B.1.4 Cerophyl® is prepared as follows:
1. Place 5.0 g of dried, powdered cereal or alfalfa
leaves, or rabbit pellets, in a blender. Cereal
leaves are available as "Cereal Leaves" from
Sigma Chemical Company, P.O. Box 14508, St.
Louis, MO, 63178 (800/325-3010); or as
Cerophyl®, from Ward's Natural Science Estab-
lishment, Inc., P.O. Box 92912, Rochester, NY,
14692-9012 (716/359-2502). Dried, powdered al-
falfa leaves may be obtained from health food
stores, and rabbit pellets are available at pet
shops.
2. Add 1 L of deionized 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.
B.1.5 Combined yeast-Cerophyl-trout chow (YCT) is
mixed as follows:
1. Thoroughly mix equal (e.g., 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.
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. Do not store
YCT frozen over three months.
169
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4. It is advisable to measure the dry weight of solids
in each batch of YCT before use. The food should
contain 1.7 to 1.9 g solids/L.
B.2 Algal Food
B.2.1 Starter cultures of the green algae, Selenastrum
capricornutum are available from the following sources:
American Type Culture Collection (Culture No. ATCC
22662), 12301 Parklawn Drive, Rockville, MD 10852, or
Culture Collection of Algae, Botany Department, Univer-
sity of Texas, Austin, TX 78712.
B.2.2 Algal Culture Medium for the green algae is pre-
pared as follows (USEPA, 1993a):
1. Prepare stock nutrient solutions using reagent
grade chemicals as described in Table B.1.
Table B.1 Nutrient Stock Solutions for Maintaining Algal
Stock Cultures
2. Add 1 ml of each stock solution, in the order
listed in Table B.1, to about 900 ml of deionized
water. Mix well afterthe addition of each solution.
Dilute to 1 L, mix well. The final concentration of
macronutrients and micronutrients in the culture
medium is listed in Table B.2.
3. Immediately filterthe medium through a 0.45 urn
pore diameter membrane at a vacuum of not
more than 380 mm (15 in.) mercury, or at a pres-
sure of not more than one-half atmosphere (8 psi).
Wash the filter with 500 ml deionized water be-
fore use.
4. If the filtration is carried out with sterile appara-
tus, the filtered medium can be used immedi-
ately, and no further sterilization steps are re-
quired before the inoculation of the medium. The
medium can also be sterilized by autoclaving
after it is placed in the culture vessels. Unused
sterile medium should not be stored more than
one week before use, because there may be
substantial loss of water by evaporation.
Stock Compound Amount dissolved in
solution 500 ml deionized water
1. Macronutrients
A. MgCI2-6H20
CaCI2-2H20
NaNO3
B. MgSO4-7H2O
C. K2HP04
D. NaHC03
2. Micronutrients
H3BO3
MnCI2-4H20
ZnCI2
FeCI3-6H2O
CoCL-6H,O
2 2
Na2Mo04-2H20
CuCI2-2H20
Na2EDTA-2H2O
Na2Se04
1ZnCI2— Weigh out 1 64 mg and dilute to 1 00 ml
to micronutrient stock.
2CoCI2-6H2O — Weigh out 71 .4 mg and dilute
this solution to micronutrient stock.
6.08 g
2.20 g
12.75g
7.35 g
0.522 g
7.50 g
92.8 mg
208.0 mg
l.64mg1
79.9 mg
0.714 mg2
3.63 mg3
0.006 mg4
I50.0 mg
1.196 mg5
.Add1 mLofthissolution
to 100mL. Add 1 ml of
3Na2MoO4-2H2O— Weigh out 36.6 mg and dilute to 10 ml. Add 1 ml
of this solution to micronutrient stock.
4CuCI2-2H2O — 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 micronutrient stock.
B.2.3 Algal Cultures
B.2. 3.1 Two types of algal cultures are maintained:
(1) stock cultures and (2) "food" cultures.
Table B.2 Final
Concentration
of Macronutrients and
Micronutrients in the Algal Culture
Macronutrient
NaNO3
MgCI2-6H20
CaCI2-2H20
MgSO4-7H2O
K,HPO4
2 4
NaHC03
Micronutrient
H3B03
MnCI2-4H20
ZnCI2
CoCI2-6H2O
CuCI2-2H20
Na2MoCy2H20
FeCI3-6H2O
Na2EDTA-2H2O
Na2Se04
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
Medium
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
—
0.91
5Na2SeO4—Weigh out 119.6 mg and dilute to 100 mL. Add 1 mL of this
solution to micronutrient stock.
