vvEPA
KPA600/R-94/024
June 1994
Agency
Washington DC 20460
Methods for Measuring the
Toxicity and
Bioaccumulation of
Sed i ment-associ ated
Contaminants with
Freshwater Invertebrates
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EPA 600/R-94/024
June 1994
Methods for Measuring theToxicity and
Bioaccumulation of Sediment-associated
Contaminants with Freshwater
Invertebrates
Office of Research and Development
U.S. Environmental Protection Agency
Dulutri, Minnesota 55804
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Disclaimer
This document has been reviewed in accordance with U.S. Environmental Protec-
tion Agency 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 poten-
tially pose a threat to a variety of aquatic ecosystems. Sediment functions as a
reservoir for common contaminants such as pesticides, herbicides, polychlori-
nated biphenyls (RGBs), polycyclic aromatic hydrocarbons, and metals such as
lead, mercury, and arsenic. In-place contaminated sediment can result in
depauparate bent hie communities, while disposal of contaminated dredge material
can potentially exert adverse effects on both pelagic and benthic systems.
Historically, assessment of sediment quality has been limited to chemical charac-
terizations. The United States Environmental Protection Agency {USEPA) is
developing methodologies to calculate chemical-specific sediment quality criteria
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 relationships between concentrations of contami-
nants in sediment and bioavailability 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, all Agency
programs have agreed to use the same 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 and toxic sub-
stances, Superfund site assessment, and assessment and cleanup of hazardous
waste treatment, storage, and disposal facilities. The use of uniform sediment
testing procedures by USEPA programs is expected to increase data accuracy
and precision, facilitate test replication, increase the comparative value of test
results, and, ultimately, increase the efficiency of regulatory processes requiring
sediment tests.
For additional guidance on the technical considerations in the manual, please
contact Teresa Norberg-King, USEPA, Duluth, MN.
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Abstract
Procedures are described for testing freshwater organisms in the laboratory to
evaluate the toxicity or bioaccumulation of contaminants associated 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. The toxicity tests are con-
ducted 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 endpoint in the toxicity test with H. azteca is survival
and the endpoints in the toxicity test with 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 with H. azteca.
Guidance for conducting 28-d bioaccumulation tests with the oligochaete
Lumbriculus variegatus is provided in this 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.
IV
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Contents
Foreword Hi
Abstract iv
Figures viii
Tables x
Acknowledgments xii
1 Introduction 1
1.1 Significance of Use 1
1.2 Program Applicability 2
1.3 Scope and Application 3
1.4 Performance-based Criteria 8
2 Summary of Method 10
2.1 Method Description and Experimental Design 10
2.2 Types of Tests 11
2.3 Test Endpoints 11
3 Definitions 12
4 Interferences 14
4.1 General Introduction 14
4.2 Non-Contaminant Factors 15
4.3 Changes in Bioavailability 15
4.4 Presence of Indigenous Organisms 16
5 Health, Safety, and Waste Management 17
5.1 General Precautions 17
5.2 Safety Equipment ,.... 17
5.3 General Laboratory and Field Operations 17
5.4 Disease Prevention 18
5.5 Safety Manuals 18
5.6 Pollution Prevention, Waste Management, and Sample Disposal 18
6 Facilities, Equipment, and Supplies 19
6.1 General 19
6.2 Facilities 19
6.3 Equipment and Supplies 19
7 Water, Formulated Sediment, Reagents, and Standards 22
7.1 Water 22
7.2 Formulated Sediment 23
7.3 Reagents 25
7.4 Standards 25
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Contents (continued)
8 Sample Collection, Storage, Manipulation, and Characterization 26
8.1 Collection 26
8.2 Storage 26
8.3 Manipulation 27
8.4 Characterization 28
9 Quality Assurance and Quality Control 30
9.1 Introduction 30
9.2 Performance-based Criteria 30
9.3 Facilities, Equipment, and Test Chambers 30
9.4 Test Organisms 31
9.5 Water 31
9.6 Sample Collection and Storage 31
9.7 Test Conditions 31
9.8 Quality of Test Organisms 31
9.9 Quality of Food 31
9.10 Test Acceptability 31
9.11 Analytical Methods 31
9.12 Calibration and Standardization 31
9.13 Replication and Test Sensitivity 32
9.14 Demonstrating Acceptable Performance 32
9.15 Documenting Ongoing Laboratory Performance 32
9.16 Reference Toxicants 32
9.17 Record Keeping 33
10 Collection, Culturing, and Maintaining Test Organisms 35
10.1 Life Histories 35
10.2 General Culturing Procedures 37
10.3 Culturing Procedures for Hyalella azteca 38
10.4 Culturing Procedures for Chironomus tentans 39
10.5 Culturing Procedures for Lumbriculus variegatus 43
11 Test Method 100.1: Hyalella azteca 10-d Survival Test for Sediments 44
11.1 Introduction 44
11.2 Recommended Test Method for Conducting a 10-d Sediment Toxicity Test
with Hyalella azteca 44
11.3 General Procedures 44
11.4 Interpretation of Results 48
12 Test Method 100.2: Chironomus tentans 10-d Survival and Growth Test for Sediments 51
12.1 Introduction 51
12.2 Recommended Test Method for Conducting a 10-d Sediment Toxicity Test
with Chironomus tentans 51
12.3 General Procedures 51
12.4 Interpretation of Results 55
VI
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Contents (continued)
13 Test Method 100.3: Lumbriculus variegatus Bioaccumulation Test for Sediments 57
13.1 Introduction 57
13.2 Procedure for Conducting Sediment Bioaccumulation Tests with
Lumbriculus variegatus 57
13.3 General Procedures 58
13.4 Interpretation of Results 63
14 Data Recording, Data Analysis and Calculations, and Reporting 64
14.1 Data Recording 64
14.2 Data Analysis 64
14.3 Data Interpretation 80
14.4 Reporting 81
15 Precision and Accuracy 82
15.1 Determining Precision and Accuracy 82
15.2 Accuracy 82
15.3 Replication and Test Sensitivity 83
15.4 Demonstrating Acceptable Laboratory Performance 83
15.5 Precision of Sediment Toxicity Test Methods 84
References 90
Appendices
A. Summary of USEPA Workshop on Development of Standard
Sediment Test Methods 101
B. Exposure Systems 109
C. Food Preparation 120
D. Sample Data Sheets 125
VII
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Figures
Figure 10.1 Length and relative age of Hyalelta azteca collected by sieving in comparison
with length of known-age organisms 40
Figure 10.2 Aspirator chamber (A) and reproduction and oviposit chamber (B)
for adult midges 42
Figure 11.1 Lifestage sensitivity of Hyalella azteca in 96-h water-only exposures 49
Figure 11.2 Average recovery of different age Hyalella azteca from sediment
by 7 individuals 50
Figure 12.1 Lifestage sensitivity of chironomids 56
Figure 14.1 Treatment response for a Type I and Type II error 66
Figure 14.2 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)) 67
Figure 14.3 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)) 68
Figure 14.4 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)) 69
Figure 14.5 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)) 70
Figure 14.6 Effect of alpha and beta on the number of replicates at various CVs
(assuming combined alpha + beta = 0.25) 71
Figure 14.7 Decision tree for analysis of survival and growth data
subjected to hypothesis testing 72
Figure 14.8 Decision tree for analysis of point estimate data 75
Figure 15.1 Control (cusum) charts: (A) hypothesis testing and
(B) point estimates (LC, EC, or 1C) 83
Figure B.1 Portable table top STIR system described in Benoit et al. (1993) 110
Figure B.2 Portable table top STIR system with several additional options
as described in Benoit et at. (1993) 111
Figure B.3 Tanks for the STIR system in Benoit et al. (1993) 112
Figure B.4 Water splitting chamber described in Zumwalt et al. (1994) 117
Figure D.1 Data sheet for the evaluation of a Chironomus tentans culture 126
Figure D.2 Data sheet for performing reference toxicant tests 127
Figure D.3 Data sheet for temperature and overlying water chemistry measurements 128
VIII
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Figures (continued)
Figure D.4 Data sheet for daily checklist for sediment tests 129
Figure D.5 Data sheet for water quality parameters 130
Figure D.6 Chemistry data sheet 131
Figure D.7 Daily comment data sheet 132
Figure D.8 Weight data sheet 133
IX
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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 5
Table 1.4 Water-only, 10-d LC50 (ng/L) Values for Hyalella azteca,
Chironomus tentans, and Lumbriculus variegatus 6
Table 4.1 Advantages and Disadvantages for Use of Sediment Tests 14
Table 6.1 Equipment and Supplies for Culturing and Testing Specific Test Organisms 21
Table 7.1 Characteristics of Three Sources of Clays and Silts
Used in Formulated Sediments 24
Table 7.2 Carbon, Nitrogen, Phosphorus Levels for Various Sources of Organic Carbon... 24
Table 7.3 Sources of Components Used in Formulated Sediments 25
Table 9.1 Recommended Test Conditions for Conducting Reference-Toxicity Tests
with One Organism/Chamber 33
Table 9.2 Recommended Test Conditions for Conducting Reference-Toxicity Tests
with More Than One Organism/Chamber 34
Table 10.1 Sources of Test Organisms 37
Table 10.2 Chironomus tentans Instar and Head Capsule Widths 40
Table 11.1 Test Conditions for Conducting a 10-d Sediment Toxicity Test
with Hyalella azteca 45
Table 11.2 General Activity Schedule for Conducting a Sediment Toxicity Test
with Hyalella azteca 45
Table 11.3 Test Acceptability Requirements for a 10-d Sediment Toxicity Test
with Hyalella azteca 46
Table 12.1 Recommended Test Conditions for Conducting a
10-d Sediment Toxicity Test with Chironomus tentans 52
Table 12.2 General Activity Schedule for Conducting a Sediment Toxicity Test
with Chironomus tentans 53
Table 12.3 Test Acceptability Requirements for a 10-d Sediment Toxicity Test
with Chironomus tentans 54
Table 13.1 Recommended Test Conditions for Conducting a
28-d Sediment Bioaccumulation Test with Lumbriculus variegatus 58
Table 13.2 Recommended Test Conditions for Conducting a Preliminary 4-d Sediment
Toxicity Screening Test with Lumbriculus variegatus 59
Table 13.3 General Activity Schedule for Conducting a 28-d Sediment Bioaccumulation
Test with Lumbriculus variegatus 60
Table 13.4 Test Acceptability Requirements for a 28-d Sediment Bioaccumulation Test
with Lumbriculus variegatus 61
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Tables (continued)
Table 13.5 Grams of Lumbriculus variegatus Tissue (Wet Weight) Required for
Various Analytes at Selected Lower Limits of Detection 62
Table 13.6 Detection Limits (ng) of Individual PAHs by HPLC-FD 62
Table 14.1 Suggested a Levels to Use for Tests of Assumptions 72
Table 14.2 Estimated Time to Obtain 95 Percent of Steady-State Tissue Residue 79
Table 15.1 Intralaboratory Precision for Survival of Hyalella azteca and
Chironomus tentans in 10-d Whole-Sediment Toxicity Tests, June 1993 84
Table 15.2 Participants in Round Robin Studies 85
Table 15.3 Interlaboratory Precision for Hyalella azteca 96-h LC50s from Water-only
Static Acute Toxicity Tests Using a Reference Toxicant (KCI) (October 1992).... 85
Table 15.4 Interlaboratory Precision for Survival of Hyaiella azteca in 10-d
Whole-Sediment Toxicity Tests Using Four Sediments (March 1993) 86
Table 15.5 Interlaboratory Precision for Chironomus tentans 96-h LC50s from
Water-only Static Acute Toxicity Tests Using a Reference Toxicant
(KCI) (December 1992) 87
Table 15.6 Interlaboratory Precision for Chironomus tentans 96-h LC50s from
Water-only Static Acute Toxicity Tests Using a Reference Toxicant
(KCI) (May 1993) 87
Table 15.7 Interlaboratory Precision for Survival of Chironomus tentans in 10-d
Whole-Sediment Toxicity Tests Using Three Sediments (May 1993) 88
Table 15.8 Interlaboratory Precision for Growth of Chironomus tentans in 10-d
Whole-Sediment Toxicity Tests Using Three Sediments (May 1993) 89
Table A.1 List of Laboratories Responding to the Survey 101
Table A.2 Summary of Testing Procedures Used to Evaluate the Toxicity of
Whole Sediments with Hyalella azteca 102
Table A.3 Summary of Testing Procedures Used to Evaluate the Toxicity of
Whole Sediments with Chironomus tentans 104
Table A.4 Summary of Testing Procedures Used to Conduct Whole-Sediment
Bioaccumulation Tests with Lumbriculus variegatus 106
Table B.1 Sediment Copper Concentrations and Organism Survival and Growth
at the End of a 10-d Test 115
Table B.2 Sediment Dieldrin Concentrations and Organism Survival and Growth
at the End of a 10-d Test 116
Table B.3 Materials Needed for Constructing a Zumwalt et al. (1994) Delivery System .... 118
Table C.1 Nutrient Stock Solutions for Maintaining Algal Stock Cultures 121
Table C.2 Final Concentration of Macronutrients and Micronutrients in the
Algal Culture Medium 121
XI
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Acknowledgments
This document is a general purpose testing manual for freshwater sediments. The
approaches have also been described in ASTM (1994a) ASTM (1994b), Ankley et
al. (1993), Phipps et at. (1993), Brooke et al. (1993), Call et al. (1993a), Call et al.
(1993b), Lee et al. (1994), and USEPA (1994a).
This manual reflects the consensus of the Freshwater Sediment Toxicity Assess-
ment Committee and the U.S. Environmental Protection Agency (USEPA) Pro-
gram Offices. Members of the Freshwater Sediment Toxicity Assessment Commit-
tee are G.T. Ankley, USEPA, Duluth, MN; D.A. Benoit, USEPA, Duluth, MN; G.A.
Burton, Wright State University, Dayton, OH; F.J. Dwyer, National Biological
Survey (NBS; formerly U.S. Fish and Wildlife Service), Columbia, MO; I.E. Greer,
NBS, Columbia, MO; R.A. Hoke, SAIC, Hackensack, NJ; C.G. Ingersoll, NBS,
Columbia, MO; P. Kosian, USEPA, Duluth, MN; P.F. Landrum, NOAA, Ann Arbor,
Ml; J.M. Lazorchak, USEPA, Cincinnati, OH; T.J. Norberg King, USEPA, Duluth,
MN; and P.V. Winger, NBS, Athens, GA.
The principal authors of this document are C.G. Ingersoll, G.T. Ankley, G.A.
Burton, F.J. Dwyer, R.A. Hoke, T.J. Norberg-King, and P.V. Winger. Contributors
to specific sections of this manual are listed below.
1. Sections 1-9; General Guidelines
G.T. Ankley, USEPA, Duluth, MN
G.A. Burton, Wright State University, Dayton, OH
F.J. Dwyer, NBS, Columbia, MO
R.A. Hoke, SAIC, Hackensack, NJ
C.G. Ingersoll, NBS, Columbia, MO
T.J. Norberg-King, USEPA, Duluth, MN
C.E. Schlekat, SAIC, Narragansett, Rl
K.J. Scott, SAIC, Narragansett, Rl
2. Sections 10-13; Culture and Test Methods
G.T. Ankley, USEPA, Duluth, MN
D.A. Benoit, USEPA, Duluth, MN
E.L. Brunson, NBS, Columbia, MO
F.J. Dwyer, NBS, Columbia, MO
I.E. Greer, NBS, Columbia, MO
R.A. Hoke, SAIC, Hackensack, NJ
C.G. Ingersoll, NBS, Columbia, MO
P.F. Landrum, NOAA, Ann Arbor, Ml
H. Lee, USEPA, Newport, OR
T.J. Norberg-King, USEPA, Duluth, MN
P.V. Winger, NBS, Athens, GA
XII
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Acknowledgments (continued)
3. Section 14; Statistical Analysis
J. Heltshe, SAIC, Narragansett, Rl
R.A. Hoke, SAIC, Hackensack, NJ
H. Lee, USE PA, Newport, OR
T.J. Norberg-King, USEPA, Duluth, MN
C. Schlekat, SAIC, Narragansett, Rl
4. Section 15; Precision and Accuracy
G.T. Ankley, USEPA, Duluth, MN
G.A. Burton, Wright State University, Dayton, OH
C.G. Ingersoll, NBS, Columbia, MO
T.J. Norberg-King, USEPA, Duluth, MN
Review comments from the following individuals are gratefully acknowledged: C.
Philbrick Barr and P. Nolan, Region 1, Lexington, MA; D. Reed, Permits Division,
OWEC, Washington, D.C.; P. Crocker, Technical Section and S. McKinney,
Marine and Estuarine Section, Region 6, Dallas, TX; F. Schmidt, Monitoring
Branch, OWOW, Washington, D.C.; T. Armitage, Standards and Applied Science
Division, OST, Washington, D.C., T. Bailey, Environmental Effects Branch and J.
Smrchek, OPPT, Washington, D.C.; D. Klemm, EMSLand L Cast, TAI, Newtown,
OH; G. Hanson, OSW, Washington, D.C., S. Ferraro and R. Swartz, ERL-N,
Newport, OR; J. Arthur, R. Spehar, and C. Stephan, ERL-D, Duluth, MN; J.
Thompson, T. Dawson, J. Jenson, S. Collyard, and J. Juenemann, ILS, Duluth,
MN; J. Scott and C. Scheklat, SAIC, Narragansett, Rl.
Participation by the following laboratories in the round-robin testing is greatly
appreciated: ABC Laboratories, Columbia, MO; Environment Canada, Burlington,
Ontario; EVS Consultants, Vancouver, BC; Michigan State University, East Lan-
sing, 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. Juenenmann, J. Jenson, and J. Denny.
USEPA's Office of Science and Technology provided support for the development
of this manual.
XIII
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Section 1
Introduction
1.1 Significance of Use
1.1.1 Sediment provides habitat for many aquatic or-
ganisms 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
organisms 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 con-
taminants in sediment may be several orders of magni-
tude higher than in the overlying water; however, bulk
sediment concentrations have not been strongly corre-
lated to bioavailability (Burton, 1991). Partitioning or
sorption of a compound between water and sediment
may depend on many factors including: aqueous solubil-
ity, pH, redox, affinity for sediment organic carbon and
dissolved organic carbon, grain size of the sediment,
sediment mineral constituents (oxides of iron, manga-
nese, and aluminum), and the quantity of acid volatile
sulfides in sediment (Di Toro et al., 1990, 1991). Al-
though 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 contaminants for bioaccumula-
tion 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
contaminant 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. In
order to obtain a direct measure of sediment toxicity or
bioaccumulation, laboratory tests have been developed
in which surrogate organisms are exposed to sediments
under controlled conditions. Sediment toxicity tests have
evolved into effective tools providing direct, quantifiable
evidence of biological consequences of sediment con-
tamination that can only be inferred from chemical or
benthic community analyses. The USEPA is developing
a national inventory of contaminated sediment sites.
This inventory will be used to develop a biennial report
to Congress on sediment quality in the United States
required under the Water Resources Development Act
of 1992. The use of consistent sediment testing meth-
ods will provide high quality data needed for the national
inventory and for regulatory programs to prevent, reme-
diate, and manage contaminated sediment (Southerland
etal., 1991).
1.1.3 The objective of a sediment test is to determine
whether contaminants in sediment are harmful to or are
bioaccumulated by benthic organisms. The tests can be
used to measure interactive toxic effects of complex
contaminant mixtures in sediment. Furthermore, knowl-
edge of specific pathways of interactions among sedi-
ments and test organisms is not necessary in order 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 contaminants, (3) compare the sen-
sitivities of different organisms, (4) determine spatial
and temporal distribution of contamination, (5) evaluate
hazards of dredged material, (6) for measuring toxicity
as part of product licensing or safety testing or chemical
approval, (7) rank areas for clean up, and (8) set cleanup
goals and estimate the effectiveness of remediation or
management practices.
1.1.4 Results of toxicity tests on sediments spiked at
different concentrations of contaminants 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
(lowest observed effect concentration). In some cases,
results of bioaccumulation tests may also be reported in
terms of a Biota-sediment Accumulation Factor (BSAF)
(Ankley et al., 1992a; Ankley et al., 1992b).
1.1.5 Evaluating effect concentrations for chemicals in
sediment requires knowledge of factors controlling their
bioavailability. Similar concentrations of a chemical in
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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
correlated to interstitial water concentrations, and effect
concentrations in interstitial water are often similar to
effect concentrations in water-only exposures. The bio-
availability of nonionic organic compounds in sediment
is often inversely correlated with the organic carbon
concentration. Whatever the route of exposure, these
correlations of effect concentrations to interstitial water
concentrations indicate that predicted or measured con-
centrations 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 concentrations (Di Toro et al., 1991).
1.1.6 Field surveys can be designed to provide either a
qualitative reconnaissance of the distribution of sedi-
ment contamination or a quantitative statistical compari-
son of contamination among sites. Surveys of sediment
toxicity or bioaccumulation are usually part of more
comprehensive analyses of biological, chemical, geo-
logical, and hydrographic data. Statistical correlations
may be improved and sampling costs may be reduced if
subsamples are taken simultaneously for sediment tests,
chemical analyses, and benthic community structure.
1.1.7 Table 1.1 lists several approaches the USEPA
has considered for the assessment of sediment quality
(USEPA, 1992c). These approaches include (1) equilib-
rium partitioning, (2) tissue residues, (3) interstitial water
toxicity, (4) whole-sediment toxicity and sediment-spiking
tests, (5) benthic community structure, and (6) Sediment
Quality Triad and Effects Range Median (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 parti-
tioning), descriptive (e.g., whole-sediment toxicity tests),
or a combination of numeric and descriptive approaches
(e.g., Effects Range Median; USEPA, 1992c). Numeric
methods can be used to derive chemical-specific sedi-
ment quality criteria (SQC). Descriptive methods such
as toxicity tests with field-collected sediment cannot be
used alone to develop numerical SQC for individual
chemicals. Although each approach can be used to
make site-specific decisions, no one single approach
can adequately address sediment quality. Overall, an
integration of several methods using the weight of evi-
dence is the most desirable approach for assessing the
effects of contaminants associated with sediment {Long
and Morgan, 1991). Hazard evaluations integrating data
from laboratory exposures, chemical analyses, and
benthic community assessments provide strong comple-
mentary evidence of the degree of pollution-induced
degradation in aquatic communities (Chapman et al.,
1992; Burton, 1991).