170
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B.2.3.2 Establishing and Maintaining Stock Cultures
of Algae
1. Upon receipt of the "starter" culture of
S. capricornutum (usually about 10 ml), a stock
culture is started by aseptically transferring 1 ml
to each of several 250-mL culture flasks contain-
ing 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. The volume of stock
culture maintained at any one time will depend on
the amount of algal food required for culture.
Stock culture volume may be rapidly "scaled up"
to several liters using 4-L serum bottles or similar
vessels containing 3 L of growth medium.
3. Culture temperature is not critical. Stock cultures
may be maintained at 25°C in environmental
chambers with cultures of other organisms if the
illumination is adequate (continuous "cool-white"
fluorescent lighting of about 4300 lux).
4. Cultures are mixed twice daily by hand.
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 3 ml of 7-d-old
algal stock culture, containing about 1.5 X
106 cells/ml are transferred to each 100 ml of
fresh culture medium. The inoculum should pro-
vide an initial cell density of about 10,000 to
30,000 cells/ml in the new stock cultures. Asep-
tic techniques should be used in maintaining the
stock algal cultures, and care should be exer-
cised to avoid contamination by other
microorganisms.
6. Stock cultures should be examined microscopi-
cally weekly at transfer for microbial contamina-
tion. Reserve quantities of culture organisms can
be maintained for 6 to 12 months if stored in the
dark at 4°C. It is advisable to prepare new stock
cultures from "starter" cultures obtained from es-
tablished outside sources of organisms every
four to six months.
B.2.3.3 Establishing and Maintaining
"S. capricornutum Food" Cultures
1. "S. capricornutum food" cultures are started 7 d
before use. About 20 ml of 7-d-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 (e.g., 3 L of
medium in a 4-L bottle or 18 L in a 20-L bottle).
The inoculum should provide an initial cell den-
sity of about 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. How-
ever, sterility of food cultures is not as critical as
in stock cultures because the food cultures are
used in 7 to 10 d. A one-month supply of algal
food can be grown at one time and stored in the
refrigerator.
2. Food cultures may be maintained at 25°C in
environmental chambers with the algal stock cul-
tures or cultures of other organisms if the illumi-
nation is adequate (continuous "cool-white" fluo-
rescent lighting of about 4300 lux).
3. Cultures are mixed continuously on a magnetic
stir plate (with a medium size stir bar), in a
moderately aerated separatory funnel, or are manu-
ally mixed twice daily. If the cultures are placed
on a magnetic stir plate, heat generated by the
stirrer might elevate the culture temperature sev-
eral degrees. Caution should be taken to prevent
the culture temperature from rising more than 2 to
3°C.
B.2.3.4 Preparing Algal Concentrate of
S. capricornutum for Use as Food
1. An algal concentrate of S. capricornutum con-
taining 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 at least
one week and siphoning off the supernatant.
2. The cell density (cells/ml) in the concentrate is
measured with an electronic particle counter, mi-
croscope and hemocytometer, fluorometer, or
spectrophotometer and used to determine the
dilution (or further concentration) required to
achieve a final cell count of 3.0 to 3.5 X 107
eel Is/ml.
3. Assuming a cell density of about 1.5 X 106 cells/
ml in the algal food cultures at 7 d, and 100%
recovery in the concentration process, a 3-L cul-
ture at 7 to 10 d will provide 4.5 X 109 algal cells.
4. Algal concentrate can be stored in the refrigerator
for one month.
5. Cultures of Hyalella azteca are fed 10 mL/L on
renewal/harvest days and 5 mL/L on all other
days(USEPA, 1993c).
B.2.3.5 Cell Counts
1. Several types of automatic electronic and optical
particle counters are available to rapidly count
cell number (cells/mL) and mean cell volume
(MCV; um3/cell). The Coulter Counter is widely
used and is discussed in detail in USEPA (1978).
When the Coulter Counter is used, an aliquot
171
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(usually 1 ml) of the test culture is diluted 10X to
20X with a 1% sodium chloride electrolyte solu-
tion, such as Coulter ISOTON®, to facilitate
counting. The resulting dilution is counted using
an aperture tube with a 100-um diameter aper-
ture. Each cell (particle) passing through the
aperture causes a voltage drop proportional to its
volume. Depending on the model, the instrument
stores the information on the number of particles
and the volume of each, and calculates the mean
cell volume. The following procedure is used:
A. Mix the algal culture in the flask thoroughly
by swirling the contents of the flask about six
times in a clockwise direction, and then six
times in the reverse direction; repeat the two-
step process at least once.
B. At the end of the mixing process, stop the
motion of the liquid in the flask with a strong
brief reverse mixing action, and quickly re-
move 1 ml of cell culture from the flask with
a sterile pipet.
C. Place the aliquot in a counting beaker, and
add 9 ml (or 19 ml) of electrolyte solution
(such as Coulter ISOTON®).
D. Determine the cell density (and MCV, if de-
sired).
2. Manual microscope counting methods for cell
counts are determined using a Sedgwick-Rafter,
Palmer-Maloney, hemocytometer, inverted mi-
croscope, or similar methods. For details on mi-
croscope counting methods, see APHA (1992)
and USEPA (1973). Whenever feasible, 400 cells
per replicate are counted to obtain ±10% preci-
sion at the 95% confidence level. This method
has the advantage of allowing for the direct ex-
amination of the condition of the cells.
B.3 Tetrafin® Food (or Other Fish Flake
Food) for Culturing and Testing
Chironomus tentans
B.3.1 Food should be stored at 4°C and used within two
weeks from preparation or can be frozen until use. If it is
frozen, it should be reblended, once thawed, to break up
any clumps
1. Blend the Tetrafin® food in deionized water for 1
to 3 min or until very finely ground.
2. Filter slurry through an #110 Nitex screen to re-
move large particles. Place aliquot of food in
100- to 500-mL screw-top plastic bottles. It is
desirable to determine dry weight of solids in
each batch of food before use. Food should be
held for no longer than two weeks at 4°C. Food
can be frozen before use, but it is desirable to
use fresh food.
3. Tetrafin® food is added to each culture chamber
to provide about 0.04 mg dry solids/mL of culture
water. A stock suspension of the solids is pre-
pared in culture water such that a total volume of
5.0 ml of food suspension is added daily to each
culture chamber. For example, if a culture cham-
ber volume is 8 L, 300 mg of food would be added
daily by adding 5 ml of a 56 g/L stock suspen-
sion (USEPA, 1993).
4. In a sediment test, Tetrafin® food (4.0 g/L) is
added at 1.5 ml daily to each test chamber.
172
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Appendix C
Supplies and Equipment for Conducting the
Chironomus tentans Life-cycle Sediment Toxicity Test
C.1 General
C.1.1 Section 15 outlines the methods for conducting a
Chironomus tentans life-cycle sediment toxicity test. This
Appendix describes the equipment needed to conduct
this test.
C.2 Emergence Traps (Figure C.1)
C.2.1 These traps are needed from Day 20 to the end of
the test. These traps fit on the top of the lipless glass
beakers with the narrow end up. These are 5-ounce
plastic cups with 14-mesh nylon screen glued to the cup
in place of the plastic bottom.
C.3 Reproduction/Oviposit Chambers
(R/O; Figure C.2)
C.3.1 These R/O chambers use emergence traps and are
needed once adults begin to emerge. Emergence traps
are used to store adults collected daily, and are placed in
a 100- X 20-mm petri dish that contains about 50 ml of
overlying water. When emergence occurs, the emer-
gence traps containing adults are removed and placed
onto a petri dish. At least one male for each emergent
female is added, and the R/O chamber (Figure C.2) is
placed back into the test system or into environmental
chambers maintained at the appropriate temperature and
lighting. A new emergence trap is then placed on top of
the lipless beaker. The R/O chambers are kept in this
manner to collect the egg masses and track mortality of
adults. If space is not a limiting factor, maintaining one
R/O chamber per pair of organisms is encouraged. Where
space is limited, many adults may be kept in a single R/O
chamber, and the chambers may be double stacked
(Double Stack Support Stand described in Section C.8)
using a larger plastic (9-ounce) cup that serves as a stand
for the second level of the emergence trap. The egg
masses are removed by lifting the edge of the cup enough
to permit transfer with a pipet.