1.2 Program Applicability
1.2.1 The USEPA has authority under a variety of
statutes to manage contaminated sediment. Until re-
cently, the USEPA has not addressed sediment quality
except in relation to disposal of material removed during
navigational dredging (Table 1.2). Southerland et al.
(1992) outlined four goals of an USEPA management
strategy for contaminated sediments: (1) in-place sedi-
ment should be protected from contamination to ensure
beneficial uses of surface waters, (2) protection of in-place
sediment should be achieved through pollution preven-
tion and source control, (3) in-place remediation should
be limited to locations where natural recovery will not
occur in an acceptable period of time, and (4) consistent
methods should be used to trigger regulatory decisions.
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 goal of the CWA is
to restore and maintain physical, chemical, and biologi-
cal integrity of the nation's waters (Southerland et al.,
1992). Federal and state monitoring programs tradition-
ally have focused on evaluating water column problems
caused by point-source dischargers. The USEPA is
developing a national inventory of contaminated sedi-
ment sites. This inventory will be used to develop a
biennial report to Congress on sediment quality in the
United States required under the Water Resources De-
velopment Act of 1992. The use of consistent sediment
testing methods will provide high quality data needed for
the national inventory and for regulatory program to
prevent, remediate, and manage contaminated sedi-
ment (Southerland et al., 1992).
1.2.3 The Office of Water (OW), the Office of Pesticide
Programs (OPP), the Office of Pollution Prevention and
Toxics (OPPT), the Office of Solid Waste (OSW), and
the Office of Emergency and Remedial Response
(OERR) are all committed to the principle of consistent
tiered testing outlined in the Agency-wide Contaminated
Sediment Strategy (Southerland et al., 1992).
Agency-wide consistent testing is desirable because all
USEPA programs will use similar methods to evaluate
whether a sediment poses an ecological or human
health risk, and comparable data would be produced. It
will also provide the basis for uniform cross-program
decision-making within the USEPA. Each program will,
however retain the flexibility of deciding whether identi-
fied risks would trigger regulatory actions.
1.2.4 Tiered testing should include a hierarchy of tests
with the tests in each successive tier becoming progres-
sively more rigorous, complex, and costly (Southerland
et al, 1992). Guidance needs to be developed to explain
how information within each tier would trigger regulatory
action. The guidance could be program specific, de-
scribing decisions based on a weight of evidence ap-
proach, a pass-fail approach, or comparison to a refer-
ence site depending on statutory and regulatory require-
ments. There are now two approaches for sediment
-------
Table 1.1 Sediment Quality Assessment Procedures'
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
Apparent Effects Threshold "
-------
Table 1.2 Statutory Needs for Sediment Quality Assessment'
Law Area of Need
CERCLA • Assess need for remedial action with contaminated sediments; assess degree of cleanup required, disposition of
sediments
CWA
FIFRA
MPRSA
NEPA
TSCA
RCRA
NPDES permitting, especially under Best Available Technology (BAT) in water-quality-hmited 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 Corps of Engineers)
Review uses of new and existing chemicals
Pesticide labeling and registration
Permits for ocean dumping
Preparation of environmental impact statements for projects with surface water discharges
Section 5: Premanufacture notice reviews for new chemicals
Sections 4, 5, and 6: Reviews for existing chemicals
Assess suitability (and permit) on-land disposal or beneficial use of contaminated sediments considered "hazardous"
Modified from Dickson et al. 1984, and Southerland et al. 1992.
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.
lation kinetics of different classes of compounds during
28-d exposures with L variegatus.
1.3.2 Additional research and methods development
are now in progress to (1) develop standard chronic
sediment toxicity tests (e.g., 28-d exposures with H.
azteca}, (2) refine formulated sediment, (3) refine sedi-
ment dilution procedures, (4) refine sediment Toxicity
Identification Evaluation (TIE) procedures (Ankley and
Thomas, 1992), (5) refine sediment spiking procedures,
and (6) produce additional data on confirmation of re-
sponses in laboratory tests with natural populations of
benthic organisms. This information will be described in
future editions of this manual.
1.3.3 This methods manual serves as a companion to
the marine sediment testing method manual (USEPA,
1994a).
1.3.4 Procedures described in this manual are based on
the following documents: ASTM (1994a), Ankley et al.
(1993), Phipps et al. (1993), Call et al. (1994), Lee et al.
(1994), and USEPA (I993a). This manual outlines spe-
cific test methods for evaluating the toxicity of sediments
with H. azteca and C. tentans. Because additional re-
search is still needed on the standardization of bioaccu-
mulation procedures with sediments, this manual out-
lines only general guidance on procedures for evaluat-
ing the bioaccumulation of contaminants in sediment
with L variegatus. Many of the critical issues necessary
for interpretation of test results are the subject of con-
tinuing research including the influence of feeding on
bioavailability, nutritional requirements of the test organ-
isms, additional performance criteria for organism health,
and confirmation of responses in laboratory tests with
natural benthos populations. See Section 4 for addi-
tional details.
1.3.5 General procedures described in this manual
might be useful for conducting tests with other aquatic
organisms; however, modifications may be necessary.
Altering the procedures described in this manual may
alter bioavailability and produce results that are not
directly comparable with results of acceptable proce-
dures. Comparison of results obtained using modified
versions of these procedures might provide useful infor-
mation concerning new concepts and procedures for
conducting sediment tests with aquatic organisms (e.g.,
Diporeia spp., Tubifex tubifex, Hexagenia spp.). If tests
are conducted with procedures different from those de-
scribed in this manual, additional tests are required to
determine comparability of results.
1.3.6 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 re-
quire their use or prohibit importation of the recom-
mended species. Where state regulations prohibit im-
-------
portation or use of the recommended test species, per-
mission should be requested from the appropriate regu-
latory agency before their using indigenous species.
1.3.7 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 se-
lected are at least as sensitive and appropriate as the
recommended species.
1.3.8 Selection of Test Organisms
1.3.8.1 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 organ-
ism should (1) have a lexicological database demon-
strating relative sensitivity and discrimination to a range
of contaminants of interest in sediment; (2) have a
database for interlaboratory comparisons of procedures
(e.g., round-robin studies); (3) be in direct contact with
sediment; (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 similar to
organisms of concern (e.g., similar feeding guild or
behavior to the indigenous organisms); (9) be tolerant of
a broad range of sediment phsyico-chemical character-
istics (e.g., grain size); and (10) be compatible with
selected exposure methods and endpoints (Table 1.3,
ASTM, 1993a). 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 1.3.8.8 and 1.3.9.6).
Table 1.3 Rating of Selection Criteria for Freshwater Sediment Toxicity Testing Organisms1
Criterion Hyalella Diporeia Chironomus Chironomus Lumbriculus Tubifex Hexagenia Mollusks Daphma spp. and
azfeca spp. tentans ripanus variegatus tubifex spp. Cenodaphnia spp.
Relative
sensitivity
toxicity + - + +
database
Round-robin
studies + - + -----
conducted
Contact with +++ + + + + +
sediment
Laboratory +-+ + + + .. +
culture
Taxonomic +/- */- +/- +/- + + + + +
identification
Ecological +++ + + + + .,. +
importance
Geographical + +/- + + + + + + +/-
distribution
Sediment
physico-
chemical
tolerance
NA
Response
confirmed + +
with benthos
populations
Peer reviewed + +
Endpoints2 S. G,M S, B, A
monitored
S,G, E
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
-------
1.3.8.2 Of these criteria (Table 1.3), a database demon-
strating relative sensitivity to contaminants, contact with
sediment, ease of culture in the laboratory, interlabora-
tory comparisons, tolerance to varying sediment phsyico-
chemical characteristics, and confirmation with responses
of natural benthos 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 standard-
ized testing procedures for benthic organisms. A similar
database must be developed in order for other organ-
isms to be included in future editions of this manual
(e.g.. mayflies (Hexagenia spp.), other midges (C.
riparius). other amphipods (Diporeia spp.), cladocerans
(Daphnia magna, Ceriodaphnia dubia), or mollusks).
1.3.8.3 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. (1991b) 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 am-
phipod or the cladoceran. West et al. (1993) found the
rank sensitivity of the three species to the lethal effects
of copper-contaminated 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 sensitive than H. azteca, C. dubia, or
Pimephales promelas to cadmium, nickel, zinc, copper,
and lead (Schubauer-Berigan et al., 1993). Of the latter
Table 1.4
Cnemical
Water-only, 10-d LC50 (ng/L) Values for Hyalella
azteca, Chironomus tentans, and Lumbriculus
variegatus'
H. azteca
C. tentans L variegatus
Copper
Zmc
Cadmium
N'Ckel
Lead
P.P -DDT
p.p-DDD
p.p-DDE
Dieidnn
Chlorpyrifos
35
73
2.83
780
<16
0.07
0.17
1.39
7.6
0.086
54
1.1252
NT-
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 CaCO, at pH 7.8 to 8.2 (Phipps et al., 1994).
50°/b mortality at highest concentration tested.
70% mortality at lowest concentration tested.
NT = rot tested.
three species, no one was consistently the most sensi-
tive to all five metals.
1.3.8.3.1 In a study of contaminated Great Lakes sedi-
ment, H. azteca, C. tentans, and C. riparius were among
the most sensitive and discriminatory of 24 organisms
tested (Burton and Ingersoll, 1994; Ingersoll et al., 1993).
Kemble et al. (1993) found the rank sensitivity of four
species to 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.
1.3.8.3.2 In 10-d water-only and whole-sediment tests,
H. azteca and C. tentans were more sensitive than D.
magnatefluoranthene (Suedel et al. 1993).
1.3.8.3.3 Ten-day, water-only tests also have been
conducted with a number of chemicals using the three
species described in this manual (Phipps et al., 1994;
Table 1.4). All tests were flow-through exposures using
a soft natural water (Lake Superior) with measured
chemical concentrations that, other than the absence of
sediment, were conducted under conditions (e.g., tem-
perature, photoperiod, feeding) similar to those being
described for the standard 10-d sediment test. In gen-
eral, H. azteca was more sensitive to copper, zinc,
cadmium, nickel and lead than either C. tentans or L.
variegatus. Conversely, the midge was usually compa-
rable to or more sensitive than the amphipod to the
pesticides tested. Lumbriculus variegatus was not tested
with several of the pesticides; however, in other studies
with whole sediments contaminated by DDT and associ-
ated 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 multiple
test organisms when performing sediment assessments.
Second, L. variegatus appears to be relatively insensi-
tive to most of the test chemicals, which perhaps is a
positive attribute for an organism used in bioaccumula-
tion tests.
1.3.8.3.4 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 infor-
mation from the AQUIRE database (AQUIRE, 1992) for
DDD and DDE, were compared to assay results for the
three species (Phipps et al., 1994). The sensitivity of H.
azteca to metals and pesticides, and C. tentans to
pesticides was comparable to chronic toxicity data gen-
erated 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. Inter-
estingly, in some instances (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 obser-
vation 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.
1.3.8.4 Relative species sensitivity frequently varies
among contaminants; consequently, a battery of tests
including 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; USEPA, 1989b). 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 organisms was the
most sensitive to all of the metals.
1.3.8.5 The sensitivity of an organism to contaminants
should be balanced with the concept of discrimination
(Burton and Ingersoll, 1994). The response of a test
organism should provide discrimination between differ-
ent levels of contamination.
1.3.8.6 The sensitivity of an organism is related to the
route of exposure and biochemical response to contami-
nants. Sediment-dwelling organisms can receive expo-
sure via three primary sources: interstitial water, sedi-
ment particles, and overlying water. Food type, feeding
rate, assimilation efficiency, and clearance rate will con-
trol the dose of contaminants from sediment. Benthic
invertebrates often selectively consume different par-
ticle sizes (Harkey et al., 1994) or particles with higher
organic carbon concentrations that may have higher
contaminant concentrations. Grazers and other
collector-gatherers that feed on aufwuchs and detritus
may receive most of their body burden directly from
materials attached to sediment or from actual sediment
ingestion. In amphipods (Landrum, 1989) and clams
(Boese et al., 1990) uptake through the gut can exceed
uptake across the gills of certain hydrophobic com-
pounds. Organisms in direct contact with sediment may
also accumulate contaminants by direct adsorption to
the body wall or by absorption through the integument
(Knezovichetal., 1987).
1.3.8.7 Despite the potential complexities in estimating
the dose that an animal receives from sediment, the
toxicity and bioaccumulation of many contaminants in
sediment such as Kepone®, fluoranthene, organochlo-
rines, and metals have been correlated with either the
concentration of these chemicals in interstitial water or
in the case of nonionic organic chemicals, concentra-
tions in sediment on an organic carbon normalized basis
(Di Toro et al., 1990; Di Toro et at., 1991). The relative
importance of whole sediment and interstitial water routes
of exposure depends on the test organism and the
specific contaminant (Knezovich et al., 1987). Because
benthic communities contain a diversity of organisms,
many combinations of exposure routes may be impor-
tant. Therefore, behavior and feeding habits of a test
organism can influence its ability to accumulate con-
taminants from sediment and should be considered
when selecting test organisms for sediment testing.
1.3.8.8 The use of H. azteca and C. tentans in labora-
tory toxicity studies has been confirmed with natural
benthos populations.
1.3.8.8.1 Chironomids were not found in sediment
samples that decreased growth of C. tentans by 30% or
more in 10-d laboratory toxicity tests (Giesy et al., 1988).
Wentsel et al. (1977a, I977b, 1978) reported a correla-
tion between effects on C. tentans in laboratory tests
and the abundance of C. tentans in metal-contaminated
sediments.
1.3.8.8.2 Benthic community evaluations and laboratory
tests with H. azteca both provided evidence of
metal-induced degradation of aquatic communities in
the Clark Fork River (Canfield et al., 1994). Total abun-
dance of benthic organisms did not follow a consistent
pattern when compared to metals in sediment samples.
The number of chironomid genera was higher at stations
that showed reduced growth or sexual maturation of H.
azteca in laboratory sediment tests and had higher
concentrations of metals in sediment.
1.3.8.8.3 The results from laboratory sediment toxicity
tests were compared to colonization of artificial sub-
strates exposed in situ to contaminated Great Lakes
sediment (Burton and Ingersoll, 1994). Survival or growth
of H. azteca and C. tentans in 10- to 28-d laboratory
exposures were negatively correlated to percent chi-
ronomids and percent tolerant taxa colonizing artificial
substrates in the field. Scklekat et al. (1994) reported
generally good agreement between sediment tests with
H. azteca and benthic community responses in the
Anacostia River, Washington, D.C.
1.3.8.8.4 Sediment toxicity to amphipods in 10-d toxicity
tests, field contamination, and field abundance of benthic
amphipods were examined along a sediment contami-
nation gradient of DDT (Swartz et al., 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 H. azteca, was negatively correlated
to DDT concentrations. The threshold for 10-d sediment
toxicity in laboratory studies was about 300 pg DDT
(+metabolites)/g organic carbon. The threshold for abun-
dance of amphipods in the field was about 100 pg DDT
(+metabolites)/g organic carbon. Therefore, correlations
between toxicity, contamination, and field populations
indicate that acute sediment toxicity tests can provide
reliable evidence of biologically adverse sediment con-
-------
tamination in the field, but may be underprotective of
chronic effects.
1.3.9 Selection of Organisms for
Sediment Bioaccumulation Testing
1.3.9.1 Several studies have demonstrated that hydro-
phobic organic compounds are bioaccumulated from
sediment by freshwater infaunal organisms including
larval insects (C. tentans, Adams et al., 1985; Adams,
1987; Hexagenia limbata, Gobas et al., 1989), oligocha-
etes (Tubifex tubifexand Limnodritus hoffmeisteri; Oliver,
1984, Oliver, 1987; Connell et al., 1988), and by marine
organisms (polychaetes, Nephtys incisa; mollusks,
Mercenaria mercenaria, Yoldia limatula; Lake et al.,
1990). Consumers of these benthic organisms may
bioaccumulate or biomagnify contaminants. Therefore,
in addition to sediment toxicity, it may be important to
examine the uptake of chemicals by aquatic organisms
from contaminated sediments.
1.3.9.2 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 organ-
isms throughout the year, (2) known chemical exposure
history, (3) adequate tissue mass for chemical analyses,
(4) ease of handling, (5) tolerance of a wide range of
sediment phsyico-chemical characteristics (e.g., particle
size), (6) low sensitivity to contaminants associated with
sediment (e.g., metals, organics), (7) amenability to
long-term exposures without adding food, (8) and ability
to accurately reflect concentrations of contaminants in
field-exposed organisms (e.g., exposure is realistic).
With these criteria in mind, the advantages and disad-
vantages of several potential freshwater taxa for bioac-
cumulation testing are discussed below.
1.3.9.3 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
contaminants compared to 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, chirono-
mids 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 continu-
ously cultured in the laboratory and consequently are
not always available for testing. Furthermore, the back-
ground concentrations of contaminants 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 expo-
sure. However, their size may be insufficient for residue
analysis and H. azteca are sensitive to contaminants in
sediment. Fish (e.g., fathead minnows) provide an ad-
equate tissue mass, are readily available, are easy to
handle, and can be used in long-term exposures. How-
ever, the route of exposure is not appropriate for evalu-
ating the bioavailability of sediment-associated contami-
nants to benthic organisms.
1.3.9.4 Oligochaetes are infaunal benthic organisms
that meet many of the test criteria listed above. Certain
oligochaete species are easily handled and cultured,
provide reasonable biomass for residue analyses, and
are tolerant of varying sediment physical and chemical
characteristics. Oligochaetes are exposed to contami-
nants via all appropriate routes of exposure including
pore water and ingestion of sediment particles. Oli-
gochaetes need not be fed during long-term bioaccumu-
lation exposures (Phipps et al., 1993). Various oligocha-
ete species have been used in toxicity and bioaccumula-
tion evaluations (Chapman et al., 1982a, Chapman et
al., 1982b; Wiederholm, 1987; Kielty et al., 1988a, Kielty
et al., !988b; 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; Robbins et al., 1989; Ankley et
al., 1992b; E.L Brunson, NBS, Columbia, MO, unpub-
lished data).
1.3.9.5 Lumbriculus variegatus does not biotransform
PAHs (Harkey et al., 1994b).
1.3.9.6 The response of L. variegatus in laboratory
bioaccumulation studies has been confirmed with natu-
ral populations of Oligochaetes.
1.3.9.6.1 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 simi-
lar between laboratory-exposed and field-collected Oli-
gochaetes. The more highly chlorinated PCBs tended to
have greater bioaccumulation in the field-collected or-
ganisms. In contrast, total PCBs in laboratory-exposed
(Pimephales promelas) and field-collected (Ictalurus
melas) fish revealed poor agreement in bioaccumulation
relative to the sediment concentrations at the same
sites.
1.3.9.6.2 Bioaccumulation of laboratory-exposed L
variegatus and field-collected Oligochaetes from the same
sites were also compared (E.L. Brunson, NBS, Colum-
bia, MO, unpublished data). Select PAH and DDT peak
concentrations were similar in field-collected Oligocha-
etes and L. variegatus exposed for 28 d in the labora-
tory.
1.4 Performance-based Criteria
1.4.1 USEPA's Environmental Monitoring Man-
agement Council (EMMC) recommended the use
of performance-based methods in developing
8
-------
chemical analytical standards (Williams, 1993).
Performance-based methods were defined by EMMC as
a monitoring approach that permits the use of appropri-
ate methods that meet pre-established demonstrated
performance standards (Section 9.2).
1.4.2 The USEPA Office of Water, Office of Science
and Technology, and Office of Research and Develop-
ment held a workshop on September 16-18, 1992 in
Washington, 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 development of standard freshwater and marine
sediment testing procedures (USEPA, I992a and Ap-
pendix A). Workgroup participants reached a consensus
on several culturing and testing methods. In developing
guidance for culturing freshwater 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 ap-
proach 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, and 13.4 for a listing of perfor-
mance criteria for culturing and testing.
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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
toxicity or bioaccumulation of contaminants associated
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. The 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 renewed
daily and test organisms are fed during the toxicity tests.
The endpoint in the toxicity test with H. azteca is survival
and the endpoints in the toxicity test with 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 with
H. azteca. Guidance for conducting 28-d bioaccumula-
tion tests with the oligochaete Lumbriculus variegatus is
provided in this manual. The overlying water is renewed
daily and the test organisms are not fed during bioaccu-
mulation tests. This guidance describes are also de-
scribed for determining bioaccumulation kinetics of dif-
ferent classes of compounds during 28-d exposures
with L variegatus.
2.1.2 Experimental Design
The following section is a general summary of experi-
mental design. See Section 14 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 and 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 (Lee et al., 1994). 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 collected near an area of concern and is used to
assess sediment conditions exclusive of material(s) of
interest. Testing a reference sediment provides a
site-specific basis for evaluating toxicity.
2.1.2.1.2 Natural geomorphological and physicochemi-
cal characteristics such as sediment texture may influ-
ence the response of test organisms (DeWitt et al.,
1988). The physicochemical characteristics of test sedi-
ment 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
including grain size and organic carbon can be evalu-
ated if the limits are exceeded in a test sediment. See
Section 10.1 for information on physicochemical re-
quirements of test organisms. If the physicochemical
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 characteris-
tics. The effects of sediment characteristics on the re-
sults of sediment tests may be able to be addressed with
regression equations (DeWitt et al., 1988; Ankley et al.,
I994a). The use of formulated sediment can also be
used to evaluate physicochemical characteristics of sedi-
ment on test organisms (Walsh et al., 1991; Suedel and
Rodgers, 1994).
2.1.2.2 The experimental design depends on the pur-
pose 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
characteristics. For instance, 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 chemical. A useful summary of
field sampling design is presented by Green (1979). See
Section 14 for additional guidance on experimental de-
sign 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.,
ANOVA), 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).