C.4 Adult Collector Dish (Figure C.3)
C.4.1 This is used as a tray which is placed under the
emergence trap or reproduction/oviposit (R/O) chambers
to provide access to adults and to facilitate transfer of the
males and females as needed. This dish is constructed
of large petri dishes, i.e., 100- X 20-mm glass dishes or
100- X 20-mm plastic dishes. A 2.54-cm hole is cut in the
middle and covered with 58-mesh opening nylon screen.
Two slits are cut within the screen at 90 degree angles to
each other. This facilitates insertion of the aspirator tube
without risk of the adults flying away.
C.5 Aspirator (Figure C.3)
C.5.1 This is used to collect and transfer adults from the
reproduction/oviposit (R/O) chambers. A 60-cc syringe is
modified by cutting the end with the tip off and adding a
retainer to hold the emergence traps and reproductive/
oviposit chambers. The retainer is a 7-cm diameter
plastic lid (from 270-mL wide mouth glass jar) and a large
stopper is used to hold the syringe. The stopper and the
lid is drilled with a hole saw of about 1 inch. The large
stopper is glued to the lid. This retainer is then attached
to the syringe. To facilitate transferring the animals,
prepare two tubes, one about 16 cm in length and one
about 4 cm (6-mm ID) and place these in a stopper (i.e.,
No. 5, 5.5 or 6) that has been drilled with two holes.
Fasten a section (about 70 cm) of tygon tubing onto the
short piece of glass and cover the tube with a piece of thin
stainless steel screen (250-um mesh) before inserting the
tube into the rubber stopper. Adults should be stationary
in trap to minimize the possibility of escape.
C.6 Auxiliary Male Holding Dish
C.6.1 When emergence begins in the auxiliary beakers,
the males are transferred individually to inverted 60- X
15-mm plastic petri dishes with several small holes (3 mm
in diameter) drilled in the top. A thin layer of overlying
water (about 5 ml) is added and renewed until the males
are needed forthe reproduction chambers. These males
are held in the test system for temperature control, and
can be used for up to 5 d after collection.
C.7 Egg Hatching Chamber
C.7.1 Petri dishes, 60- X 15-mm plastic, are used to
incubate (23°C) egg masses in approximately 15 ml of
water. Hatch is monitored for 6 d. Hatch success is
determined by subtracting the number of unhatched eggs
at the end of 6 d from the initial estimate of the egg mass.
173
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Figure C.1.
Emergence trap used in the life-cycle Chironomus
tentans sediment test. A: the nylon screen; B: the
inverted plastic cups; C: the 300-mL lipless expo-
sure beaker; D: the water exchange screen ports;
E: test sediment.
Figure C.2.
The reproduction/oviposit chamber with the
double stack support stand. A: the notched,
inverted 270-ml (9-oz) plastic cup used to allow
double stacking; B: the reproduction/oviposit (R/
O) unit (C and D); C: inverted, 120-mL (4-oz) plastic
cup with nylon screen; D: one-half of petri dish
(100 X 20 mm) with 50 mL of overlying water; E: the
reproduction/oviposit (RIO) chamber.
C.8 Supplies and Sources
A. Emergence Trap/Reproduction Oviposit Chamber.
1. 120-mL (5-ounce) plastic cups, Plastics Inc.,
St. Paul, MN 55164.
2. 1400-mesh opening (micron) nylon screen
(mesh count = 14/inch), Monodur® 1400
Farbric Corporation, 7160 Northland Circle,
Minneapolis, MN 55428,612/535-3220.
B. Double Stack Support Stand: 270-mL (9-ounce)
plastic cups, Solo Inc, Urbana, 11,61801-2895.
C. Aspirator.
1. 60-cc syringe, 1 each, B-D® No. 309663,
Becton and Dickinson & Company, Franklin
Lakes, NJ 07417-1884.
2. 7-cm diameter plastic lid, 1 each.
3. Rubber stopper, 1 each, size 10,10.5, or 11.
4. Rubber stopper, 1 each, size 5.5 or 6.
5. Glass tubing, 6-mm I.D., 1- 16 cm long, 1-
4 cm long.
6. Nalgene 6-mm plastic connector for mouth
piece.
7. Stainless-steel screen, 250-um mesh.
D. Auxiliary Male Holding Chamber: 60- X 15-mm
petri dish with 3-mm holes drilled, Falcon 1007 B-
D®, Becton and Dickinson and Company, Franklin
Lakes, NJ 07417-1884.
E. Egg Hatching Chambers: 60- X 15-mm petri dish,
Falcon 1007 B-D®, Becton and Dickinson and
Company, Franklin Lakes, NJ 07417-1884.
F. Adult Collector Dish:
1. 100- X 20-mm glass petri dish with a 2.54-cm
access hole, Corning Glassware Corning, New
York or 100- X 20-mm plastic petri dish with a
2.54-cm access hole, Falcon 1005 B-D®,
Becton and Dickinson and Company, Franklin
Lakes, NJ 07417-1884.
2. 58-mesh opening nylon screen, cut with slits
at 90° angles to each other, Monodur®,Farbric
Corporation, 7160 Northland Circle,
Minneapolis, MN 55428,612/535-3220.
174
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Figure C.3. Adult collection/transfer equipment. A: transfer retainer unit showing inverted plastic cover and rubber stopper glued
inside of it; B: 60-cc syringe; C: plunger; D: detachable aspirator unit; E: long glass collector tube; F: short glass
tube to serve as connector for inhaler tube; note stainless steel screen attached to end through stopper; G: 2-hole
rubber stopper; H: nalgene plastic connector attached to tygon tubing and used as a mouthpiece to provide slight
suction; I: collector dish, one-half of glass or plastic petri dish; J: petri dish with hole access that is screen covered
and slotted; K: tygon tubing attached to glass tubing (F).
C.9 Construction of an Adult Midge
Emergence Trap for Use in a
"Zumwalt" Exposure System in
Life-cycle Sediment Tests
C.9.1 The construction of the emergence trap described
in Figure C.4 is an alternate design to the trap illustrated in
Figures C.1 and C.2. The emergence trap illustrated in
Figure C.4 is designed to fit under the exposure system
described by Zumwalt et al. (1994; Section A.4). The
level of the syringes will need to be raised about 1 1/2
inches using the threaded steel rods supporting the upper
chamber.
C.9.2 Cut a 2 1/2-inch plexiglass tube into 1 1/4-inch-long
pieces using a bandsaw or miter box and a handsaw.
C.9.3 Drill a 1/2-inch hole in the side (middle) of the
1 1/4-inch ring of plexiglass. Cut a small board to fit inside
of the 1 1/4-inch ring to help support the plexiglass when
drilling. The 1/2-inch drill bit should be dulled to help
prevent the bit from digging in too fast.
C.9.4 Drill three 1/16-inch holes in the plexiglass ring
spaced evenly around the ring and 1/4 inch off the bottom
of the ring.
C.9.5 Trace around the stainless-steel screen. Cut out
screen and place on top of the plexiglass ring. Use a
propane-soldering torch or glass-blowing torch to heat up
one end of a 1/4-inch or 3/8-inch threaded steel rod (about
12 to 15 inches long so that one end remains cool). Press
the hot end of the steel rod against the screen and
plexiglass until the screen melts into the plexiglass (usu-
ally a few seconds). Repeat the process until the screen
is completely melted to the top of the plexiglass ring.
C.9.6. Bend 4-mm glass tubing (outer diameter) over a
propane-soldering torch orglass-blowing torch and cut the
tubing with a glass wheel or etch the tubing with a file to
break. This glass tube is only to be used if beakers need
175
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Figure C.4 Emergence traps that can be used with the Zumwalt water-delivery system described in Section A.4.
to be aerated during the midge exposure. An air line is
connected to each tube and a gang valve is used to
regulate air flow (about 1 bubble/second). The glass tube
extends below the bottom of the plexiglass tube into the
surface of the overlying water. A 4-mm slot will need to
be cut in the petri dish in orderto slide the petri dish under
the emergence trap to remove adult midges from the test
beakers (Figure C.2). The emergence trap capped with
this petri dish can then be set on a 300-mL beaker to
remove the adults with an aspirator as illustrated in
Figure C.3.
C.9.7 Press 3/8-inch-long pins into the three 1/16-inch
holes drilled in the side of the plexiglass tube. These pins
make the plexiglass tube stable on the top of the beaker.