10
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2.1.2.4 In other instances, the purpose of the study
might be to conduct a quantitative sediment survey to
determine statistically significant differences between
effects among control and test sediments from several
sites. The number of replicates/site should be based on
the need for sensitivity or power (Section 14). In a
quantitative survey, replicates (separate samples from
different grabs collected at the same site} would need to
be taken at each site. Chemical and physical character-
izations of each of these grabs would be required for
each of these replicates used in sediment testing. Sepa-
rate 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, 1994a).
2.1.2.5 Sediments often exhibit high spatial and tempo-
ral variability (Stemmer et al., 1990a). Therefore, repli-
cate samples may need to be collected to determine
variance in sediment characteristics. Sediment should
be collected with as little disruption as possible; how-
ever, subsampling, compositing, or homogenization of
sediment samples may be necessary for some experi-
mental 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. Comparisons can be made
in both space and time. In pre-dredging studies, a
sampling design can be prepared to assess the con-
tamination of samples representative of the project area
to be dredged. Such a design should include subsampling
cores taken to the project depth.
2.1.2.7 The primary focus of the physical and experi-
mental 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 independently assigned (Steel and Torrie, 1980) and
to which air and water exchange between test chambers
are kept to a minimum. As the number of test chambers/
treatment increases, the number of degrees of freedom
increases, and, therefore, the width of the confidence
interval on a point estimate, such as an LC50, de-
creases, and the power of a significance test increases
(Section 14). Because of factors that might affect results
within test chambers and results of a test, all test cham-
bers should be treated as similarly as possible. Treat-
ments should be randomly assigned to individual test
chamber locations. Assignment of test organisms to test
chambers should be non-biased.
2.2 Types of Tests
2.2.1 Toxicity methods are outlined for two organisms,
the amphipod H. azteca (Section 11) and the midge C.
tentans (Section 12). This manual primarily describes
methods for testing freshwater sediments; however, the
methods described can also be used for testing H.
azteca in estuarine sediments (up to 15 %o salinity).
2.2.2 Guidance for conducting 28-d bioaccumulation
tests with the oligochaete L variegatus is described in
Section 13. Methods are also described for determining
bioaccumulation kinetics of different classes of com-
pounds during 28-d exposures with L. variegatus.
2.3 Test Endpoints
2.3.1 The endpoints measured in the toxicity tests are
survival or growth (the growth endpoint is optional in the
H. azteca test). 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).
11
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Section 3
Definitions
3.1 Terms
The following terms were defined in Lee (1980), NRC
(1989), USEPA (1989C), USEPA-USCOE (1991),
USEPA-USCOE (1994), Leeetal. (1994), ASTM(1993b),
or ASTM (1994a).
3.1.1 Technical Terms
3.1.1.1 Sediment. Participate material that usually lies
below water. Formulated particulate material that is
intended to lie below water in a test.
3.1.1.2 Contaminated sediment. Sediment containing
chemical substances at concentrations that pose a known
or suspected threat to environmental or human health.
3.1.1.3 Whole sediment. Sediment and associated
pore water that have had minimal manipulation. The
term bulk sediment has been used synonymously with
whole sediment.
3.1.1.4 Control sediment. A sediment that is essen-
tially free of contaminants and is used routinely to as-
sess the acceptability of a test. Any contaminants in
control sediment may originate from the global spread of
pollutants and does 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.
3.1.1.5 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 near the site
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 dredge material evalua-
tions.
3.1.1.6 interstitial water or pore water. Water occupy-
ing space between sediment or soil particles.
3.1.1.7 Spiked sediment. A sediment to which a mate-
rial has been added for experimental purposes.
3.1.1.8 Reference toxicity test A test with a high-grade
reference material conducted in conjunction with sedi-
ment tests to determine possible changes in condition of
the test organisms. Deviations outside an established
normal range indicate a change in the condition of the
test organism population. Reference-toxicity tests are
most often performed in the absence of sediment.
3.1.1.9 Clean. Denotes a sediment or water that does
not contain concentrations of test materials which cause
apparent stress to the test organisms or reduce their
survival.
3.1.1.10 Overlying water. The water placed over sedi-
ment in a test chamber during a test.
3.1.1.11 Concentration. The ratio of weight or volume
of test material(s) to the weight or volume of sediment.
3.1.1.12 No-observable-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 significant different
from the controls).
3.1.1.13 Lowest-observable-effect concentration
(LOEC). The lowest concentration of a toxicant to which
organisms 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).
3.1.1.14 Lethal concentration (LC). The toxicant con-
centration that would cause death in a given percent of
the test population. Identical to EC when the observable
adverse effect is death. For example, the LC50 is the
concentration of toxicant that would cause death in 50%
of the test population.
3.1.1.15 Effect concentration (EC). The toxicant con-
centration that would cause an effect in a given percent
of the test population. Identical to LC when the observ-
able adverse effect is death. For example, the EC50 is
the concentration of toxicant that would cause death in
50% of the test population.
12
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3.1.1.16 Inhibition concentration (1C). The toxicant
concentration that would cause a given percent reduc-
tion in a non-quantal measurement for the test popula-
tion. For 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 Biota-sediment accumulation factor (BSAF).
The ratio of tissue residue to source concentration (e.g.,
sediment at steady state normalized to lipid and sedi-
ment organic carbon).
3.1.1.18 Bioaccumulation. The net accumulation of a
substance by an organism as a result of uptake from all
environmental sources.
3.1.1.19 Bioaccumulation factor. Ratio of tissue resi-
due to contaminant source concentration at steady-state.
3.1.1.20 Bioaccumulation potential. Qualitative as-
sessment of whether a contaminant is bioavailable.
3.1.1.21 Bioconcentration. The net assimilation of a
substance by an aquatic organism as a result of uptake
directly from aqueous solution.
3.1.1.22 Bioconcentration factor (BCF). Ratio of tis-
sue residue to water contaminant concentration at
steady-state.
3.1.1.23 Depuration. Loss of a substance from an
organism as a result of any active (e.g., metabolic
breakdown) or passive process when the organism is
placed into an uncontaminated environment. Contrast
with Elimination.
3.1.1.24 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.
3.1.1.25 fcr. Uptake rate coefficient from the aqueous
phase, with units of g-water x g-tissue'1 x time'1. Con-
trast with kc.
3.1.1.26 k . Sediment uptake rate coefficient from the
sediment pnase, with units of g-sediment x g-tissue"' x
time"1. Contrast with k..
3.1.1.27
time"1.
Elimination rate constant, with units of
3.1.1.28 Kinetic Bioaccumulation Model. Any model
that uses uptake and/or elimination rates to predict
tissue residues.
3.1.1.29
cient.
K . Organic carbon-water partitioning coeffi-
3.1.1.30 Kow. Octanol-water partitioning coefficient.
3.1.1.31 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
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Section 4
Interferences
4.1 General Introduction
4.1.1 Interferences are characteristics of a sediment or
sediment test system that can potentially affect test
organism survival aside from those related to
sediment-associated contaminants. These interferences
can potentially confound interpretation of test results in
two ways: (1) toxicity is observed in the test when
contamination is not present, or there is more toxicity
than expected; and (2) no toxicity or bioaccumulation is
observed when contaminants are present at elevated
concentrations, or there is less toxicity or bioaccumula-
tion than expected.
4.1.2 There are three categories of interfering factors:
those characteristics of sediments affecting survival in-
dependent of chemical concentration (i.e.,
non-contaminant factors); changes in chemical bioavail-
ability as a function of sediment manipulation or storage;
and 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 sedi-
ments, extrapolation from laboratory studies to the field
can sometimes be difficult (Table 4.1; Burton, 1991).
Sediment collection, handling, and storage may alter
bioavailability and concentration by changing the physi-
cal, chemical, or biological characteristics of the sedi-
ment. Maintaining the integrity of a field-collected sedi-
ment during removal, transport, mixing, storage, and
testing is extremely difficult and may complicate the
interpretation of effects. Direct comparisons of organ-
isms exposed in the laboratory and in the field would be
useful to verify laboratory results. However, spiked sedi-
ment may not be representative of contaminated sedi-
ment in the field. Mixing time (Stemmer et al., 1990a)
and aging (Word et al., 1987; Landrum, 1989; Landrum
and Faust, 1992) of spiked sediment can affect re-
sponses of organisms.
4.1.3.1 Laboratory sediment testing with field-collected
sediments may be useful in estimating cumulative ef-
lects and interactions of multiple contaminants in a
sample. Tests with field samples usually cannot dis-
criminate between effects of individual chemicals. Most
sediment samples contain a complex matrix of inorganic
Table 4.1 Advantages and Disadvantages for Use of Sediment
Tests'
Advantages
Measure bioavailable fraction of contaminant(s).
Provide a direct measure of benthic effects, assuming no field
adaptation or amelioration of effects.
Limited special equipment is required.
Methods are rapid and inexpensive.
Legal and scientific precedence exists for use; ASTM standard
guides are available.
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 sediments.
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 ecologi-
cal effects.
' Modified from Swartz (1989}
and organic contaminants with many unidentified com-
pounds. The use of Toxicity 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). Sedi-
14
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ment spiking can also be used to investigate additive,
antagonistic, or synergistic effects of specific contami-
nant mixtures in a sediment sample (Swartz et al.,
1988).
4.1.4 Methods that measure sublethal effects are either
not available or have not been routinely used to evaluate
sediment toxicity {Craig, 1984; Dillon and Gibson, 1986;
Ingersoll and Nelson, 1990; Ingersoll, 1991; Burton et
al., 1992). Most assessments of contaminated sediment
rely on short-term-lethality testing methods (e.g., <10 d;
USEPA-USCOE, 1977; USEPA-USCOE, 1991). Short-
term-lethality tests are useful in identifying "hot spots" of
sediment contamination but may not be sensitive enough
to evaluate moderately contaminated areas. However,
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). Additional
methods development of chronic sediment testing pro-
cedures and culturing of infaunal organisms with a vari-
ety of feeding habits including suspension and deposit
feeders is needed.
4.1.5 Despite the interferences discussed in this sec-
tion, existing sediment testing methods can be used to
provide a rapid and direct measure of effects of contami-
nants on benthic communities. Laboratory tests with
field-collected sediment can also be used to determine
temporal, horizontal, or vertical distribution of contami-
nants in sediment. Most tests can be completed within
two to four weeks. Legal and scientific precedents exist
for use of toxicity and bioaccumulation tests in regula-
tory decision-making (e.g., USEPA, 1986a). Further-
more, sediment tests with complex contaminant mix-
tures are important tools for making decisions about the
extent of remedial action for contaminated aquatic sites
and for evaluating the success of remediation activities.
4.2 Non-Contaminant 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.
Photoinduced toxicity caused by ultraviolet (UV) light,
may be important for some compounds associated with
sediment (e.g., polycyclic aromatic hydrocarbons (PAHs);
Davenport and Spacie, 1991; Ankley et al., I994b).
Fluorescent light does not contain UV light, but natural
sunlight does. Lighting can therefore affect lexicological
responses and is an important experimental variable for
photoactivated chemicals. However, lighting typically
used to conduct laboratory tests does not include the
appropriate spectrum of ultraviolet radiation to photoac-
tivate compounds (Oris and Giesy, 1985), and thus
laboratory tests may not account for toxicity expressed
by this mode of action.
4.2.2 Natural geomorphological and physicochemical
characteristics such as sediment texture may influence
the response of test organisms (DeWitt et al., 1988).
The physicochemical characteristics of test sediment
need to be within the tolerance limits of the test organ-
ism. Ideally, the limits of the test organism should be
determined in advance; however, control samples re-
flecting differences in factors such as grain size and
organic carbon can be evaluated if the limits are ex-
ceeded 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..
I994a). The use of formulated sediment can also be
used to evaluate physicochemical characteristics of sedi-
ment on test organisms (Walsh et al., 1991; Suedel and
Rodgers, 1994).
4.2.3 Interferences of tests with each specific species
are described in Tables 11.3, 12.3, and 13.4.
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. Although the
tests are not designed to simulate natural conditions, m
some cases contaminant availability in laboratory toxic-
ity test may be different from what it is representative of
in-place sediments in the field.
4.3.2 Sediment collection, handling, and storage may
alter contaminant bioavailability and concentration by
changing the physical, chemical, or biological character-
istics of the sediment. These manipulation processes
are generally thought to increase availability of organic
compounds because of disruption of the equilibrium with
organic carbon in the pore water/particle system. Simi-
larly, oxidation of anaerobic sediments increases the
availability of certain metals (Di Toro et al., 1990).
Because the availability of contaminants may be a func-
tion of the degree of manipulation, this manual recom-
mends that handling, storage, and preparation of the
sediment for actual testings be as consistent as pos-
sible. If sieving is performed, it is done primarily to
remove predatory organisms and large debris. This
manipulation most likely results in a worst-case condi-
tion of heightened bioavailability yet eliminates preda-
tion 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 sediments on a routine basis.
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 may influ-
ence bioavailability (Stemmer et al., 1990b).
15
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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
bacterial or fungal growth. Without addition of food, the
test organisms may starve during exposures (Ankley et
al., 1994). However, the addition of food may alter the
availability of the contaminants in the sediment
(Wiederholm et al., 1987, Harkey et al., 1994) depend-
ing 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 test
chamber may also influence availability. In most cases,
the organism is a minor sink for contaminants relative to
the sediment. However, within the burrow of an organ-
ism, sediment desorption kinetics may limit uptake rates.
Within minutes to hours, a major portion of the total
chemical may 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 or advection 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 sedi-
ment are unknown. Bulk-sediment chemical concentra-
tions may be normalized to factors other than dry weight.
For example, concentrations of nonionic organic com-
pounds 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 dis-
solved 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
(ASTM, 1994b; K.E. Day Environment Canada,
Burlington, Ontario, personal communication). However,
further research is needed to determine effects on con-
taminant bioavailability or other modifications of sedi-
ments from treatments such as those used to remove or
destroy indigenous organisms.
16
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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
includes (1) the appointment of a laboratory health and
safety officer with the responsibility and authority to
develop and maintain a safety program, (2) the prepara-
tion of a formal written health and safety plan, which is
provided to each laboratory staff member, (3) an ongo-
ing training program on laboratory safety, and (4) regu-
lar safety inspections.
5.1.2 This manual addresses procedures that may
involve hazardous materials, operations, and equipment,
and it does not purport to address all of the safety
problems associated with their use. It is the responsibil-
ity of the user to establish appropriate safety and health
practices, and determine the applicability of regulatory
limitations before use. While some safety considerations
are included in this manual, it is beyond the scope of this
manual to encompass all safety requirements neces-
sary to conduct sediment tests.
5.1.3 Collection and use of sediments may involve
substantial risks to personal safety and health. Contami-
nants in field-collected sediment may include carcino-
gens, mutagens, and other potentially toxic compounds.
Inasmuch as sediment testing is often begun before
chemical analyses can be completed, worker contact
with sediment needs to be minimized by (1) using gloves,
laboratory coats, safety glasses, face shields, and respi-
rators as appropriate, (2) manipulating sediments under
a ventilated hood or in an enclosed glove box, and (3)
enclosing and ventilating the exposure system. Person-
nel 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, inhala-
tion or absorption of corrosive or toxic substances through
skin contact, and asphyxiation because of lack of oxy-
gen or presence of noxious gases.
5.1.4 Before sample collection and laboratory work,
personnel should determine that all required safety equip-
ment 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 safety equipment, such
as rubber aprons, laboratory coats, respirators, 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 fountains.
5.2.2.2 Mobile laboratories should be equipped with a
telephone to enable personnel to summon help in case
of emergency.
5.3 General Laboratory and Field
Operations
5.3.1 Special handling and precautionary guidance in
Material Safety Data Sheets should be followed for
reagents and other chemicals purchased from supply
houses.
5.3.2 Work with some sediments may require compli-
ance with rules pertaining to the handling of hazardous
materials. Personnel collecting samples and performing
tests should not work alone.
5.3.3 It is advisable to wash exposed parts of the body
with soap and water immediately after collecting 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.
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 in a fume hood.
17
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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, teta-
nus, typhoid fever, and polio.
5.5 Safety Manuals
5.5.1 For further guidance on safe practices when
handling 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
Management, 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 iden-
tification rules and land disposal restrictions, and to
protect the air, water and land by minimizing and con-
trolling all releases from fume hoods and bench opera-
tions. Also, compliance is required with any sewage
discharge permits and regulations. For further informa-
tion on waste management, consult "The Waste Man-
agement 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 haz-
ardous 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
regulations. As a handler of hazardous materials, it is
your responsibility to know and comply with the pertinent
regulations applicable in the state in which you are
operating. Refer to The Bureau of National Affairs Inc.,
(1986) for the citations of the federal requirements.
18
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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 "non-toxicant" test
with each potential test species, in which all test cham-
bers contain a controf sediment (sometimes called the
negative control), and clean overlying water for each
organism to be tested. Survival, growth, or reproduction
of the test organism will demonstrate whether facilities,
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.
6.2 Facilities
6.2.1 The facility must include separate areas for cultur-
ing and testing to reduce the possibility of contamination
by test materials and other substances, especially vola-
tile compounds. Holding and culture chambers should
not be in a room in which sediment tests are conducted,
where stock solutions or sediments are prepared, or
equipment is cleaned. Test chambers may be placed in
a temperature-controlled recirculating water bath or a
constant-temperature area. An enclosed test system is
desirable to provide ventilation during tests to limit expo-
sure of laboratory personnel to volatile substances.
6.2.2 Light of the quality and luminance normally ob-
tained in the laboratory is adequate (about 500 to 1000
lux using wide-spectrum fluorescent lights; e.g.,
cool-white or daylight) for culturing and testing. Lux is
the unit selected for reporting luminance 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 nE/rrvVs1 (assuming an average wavelength of
550 nm (umo! -2 S"' = W m x X(nm) x 8.36 x 10'3) (ASTM,
1994c). Luminance should be measured at the surface
of the water. A uniform photoperiod of 16L:8D can be
achieved in the laboratory or in an environmental cham-
ber using automatic timers.
6.2.3 During phases of rearing, holding, and testing,
test organisms should be shielded from external distur-
bances such as rapidly changing light or pedestrian
traffic.
6.2.4 The test facility should be well ventilated and free
of fumes. Air used for aeration should be free of oil and
fumes. Filters to remove oil, water, and bacteria are
desirable. Oil-free air pumps should be used where
possible. Particulates can be removed from the air using
filters such as BALSTON® Grade BX or equivalent
(Balston, Inc., Lexington, MA), and oil and other organic
vapors can be removed using activated carbon filters
(e.g., BALSTON®, C-1 filter, or equivalent). Laboratory
ventilation systems should be checked to ensure that
return air from chemistry laboratories or sample han-
dling 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 potentially contami-
nated air to sensitive areas through open or loosely
fitting doors.
6.3 Equipment and Supplies
6.3.1 Equipment and supplies that contact stock solu-
tions, sediments, 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, polycarbonate, and fluoro-
carbon plastics should be used whenever possible to
minimize leaching, dissolution, and sorption. Concrete
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 deter-
gent, 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 should 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.
19
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6.3.3 General Equipment
6.3.3.1 Environmental chamber or equivalent facility
with photoperiod and temperature control (20 to 25°C).
6.3.3.2 Water purification system capable of producing
at least 1 mega-ohm water (USEPA, 1993a).
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,
instrument electrodes and probes.
6.3.3.11 Glass or electronic thermometers—for mea-
suring 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 for a list of additional equipment
and supplies.
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. 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). The water-delivery system should be cali-
brated before the test by determining 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.
Therefore, 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 silicone 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, and 13.
6.3.6 Cleaning
6.3.6.1 All non-disposable sample containers, test cham-
bers, and other equipment mat 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 ac-
etone to remove organic compounds (use a fume
hood or canopy).
6. Rinse three times with deionized water.
20
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Table 6.1 Equipment and Supplies for Culturing and Testing Specific Test Organisms
A. Biological Supplies
Brood stock of test organisms
Active dry yeast (HA)
Cerophyl® (dried cereal leaves; HA)
Trout food pellets (HA)
Tetrafin® goldfish food (CT)
Trout starter (LV)
Helisoma sp. snails (optional; LV)
Algae (e.g., Selenastrum capricornutum, Chlorelta;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 oishes or beakers (200- to 300-mL; CT)
Erlenmeyer flasks (250 and 500 mL; CT)
Larval rearing chambers (e.g., I9-L capacity; CT)
1/4" glass tubing (for aspirating flask; CT)
Glass bowls (20-cm diameter; LV)
Glass vials (10mL; 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)
Photopenod 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, cutler, 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 jim mesh opening; Nytex® 110; HA)
Aluminum-weighing pans
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, MM; 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 (non-phosphate)
Acetone (reagent grade)
Hexane (reagent grade)
Hydrochloric acid (reagent grade)
Chloroform (LV)
Methanol (LV)
Copper Sulfate
Potassium Chloride
Reagents for reconstituting water
Formalin (or Notox®)
Sucrose
HA = Hyalella azteca
CT = Chironomus tentans
LV = Lumbriculus variegatus
6.3.6.2 All test chambers and equipment should be
thoroughly rinsed with the dilution water immediately
before use in a test.
6.3.6.3 Many organic solvents 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, 1988a), but the solution might
attack silicone adhesive and leave chromium residues
on glass. A alternative to use of dichromate-sulfuric acid
could be to heat glassware for 8 h at 450°C.
21
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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
characteristics of the water. See USEPA (1993a) and
ASTM (1994a) 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 or spring, if possible, or from a
surface-water source. If surface water is used, the in-
take should be positioned to (1) minimize fluctuations in
quality and contamination, (2) maximize the concentra-
tion of dissolved oxygen, and (3) ensure low concentra-
tions of sulfide and iron. Municipal-water supplies may
be variable and may contain unacceptably high concen-
trations of materials such as copper, lead, zinc, fluoride,
chlorine, or chloramines. Chlorinated water should not
be used for culturing or testing because residual chlo-
rine 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, I993a).
7.1.2.2 For site-specific investigations, it is desirable to
have the water-quality characteristics of the overlying
water as simitar 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 Water that might be contaminated with faculta-
tive pathogens may be passed through a properly main-
tained 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 oxy-
gen and other gases into equilibrium with air, and mini-
mize oxygen demand and concentrations of volatiles.
Excessive aeration may reduce hardness and alkalinity
of hard water (e.g., 280 mg/L hardness as CaCO3; E.L.