C.9.8 If the plexiglass tubes are used in beakers with a
notch at the top (i.e., the beakers described in Zumwalt et
al., 1994; Section A.4), a 2-cm length of 1/8-inch inner
diameter latex tubing will need to be slit lengthwise and
then slipped onto the bottom of the plexiglass tube. This
tubing is then lined up with the notch in the beakers to
prevent emerging midges from escaping. This piece of
tubing is not needed if beakers described in Benoit et al.
(1993) are used (i.e., beakers with holes drilled in the
side).
C.9.9 Supplies
A. McMaster Carr, P.O. Box 4355, Chicago, IL
60680-4355, 708/833-0300 (part number and ma-
terials).
1. 8486 K 115, Acrylic tube 2 1/2-inch outer
diameter and 1/8-inch wall.
2. 9226 T 84, 16- X 16-inch stainless wire cloth
(0.018-wire diameter).
3. 90145 A 417, 1/16-inch diameter stainless
dowel pins 3/8 inch long.
B. Thomas Scientific, P.O. Box 99, Swedesboro,NJ
08085-0099, 609/467-2000: 8747-E17, #00 sili-
cone stopper.
176
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Appendix D
Sample Data Sheets
177
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Culture
Aquarium
A
B
C
D
E
F
Date of Egg
Mass
Deposition
Date 4th
Instar
Larvae
Were
Weighed
Age of
Weighed
4th Instar
Larvae
Mean Dry
Weight of
4th Instar
Larvae
(n = 10)
Date of
Observed
First
Emergent
Adult
Total
Number of
ME"
Masses
Produced
General
Comments
Initials of
Culturist
Figure D.1 Data sheet for the evaluation of a Chironomus tentans culture.
178
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Position #
Embryo Dt
Embryo He
Number of
or number
Date 10 D
First Emer
Substrate
Food Type
^position Date
atch Date (da
larvae used t
of egg cases
ays Old Post
gence Date
Tvpe
Tank #
5 / /
/ 01 / /
o initiate tank
used
Hatch /
/ /
. Cone.
Set up Date
__•
/
/ / . Init.
. Date Made
Emergence Data (Performed 3 x Per Week)
Date
# of
# of
Total
Chemistries (Performed
Date
PH
D.O.
Ammonia
Comments
Weekly)
Temperature
Init.
Init.
Figure D.2 QA/QC data sheet for Chironomus tentans culture.
179
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10-d Old c- tentans Bo(jy Lengths
Date:
Tank #
Mean
Length
Init.
10-d Old c- tentans Head Capsule Widths
Date:
Tank #
% 2nd Instar
% 3rd Instar
% 4th Instar
Width
Init.
C. tentans Dry Weight Data
Date
Mean
Tank #
Pan + 10
Organisms
Pan Only
Difference
Weight/
Organism (mg)
Init.
Figure D.3 QA/QC data sheets for Chironomus tentans culture.
180
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Brood Stock Source
Test Type (circle one)1: SU SM RU RM FU FM
No. of Animals Tested Per Replicate
No. of Replicates
Method of LC50 Estimate
Reference Toxicant (CuSO4 or KCI)
Reference Toxicant Supplier and Lot No.
Reference Toxicant Purity
Test Initiation Date
Toxicologist
Exposure Duration (Hr)
0
24
48
72
96
Number of Mortalities
Control
A B
Exp. 1
A B
Exp. 2
A B
Exp. 3
A B
Exp. 4
A B
Exp. 5
A B
Current Test 96-h LC50 =
Number of Reference-toxicity Test Used
to Determine Cumulative Mean 96-h LC50_
Mean 96-h LC50 for All Tests to Date
Acceptability of Current Test2 Yes
No
1 SU = Static unmeasured
SM = Static measured
RU = Renewal unmeasured
RM = Renewal measured
FU = Flow-through unmeasured
FM = Flow-through measured
2 Based on two standard deviations around the cumulative mean 96 h-LC50
Figure D.4 Data sheet for performing reference-toxicity tests.
181
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Sediment Sample Source_
Date of Test Initiation
Toxicologist Conducting Test_
Test Dav
0
1
2
3
4
5
6
7
8
9
10
Test
Replicate
Samoled
Temperature
(°C}
Dissolved
Oxygen
(ma/L.}
DH
Hardness
(ma/L.}
Alkalinity
(ma/L.}
Specific
Conductance
(umnos/crrO
Total
Ammonia
(ma/L.}
Figure D.5 Data sheet for temperature and overlying water chemistry measurements.