Brunson, NBS, Columbia, MO, personal communica-
tion). The concentration of dissolved oxygen in source
water should be between 90 to 100% saturation to help
ensure that dissolved oxygen concentrations are ac-
ceptable in test chambers.
7.1.3 Reconstituted Water
7.1.3.1 Ideally, reconstituted water should be prepared
by adding specified amounts of reagent-grade chemi-
cals to high-purity distilled or deionized water (ASTM,
1988a; USEPA, 1993a). In some applications, accept-
able high-purity water can be prepared using deioniza-
tion, distillation, or reverse-osmosis units (Section 6.3.3.2;
USEPA, 1993a). In some applications, test water can be
prepared by diluting natural water with deionized water
(Kemble et al., 1993).
7.1.3.2 Deionized water should be obtained from a
system capable of producing at least 1 mega-ohm wa-
ter. 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.
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
(1993a) recommends using a batch of reconstituted
water for two weeks.
7.1.3.4 Reconstituted Fresh Water
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.
22
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2. Add 5 g of CaSO4 and 5 g of CaCI? 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 100
mg/L as CaCO alkalinity 50 to 70 mg/L as CaCO3,
conductivity 330 to 360 uS/cm, and pH 7.8 to 8.2.
7.1.3.4.2 This reconstituted fresh water was developed
by USEPA EMSL-Cincinnati (J.M. Lazorchak, USEPA,
Cincinnati, OH, personal communication) and has been
used successfully in round-robin testing with H. azteca,
C. tentans, and C. riparius (Section 15). This reconsti-
tuted water has a higher proportion of chloride to sulfate
compared to the reconstituted waters described in ASTM
(1988a) and USEPA (1993a). Variable success has
been reported using USEPA or ASTM reconstituted
waters (USEPA, 1993a) with H. azteca. Research is
ongoing to develop additional types of reconstituted
waters suitable for these test organisms.
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, 1994c).
7.1.3.5.3 Ingersoll et al. (1992} describe procedures for
culturing H. azteca at salinities up to 15 %o. Reconsti-
tuted 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 (Nitromonas sp. and Nitrobacter
sp.; Fritz Chemical Company, Dallas, TX) to each liter of
water. The cultures were maintained by using static
renewal procedures; 25% of the culture water was re-
placed weekly. Hyalella azteca have been used to evalu-
ate the toxicity of estuarine sediments up to 15 %o
salinity (Nebeker and Miller, 1988; Roach et al., 1992;
Winger et al., 1993).
7.2 Formulated Sediment
7.2.1 General Requirements
7.2.1.1 Formulated sediments are mixtures of materials
that mimic natural sediments. Formulated sediments
have not been routinely applied to evaluate sediment
contamination. A primary use of formulated sediment
could be as a control sediment. Formulated sediments
allow for standardization of sediment testing or as a
basis for conducting sediment 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 cri-
teria necessary for test acceptability. The use of formu-
lated sediment eliminates interferences 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 characteristics
and provides a consistent method for evaluating the fate
of chemicals in sediment. However, additional research
is needed before formulated sediments are used rou-
tinely for sediment spiking procedures (e.g., identifying
standardized and representative sources of organic car-
bon).
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 biologi-
cal endpoints for a variety of species, and (3) the mate-
rials used in formulation of the sediment should have
consistent characteristics. Consistent material charac-
teristics include (1) consistency of materials from batch
to batch, (2) contaminant concentrations below concen-
trations of concern, and (3) availability to all individuals
and facilities.
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
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, none of the procedures have been subjected to
standardization and consensus approval or round-robin
(ring) testing.
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®
23
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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 character-
istics should be evaluated when considering the materi-
als 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 is list of suppliers.
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. Other sources
of organic carbon listed in Table 7.2 have been evalu-
ated including humus, potting soil, maple leaves,
composted cow manure, rabbit chow, cereal leaves,
chlorella, trout chow, Tetramin® and Tetrafin®. Only
peat, humus, potting soil, and composted cow manure
have been used successfully without fouling the overly-
ing water. The other sources of organic carbon (Table
7.2) cause dissolved oxygen concentrations to fall to
unacceptable levels (F.J. Dwyer, NBS, Columbia, MO,
personal communication). If appropriate conditioning pro-
cedures can be determined these other sources of
organic carbon may be acceptable. An important consid-
eration in the selection of an organic carbon source is the
ratio of carbon:nitrogen:phosphorus. As demonstrated in
Table 7.2, percentage carbon ranged from 30 to 47,
nitrogen ranged from 3 to 45 mg/g, and phosphorus
ranged from below detection to 11 ^ig/g for several
different carbon sources. These characteristics should
be evaluated when considering the materials to use in a
formulated sediment.
Table 7.2. Carbon, Nitrogen, Phosphorus Levels for Various
Sources of Organic Carbon1
Organic carbon
Source
Peat
Maple leaves 1
Maple leaves 2
Cow manure
Rabbit chow
Humic acid
Cereal leaves
Chlorella
Trout chow
Tetramin®
Tetrafin®
Carbon
47
42
47
30
40
40
47
40
43
37
36
Nitrogen
(mg/g)
4
6
3
11
18
3
4
41
36
45
29
Phosphorus
0.4
1.3
1.7
8.2
0.2
—
0.4
5.7
11
9.6
8.6
7.2.3 Procedure
7.2.3.1 A summary of procedures that have been used
to formulate sediment are listed below. Suppliers of
various components are listed in Table 7.3.
1. Walsh et al. (1981): (1) Wash sand (Mystic White No
85, 45, and 18—New England Silica Inc) and sieve
into three grain sizes: coarse (500 to 1500 \im);
medium (250 to 499 um); and fine {63 to 249 urn).
(2) Clay and silt were obtained from Engelhard
Corp.; (3) Peat moss is milled and sieved through an
840 jim screen. (4) Constituents are mixed dry in
the following quantities: coarse sand (0.6%); me-
dium sand (8.7%); fine sand (69.2%); silt (10.2%);
clay (6.4%); and organic matter (4.9%).
2. Clements, W.H. (Colorado State University, Ft.
Collins, CO, personal communication): (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 um. (5) Wash sand (Mystic white #45)
and dry. (6) Clay and silt are obtained using ASP
400 (Englehard Corp). (7) Constituents are mixed
dry in the following quantities: sand (1242 g); silt and
clay (219 g); dolomite (7.5 g); peat moss (31.5 g);
and humic acid (0.15 g). (8) Sediment is mixed for
an hour on a rolling mill and stored dry until ready for
use.
3. Hanes et al. (1991): (1) Sieve sand and retain two
particle sizes (90 to 180 um and 180 to 250 um)
which are mixed in a ratio of 2:1. (2) Potting soil is
dried for 24 h at room temperature and sieved
through a 1-mm screen. Clay is commercially avail-
able 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 materi-
als). (4) Constituents are mixed by weight in the
following ratios: sand mixture (42%); clay (42%);
and soil (16%). (5) After mixing, autoclave in a foil-
covered container for 20 min. (6) Mixture can be
stored indefinitely if kept covered after autoclaving.
4. Naylor (1993): (1) Sand is acid-washed and sieved
to obtain a 40 to 100 mm size. (2) Clay is kaolin light.
(3) Peat moss is ground and sieved using a 2-mm
screen (peat moss which is allowed to dry out will
not rehydrate 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) Constituents are
mixed by weight in the following percentages: sand
(69%); kaolin (20%); peat (10% [adjust for moisture
F.J. Dwyer, NBS, Columbia, MO, personal communication
24
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Table 7.3 Sources of Components Used in Formulated Sediments
Component Sources
Sand • Mystic White #18, #45, #85, #90—New England Silica, Inc., South Windsor, CT
• Product No. 33094, BDH Chemical, Ltd., Poole, England
Kaohnite • 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 • Lewiscratt Sculptor's Clay, available in hobby and artist supply stores
Humus • Sims Bark Co., Inc., Tuscumbia, AL
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
• Southern Agri-minerals Corp., Hartford, AL
content]); and CaCO3 (1%). (6) Mix for 2 h in a soil
shaker and store in sealed containers.
3. Organic matter (peat, humus, cow manure) should
be dried, milled, and passed through a 0.84 mm
sieve.
5. Suedel and Rodgers (1994): (1) Sand (Mystic White
#18 and 90) is sieved 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) Silt (ASP
400), clay (ASP 600 and 900), montmorillonite clay,
and dolomite are ashed at 550°C. for 1 h to remove
organic matter. (3) Humus is dried (70°C) and milled
to 2.0 mm. (4) Dolomite is added as 1% of the silt
requirement. (5) Materials are aged for 7 d in flowing 7.3 Reagents
water before mixing. (6) Constituents are mixed to
mimic the desired characteristics of the sediment of
concern.
4. Either CaMg{CCg,, or CaC03 should be added to
buffer the sediments.
5. All constituents are mixed on a percent dry weight
basis. Mix in the following ratios: sand (77%); silt/
clay (17%); organic matter (5%); buffer (1%).
7.2.3.2 The procedure for formulating a sediment is a
combination of methods outlined in Section 7.2.3.1. The
characteristics of this formulation would be sand 77%,
silt/clay 17%. The organic matter would depend on the
source of organic carbon. This approach could be modi-
fied to mimic specific characteristics of a sediment. If a
formulated sediment is to be used as a control sediment,
the physico-chemical characteristics of the formulated
sediment should be within the tolerance limits of the test
organism.
1. Wash sand, sieve, and retain the following two size
groups: medium (0.5 to 0.25 mm) and fine (0.25 to
0.05 mm). Sand should be mixed at a ratio of 2:1,
fine: medium.
2. Clay and silt fractions are obtained using ASP®
400. Other clays or silts (e.g., Attagel® 50, ASP®
400P, ASP® 600, ASP® 900, montmorillonite) might
be used if specific characteristics are required.
7.3.1 Data sheets should be followed for reagents and
other chemicals purchased from supply houses. The
test material(s) should be at least reagent grade, unless
a test on formulation commercial product, technical-grade,
or use-grade material is specifically needed. Reagent
containers should be dated when received 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.
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.
25
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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
sediments that might contain unknown quantities of
toxic contaminants (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 disrup-
tion as possible; however, subsampling, compositing, or
homogenization of sediment samples may be necessary
for some experimental designs. Sampling may cause
loss of sediment integrity, change in chemical specia-
tion, or disruption of chemical equilibrium (ASTM, 1994b).
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 expected exposure. For example, oligocha-
etes may burrow 4 to 15 cm into sediment. Samples
collected for evaluations of dredged material should
include all sediment to project depth. 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, 1994a). Dry
ice can be used to cool samples in the field; however,
sediments should never be frozen. Monitors can be
used to measure temperature during shipping (e.g.,
TempTale Temperature Monitoring and Recording Sys-
tem, Sensitech, Inc., Beverly, MA).
8.1.4 For additional information on sediment collection
and shipment see ASTM (1994b).
8.2 Storage
8.2.1 Manipulation or storage can alter bioavailability of
contaminants in sediment (Burton and Ingersoll, 1994);
however, the alterations that occur may not substantially
affect toxicity. Storage of sediment samples for several
months at 4°C did not result in significant changes in
chemistry or toxicity (T. Dillon and H. Tatem, USCOE,
Vicksburg, MS, personal communication; G.T. Ankley
and D. DeFoe, USEPA, Duluth, MN, unpublished data);
however, others have demonstrated changes in spiked
sediment within days to weeks (e.g., Burton, 1991;
Stemmer et al., 1990a). Sediments primarily contami-
nated with nonionic, nonvolatile organics will probably
change little during storage because of their relative
resistance to biodegradation and sorption to solids. How-
ever, metals and metalloids may be affected by chang-
ing redox, oxidation, or microbial metabolism (such as
with arsenic, selenium, mercury, lead, and tin; all of
which are methylated by a number of bacteria and
fungi). Metal-contaminated sediments may need to be
tested relatively soon after collection with as little ma-
nipulation as possible (Burton and Ingersoll, 1994).
8.2.2 Given that the contaminants of concern and the
influencing sediment characteristics are not always known
a priori, it is desirable to hold sediments in the dark at
4°C and start tests soon after collection from the field.
Recommended sediment holding time ranges from less
than two (ASTM, 1994a) to less than eight weeks
(USEPA-USCOE, 1994). If whole-sediment tests are
started after two weeks of collection, it may be desirable
to conduct additional characterizations of sediment to
evaluate possible effects of storage on sediment. For
example, concentrations of contaminants of concern
could be measured in pore water within two weeks from
sediment collection and at the start of the sediment test
(Kemble et al., 1993). Ingersoll et al. (1993) recommend
conducting a toxicity test with pore water within two
weeks from sediment collection and at the start of the
sediment test. Freezing and longer storage might further
change sediment properties such as grain size or con-
taminant partitioning and should be avoided (ASTM,
1994b; Schuytema et al., 1989; K.E. Day Environment
Canada, Burlington, Ontario, personal communication).
Sediment should be stored with no air over the sealed
samples (no head space) at 4°C before the start of a test
(Shuba et al., 1978; ASTM, 1994b). Sediment may be
stored in containers constructed of suitable materials as
outlined in Section 6. It is desirable to avoid contact with
metals, including stainless steel and brass sieving
screens, and some plastics.
26
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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 dis-
carded but should be mixed back into the sediment
during homogenization. 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 indigenous organisms and large debris can be
removed using forceps. Reynoldson et al. (1994) ob-
served reduced growth of amphipods, midges, and may-
flies in sediments with elevated numbers of oligochaetes
and recommended sieving sediments suspected to have
high numbers of indigenous 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 using
stirring or a rolling mill, feed mixer, or other suitable
apparatus (see ASTM, 1994b). Homogenization of sedi-
ment can be accomplished using a modified 30-cm
bench-top drill press (Dayton Model 3Z993) or a
variable-speed hand-held drill outfitted 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 et a!., 1994). These pro-
cedures could also be used to mix lest sediment with a
control sediment in dilution experiments.
8.3.2 Sediment Spiking
8.3.2.1 Test sediment can be prepared by manipulating
the properties of a control sediment. Additional research
is needed before formulated sediments are used rou-
tinely for sediment spiking procedures (e.g., identifying
standardized and representative sources of organic car-
bon). Mixing time (Stemmer et al., 1990a) and aging
(Word et al., 1987; Landrum, 1989; Landrum and Faust,
1992) of spiked sediment can affect responses. Many
studies with spiked sediment are often started only a few
days after the chemical has been added to the sedi-
ment. This short time period may not be long enough for
sediments to equilibrate with the spiked chemicals. Con-
sistent spiking procedures should be followed in order to
make interlaboratory comparisons. It is recommended
that spiked sediment be aged at least one month before
starting a test; however equilibration for some chemicals
may not be achieved for long periods of time.
8.3.2.1.1 The cause of sediment toxicity and the magni-
tude of interactive effects of contaminants can be esti-
mated by spiking a sediment with chemicals or complex
waste mixtures (Lamberson and Swartz, 1992). Sedi-
ments spiked with a range of concentrations can be
used to generate either point estimates (e.g., LC50) or a
minimum concentration at which effects are observed
(lowest-observable-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 deter-
mined 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 on formulation commercial product,
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 con-
centration of major ingredients and impurities, (2) water
solubility in test water, (3) estimated toxicity to the test
organism and to humans, (4) if the test concentration (s)
are to be measured, the precision and bias of the
analytical method at the planned concentration(s) of the
test material, and (5) recommended handling and dis-
posal procedures.
8.3.2.2.1 Organic compounds have been added in the
dry form or coated on the inside walls of the container
(Ditsworth et al., 1990). Metals are generally added in
an aqueous solution (ASTM, 1994b; Carlson et al.,
1991; Di Toro et at., 1990). If an organic solvent is used,
the solvent in the sediment should be at a concentration
that does not affect the test organism. Concentrations of
the chemical in the pore water and in the whole sedi-
ment should be monitored at the beginning and the end
of a test.
8.3.2.3 Use of a solvent other than water should be
avoided if possible. Addition of organic solvents may
dramatically influence the concentration of dissolved
organic carbon in pore water (G.T. Ankley, USEPA,
Duluth, MN, personal communication). If an organic
solvent must be used, both a solvent-control and a
negative-control sediment must be included in a test.
The solvent control must contain the highest concentra-
tion of solvent present and must be from the same batch
used to make the stock solution (see ASTM, 1988a).
The same concentration of solvent should be used in all
treatments. If an organic solvent is used as a carrier, it
may be possible to perform successive washes of sedi-
ment to remove most of the solvent while leaving the
compound of study (Harkey et al., 1994).
8.3.2.4 If the concentration of solvent is not the same in
all test solutions that contain test material, a solvent test
should be conducted to determine whether survival,
growth, or reproduction of the test organisms is related
to the concentration of the solvent.
8.3.2.4.1 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 meet-
ing the acceptability of the test and as the basis for
calculating results. The negative control might provide
additional information on the general health of me or-
ganisms tested. If no statistically significant difference is
detected, the data from both controls should be used for
27
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meeting the acceptability of the test and as the basis for
calculating the results (ASTM, 1992).
8.3.2.5 Test Concentration(s) for Laboratory
Spiked Sediments
8.3.2.5.1 If a test is intended to generate an LC50, the
selected test concentrations should bracket the pre-
dicted LC50. The prediction might be based on the
results of a test on the same or a similar test material
with the same or a similar test organism. The LC50 of a
particular compound may vary depending on physical
and chemical sediment characteristics. If a useful pre-
diction is not available, it is 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 of ten. Results from
water-only tests could be used to establish concentra-
tions to be tested in a whole-sediment test based on
predicted pore-water concentrations (Di Toro et al., 1991}.
8.3.2.5.2 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 con-
tent, and simultaneously extracted metals might be nor-
malized to acid volatile sulfrdes (Di Toro et al, 1990; Di
Toroetal., 1991).
8.3.2.5.3 In some situations it might be necessary to
only 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.
8.3.2.6 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,
1994b). Modifications of the mixing techniques might be
necessary to allow time for a test material 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 characteristics of the
sediment (ASTM, 1994b). Duration of contact between
the chemical and sediment can affect partitioning and
bioavailability (Word et al., 1987). Care should be taken
to ensure that the chemical is thoroughly and evenly
distributed in the sediment. Analyses of sediment sub-
samples are advisable to determine the degree of mix-
ing 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).
8.4 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, 1994a; Plumb, 1981).
8.4.2 Other analyses on sediments might include bio-
logical oxygen demand, chemical oxygen demand, cat-
ion exchange capacity, Eh, total inorganic carbon, total
volatile solids, acid volatile sulfides, metals, synthetic
organic compounds, oil and grease, petroleum hydro-
carbons, 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
collected from the same sample for analysis of sediment
physical and chemical characterizations. Qualitative de-
scriptions of the sediment may include color, texture,
presence of macrophytes or animals. Monitoring the
odor of sediment samples should be avoided because of
potential hazardous volatile contaminants.
8.4.4 A nalytical 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
analytical method used should be determined in the
appropriate matrix: that is, sediment, water, tissue. Re-
agent blanks and analytical standards should be ana-
lyzed, and recoveries should be calculated.
8.4.4.3 Concentration of spiked test material(s) in sedi-
ment, interstitial water, and overlying water should be
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
material(s) degradation products might also be desir-
able.
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 might be added to these extra chambers depend-
ing on the objective of the study.
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-
28
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tain any surface debris, ^ material fro. the sides of %£
the test chamber, or any sed.ment. ^ pressure> Qr by usjng ap .^^ ^ samp|er;
^
the surface of the sediment, then removing appropriate
ahquots of the sediment for chem,cal analysis. degrade Qr vo|ati|ize durjng Jso|atjon Qr s,orage Qf ,he
interstitial water sample.
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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 labora-
tory Standard Operating Procedures (SOPs) for test
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).
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 condi-
tions, (5) instrument 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 activi-
ties carried out within the scope of the overall QA
program. For more detailed discussion of quality assur-
ance, and general guidance on good laboratory prac-
tices related to testing see FDA (1978), USEPA (1979a),
USEPA (1980a), USEPA (1980b), USEPA (1993a),
USEPA (1994b), USEPA (1994c), DeWoskin (1984),
and Taylor (1987).
9.2 Performance-based Criteria
9.2.1 USEPA Environmental Monitoring Management
Council (EMMC) recommended the use of
performance-based methods in developing standards
for chemical analytical methods (Williams, 1993).
Performance-based methods were defined by EMMC
as a monitoring approach that permits the use of appro-
priate methods 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 perfor-
mance standards.
9.2.2 Participants at a September 1992 USEPA sedi-
ment toxicity workshop arrived at a consensus on sev-
eral culturing and testing methods for freshwater organ-
isms (Appendix A, Section S.4). 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
organisms are taken for testing. Having healthy organ-
isms 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 criteria were chosen to al-
low each laboratory to optimize culture methods while
providing organisms that produce reliable and compa-
rable test results. See Tables 11.3, 12.3, and 13.4 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 cultur-
ing or sediment testing areas, and from sediment testing
laboratories and sample preparation areas into culture
rooms.
9.3.2 Equipment for temperature control should be
adequate to maintain recommended test-water tem-
peratures. 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 "non-contaminant" test should be conducted in
which all test chambers contain a control sediment and
30
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overlying water. This information is used to demonstrate
that the facility, control sediment, water, and handling
procedures provide acceptable responses of test organ-
isms (Section 9.14).
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
satisfactory survival, growth, or reproduction of the test
organisms. Test organisms should not show signs o!
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 continu-
ously in at least one chamber during each test. Tem-
peratures should be maintained within the limits speci-
fied for each test. Dissolved oxygen, alkalinity, water
hardness, conductivity, ammonia, and pH should be
checked as prescribed in Sections 11.3, 12.3, and 13.3.
9.8 Quality of Test Organisms
9.8.1 Monthly reference-toxicity tests should be con-
ducted on all test organisms using procedures outlined
in Section 9.16. If reference-toxicity tests are not con-
ducted monthly, the lot of organisms used to start a
sediment test must be evaluated using a reference
toxicant. Physiological measurements such as lipid con-
tent might also provide useful information regarding the
health of the cultures.
9.8.2 The quality of test organisms obtained from an
outside source must be verified by conducting a
reference-toxicity test concurrently with the sediment
test. The supplier should provide data with the shipment
describing the history of the sensitivity of organisms
from the same source culture. If the supplier has not
conducted five reference toxicity tests with the test
organism, it is the responsibility of the testing laboratory
to conduct these five reference toxicity tests before
starting a sediment test (Section 9.14.1).