182
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Daily Checklist for Sediment Tests
oo
CO
en
D
a>
s
in
o
-1
Q.
O
I
g.
(D
3
Study Code
Study Name
Building
Study Director
Lead Technician
Diluter
Waterbath
Target temperature
Acceptable Range _
°Cto
Month
Dissolved Oxygen
Minimum Acceptable Concentration
(40% of Saturation at Target Temp)
_mg/L
Day of Month
Day of Study
Diluter
Operation
Number of
Cycles
Time of Day
Temperature
Air Pressure
Aeration
Brush
Screens
Clean
Needles
Feeding
Total Water
Quality
Partial Water
Quality
Initials
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
26
27
28
29
30
31
Comments
Approved by
Date
-------
Water Quality Data Sheet
Study Code _
Study Name
Date
Test Day
Study Director .
Investigator
2!
-------
CHEMISTRIES
Page.
of
Test Type _
Organism
Test Dates,
Sample Info
CO
I
o
o
oo 2.
en w
a
a
S
(D
2
Water Type
Experimenter.
Test System
I.D.
pH
DO (mall)
Temo°C
Hard/Alk
oH
DO (ma/L)
TemD°C
Hard/Alk
pH
DO (mall)
Temo°C
Hard/Alk
oH
DO (mall)
TemD°C
Hard/Alk
Dav
-1
0
1
2
3
4
5
6
7
8
9
10
Remarks
-------
Study Director
Study Code
Study Name
Daily Comment Sheet
Day Date - - Initials.
Day Date - - Initials,
Day Date - - Initials,
Day Date - - Initials,
Day Date - - Initials,
Figure D.9 Daily comment data sheet.
186
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Weiaht Data Form
Test Dates
Test Material
Location
Analyst
Sample
Rppliratp
Wt. of
Oven
Dried Pan (mg)
Species
Weighing Date
Oven Temp (°C)
Drying Time (h)
Wt. of
Pan + Oven
Dried
Organisms
tmn\
Dried Wt. of
Organisms
(mg)
Number of
Survivors
Food
Age Organisms
Initial No/Rep
Mean wt per
Survivor
Sample Mean
Figure D.10 Weight data sheet.
187
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Date:
Test:
Species:
Facility:
Investigator:
Treatment
(Site)
Rep
1
2
3
4
5
6
7
8
1
2
3
4
5
6
7
8
1
2
3
4
5
6
7
8
Number
Surviving
Pan
Weight
Pan +
Larvae
Dry Weight
Total Indiv.
Pan +
Ash
Ash-free Dry Wt
Total Indiv.
Figure D.11 Data sheets for Chironomus tentans tests.
188
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At termination of test:
1. Sieve sediment from each beaker and record the number of recovered larvae in the
"survival" column.
2. Place all larvae from one replicate in a pre-ashed and pre-weighed aluminum weigh pan.
3. Dry larvae at 60°C for at least 24 hr.
4. Weigh pan + larvae and record weight under appropriate column of data sheet.
5. Ash pan + larvae at 550°C for 2 hr. Let cool to room temperature.
6. Weigh pan + ashed material.
7. Remove ash (e.g. with a small brush) and weigh pan.
8. Calculate dry weight as the difference between the pan+larvae weight and the pan weight.
9. Calculate ash-free dry weight as the difference between the pan+larvae weight and the
pan+ash weight.
Figure D.12 Instructions for terminating a Chironomus tentans test.
189
-------
Date:
Test:
Species:
Investigator:
Facility:
Treatment
Rep
Larvae
Number
1
7
3
4
5
6
7
8
9
10
11
12
Dead Pupae
No
ID rf ?
Date of Emergence
Partial Complete
rf ? rf ?
Date of
Egg
Mass
Egg Counts
Uve Acid
Number
Eggs Not
Hatched
Date
Adult
Died
3
IQ
C
3
o
O
B>
s
8 !
o
i
Comments (Adult transfers, mate pairings etc.)
o.