9.8.3 The supplier should also certify the species iden-
tification of the test organisms and provide the taxo-
nomic references or name(s) of the taxonomic expert(s)
consulted.
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 bioaccu-
mulation tests must be analyzed for compounds to be
measured in the bioaccumulation tests.
9.10 Test Acceptability
9.10.1 For the test results to be acceptable, survival at
10 d must equal or exceed 80% for H. azteca and 70%
for C. tentans in the control sediment. Numbers of L
variegatus should not be reduced in test sediments
relative to the control sediment and organisms should
burrow into the test sediment. Avoidance of test sedi-
ment by L variegatus will decrease bioaccumulation.
See Table 11.3, 12.3, and 13.4 for additional require-
ments for acceptability of the tests.
9.10.2 An individual test may be conditionally accept-
able if temperature, dissolved oxygen, and other speci-
fied conditions fall outside specifications, depending on
the degree of the departure and the objectives of the
tests (see test condition summaries). 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, 1993a; USEPA, 1994b).
9.11.2 Reagent containers should be dated when re-
ceived from the supplier, and the shelf Ufe o1 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
general section on quality assurance (see USEPA Meth-
ods 150.1, 360.1, 170.1, and 120.1; USEPA, 1979b).
Calibration data should be recorded in a permanent log.
31
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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).
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 var-
ies with the objectives of the test and the statistical
method used for analysis of the data (Section 14).
9.14 Demonstrating Acceptable
Performance
9.14.1 It is the responsibility of a laboratory to demon-
strate its ability to obtain consistent, precise results with
reference toxicants before it performs sediment tests
(see Section 9.16). Intralaboratory precision, expressed
as a coefficient of variation (CV) of the range in re-
sponse for each type of test to be used in a laboratory,
should be determined by performing five or more tests
with different batches of test organisms using the same
reference toxicant at the same concentrations with the
same test conditions (e.g., the same test duration, type
of water, age of test organisms, feeding) and the same
data analysis methods. This should be done to gain
experience for the toxicity tests and as a point of refer-
ence for future testing. A reference toxicant concentra-
tion series (0.5 or higher) should be selected that will
consistently provide partial mortalities at two or more
concentrations of the test chemical (Section 15).
9.14.2 Before conducting tests with contaminated sedi-
ment, the laboratory should demonstrate its ability to
conduct tests by conducting five exposures in control
sediment as outlined in Table 11.1, 12.1, or 13.1. It is
recommended that these five exposures with control
sediment be conducted concurrently with the five refer-
ence toxicity tests described in Section 9.14.1.
9.14.3 Laboratories should demonstrate that their per-
sonnel 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 sediment or test
sediments and recovery could be determined after 1 h
(Tomasovic et al., 1994).
9.15 Documenting Ongoing Laboratory
Performance
9.15.1 Satisfactory laboratory performance on a con-
tinuing basis is demonstrated by conducting monthly
water-only 96-h reference-toxicity tests with each test
organism. For a given test organism, successive tests
should be performed with the same reference toxicant at
the same concentrations in the same type of water using
the same data analysis method (Section 15).
9.15.2 Outliers, which are data falling outside the con-
trol limits and trends of increasing or decreasing sensi-
tivity are readily identified. If the reference toxicity datum
from a given test falls outside the "expected" range (e.g.,
±2 SD), the sensitivity of the organisms and the credibil-
ity of the test results are suspect. In this case, the test
procedure should be examined for defects and should
be repeated with a different batch of test organisms.
9.15.3 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 automatically be judged unaccept-
able 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%. All the perfor-
mance criteria outlined in Tables 11.3, 12.3, and 13.4
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.
9.15.4 Performance should improve with experience,
and the control limits should gradually narrow, as the
statistics stabilize. However, control limits of ±2 SD, by
definition, will be exceeded 5% of the time, regardless of
how well a laboratory performs. For this reason, good
laboratories that develop very narrow control limits may
be penalized if a test result that falls just outside the
control limits is rejected de facto. The width of the
control limits should be considered in decisions regard-
ing rejection of data (Section 15).
9.16 Reference Toxicants
9.16.1 Ideally, reference-toxicity tests should be con-
ducted in conjunction with sediment tests to determine
possible changes in condition of a test organism (Lee,
1980). Water-only reference-toxicity tests should be con-
ducted monthly. Deviations outside an established nor-
mal range may indicate a change in the condition of the
test organism population. Results of reference-toxicity
tests also enable interlaboratory comparisons of test
organism sensitivity.
9.16.2 Reference toxicants such as sodium chloride
(NaCI), potassium chloride (KCI), cadmium chloride
(CdCL), and copper sulfate (CuSOj are suitable for
use. No one reference toxicant can be used to measure
the condition 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 or
two reference toxicants routinely. KCI has been used
successfully in round-robin water-only exposures with
H. azteca and C. tentans (Section 15).
9.16.3 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
32
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have observed low control survival when more than one
midge/chamber is tested in water-only exposures.
9.17 Record Keeping
9.17.1 Proper record keeping is important. A complete
file should be maintained for each individual sediment
test or group of tests on closely related samples. This
file should contain a record of the sample
chain-of-custody; a copy of the sample log sheet; the
original bench sheets for the test organism responses
during the sediment test(s); chemical analysis data on
the sample(s); control data sheets for reference toxi-
cants; 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 instruments; test conditions used; and results of
reference toxicant tests. Laboratory data should be re-
corded immediately to prevent the loss of information or
inadvertent introduction of errors into the record. Origi-
nal data sheets should be signed and dated by the
laboratory personnel performing the tests. For additional
detail see Section 14.
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.
17.
Aeration:
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 SOOto 1000 lux
16L:8D
None
H. azteca: 7- to 14-dold
C. tentans: third instar larvae1
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 instar or younger with at least 50% of the organisms at third instar.
33
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Table 9.2 Recommended Test Conditions for Conducting Reference-Toxicity Tests with More Than One Organism/Chamber
Parameter Conditions
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
Test type:
Dilution series:
Toxicant:
Temperature:
Light quality:
Illuminance:
Photopenod:
Renewal of water:
Age of organisms:
Test chamber:
Volume of water:
Number of organisms/chamber:
Number of replicate chambers/treatment:
Feeding:
Water-only test
Control and at least 5 test concentrations (0.5 dilution factor)
NaCI, KCI.Cd, orCu
23 ± 1°C
Wide-spectrum fluorescent lights
About 500 to 1000 lux
16L:8D
None
H. azteca: 7- lo 14-dold
C. tentans: third instar
L. variegatus: adults
250-mL glass beaker (covered with glass or plastic)
100 rnL (minimum)
10 minimum
3 minimum
H. azteca: 0.5 mL YCT (1 800 mg/L stock) on Day 0 and 2
15. Substrate:
16. Aeration:
17. Dilution water:
18. Test chamber cleaning:
19. Water quality:
20. Test duration:
21. Endpoint:
22. Test acceptability:
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 reconsti-
tuted 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 instar or younger with at least 50% of the organisms at third instar.
34
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Section 10
Collection, 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 H. azteca is
most common in warm (20 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 (1970) re-
ported that H. azteca selectively ingest bacteria and
algae. The behavior and feeding habits of H. azteca
make them excellent test organisms for sediment as-
sessments.
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 near the 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 marsu-
pium where they are fertilized. Hyalella azteca average
about 18 eggs/brood (Pennak, 1989) with larger organ-
isms 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 to 28°C, hatching ranges from 5 to 10 d after fertiliza-
tion (Embody, 1911; Bovee, 1950; Cooper, 1965). The
time between molts for females is 7 to 8 d at 26 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 indefi-
nite 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 cul-
tured at illuminance of about 500 to 1000 lux (Ingersoll
and Nelson, 1990; Ankley et al., 1991 a; Ankley et al.,
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). Al temperatures less than 10°C the organisms
rest and are immobile (de March, 1977; de March,
1978). At temperatures of 10 to 18°C, reproduction can
occur. Juveniles grow more slowly at colder tempera-
tures and become larger adults. Smaller adults with
higher reproduction are typical when organisms are
grown at 18 to 28°C. The highest rates of reproduction
occur at 26 to 28°C (de March, 1978) while lethality
occurs at 33 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 %o; 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 nS/cm, but
only up to 12,000 jiS/cm in Na-dominated waters
(Ingersoll et al., 1992). De March (1981) reported H.
azteca were not collected from locations where calcium
was less than 7 mg/L. Hyalella azteca have been cul-
tured in water with a salinity up to 15 %o in reconstituted
salt water (Ingersoll et al., 1992; Winger and Lasier,
1993). In laboratory studies, Sprague (1963) reported a
24-h LC50 for dissolved oxygen at 20°C of 0.7 mg/L.
Pennak and Rosine (1976) reported similar findings.
Nebeker et 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 H. azteca were reduced
after 30-d exposure to 1.2 mg/L dissolved oxygen.
35
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10.1.1.8 Hyalella azteca tolerate a wide range of sub-
strates. Ingersoli and Nelson (1990) and Ingersoll et al.
(1993) reported that H. azteca tolerated sediments rang-
ing 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-d tests
with formulated sediment (Suedel and Rodgers, 1994).
Ankley et al. (I994a) evaluated the effects of natural
sediment physico-chemical characteristics on the re-
sults 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.
10.1.2 Chironomus tentans
10.1.2.1 Chironomus tentans have a holarctic distribu-
tion (Townsend et al., 1981) and are commonly found in
eutrophic ponds and lakes (Flannagan, 1971; Driver,
1977). Midge larvae are important in the diet of fish and
waterfowl {Sadler, 1935; Siegfried, 1973; Driver et al.,
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 2.0 mm, 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 nmhos/cm, and sediment
organic carbon 1.9 to 15.5%. Larvae were absent from
lakes if hydrogen sulfide concentration in overlying wa-
ter exceeded 0.3 mg/L. Abundance of larvae was posi-
tively correlated with conductivity, pH, amount of food,
percentages of particles in the 0.59 to 1.98 mm size
range, and concentrations of Na, K, Mg, Cl, SO , 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
described by Sadler (1935) and others (ASTM, 1994a).
Males are easily distinguished from females because
males have large, plumose antennae and a much thin-
ner abdomen with visible genitalia. The male has paired
genital claspers on the posterior tip 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 directly into the water. At ERL-D, the females
oviposit eggs within 24 h after emergence. An egg mass
contains about 2,300 eggs (Sadler, 1935) 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 sec-
ond 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 masses (2 to 3 d post-emergence;
Sadler, 1935).
10.1.2.3 Chironomus tentans tolerate a wide range of
substrates. Survival or growth of C. tentans was not
reduced over a wide range in sediment grain sized in
10-d tests with formulated sediment; however, survival
was reduced in artificial sediments below 0.91% organic
matter when organisms were not fed (Suedel and
Rodgers, 1994). Ankley et al. (1994a) evaluated the
effects of natural sediment physico-chemical character-
istics 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 growth of organisms was decreased
in tests without added food. Physico-chemical sediment
characteristics including grain size and TOC were not
significantly correlated to survival of C. tentans in tests
in which organisms were fed. However, linear modeling
indicated growth of C. tentans was influenced by grain
size distribution of the test sediments (growth slightly
increased in coarser sediment).
10.1.3 Lumbriculus variegatus
10.1.3.1 Lumbriculus variegatus inhabit a variety of
sediment types throughout the United States and Eu-
rope (Chekanovskaya, 1962; Cook, 1969; Spencer, 1980;
Brinkhurst, 1986). Lumbriculus variegatus typically tun-
nel 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 et al., 1991; Phipps et al., 1993). Lipid
content is about 1.0% (wet weight, Ankley et al., 1992b).
Lumbriculus variegatus most commonly reproduce
asexually, although sexual reproduction has been re-
ported (Chekanovskaya, 1962). Newly hatched worms
have not been observed in cultures (Call et al., 1991;
Phipps et al., 1993. Cultures consist of adults of various
sizes. Populations of laboratory cultures double (num-
ber of organisms) every 10 to 14 d at 20°C (Phipps et al.,
1993).
36
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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. Sur-
vival and reproduction of organisms was decreased in
tests without added food. Physico-chemical sediment
characteristics including grain size and TOC were not
significantly correlated to reproduction or growth of L.
variegatus in either fed or unfed tests.
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 H. 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, or 13.4.
10.2.2 All organisms in a test must be from the same
source. Organisms may be obtained from laboratory
cultures, from commercial, or government sources (Table
10.1). The test organism used should be identified using
Table 10.1 Sou rces of Test Organ isms
Source
Species
U.S. Environmental Protection Agency H. azteca
Environmental Research Laboratory-Duluth C. tentans
6201 Congdon Boulevard L. variegatus
Duluth, MN 55804
Teresa Norberg-King (218/720-5500)
U.S. Environmental Protection Agency H. azteca
Environmental Monitoring System Laboratory C. tentans
3411 Church Street
Cincinnati, OH 45244
Jim Lazorchak (513/569-7076)
Midwest Science Center H. azteca
National Biological Survey C. tentans
4200 New Haven Road L. variegatus
Columbia, MO 65201
Eugene Greer (314/875-5399)
Great Lakes Environmental Research L variegatus
Laboratory, NOAA
2205 Commonwealth Boulevard
Ann Arbor, Ml 48105-1593
Peter Landrum (313/741-2276)
Wright State University H. azteca
Department of Biological Sciences C. tentans
Dayton, OH 45435 L. variegatus
Allen Burton (513/873-2201)
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)
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 addi-
tion, the ability of the wild population of sexually repro-
ducing organisms to cross-breed with the existing labo-
ratory population must be determined. Sensitivity of the
wild population to select contaminants (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.
Temperature 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 dissolved oxygen or temperature fluctuations.
The temperature of the shipped water should be gradu-
ally adjusted to the desired culture temperature at a rate
not exceeding 2°C per 24 h. Additional reference-toxicity
testing is required if organisms are not cultured at the
testing laboratory (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 other-
wise stressed (e.g., >20% mortality for 48 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 organ-
isms 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 suffi-
cient to culture fathead minnows (Pimephales promelas)
or cladocerans will generally be adequate.
10.2.5.1 Variable success has been reported using
reconstituted waters described in ASTM (I988a) or
USEPA (1993a) to culture or test H. azteca (USEPA.
1992). However, the reconstituted water described in
Section 7.1.3.4 has been used to successfully culture H.
azteca (J.M. Lazorchak, USEPA, Cincinnati, OH, per-
sonal communication). The reconstituted water described
in Section 7.1.3.4 has a higher proportion of chloride to
sulfate compared to the reconstituted waters described
in ASTM (1988a) and USEPA (1993a). H. aztecacan be
cultured and tested at salinities up to 15 %o (Ingersoll et
al., 1992; Winger et al., 1993).
10.2.5.2 Organisms can be cultured using either static
or renewal procedures. Renewal of water is recom-
mended to limit loss of the culture organisms from a
drop in dissolved 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
37
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maintained in static systems. For example, aeration is
needed in static systems to maintain dissolved oxygen
at >40% of saturation.
10.2.5.3 A recirculating system using an under-gravel
filter has been used to culture amphipods and midges
(P.V. Winger. NBS, Athens, GA, personal communica-
tion). The approach for using a recirculating system to
culture organisms has been described by New et al.
(1974), Crandall et al. (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 an artificial substrate (e.g., plastic
balls or multi-plate substrates) are 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®
should include limestone gravel to help avoid depres-
sion in pH. Recirculating systems require less mainte-
nance than static systems.
10.2.6 Cultures should be maintained at 23°C with a
16L8D photoperiod at a illuminance of about 500 to
1000 lux (ASTM, (!994a) and Appendix A). Cultures
should be observed daily. Water temperature should be
measured daily or continuously, and dissolved oxygen
should be measured weekly. Reference-toxicity tests
should be conducted at least monthly. If reference-toxicity
tests are not conducted monthly, the lot of organisms
used to start a sediment test must be evaluated using a
reference toxicant. Culture water hardness, alkalinity,
ammonia, and pH should be measured at least quarterly
and the day before the start of a sediment test. If
reconstituted water is used to culture organisms, water
quality should be measured on each batch of reconsti-
tuted water. Culture procedures should be evaluated
and adjusted as appropriate to restore or maintain the
health of the culture.
10.3 Culturing Procedures for Hyalella
azteca
10.3.1 The culturmg procedures described below are
based on methods described in USEPA (1993), Ankley
et al. (1994a), Call et al. (1994), Tomasovic et al. (1994),
Greer (1993), Ingersoll and Nelson (1990), ASTM (1993a)
and Appendix A. The culturing procedure must produce
7- to 14-d old amphipods to start a sediment test (Table
11.3). A narrower age range of organisms used to start a
test may be desirable when growth is measured as an
endpoint. Amphipods within a range of 1- to 2-d old will
be more uniform in size than organisms within a range of
7-d old.
10.3.2 The following procedure described by Call et al.
(1994) and USEPA (1993) 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. Cotton gauze should be
soaked in water for 24 h before use and should be
renewed on a weekly basis. Amphipods are fed 10 ml of
a the yeast-Cerophyl®-trout chow (YCT) mixture (Ap-
pendix C), 10 mL of the green algae Selenastrum
capricornutum (about 35 x 106 cells/ml), and 10 mL of
the diatom, Navicula spp. (1.0 x 109 cells/ml) on Mon-
day. Five mL of each food is added to cultures on
Wednesdays and Fridays.
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
translucent white plastic or white enamel pan. After the
adults are removed, the remaining amphipods will range
in age from <1- to 7-d old. Young amphipods are trans-
ferred with a pipet into a 1-L beaker containing culture
water and are held for one week before starting a toxicity
test. Presoaked cotton gauze is placed in the beakers,
and organisms are fed 10 mL of YCT and 10 mL of
green algae, and 10 mL of diatoms with renewal of
water, and 5 mL of each food on Wednesdays and
Fridays (Appendix C). 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 which are useful in determining if 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 repro-
duce for several months; however, young production
gradually decreases after about three 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 organisms 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.
38
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10.3.4 A third procedure for obtaining known-age am-
phipods is described by Greer (1993) and Tomasovic et
al. (1994). Mass cultures of mixed-age amphipods are
maintained in 80-L glass aquaria containing about 50 L
of water (Ingersoll and Nelson, 1990). Tetramin® 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
Tetramin® is added when most of the Tetramin® has
been consumed. Laboratories using static systems should
develop lower feeding rales 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 "coiled-web material"; 3-M, St. Paul,
MN). Before use, leaves are soaked in water for about
30 d. The leaves are then flushed with water to remove
residuals of naturally occurring tannic acid before place-
ment in the cultures.
10.3.4.1 To obtain known-age amphipods, a U.S. Stan-
dard Sieve #25 (710 ^m 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 under water. The artificial
substrates or leaves are then shaken under water sev-
eral 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 water to 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 amphipods 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 removed from the pan, and the mature amphi-
pods are placed back into their culture chamber or
placed in a second pan containing culture water if addi-
tional organisms are needed for testing. The newborn
amphipods are moved with a pipet and placed in a
culture chamber with flowing water during a grow-out
period. The newborn amphipods should be counted to
determine if adequate numbers 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 test. A
few maple leaves and a small amount of Tetramin® is
placed into the grow-out culture chamber to provide
food.
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, NBS, Athens, Ga, unpublished
data). The mean length of these sieved organisms cor-
responds to that of 6-d old amphipods (Figure 10.1).
After holding for 3 d before testing to eliminate organ-
isms injured during sieving, these amphipods were 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) and ASTM (1994a)
describe the following procedure for obtaining mixed-age
amphipods of a similar size to start a test. Smaller
amphipods are isolated from larger amphipods using a
stack of U.S. Standard sieves: #30 (600 |im), #40 (425
jim), and #60 (250 jam). Sieves should be held under
water to isolate the amphipods. 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 bea-
ker) with gentle aeration.
10.3.5.2 Amphipods should be held and fed at a rate
similar to the mass cultures for 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
evaluate 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 (1993), Ankley
et al. (1994a), Call et al. (1994), Greer (1993), ASTM
(1994a), and Appendix A. Sediment tests must be started
with third instar larvae (at least 50% of the larvae must
be third instar with the remaining larvae second instar;
Table 12.3). 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 must be determined using head
capsule width (Table 10.2). It is also desirable to monitor
the weight or length of midges at the start of a sediment
test.
39
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E
£
a>
Size retained on 355um sieve after passing 500um sieve.
4U-B--H-
8
Day
10 11 12 13
14
15 16
Mean (+/- 2SD)
Figure 10.1 Length and relative age of Hyalella azteca collected by sieving in comparison with length of known-age organisms.
P.V. Winger, NBS, Athens, GA, unpublished data.
Table 10.2 Chironomus tentans Instar and Head Capsule
Widths2
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
TJ. Norberg-King, USEPA, Duluth, MM, unpublished data.
10.4.2 Recent research has indicated that the third
instar C. tentans were frequently referred to as the
second instar in previous literature (T.J. Norberg-King,
USEPA, Duluth, MM, unpublished data). When C. tentans
larvae were measured daily, the C. tentans raised at 22
to 24°C were third instar, not second instar, by 9 to 11 d
after hatching.
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 ease in removing larvae for testing. Sources of sand
are listed in Section 7.
10.4.3.1 Paper towels are prepared according to a
procedure adapted from Batac-Catalan and White (1982).
Plain white kitchen paper towels are cut into strips. Cut
toweling is loosely packed into a 2-L beaker, submersed
in acetone, covered and placed in a fume hood, and
soaked overnight to solubilize organic contaminants.
The acetone is drained completely, and deionized water
is added, brought to a boil, and stirred to drive off any
remaining acetone vapors. This process is repeated two
more times. Finally, the toweling is rinsed three times
with cold deionized water. A mass of the toweling suffi-
cient 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 jim sieve and
rinsed well with deionized water to remove the shortest
fibers.
10.4.3.2 Dry shredded paper toweling loosely packed
into a 2-L beaker will provide sufficient substrate for
about ten 19-L chambers (USEPA, 1993). 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 masses will provide a sufficient number
of organisms to start a new culture chamber. Egg masses
should be held at 23°C in a glass beaker or crystallizing
40
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dish containing about 100 to 150 mL of culture water
(temperature change should not exceed 2°C per d).