ID
Data Summary
No. of larvae recovered at end of test:
No. Dead/Escaped Adults:
Total Larvae:
Total Emerged Adults:
Number Dead Pupae:
Total Egg Mass: •
-------
Figure D.14 Example entries for a Chironomus tentans life-c
O
(D
m
a.
S
in
>ate: 01/28/96
nvestigator:
7RR-A
Rep
A
Larvae
Number
1
2
3
4
5
6
7
8
9
10
11
12
Dead Pupa
No ID *
3/4
3/15
Co
a Fully emerged: dead on water
b 2/24
-------
Copy of a sample data sheet that will be used to record all information pertaining to emergence and reproduction of
C. tentans during the life-cycle test. For clarity, consistency, and ease of data interpretation, it is important that
each lab fill out this sheet as illustrated. A brief interpretation of each recording (column) is provided below.
I Data Sheet Requirement. One data sheet is needed for each replicate. Thus, a treatment having 8 reproduction
replicates will have 8 data sheets (survival and growth data are recorded on separate sheets). All emergence and
reproduction data for a replicate are recorded on the corresponding data sheet.
II Recording Pupae, Emergence, and Egg Mass Data. Record all pupae, emergence, and egg mass data as dates.
Ill Column Heading Interpretation
Station/Site and Replicate. Enter name of sample and corresponding replicate (e.g., 7RR-A).
Larvae #. These numbers correspond to the 12 larvae placed in each replicate.
Dead Pupae. If it is not possible to determine the gender of the dead pupae, enter the date found in the "No ID"
column. Otherwise, enter the date found in either the male or female column.
Date of Emergence. If an adult has not completely shed the pupal exuviae, enter the date found under the "partial
emergence" category as a male or female. If emergence is complete but the adult is dead (typically floating on
the water surface), record date under "complete emergence" category as a male or female and enter a footnote
as indicated in "footnote a" in comments section of data sheet.
Partially emerged adults, and those that have emerged completely but were unable to escape the surface tension
of the water, usually die within 24 hr. In both cases, the date of death should be recorded as one day later under
the "Date Adult Died" column.
Date of Egg Mass. Record the date on which the egg mass was collected from the replicate.
Egg Counts. Enter number of eggs counted using either the acid-digestion (direct count) or ring method (indirect
count).
Number Eggs Not Hatched. Enter the number of unhatched eggs from each oviposited egg mass for which an
indirect count (ring method) was determined.
Date Adult Died. Enter the date that the adult died (be sure to follow transferred adults).
IV Comments Section. All comments concerning adult transfers and emergence patterns should be recorded in this
section as footnotes (see footnotes a-e on sample data sheet).
V Data Summary Section. At termination of each replicate, record the Number of Larvae Recovered at End of Test
after sieving and determine the number of Total Larvae alive during the test. Also record the Number Dead Pupae,
Number Dead/Escaped Adults, Total Emerged Adults, and number of Total Egg Masses by summing the
appropriate columns.
VI Example Entries for C. tentans Data-Sheet 7RR-A
Example #1. On 2/23/95 a male emerged from this replicate. This is recorded under the "Male" category of the
"Complete Emergence" column on the first line. This male was fully emerged but was dead and floating
on the water surface. This is recorded as footnote "a" in the "Comments" section and the date of death
recorded under the "Date Adult Died" column.
Example #2. A female emerged from this replicate on 2/26/95 which is recorded under the "Female" heading of the
"Complete Emergence" column. This female produced an egg mass on 2/28/96 which is recorded under
the "Date of Egg Mass" column.
Example #3. A dead pupae was recorded on 3/4/95. Since the sex was not determined, it was recorded under the
"No ID" heading of the "Dead Pupae" column. Pupal sex may be determined by examining the genitalia
under a dissecting microscope (the genitalia can be seen through the pupal exuviae which is usually,
but not always, transparent).
Example #4. A male emerged on 2/24/95 in 7 RR-A and was transferred to replicate 7RR-B. This is shown as foot-
note "b". Recording this type of data helps to keep track of where males are and the number of times
they have reproduced.
A male from 7SR-A (one of the stand-by replicates) was transferred to 7 RR-A on 3/8/95. This is
recorded as footnote "e" on the 7RR-A data sheet. For completeness, a corresponding footnote on
the 7 SR-A data sheet should be made regarding this transfer.
D.15 Instructions for completing the Chironomus tentans life-cycle test data sheet.
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