Food is not added until the embryos start to hatch (in
about 2 to 4 days at 23°C) to reduce the risk of oxygen
depletion. A small amount of green algae (e.g., a thin
green layer) is added to the water when embryos start to
hatch. When most of the larvae have left the egg mass,
150 to 200 larvae should be placed into a culture cham-
ber. Crowding of larvae will reduce growth. Larvae that
have formed cases can be transferred to aquaria or
culture chambers using a gentle stream of water from a
squeeze bottle. See Section 10.4.5.1 or 10.4.6.1 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).
10.4.5 Chironomus tentans are cultured in soft water at
the USEPA laboratory in Duluth in glass aquaria (19.0-L
capacity, 36- x 21- x 26-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 sand is placed
in each chamber.
10.4.5.1 Tetrafin® food is added to each culture cham-
ber to provide a final food concentration of about 0.04
mg dry solids/mL of culture water. A stock suspension of
the solids is prepared 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 suspension. 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 C). Laboratories using static systems
should develop lower feeding rales specific to their
systems.
10.4.6 Chironomus tentans are cultured by Greer (1993)
in Rubbermaid® 5.7-L polyethylene cylindrical contain-
ers. 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 2 mL/d of a
100 g/L Tetrafin® suspension on Tuesday, Wednesday,
Thursday, Friday, and Sunday. A 2-mL chlorella sus-
pension (deactivated "Algae-Feast® ChlorellsC, Earth-
rise Co., Callpatria, CA) is added to each chamber on
Saturday and on Monday. The chlorella suspension 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
aspirator. 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. After the
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 sections of 0.25 inch glass tubing, and Tygon®
tubing for collecting and providing suction (Figure 10.2).
Adults should be aspirated with short inhalations to
avoid injuring the organisms. The mouth piece 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 mat-
ing and fertilization. One male may fertilize more than
one female. However, a ratio of one female to three
males ensures good fertilization.
10.4.7.1 A reproduction and oviposit chamber may be
prepared in several different ways (Figure 10.2). Culture
water (about 50 to 75 mL) can be added to the aspiration
flask in which the adults were collected (Figure 10.2;
Batac-Catalan and White, 1982). ERL-Duluth (USEPA,
1993) uses a 500-mL collecting flask with a length of
Nitex® screen positioned vertically and extending into
the culture water (Figure 10.2). 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) used an oviposition box to hold
emergent adults. The box is constructed of a 5.7-L
chamber 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
41
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Tygon Tubing
500 ml Erlenmeyer
Mesh Cover
Nitex Screen
Water
Figure 10.2 Aspirator chamber (A) and reproduction and oviposit chamber (B) for adult midges.
42
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chamber and a #11 stopper is used to close the hole.
Egg masses are removed by first sliding a piece of
plexiglass between the top and bottom chambers. Adult
midges are then aspirated from the bottom chamber.
The top chamber with plexiglass is removed from the
bottom chamber and a forceps is used to remove the
egg masses. The top chamber is put back on top of the
bottom chamber, the plexiglass is removed, and the
aspirated adults are released from the aspirator into the
chamber through the 5-cm hole.
10.4.8 About two to three weeks before the start of a
test, at least 3 to 5 egg masses should be isolated for
hatching using procedures outlined in Section 10.4.3.
10.4.9 Records should be kept on the time to first
emergence and the success of emergence for each
culture chamber. It is also desirable to monitor growth
and head capsule width periodically in the cultures. See
Section 10.2.6 for additional detail on procedures for
evaluating 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 (1993), Call et al. (1994), E.L. Brunson (NBS,
Columbia, MO, unpublished data), and Appendix A.
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 contain-
ing 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 a scissors.
Cut toweling is loosely packed into a 2-L beaker, sub-
mersed in acetone, covered and placed in a fume hood,
and soaked overnight to solubilize organic contami-
nants. The acetone is discarded, and the towels are
rinsed several times with deionized water. Deionized
water is added, brought to a boil, and stirred three times
to drive off the acetone vapors. This is repeated two
more times. The strips are conditioned for at least one
week by placing 4 L of strips into an aquarium equipped
with two water lines each having a flow capacity of about
100 mL/min. One line is placed below and one line is
placed above the towel mass. Glass weights (several
2.5-cm x 25.4-cm glass strips standing on edge and
glued on both ends to glass strips about 50 cm in length)
can be placed on the strips to prevent floating. This
approach creates a uniform water flow throughout the
strips and minimizes fouling. Following conditioning, the
strips are removed and evenly placed on the bottom of
the culture chamber. Glass weights keep the strips in
place. Conditioned strips can be frozen in deionized
water for later use. The substrate is renewed with condi-
tioned towels when thin or bare areas appear. 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 dis-
perse on the surface film, break through the surface
tension, and settle out over the substrate. Laboratories
using static systems should develop lower feeding rates
specific to their systems. Food and substrate used to
culture oligochaetes should be analyzed tor compounds
to be evaluated in bioaccumulation tests. If the concen-
tration of the test compound is above the detection level
and the food is not measured, the test may be invali-
dated. Recent studies in other laboratories, for example.
have indicated elevated concentrations of PCBs in sub-
strate and/or food used for culturing the oligochaete (J.
Amato, AScI Corporation, Duluth, MN, personal commu-
nication).
10.5.5 Phipps et al. (1993) recommend starting a new
culture with 500 to 1000 worms. Conditioned paper
toweling 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 cul-
tures 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 et al., 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 remain-
ing 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 appropri-
ate test chamber. Organisms should not be blotted with
a paper towel to remove excess water.
10.5.7 The culture population generally doubles (num-
ber of organisms) in about 10 to 14 d. See Section
10.2.6 for additional detail on procedures for evaluating
the health of the cultures.
43
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Section 11
Test Method 100.1
Hyalella azteca 10-d Survival Test for Sediments
11.1 Introduction
11.1.1 Hyalella azteca (Saussure) have many desirable
characteristics of an ideal sediment toxicity testing or-
ganism including relative sensitivity to contaminants as-
sociated 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 inter-
laboratory studies and has been confirmed with natural
benthos populations. Many investigators have success-
fully used H. azteca to evaluate the toxicity of freshwater
sediments (e.g., Nebeker et al., I984a; Borgmann and
Munwar, 1989; Ingersoll and Nelson, 1990; Ankley et
al., 1991 a, Ankley et al., 1991b; Burton et al., 1989;
Winger and Lasier, 1993; Kemble et al., 1994). H.
azteca has been used for a variety of sediment assess-
ments (Ankley et al., 1991; West et al., 1993; Hoke et
al., 1994; West et al., 1994; and Hoke et al., 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 sediment toxicity
tests with H. azteca include survival and growth.
11.1.2 A specific test method for conducting a 10-d
sediment toxicity test is described in Section 11.2 for H.
azteca. Methods outlined in Appendix A and the litera-
ture cited in Table A.2 were used for developing test
method 100.1. Results of tests using procedures differ-
ent from the procedures described in Section 11.2 may
not be comparable, and these different procedures may
alter contaminant bioavailability. Comparison of results
obtained using modified versions of these procedures
might provide useful information concerning new con-
cepts 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 com-
parability 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 14). 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 500 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 number of replicates/treatment de-
pends on the objective of the test. Eight replicates are
recommended for routine testing (Section 14). Amphi-
pods in each test chamber are fed 1.5 mL of YCT food
daily (Appendix C). Each chamber receives 2 volume
additions/d of overlying water. Water renewals may be
manual or automated, and Appendix B describes
water-renewal systems that can be used to deliver over-
lying 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 sediment is collected. Requirements for test
acceptability are summarized in Table 11.3.
11.3 General Procedures
11.3.1 Sediment into Test Chambers: The day before
the sediment test is started (Day -1) each sediment
should be thoroughly mixed and added to the test cham-
bers (Section 8.3.1). Sediment should be visually in-
spected to judge the degree of homogeneity. Excess
water on the surface of the sediment can indicate sepa-
ration of solid and liquid components. If a quantitative
measure of homogeneity is required, replicate sub-
samples should be taken from the sediment batch and
analyzed for TOC, chemical concentrations, and particle
size.
11.3.1.1 Each test chamber should contain the same
amount of sediment, determined either by volume or by
44
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Table 11.1 Test Conditions for Conducting a 10-d Sediment Toxicity Test with Hyalella azteca
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 organisms:
11. Number ot 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 500 to 1000 lux
16L:8D
300-mL high-form lipless beaker
100 ml
175mL
2 volume additions/d; continuous or intermittent (e.g., one volume
addition every 12h)
7- to 14-d old at the start of the test
10
Depends on the objective
of the test. Eight replicates are recommended for routine testing (see
Section 14)
YCT food, fed 1.5 ml daily to each test chamber
None, unless dissolved oxygen in overlying water drops below 40% ot
saturation
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 B)
Hardness, alkalinity, conductivity, pH, and ammonia at the beginning
and end of a test. Temperature and dissolved oxygen daily
10d
Survival (growth optional)
Minimum mean control survival of 80% and performance-based criteria
specifications outlined in Table 11.3
Table 11.2 General Activity Schedule for Conducting a Sediment Toxicity Test with Hyalella azteca'
Day Activity
-7 Separate known-age amphipods from the cultures and place in holding chambers. Begin preparing food for the test.
-6 to -2 Feed and observe isolated amphipods, monitor water quality (e.g., temperature and dissolved oxygen).
-1 Feed and observe isolated amphipods, monitor water quality. Add sediment into each test chamber, place chambers into
exposure system, and start renewing overlying water.
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 1.5 mL of YCT into
each test chamber. Archive 20 test organisms for weight or length determination. Observe behavior of test organisms.
1 to 8 Add 1.5 mL of YCT food to each test chamber. Measure temperature and dissolved oxygen. Observe behavior of test
organisms.
9 Same as Day 1. Measure total water quality.
10 Measure temperature and dissolved oxygen. End the test by collecting the amphipods with a sieve. Count survivors and set
aside organisms for weight or length measurements.
1 Modified from Call et al., 1994
45
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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 Hyatella azfecalhat the following performance criteria be net:
1 Age of H. azteca at the start of the test must be between 7- to 14-d old.
2. Average survival of H. a/feca in the control sediment must be greater than or equal to 80% at the end of the test.
3. Hardness, alkalinity, pH, anc! ammonia in the overlying water within a treatment should not vary by more than 50% during the test.
B. Performance-based criteria for culturing H. azteca include
1. Laboratories should perform monthly 96-h water-only reference-toxicity tests to assess the sensitivity of culture organisms. If refer-
ence-toxicity tests are no', conducted monthly, the lot of organisms used to start a sediment test must be evaluated using a reference
toxicant (Section 9.16).
2. Laboratories should track parental survival in the cultures and record this information using control charts if known-age cultures are
maintained. Records shojld 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 and the day before the start of a
sediment test: pH, hardness, alkalinity, and ammonia. Dissolved oxygen should be measured weekly. Temperature should be
recorded daily.
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. It is desirable to start tests soon after collection of sediment from the field (see Section 8.2 for additional detail).
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
aoversely affect test organisms.
5 Test organisms must be cultured and tested at 23°C.
6. The daily mean test temperature must be within ±1°C o1 the desired temperature. The instantaneous temperature must always be
within -3-C of the desired temperature.
7. Natural physico-chemical characteristics of test sediment collected from the field should be within the tolerance limits of the test
organisms.
weight. Overlying water is added to the chambers in a
manner that 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 at-
tached) 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).
11.3.2 Renewal of overlying water: Renewal of over-
lying 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%. 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). 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 B describes water-renewal
systems that can be used for conducting sediment
tests.
11.3.2.1 In water-renewal tests with one to four volume
additions of overlying water/d, water-quality characteris-
tics generally remain similar to the inflowing water
(Ingersoll and Nelson, 1990; Ankley et al., 1993); how-
ever, in static tests, water quality may change pro-
foundly 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 expo-
sure (Ingersoll and Nelson, 1990). Additionally, concen-
trations 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 char-
acteristics such as hardness may influence the toxicity
of many inorganic {Gauss et al., 1985) and organic
(Mayer and Ellersieck, 1986) contaminants. Although
contaminant concentrations 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 interstitial water.
46
-------
11.3.3 Acclimation: 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, acclimation of test organisms to the test water
is not required.
11.3.4 Placing Organisms in Test Chambers: Test
organisms should be handled as little as possible. Am-
phipods should be introduced into the overlying water
below the air-water interface. Test organisms can be
pipetted directly into overlying water (Ankley et al., 1993).
Alternatively, test organisms can be placed into 30-mL
counting cups that are floated in the test chambers for
15 min before organisms are introduced into the overly-
ing water (Ingersoll and Nelson, 1990). Length or weight
should be measured on a subset of at least 20 organ-
isms used to start the test.
11.3.5 Monitoring a Test: All chambers should be
checked daily and observations made to assess test
organism behavior such as sediment avoidance. How-
ever, monitoring effects on burrowing activity of test
organisms may be difficult because the test organisms
are often not visible during the exposure. The operation
of the exposure system should be monitored daily.
11.3.5.1 Measurement of Overlying Water-quality
Characteristics: 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 pool water samples from individual repli-
cates. The pipet should be checked to make sure no
organisms are removed during sampling of overlying
water. Hardness, alkalinity, pH, conductivity, and ammo-
nia in the overlying water with a treatment should not
vary by more than 50% during a test,
11.3.5.1.1 Dissolved oxygen should be measured daily
and should be between 40 and 100% saturation (ASTM,
1988a). 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 contamination. Aeration can be used to
maintain dissolved oxygen in the overlying water above
40% saturation. Dissolved oxygen and pH can be mea-
sure directly in the overlying water with a probe.
11.3.5.1.2 Temperature should be measured at least
daily in at least one test chamber from each treatment.
The temperature of the water bath or the exposure
chamber should be continuously monitored. The daily
mean test temperature must be within ±1°C of the
desired temperature. The instantaneous temperature
must always be within ±3°C of the desired temperature.
11.3.6 Feeding: 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 de-
velop on the sediment surface. Therefore, the amount of
food added to the test chambers is kept to a minimum.
11.3.6.1 Suspensions of food should be thoroughly
mixed before aliquots are taken. If excess food collects
on the sediment, a fungal or bacterial growth may de-
velop on the sediment surface, in which case feeding
should be suspended for one or more days. A drop in
dissolved oxygen below 40% of saturation during a test
may indicate that the food added is not being consumed.
Feeding should be suspended for the amount of time
necessary to increase the dissolved oxygen concentra-
tion (ASTM, 1994a). If feeding is suspended in one
treatment, it should be suspended in all treatments.
Detailed records of feeding rates and the appearance of
the sediment surface should be made daily.
11.3.7 Ending a Test: 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 sedi-
ment surface or from sieved material should be consid-
ered dead. Ankley et al. (1994a) recommend using a
#25 sieve (710 |um mesh) to remove amphipods from
sediment. Alternatively, Kemble et al. (1994) recom-
mend sieving sediment using the following procedure:
(1) pour about half of the overlying water through a #50
(300 urn) U.S. Standard mesh sieve, (2) swirl the re-
maining water to 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 urn) mesh sieve and wash
the contents of this second sieve into a second exami-
nation pan. Surviving test organisms can be removed
from the two pans and preserved in 8% sugar formalin
solution for growth measurements (Ingersoll and Nelson,
1990). NoTox® (Earth Safe Industries, Belle Mead. NJ)
can be used as a substitute for formalin (Linger et al..
1993).
11.3.7.1 A consistent amount of time should betaken to
examine sieved material for recovery of test organisms
(e.g., 10 min/replicate). Laboratories should demon-
strate that their personnel are able to recover an aver-
age of at least 90% of the organisms from whole sedi-
ment. For example, test organisms could be added to
control or test sediments, and recovery could be deter-
mined after 1 h (Tomasovic et al., 1994).
11.3.8 Test Data: Survival is the primary endpoint
recorded at the end of the 10-d sediment toxicity test
with H. azieca. Measuring growth is optional; however.
growth of amphipods may be a more sensitive toxicity
endpoint compared to survival (Burton and Ingersoll.
1994; Kemble etal.. 1994). The duration of the 10-d test
started with 7- to 14-d old amphipods is not long enough
to determine sexual maturation or reproductive effects.
47
-------
11.3.8.1 Amphipod body length (±0.1 mm) can be
measured from the base of the first antenna to the tip of
the third uropod along the curve of the dorsal surface.
Ingersoll and Nelson (1990) describe the use of a digitiz-
ing system and microscope to measure lengths of H.
azteca. Antennal segment number can also be used to
estimate length or weight of amphipods (E.L Brunson,
NBS, Columbia, MO, personal communication). Wet or
dry weight measurements have also been used to esti-
mate growth of H. azteca (ASTM, 1994a). Dry weight of
amphipods should be determined by pooling all living
organisms from a replicate and drying the sample at
about 60 to 90°C to a constant weight. The sample is
brought to room temperature in a desiccator and weighed
to the nearest 0.01 mg to obtain mean weight per
surviving organism per replicate.
11.4 Interpretation of Results
11.4.1 Section 14 describes general information for
interpretation of test results. The following sections de-
scribe species-specific information that is useful in help-
ing to interpret the results of sediment toxicity tests with
H. azteca.
11.4.2 Age Sensitivity: The sensitivity of H. azteca
appears to be relatively similar up to at least 24- to 26-d
old organisms (Collyard et al., 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 organ-
isms through 24- to 26-d old organisms (Figure 11.1).
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 repre-
sentative of the sensitivity of H. azteca up to at least the
adult life stage.
11.4.3 Grain Size: Hyalella azteca are tolerant of a
wide range of substrates. Physico-chemical characteris-
tics (e.g., grain size or TOG) 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., !994a).
11.4.4 Isolating Organisms at the End of a Test:
Quantitative recovery of young amphipods (e.g., 0- to
7-d old) is difficult given their small size (Figure 11.2,
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: Survival
of H. azteca in 28-d tests was not reduced in the
presence of oligochaetes in sediment samples
(Reynoldson el al., 1994). However, growth of amphi-
pods was reduced when high numbers of oligochaetes
were placed in a sample. Therefore, it is important to
determine the number and biomass of indigenous or-
ganisms in field-collected sediment in order to better
interpret growth data (Reynoldson et al., 1994). Further-
more, presence of predators may also influence the
response of test organisms in sediment (Ingersoll and
Nelson, 1990).
48
-------
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0-2 2-4 6-8 8-10 12-14 16-18 20-22 24-26
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0-2 2-4 4-6 6-8 10-12 18-20 20-22
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400-
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12-14 24-26
Figure 11.1 Lifestage sensitivity of Hyalella azteca in 96-h water-only exposures.
49
-------
100
80
60
> 40
•84
63
193
1
197
7 14
Age (days)
21
Figure 11.2 Average recovery of different age Hyalella azteca from sediment by 7 individuals.
50
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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 de-
sirable characteristics of an ideal sediment toxicity test-
ing organism including relative sensitivity to contami-
nants associated with sediment, contact with sediment,
ease of culture in the laboratory, toterance to varying
physico-chemical characteristics of sediment, and short
generation time. Their response has been evaluated in
interlaboratory studies and has been confirmed with
natural benthos populations. Many investigators have
successfully used C. tentans to evaluate the toxicity of
freshwater sediments (e.g., Wentsel et al., 1977; Nebeker
et al., 1984a; Nebeker et al., 1988; Adams et al., 1985;
Giesy et al., 1988; Hoke et al., 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 (West et al., 1993; Hoke et al.,
1994; West et al., 1994; and Ankley et al., 1994c).
Endpoints typically monitored in sediment toxicity tests
with C. tentans include survival and growth (ASTM,
1994a).
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 and the litera-
ture cited in Table A.3 were used for developing test
method 100.2. Results of tests using procedures differ-
ent from the procedures described in Section 12.2 may
not be comparable and these different procedures may
alter contaminant bioavailability. Comparison of results
obtained using modified versions of these procedures
might provide useful information concerning new con-
cepts 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 com-
parability 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 14). 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 500 to
1000 lux (Table 12.1). Test chambers are 300-mL
high-form lipless beakers containing 100 ml of sedi-
ment and 175 mL of overlying water. Ten third-instar
midges are used to start a test. All organisms must be
third instar or younger with at least 50% of the organ-
isms at third instar. The number of replicates/treatment
depends on the objective of the test. Eight replicates are
recommended for routine testing (see Section 14). Midges
in each test chamber are fed 1.5 mL of a 4-g/L Tetrafin®
suspension daily. Each chamber receives 2 volume
additions/d of overlying water. Water renewals may be
manual or automated, and Appendix B describes
water-renewal systems that can be used to deliver over-
lying 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 sediment is collected. Requirements for test
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 mixed and added to the test cham-
bers (Section 8.3.1). Sediment should be visually in-
spected to judge the extent of homogeneity. Excess
water on the surface of the sediment can indicate sepa-
ration of solid and liquid components. If a quantitative
measure of homogeneity is required, replicate sub-
samples should be taken from the sediment batch and
analyzed for TOC, chemical concentrations, and particle
size.
51
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Table 12.1 Recommended Test Conditions for Conducting a 10-d Sediment Toxicity Test with Chironomus tentans
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 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 500 to 1000 lux
16L8D
high-form lipless beaker
100 mL
175 ml
2 volume additions/d; continuous or intermittent (e.g., one volume addition
every 12 h)
Third instar larvae (All organisms must be third instar or younger with at
least 50% of the organisms at third instar)
10
Depends on the objective of
the test. Eight replicates are recommended for routine testing (see
Section 14)
Tetrafin® goldfish food, fed 1.5 mL daily to each test chamber (1.5 mL
contains 4.0 rng of dry solids)
None, unless dissolved oxygen in overlying water drops below 40% of
saturation
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 B)
Hardness, alkalinity, conductivity, pH, and ammonia at the beginning and
end of a test. Temperature and dissolved oxygen daily
10d
Survival and growth (dry weight)
Minimum mean control survival of 70% and mean weight per surviving
control organism of 0.6 mg. Performance-based criteria specifications
outlined in Table 12.3
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
manner that 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
chambers (Day 0).
12.3.2 Renewal of overlying water: Renewal of over-
lying 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%. Mount
and Brungs (1967) diluters have been modified for sedi-
ment testing, and other automated water delivery sys-
tems have also been used (Maki, 1977; Ingersoll and
Nelson, 1990; Benoit et al., 1993; Zumwalt et al., 1994).
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 B describes water-renewal systems that
can be used for conducting sediment tests.
12.3.2.1 In water-renewal tests with one to four volume
additions of overlying water/d, water-quality characteris-
tics generally remain similar to the inflowing water
(Ingersoll and Nelson, 1990; Ankley et al., 1993); how-
ever, in static tests, water quality may change pro-
foundly 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 expo-
sure (Ingersoll and Nelson, 1990). Additionally, concen-
trations 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
52
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Table 12.2 General Activity Schedule (or Conducting a Sediment Toxicity Test with Chironomus tentans'
Day Activity
-14 Isolate adults for production of egg masses.
-13 Place newly deposited egg masses into hatching dishes.
-12 A larval rearing chamber is prepared with new substrate.
-11 Examine egg masses for hatching success. If egg masses 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 Feeo and observe midges. Measure water quality (e.g., temperature and dissolved oxygen).
-1 Add ;ood 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 lor 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 Same as Day 1. 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.
' Modified from Call et al.. 1994
contribute to the toxicity of the contaminants in the
sediment. Furthermore, changes in water-quality char-
acteristics such as hardness may influence the toxicity
of many inorganic (Gauss et al., 1985) and organic
(Mayer and Ellersieck, 1986) contaminants. Although
contaminant concentrations 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 interstitial water.
12.3.3 Acclimation: 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, acclimation of test organisms to the test water
is not required.
12.3.4 Placing Organisms in Test Chambers: 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 di-
rectly into overlying water (Ankley et al., 1993). Alterna-
tively, test organisms can be placed into 30-mL counting
cups that are floated in the test chambers for 15 min
before organisms are introduced into the overlying water
(Ingersoll and Nelson, 1990). Length or weight should
be measured on a subset of at least 20 organisms used
to start the test. Head capsule width of midges must be
measured on this subset of test organisms to determine
the instar used to start the test (Table 10.2).
12.3.5 Monitoring a Test: All chambers should be
checked daily and observations made to assess test
organism behavior such as sediment avoidance. How-
ever, monitoring effects on burrowing activity of test
organisms may be difficult because the test organisms
are often not visible during the exposure. The operation
of the exposure system should be monitored daily.
12.3.5.1 Measurement of Overlying Water-quality Char-
acteristics: 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 pool water samples from individual repli-
cates. The pipet should be checked to make sure no
organisms are removed during sampling of overlying
water. Hardness, alkalinity, pH, conductivity, and ammo-
nia in the overlying water within a treatment should not
vary by more than 50% during a test.
12.3.5.1.1 Dissolved oxygen should be measured daily
and should be between 40 and 100% saturation (ASTM,
1988a). 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 contamination. Aeration cart be used to
maintain dissolved oxygen in the overlying water above
40% saturation. Dissolved oxygen and pH can be mea-
sured directly in the overlying water with a probe.
12.3.5.1.2 Temperature should be measured at least
daily in at least one test chamber from each treatment.
The temperature of the water bath or the exposure
53
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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 third-instar and younger larvae. At least 50% of the larvae must be in the third instar at the start of the test.
2. Average survival of C. tenians 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 0.6 mg at the end of the test.
4. Hardness, alkalinity, pH, and ammonia in the overlying water within a treatment should not vary by more than 50% during the test.
B. Performance-based criteria tor culturing C. tentans include
1. Laboratories should perform monthly 96-h water-only reference-toxicity tests to assess the sensitivity of culture organisms. If refer-
ence-toxicity tests are not conducted monthly, the lot of organisms used to start a sediment test must be evaluated using a reference
toxicant (Section 9.16).
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 trie cultures at least quarterly and the day before the start of a
sediment test: pH, hardness, alkalinity, and ammonia. Dissolved oxygen should be measured weekly. Temperature should be
recorded daily.
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. It is desirable to start tests soon after collection of sediment from the field (see Section 8.2 for additional detail).
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.
6. The daily mean test temperature must be within ±1 °C of the desired temperature. The instantaneous temperature must always be
within ±3°C of the desired temperature.
7. Natural physico-chemical characteristics of test sediment collected from the field should be within the tolerance limits of the test
organisms.
chamber should be continuously monitored. The daily dissolved oxygen below 40% of saturation during a test
mean test temperature must be within ±1°C of the may indicate that the food added is not being consumed.
desired temperature. The instantaneous temperature Feeding should be suspended for the amount of time
must always be within ±3°C of the desired temperature, necessary to increase the dissolved oxygen concentra-
tion (ASTM, 1994a). If feeding is suspended in one
12.3.6 Feeding: Without addition of food, the test treatment, it should be suspended in all treatments.
organisms may starve during exposures. However, the Detailed records of feeding rates and the appearance of
addition of the food may alter the availability of the the sediment surface should be made daily.
contaminants in the sediment (Wiederholm et al., 1987;
Harkey et al., 1994). Furthermore, if too much food is 12.3.7 Ending a Test: Immobile organisms isolated
added to the test chamber or if the mortality of test from the sediment surface or from sieved material should
organisms is high, fungal or bacterial growth may de- be considered dead. Ankley et al. (1994a) recommend
velop on the sediment surface. Therefore, the amount of using a #25 sieve (710 jim mesh) to remove midges
food added to the test chambers is kept to a minimum, from sediment. Alternatively, Kemble et al. (1994) rec-
ommend sieving sediment using the following proce-
12.3.6.1 Suspensions of food should be thoroughly dure: (1) pour about half of the overlying water through a
mixed before aliquots are taken. If excess food collects #50 {300 urn) U.S. Standard mesh sieve, (2) pour about
on the sediment, a fungal or bacterial growth may de- half of the sediment through the #50 mesh sieve and
velop on the sediment surface, in which case feeding wash the contents of the sieve into an examination pan,
should be suspended for one or more days. A drop in (3) rinse the coarser sediment remaining in the test
54
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chamber through a #40 (425 nm) mesh sieve and wash
the contents of this second sieve into a second exami-
nation pan. Surviving midges can then be isolated from
these pans. See Section 12.3.8.1 and 12.3.8.2 for the
procedures for measuring weight or length of midges.
12.3.7.1 A consistent amount of time should be taken to
examine sieved material for recovery of test organisms
(e.g., 10 min/replicate). Laboratories should demon-
strate that their personnel are able to recover an aver-
age of at least 90% of the organisms from whole sedi-
ment. For example, test organisms could be added to
control sediment and recovery could be determined
after 1 h (Tomasovic et al., 1994).
12.3.8 Test Data: Dry weight and survival are the
endpoints measured at the end of the 10-d sediment
toxicity test with C. tentans. The duration 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 (Ankley et al., 1993; ASTM, 1994b;
Section 15).
12.3.8.1 Head capsule width can be measured on
surviving midges at the end of the test before dry weight
is determined. Dry weight of midges should be deter-
mined by pooling all living larvae from a replicate and
drying the sample at about 60 to 90°C to a constant
weight. The sample is brought to room temperature in a
desiccator and weighed to the nearest 0.01 mg to obtain
mean weights per surviving organism per replicate. Pu-
pae or adult organisms must not be included in the
sample to estimate dry weight.
12.3.8.2 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). 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 seg-
ment (Smock, 1980). Kemble et al. (1994) photographed
midges at magnification of 3.5x and measured the im-
ages using a computer-interfaced digitizing tablet. A
digitizing system and microscope can also be used to
measure length (Ingersoll and Nelson, 1990).
12.4 Interpretation of Results
12.4.1 Section 14 describes general information for
interpretation of test results. The following sections de-
scribe species-specific information that is useful in help-
ing to interpret the results of sediment toxicity tests with
C. tentans.
12.4.2 Age Sensitivity: Midges are perceived to be
relatively insensitive organisms in toxicity assessments
(Ingersoll, 1994). This conclusion is based on the prac-
tice of conducting short-term tests with 4th instar larvae
in water-only exposures, a procedure that may underes-
timate 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 ex-
ample, first instar C. tentans larvae were 6 to 27 times
more sensitive than 4th 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.
While 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.
12.4.3 Grain Size: Chironomus tentans are tolerant of a
wide range of substrates. Physicochemical characteris-
tics (e.g., grain size or TOC) of sediment were not
significantly correlated to the survival of C. tentans in
toxicity tests in which organisms were fed. However,
linear modeling indicated that growth of C. tentans may
have been slightly influenced by grain size distribution of
the test sediments (Section 10.1.2.3; Ankley et al.,
1994a). Survival of C. fen/answas reduced below 0.91%
organic matter in 10-d tests with formulated sediment
(Suedel and Rodgers, 1994); however these organisms
did not receive a supplemental source of nutrition.
12.4.4 Isolating Organisms at the End of a Test:
Quantitative recovery of larvae at the end of a 10-d
sediment test should not be a problem. The larvae are
red and typically greater than 5-mm long.
12.4.5 Influence of Indigenous Organisms: The influ-
ence of indigenous organisms on the response of C.
tentans in sediment tests has not been" reported. Sur-
vival of a closely related species, C. riparius was not
reduced in the presence of oligochaetes in sediment
samples (Reynoldson et al., 1994). However, growth of
C. riparius was reduced when high numbers of oligocha-
etes were placed in a sample. Therefore, it is important
to determine the number and biomass of indigenous
organisms in field-collected sediment in order to better
interpret growth data (Reynoldson et al., 1994). Further-
more, presence of predators may also influence the
response of test organisms in sediment (Ingersoll and
Nelson, 1990).
55
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A. Chironomus riparius: Cadium
2.5
1-5
S
X 1
CM
0.5
1st
1 st 2nd 3rd
INSTAR
B. Chironomus tentans: Copper
2nd 3rd
INSTAR
4th
Williams etal. (1986)
4th
Nebeker etal. (1984)
Figure 12.1 Ufestage sensitivity of chironomids.
56
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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 bioaccu-
mulation testing organism including contact with sedi-
ment, ease of culture in the laboratory, and tolerance to
varying physico-chemical characteristics of sediment.
The response of L. variegatus in laboratory exposures
has been confirmed with natural benthos populations.
Many investigators have successfully used L variegatus
in toxicity or bioaccumulation tests. Toxicity studies have
been conducted in water-only tests (Bailey and Liu,
1980; Hornig, 1980; Ewell et al., 1986; Nebeker et al.,
1989; Ankley et al., 1991 a, Ankley et al., 1991b), in
effluent tests (Hornig, 1980), and in whole-sediment
tests (Nebeker et al., 1989; Ankley etal., 1991 a, Ankley
et al., 1991b; Ankley et al., 1992a; Call et al., 1991;
Carlson et al., 1991; Phipps et al., 1993; West et al.,
1993). Several studies have reported the use of L.
variegatus to examine bioaccumulation of chemicals
from sediment (Schuytema et al., 1988; Nebeker et al.,
1989; Ankley et al., 1991b; Call et al., 1991; Carlson et
al., 1991; Ankley et al., 1993; Kukkonen and Landrum,
1994; and E.L. Brunson, NBS, Columbia, MO, unpub-
lished data). However, interlaboratory studies have not
yet been conducted with L. variegatus.
13.1.2 Additional research is needed on the standard-
ization of bioaccumulation procedures with sediment.
Therefore, Section 13.2 describes general guidance for
conducting a 28-d sediment bioaccumulation test with L.
variegatus. Methods outlined in Appendix A and the
literature cited in Table A.4 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 pro-
cedures for conducting sediment toxicity tests with L.
variegatus. A general activity schedule is outlined in
Table 13.3. 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 14). 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.
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 500 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 test-
ing (Section 14). To minimize depletion of sediment
contaminants, the ratio of total organic carbon in sedi-
ment 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. Appendix B describes water-renewal systems
that with minor modifications 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 sediment is collected. Requirements for
test acceptability are outlined in Table 13.4.
13.2.2.1 Before starting a 28-d sediment bioaccumula-
tion test with L. variegatus, a toxicity screening test
should be conducted for at least 4 d using procedures
outlined in Table 13.2 (E.L. Brunson, NBS, Columbia,
57
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Table 13.1 Recommended Test Conditions (or Conducting a 28-d Sediment Bioaccumulation Test with LumMculus 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 500 to 1000 lux
16L:8D
4- to 6-L aquaria with stainless steel screens or glass standpipes
1 L or more depending on TOC
1 L or mof e depending on TOC
2 volume additions/d; continuous or intermittent (e.g., one volume addition every 12 h)
Adults
Ratio of total organic carbon in sedi-
ment 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 14)
None
None, unless dissolved oxygen in overlying water drops below 40% of saturation
Culture water, well water, surface water, site water, or reconstituted water
It screens become clogged during the test, gently brush the outside of the screen (Appendix B)
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 outlined in Table 13.4.
MO, unpublished data). The preliminary toxicity screen-
ing test is conducted at 23°C with a 16L8D photoperiod
at a illuminance of about 500 to 1000 lux. Test chambers
are 300-mL high-form lipless beakers containing 100 mL
of sediment and 175 mL of overlying water. Ten adult
oligochaetes/replicate are used to start a test. Four
replicates are recommended for routine testing. Oli-
gochaetes are not fed during the test. Each chamber
receives 2 volume additions/d of overlying water. Ap-
pendix B describes 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 screening 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 sediment by L variegatus
may decrease bioaccumulation.
13.3 General Procedures
13.3.1 Sediment into Test Chambers: The day before
the sediment test is started (Day -1} each sediment
should be thoroughly mixed and added to the test cham-
bers (Section 8.3.1). Sediment should be visually in-
spected to judge the extent of homogeneity. Excess
water on the surface of the sediment can indicate sepa-
ration of solid and liquid components. If a quantitative
measure of homogeneity is required, replicate sub-
samples should be taken from the sediment batch and
analyzed for TOC, chemical concentrations, and particle
size.
13.3.1.1 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
manner that 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
chambers (Day 0).
13.3.2 Renewal of Overlying Water: Renewal of over-
lying water is recommended during a test. At any par-
ticular 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 automated water delivery
systems have also been used (Maki, 1977; Ingersoll and
Nelson, 1990; Benoit et al., 1993; Zumwatt et al., 1994).
Each water-delivery system should be calibrated before
a test is started to verify that the system is functioning
property. Renewal of overlying water is started on Day
58
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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±°C
Wide-spectrum fluorescent lights
about 500 to 1000 lux
16U80
300-mL high-form liptess beaker
100mL
175mL
2 volume additions/d; continuous or intermittent (e.g., one volume addition every 12 h)
Adults
10
4 minimum
None
None, unless dissolved oxygen in overlying water drops below 40% of saturation
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
4 d (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 outlined in Table 13.4
-1 before the addition of test organisms or food on Day 0
(Appendix B).
13.3.2.1 In water-renewal tests with one to four volume
additions of overlying water/d, water-quality characteris-
tics generally remain similar to the inflowing water
(Ingersoll and Nelson, 1990; Ankley et al., 1993); how-
ever, in static tests, water quality may change pro-
foundly 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 expo-
sure (Ingersoll and Nelson, 1990). Additionally, concen-
trations 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 char-
acteristics such as hardness may influence the toxicity
of many inorganic (Gauss et al., 1985) and organic
(Mayer and Ellersieck, 1986) contaminants. Although
contaminant concentrations 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 interstitial water.
13.3.3 Acclimation: 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, acclimation of test organisms to the test water
is not required.
13.3.4 Placing Organisms in Test Chambers: Isolate
oligochaetes for starting a test as described in Section
10.5.6.. A subset of L variegatus at the start the test
should be sampled to determine starting concentrations
of contaminants of concern. Mean group weights should
be measured on a subset of at least 100 organisms
used to start the test. The ratio of total organic carbon in
sediment to dry weight of organisms at the start of the
test should be no less than 50:1.
13.3.4.1 Oligochaetes added to each replicate should
not be blotted to remove excess water (Section 10.5.6).
Oligochaetes can be added to each replicate at about
1.33x of the target stocking weight (E.L. Brunson, NBS,
Columbia, MO, unpublished data). This additional 33%
should account for the excess weight from water in the
sample of nonblotted oligochaetes at the start of the
test.
59
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Table 13.3 General Activity Schedule for Conducting a 28-d Sediment Bioaccumulation Test with Lumbricutus variegatus
A. Conducting a 4-d Toxicity Screening Test (conducted before the 28-d bioaccumulation test)
Day Activity
-1 Isolate worms tor 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 -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.
1-6 Measure temperature and dissolved oxygen. Observe behavior of test organisms.
7 Same as Day 1. Measure total water quality.
8-13 Same as Day 1
14 Same as Day 7
15-20 Same as Day 1
21 Same as Day 7
22-26 Same as Day 1
27 Same as Day 1. Measure total water quality.
28 Measure temperature and dissolved oxygen. End uptake by collecting the worms with a sieve. Separate any indigenous organisms
from L. variegatus. Determine weight of survivors. Eliminate gut contents of surviving worms in water for 24 h.
29 Sample surviving worms after 24 h of elimination for chemical analysis.
13.3.5 Monitoring a Test: All chambers should be 13.3.5.1.1 Dissolved oxygen should be measured
checked daily and observations made to assess test daily and should be between 40 and 100% saturation
organism behavior such as sediment avoidance. How- (ASTM, 1988a), If a probe is used to measure dis-
ever, monitoring effects on burrowing activity of test solved oxygen in overlying water, it should be thor-
organisms may be difficult because the test organisms oughly inspected between samples to make sure that
are often not visible during the exposure. The operation organisms are not attached and should be rinsed be-
of the exposure system should be monitored daily. tween samples to minimize cross contamination. Aera-
tion can be used to maintain dissolved oxygen in the
13.3.5.1 Measurement of Overlying Water-quality overlying water above 40% saturation. Dissolved oxy-
Characteristics: Conductivity, hardness, pH, alkalinity, gen and pH can be measured directly in the overlying
and ammonia should be measured in all treatments at water with a probe.
the beginning and end of a test. Overlying water should
be sampled just before water renewal from about 1 to 2 13.3.5.1.2 Temperature should be measured at least
cm above the sediment surface using a pipet. It may be daily in at least one test chamber from each treatment.
necessary to pool water samples from individual repli- The temperature of the water bath or the exposure
cates. The pipet should be checked to make sure no chamber should be continuously monitored. The daily
organisms are removed during sampling of overlying mean test temperature must be within ±1°C of the
water. Hardness, alkalinity, pH, conductivity, and ammo- desired temperature. The instantaneous temperature
nia in the overlying water within a treatment should not must always be within ±3°C of the desired tempera-
vary by more than 50% during a test. ture.
60
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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, pH, and ammonia in the overlying water within a treatment should not vary by more than 50% during the test.
B. Performance-based criteria for culturing L. variegatus include:
1. Laboratories should perform monthly 96-h water-only reference-toxicity tests to assess the sensitivity of culture organisms. If
reference-toxicity tests are not conducted monthly, the lot of organisms used to start a sediment test must be evaluated using a
reference toxicant (Section 9.16).
2. Laboratories should monitor the frequency with which the population is doubling in the culture (number of organisms) and record tfiis
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.
3. Food used to culture organisms should be analyzed before the start of a test for compounds to be evaluated in the bioaccumulation
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 should be measured weekly. Temperature 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. It is desirable to start tests soon after collection of sediment from the field (see Section 8.2 for additional detail).
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.
6. The daily mean test temperature must be within ±1°C of the desired temperature. The instantaneous temperature must always be
within ±3°C of the desired temperature.
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 Feeding: Lumbriculus variegatus should not be maintain dissolved oxygen above 40% of saturation.
fed during a bioaccumulation test. Worms will clear more than 90% of the gut contents in
24 h (Call et a!., 1991). Following the 24-h elimination
13.3.7 Ending a Test: Sediment at the end of the test period, oligochaetes should be collected, placed in a
can be sieved through a fine-meshed screen sufficiently tared weigh boat, blotted to remove excess water, and
small to retain the oligochaetes (e.g., U.S. Standard weighed to determine wet weight. Each sample should
Sieve 35 (500 um mesh) or 40 (425 urn mesh)). The then be split into appropriate aliquots (e.g., metals,
sieved material should be quickly transferred to a shal- organics), placed in clean containers, and frozen for
low pan to keep oligochaetes from moving through the later analysis. Containers should be placed inside sec-
screen. Immobile organisms should be considered dead, ondary freezer containers to minimize "freezer burn" or
Live oligochaetes are transferred to a 1-L beaker con- dehydration during storage.
taining overlying water without sediment for 24 h to
eliminate gut contents. Oligochaetes should not be placed 13.3.7.1 Field-collected sediments may include indig-
in clean sediment to eliminate gut contents. Clean sedi- enous oligochaetes. The behavior and appearance of
ment can add 15 to 20% to the dry weight of the indigenous oligochaetes are usually different from L
oligochaetes which would result in a dilution of contami- variegatus. It may be desirable to test extra chambers
nant concentrations on a dry weight basis (Peter without the addition of L. variegatus to check for the
Landrum, NOAA, Ann Arbor, Ml, personal communica- presence of indigenous oligochaetes in field-collected
tion). The elimination beakers may need to be aerated to sediment (Phipps et al.. 1993). Bioaccumulation of con-
61
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Table 13.5 Grams of Lumbriculus variegatus Tissue (Wet
Weight) Required (or Various Analytes at Selected
Lower Limits ol Detection
Grams of Tissue
Table 13.6 Detection Limits
-------
Oliver and Niimi, 1983; de Boer, 1988). Because of the
importance of 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 14).
However, the difficulty with using this approach is that
each lipid method generates different lipid concentra-
tions (see Kates (1986) for discussion of lipid methodol-
ogy). The differences in lipid concentrations directly
translate to a similar variation in the lipid-normalized
contaminant concentrations or BSAF.
13.3.8.2.2 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 diffi-
cult because the lipid methodology is often intimately
tied in with the extraction procedure for contaminant
analysis. As an interim solution, the Bligh-Dyer lipid
method (Bligh and Dyer, 1959) is recommended as a
temporary "intercalibration standard" (Lee et al., 1994).
13.3.8.2.3 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, 1983; de Boer, 1988; Landrum, 1989). Be-
cause the technique is independent of any particular
analytical extraction procedure, it will not change when
the extraction 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 et al., 1985).
13.3.8.2.4 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
residues reported in "Bligh-Dyer equivalents." In the
interim, 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 (Lee et
al., 1994).
13.4 Interpretation of Results
13.4.1 Section 14 describes general information for
interpretation of test results. The following sections de-
scribe species-specific information that is useful in help-
ing to interpret the results of sediment bioaccumulation
tests with L variegatus.
13.4.2 Duration of Exposure: Because data from
bioaccumulation tests often will be used in ecological or
human health risk assessments, the procedures are
designed to generate quantitative estimates of
steady-state tissue residues. Eighty percent of
steady-state is used as the general criterion (Lee et al.,
1994). Because results from a single or few species
often will be extrapolated to other species, the proce-
dures are designed to maximize exposure to
sediment-associated contaminants so as not to system-
atically underestimate residues in untested species.
13.4.2.1 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; E.L. Brunson, NBS, Columbia, MO, unpublished
data; USEPA-USCOE, 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 po-
tential).
13.4.2.2 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, fluorene, phenanthrene) generally
peaked at Day 3 and tended to decline to Day 56 (E.L.
Bruson, NBS, Columbia, MO, unpublished data). 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:
Field-collected sediments may include indigenous oli-
gochaetes. Phipps et al. (1993) recommend testing
extra chambers without the addition of L. variegatus to
check for the presence of indigenous oligochaetes in
field-collected sediment.
13.4.4 Sediment Toxicity in Bioaccumulation Tests:
Toxicity or altered behavior of organisms in a sample
may not preclude use of bioaccumulation data; how-
ever, information on adverse effects of a sample should
be included in the report.
13.4.4.1 Grain Size: Lumbriculus variegatus are toler-
ant of a wide range of substrates. Physicochemtcal
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 et al., 1994a).
13.4.4.2 Sediment Organic Carbon: Reduced growth
of L variegatus may result from exposure to sediments
with low organic carbon concentrations (G.T. Ankley,
USEPA, Duluth, MM, personal communication). There-
fore, number of organisms and behavior in the 4-d
toxicity screening test should be the criteria used to
judge the acceptability of a bioaccumulation test.
63
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Section 14
Data Recording, Data Analysis and Calculations, and Reporting
14.1 Data Recording
14.1.1 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.
14.1.2 A file should be maintained for each sediment
test or group of tests on closely related samples (Sec-
tion 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 toxi-
cants; 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 instruments; test conditions used; and results of
reference toxicant tests. Original data sheets should be
signed and dated by the laboratory personnel perform-
ing the tests.
14.1.3 Example data sheets are included in Appendix
D.
14.2 Data Analysis
14.2.1 Statistical methods are used to make inferences
about populations, base^ on samples from those popu-
lations. In most sediment toxicity and bioaccumulation
tests, test organisms are exposed to contaminated sedi-
ment to estimate the response of the population of
laboratory organisms. The organism response to these
contaminated sediments is usually compared with the
response to a control or reference sediment, or in some
analyses of bioaccumulation test data, with a fixed stan-
dard such as an Food and Drug Administration (FDA)
action level. In any toxicity or bioaccumulation test,
summary statistics such as means and standard errors
for response variables (e.g., survival, contaminant levels
in tissue) should be provided for each treatment (e.g.,
pore-water concentration, sediment).
14.2.1.1 Types of Data. Two types of data can be
obtained from sediment toxicity or bioaccumulation tests.
The most common endpoint in toxicity testing is mortal-
ity, which is a dichotomous or categorical type of data.
Other endpoints commonly encountered in sublethal
evaluations are growth (e.g. in sediment toxicity tests
conducted with amphipods and midges) and tissue con-
centrations (e.g. in sediment bioaccumulation tests con-
ducted with oligochaetes or polychaetes and mollusks;
USEPA, 1994a). These types of endpoints are repre-
sentative of continuous data.
14.2.1.2 Sediment Testing Scenarios. 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 stud-
ies designed to (1) evaluate hazards of dredged mate-
rial, (2) assess site contamination in the environment
(e.g., to rank areas for cleanup), and (3) determine
effects of specific contaminants, or combinations of
contaminants, 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.
14.2.1.2.1 Dredged Material Hazard Evaluation. In
these studies, n sites are compared individually to a
reference sediment. The statistical procedures appropri-
ate for these studies are generally pairwise compari-
sons. Additional information on toxicity testing of dredged
material and analysis of data from dredged material
hazard evaluations is available in USEPA-USCOE 1994.
14.2.1.2.2 Site Assessment of Field Contamination.
Surveys of sediment toxicity or bioaccumulation often
are included in more comprehensive analyses of biologi-
cal, chemical, geological, and hydrographic data. Statis-
tical correlation can be improved and costs may be
reduced if subsamples are taken simultaneously for
sediment toxicity or bioaccumulation tests, chemical
analyses, and benthic community structure determina-
tions. There are several statistical approaches to field
assessments, each with a specific purpose. If the objec-
tive 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 compare among all sites in the study
area, then a multiple comparison procedure that em-
ploys 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 tech-
nique.
64
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14.2.1.2.3 Sediment-Spiking Experiments. Sediments
spiked with known concentrations of contaminants 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, or LOEC. Results of bioac-
cumulation tests with either field or spiked samples may
be reported in terms of a BSAF (biota-sediment accu-
mulation factor, Ankley et al. 1992b). 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 toxicant
tests.
14.2.2 The guidance outlined below on the analysis of
sediment toxicity and bioaccumulation test data is
adapted from a variety of sources including Lee et al.
(1994), USEPA (19933), USEPA (1993b), USEPA
(1993C), and USEPA-USCOE (1994). The objectives of
a sediment toxicity or bioaccumulation test are to quan-
tify contaminant effects on or accumulation in test or-
ganisms exposed to natural or spiked sediments or
dredged materials and to determine whether these ef-
fects are statistically different from those occurring in a
control or reference sediment. Each experiment con-
sists of at least two treatments: the control and one or
more test treatment(s). The test treatment(s) consist(s)
of the contaminated or potentially contaminated
sediment(s). A control sediment is always required to
ensure that no contamination is introduced during the
experiment setup and that test organisms are healthy. A
control sediment is used to judge the acceptability of the
test. Some designs will also require a reference sedi-
ment that represents an environmental condition or po-
tential treatment effect of interest.
14.2.2.1 Experimental Unit. During toxicity testing,
each test chamber to which a single application of
treatment is applied is an experimental unit. During
bioaccumulation testing, however, 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 independent and should
not differ systematically.
14.2.2.2 Replication. 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 between treatments.
14.2.2.3 Minimum Detectable Difference (MOD). As
the minimum difference between treatments which the
test is required or designed to detect decreases, the
number of replicates required to meet a given signifi-
cance level and power increases. Because no consen-
sus currently exists on what constitutes a biologically
acceptable MOD, the appropriate statistical minimum
significant difference should be a data quality objective
(DQO) established by the individual user (e.g., program
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.
14.2.2.4 Minimum Number of Replicates. Four repli-
cates per treatment or control are the absolute minimum
number of replicates for a sediment toxicity test. How-
ever, USEPA recommends five replicates for marine
testing (USEPA, 1994) or eight replicates for freshwater
testing for each control or experimental treatment, tt is
always prudent to include as many replicates in the test
design as are economically and logistically possible. A
minimum of five replicates per treatment also is recom-
mended for bioaccumulation testing. USEPA sediment
toxicity testing methods recommend the use of 20 or-
ganisms per replicate for marine testing {USEPA, 1994a)
or 10 organisms per replicate for freshwater testing. An
increase in the number of organisms per replicate in all
treatments, including the control, is allowable only if (1)
test performance criteria for the recommended number
of replicates are achieved and (2) it can be demon-
strated that no change occurs in contaminant availability
due to the increased organism loading.
14.2.2.5 Randomization. Randomization is the unbi-
ased 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), (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) random-
ize the location of exposure units.
14.2.2.6 Pseudoreplication. The appropriate assign-
ment of treatments to the replicate exposure chambers
is critical to the avoidance of a common error in design
and analysis termed "pseudoreplication" (Hurlbert, 1984).
Pseudoreplication occurs when inferential statistics are
used to test for treatment effects even though the treat-
ments are not replicated or the replicates are not statis-
tically 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, the
other with test sediment, and 10 organisms are placed in
each aquarium. Even if each organism is analyzed
individually, the 10 organisms only replicate the biologi-
cal 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 pseudoreplication is the potential sys-
tematic error due to the physical segregation of expo-
sure chambers by treatment. For example, if all the
control exposure chambers are placed in one area of a
65
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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 ran-
domization of treatment location may be necessary to
avoid this type of pseudoreplication. Pseudoreplication
can be avoided or reduced by properly identifying the
experimental unit, providing replicate experimental units
for each treatment, and applying the treatments to each
experimental unit in a manner that includes random
physical intermixing (interspersion) and independence.
However, avoiding pseudoreplication completely may
be difficult or impossible given resource constraints.
14.2.2.7 Compositing Samples. Compositing is used
primarily in bioaccumulation experiments when the bio-
mass of an individual organism is insufficient for chemi-
cal analysis. Compositing consists of combining samples
(e.g., organisms, sediment) and chemically analyzing
the mixture rather than the individual samples. The
chemical analysis of the mixture provides an estimate of
the average 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 analysis. 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 compos-
ites 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 com-
posite equivalent in size. If sample sizes vary, consult
the tables in Schaeffer and Janardan (1978) to deter-
mine if replicate composite variances will be higher than
individual sample variances, which would make
compositing inappropriate.
14.2.3 The purpose of a toxicity or bioaccumulation test
is to determine if the biological response to a treatment
sample differs from the response to a control sample.
Figure 14.1 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.
Decision
TR =Control
TR > Control
TR =Control
TR > Control
Correct
1 -«
Type I
Error
u
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
(he probability of making a Type II statistical error (talse negative).
Figure 14.1 Treatment response tor a Type I and Type II error.
The alternative hypothesis of greatest interest in sedi-
ment tests is that the treatments are toxic, or contain
concentrations of bioaccumulatable compounds, rela-
tive to the control or reference sediment.
14.2.3.1 Statistical tests of hypotheses can be designed
to control for the chances of making incorrect decisions.
In Figure 14.1, 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
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 mak-
ing a Type I error. Historically, having chosen a, environ-
mental researchers have ignored p and the associated
power of the lest (1-P).
14.2.3.2 Fairweather (1991) presents a review of the
need for, and the practical implications of, conducting
power analysis in environmental monitoring studies. This
review also includes a comprehensive bibliography of
recent 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 the most important criterion 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 degrada-
tion and the eventual cost of its rectification)."
14.2.3.3 The critical components of the experimental
design associated with the test of hypothesis 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,
66
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(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 se-
lected probabilities of Type I (a) and Type II (P) errors.
14.2.3.4 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 |1 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.
14.2.3.5 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 t-test
with its associated assumptions is the appropriate para-
metric analysis. If the desired MOD, the number of
replicates per treatment, the number of organisms per
replicate and an estimate of typical among replicate
variability, such as the coefficient of variation (CV) from
a control sample, are available, it is possible to use a
graphical approach as in Figure 14.2 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 treat-
ment 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 theCV is 15% or less (Figure 14.2). The choice of
these variables also affects the power of the test. If 5
replicates are used per treatment (Figure 14.3), 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%.
1 -r
0.9-•
10
20 30 40 50
% Reduction of Control Mean
60
70
Figure 14.2 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)).
67
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1 T
0.9-
10
20
30 40
% Reduction of Control Mean
60
70
Figure 14.3 Power of the test vs. percent reduction in treatment response relative to the control mean at various CVs (5 repli-
cates, alpha = 0.05 (one-tailed)).
14.2.3.6 Relaxing the a level of a statistical test in-
creases the power of the test. Figure 14.4 duplicates
Figure 14.2 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 14.2 illustrates that with a
CV of 15% and an a level of 0.05, there is an 80%
probability (power) of detecting a 20% reduction in the
mean treatment response relative to the control mean.
However, if a is set at 0.10 (Figure 14.4) 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 interpreta-
tion of the data is desirable.
14.2.3.7 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 (Fig-
ure 14.5). Note, however, that for less than 8 replicates
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 treat-
ment, then it may be necessary to find ways to reduce
the among replicate variability and consequently the CV.
Options that are available include selecting more uni-
form organisms to reduce biological variability or in-
creasing the a level of the test. For CVs in the range of
30% to 40%, even eight replicates per treatment is
inadequate to detect small reductions (<20%) in re-
sponse relative to the control mean.
14.2.3.8 The effect of the choice of a and (3 on number
of replicates for various CVs is illustrated in Figure 14.6
in which the combined total probability of Type I and
Type II statistical errors is fixed and assumed to be 0.25.
An a of 0.10 therefore establishes a p of 0.15. In Figure
14.6, if a = 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 repli-
cates required to detect the same 20% difference rela-
tive to the control increases. However, the curves are
68
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10
20 30 40 50
% Reduction of Control Mean
60
70
Figure 14.4 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)).
relatively flat over the range of 0.05 to 0.20 and the
curves are very dependent upon the choice of the
combined total of a + p. Limiting the total of a + p to 0.10
greatly increases the number of replicates necessary to
detect a preselected percentage reduction in mean treat-
ment response relative to the control mean.
14.2.4 Figure 14.7 outlines a decision tree for analysis
of survival and growth data subjected to hypothesis
testing. In the tests described herein, samples or obser-
vations refer to replicates of treatments. Sample size n
is the number of replicates {i.e., exposure chambers) in
an individual treatment, not the number of organisms 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
(1980) may also be helpful.
14.2.4.1 Mean. The sample mean (\) is the average
value, or £Xj In, where
n = number of observations (replicates)
x( = ith observation
Ix, = every x summed = x, + x2 + x3 + ... + xn
14.2.4.2 Standard Deviation. The sample standard
deviation (s) is a measure of the variation of the data
around the mean and is equivalent to % s2. The sample
variance, s2, is given by the following "machine" or
"calculation" formula:
69
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,. CV = 5%
I
£
0.2
8 10
No. of Replicates (n)
12
14
16
Figure 14.5 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)).
T- -
-------
0 I I I I I I I I I I I I I I I I I I I I I I I
0.01 0.03 0.05 0.07 0.09 0.11 0.13 0.15 0.17 0.19 0.21 0.23
Alpha (Beta = 0.25 - Alpha)
Figure 14.6 Effect of alpha and beta on the number of replicates at various CVs (assuming combined alpha + beta = 0.25).
metric tests, but careful laboratory practices can reduce
the frequency of outliers.
14.2.4.4.2 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
because subtracting the treatment mean removes any
differences among treatments. This scales the observa-
tions so that the mean of residuals for each treatment
and overall 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. W will gener-
ally vary between 0.3 and 1.0, with lower values indicat-
ing 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.
14.2.4.4.3 Table 14.1 provides a levels to determine
whether departures from normality are significant. Nor-
mality should be rejected when the probability associ-
ated 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 cham-
bers. 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 departures from normality have the
greatest effects on t-tests and other parametric compari-
sons. If data fail the test for normality, even after trans-
formation, nonparametric tests should be used for addi-
tional analyses.
14.2.4.4.4 Tables of quantiles of W can be found in
Shapiro and Wilk (1965), Gill (1978), Conover (1980),
USEPA (1989b) and other statistical texts. These refer-
ences also provide methods of calculating W, although
the calculations can be tedious. For that reason, com-
monly available computer programs or statistical pack-
ages are preferred for the calculation of W.
14.2.4.4.5 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 (n=5) (Conover et al., 1981), and any one of these
tests is adequate for the analyses in this section. Many
software packages for t-tests and analysis of variance
71
-------
Data—Survival, Growth, Etc.
Test for Normality
Normal
Shapiro-Wilk's Test (N<60)
Non-Normal •
Tests for Homogeneity of Variance
t_
• Transformation?
Bartlett's || Hartley's |
Heterogenous Variances
Rankits I -
No
Homogenous Variances
Yes, N>2 X
No, N-2
| >3 Replicates [
Yes
T
I Anova |
Equal Repl
1 No
cation
Yes
—+>
Comparison-Wise Alpha _^
Fisher's LSD, Duncan's *"
1 Experiment-Wise Alpha "^
Dunnett's 1
Equal Replication
JYes
Steel's
Many-One
Rank Test
r W
INO
Wilcoxon
w/
Bonferroni's
|
Figure 14.7 Decision tree for analysis of survival and growth data subjected to hypothesis testing.
(ANOVA) provide at least one of the tests. Bartlett's Test
is recommended for routine evaluation of homogeneity
of variances (USEPA, 1985; USEPA, 1994b;
USEPA,1994c).
14.2.4.4.6 If no tests for equality of variances are
included in the available statistical software, Hartley's
Fmax can easily be calculated:
Fmax = (larger of s, , s^ ) / ( smaller of s? . s; )
Table 14.1 Suggested a Levels to Use for Tests of Assumptions
Test
Normality
Equality of variances
Number of
Observations'
N = 2 to 9
N = 10 to 19
N = 20 or more
n = 2 to 9
n = 10 or more
a When
Balanced
0.10
0.05
0.01
0.10
0.05
Design Is
Unbalanced2
0.25
0.10
0.05
0.25
0.10
N * total number of observations (replicates) in all treatments
combined, n = number of observations (replicates) in an individual
treatment
n > 2 n
When F is large, the hypothesis of equal variances is
more likely to be rejected. F is a two-tailed test
because it does not matter which variance is expected
to be larger. Some statistical texts provide critical values
of Fmax (Winer, 1971; Gill, 1978; Rohlf and Sokal, 1981).
14.2.4.4.7 Levels of a for tests of equality of variances
are provided in Table 14.1. 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.
14.2.4.5 Transformations of the Data. When the
assumptions of normality or homogeneity 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 proportion 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
72
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assumption of normality and there are four or more
replicates per group, 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 F test for equality of variances
is used to test the homogeneity of variance assumption.
Failure of the homogeneity of variance assumption leads
to the use of a modified t test, and the degrees of
freedom for the lest are adjusted.
14.2.4.5.1 The arc sine-square root transformation con-
sists of determining the angle (in radians) represented
by a sine value. In this transformation, the proportion
surviving 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 example of the arc sine-square root
transformation and modification are provided below.
1. Calculate the response proportion (RP) for each
replicate within a group, where
RP = (number of surviving organisms)/(number
exposed)
2. Transform each RP to arc sine, as follows:
a. For RPs greater than zero or less than one:
Angle (in radians) = arc sine
b. Modification of the arc sine when RP = 0.
Angle (in radians) = arc sine J—
V 4n
where n = number of animals/treatment rep.
c. Modification of the arc sine when RP = 1.0.
Angle = 1.5708 radians-fradians for RP = 0)
14.2.4.6 Two Sample Comparisons (N=2). The true
population mean (n) 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 /-values account for this uncer-
tainty. 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 t-value. Student M/alues
decrease with increasing sample size because larger
samples provide a more precise estimate of n and o.
14.2.4.6.1 When using a t table, it is crucial to determine
whether the table is based on one-tailed probabilities or
two-tailed probabilities. In formulating a statistical hy-
pothesis, 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
analyzed are equal. A one-sided alternative hypothesis
(Ha) is that there is a specified relationship between the
two values (e.g., one value is greater than 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 differ-
ence is not important or cannot be assumed before
testing.
14.2.4.6.2 Since control organism mortality or tissue
residues and sediment contaminant concentrations are
presumed lower than reference or treatment sediment
values, 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). This is a critical consideration 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.
14.2.4.6.3 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 alternative 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).
14.2.4.6.4 The t-value for a one-tailed probability may
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 t-value for a = 0.05 and df = 20
is 1.725, and is found in a two-tailed table using the
column for a = 0.10.
14.2.4.7 The usual statistical test for comparing two
independent samples is the two-sample t-test (Snedecor
and Cochran, 1989). The t-statistic for testing the equal-
ity of means x 1 andx2 from two independent samples
with n1 and n2 replicates and unequal variances is
H,
where s; 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 I
73
-------
distribution with degrees of freedom (df) given by
Satterthwaite's (1946) approximation:
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 order to 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 replicates in the treatments, thet-test with Bonferroni's
adjustment can be used for data analysis (USEPA,
1993b; USEPA, 1993c). When variances are equal, an
Ftest for equality is unnecessary.
14.2.4.8 Nonparametric Tests. Tests such as the
t-test, which analyze the original or transformed data
and which rely on the properties of the normal distribu-
tion, are referred to as parametric tests. Nonparametric
tests, which do not require normally distributed data,
analyze the ranks of data and generally compare medi-
ans rather than means. The median of a sample is the
middle or 50th percentile observation when the data are
ranked from smallest to largest. In many cases, non-
parametric tests can be performed simply by converting
the data to ranks or normalized ranks (rankits) and
conducting the usual parametric test procedures on the
ranks or rankits.
14.2.4.8.1 Nonparametric tests are useful because of
their generality but have less statistical power than
corresponding parametric tests when the parametric
test assumptions are met. If parametric tests are not
appropriate for comparisons because the normality as-
sumption is not met, data should be converted to nor-
malized ranks (rankits). Rankits are simply the z-scores
expected for the rank in a normal distribution. Thus,
using rankits imposes a normal distribution over all 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 for-
mula:
rankit =
. 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).
14.2.4.8.2 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 un-
equal variances using the rankits or ranks.
14.2.4.9 Analysis of Variance (N>2). Some experiments
are set up to compare more than one treatment with a
control while others may also be interested in comparing
the treatments with one another. The basic design of
these experiments is the same as for experiments evalu-
ating pairwise comparisons. After the applicable com-
parisons are determined, the data must to be tested for
normality to determine if parametric statistics are appro-
priate 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 transfor-
mations of the data may be appropriate or nonparamet-
ric statistics can be used to test for equal means. Tests
for normality of the data should be performed on the
treatment residuals. A residual is defined as the ob-
served value minus the treatment mean, that is, rik = olk
- (kth treatment mean). Pooling residuals provides an
adequate sample size to test the data for normality.
14.2.4.9.1 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 variances. 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 treat-
ments with 0 variance may be tested against the other
treatment means using the appropriate mean compari-
son. Equality of variances among the treatments can be
evaluated with the Hartley F^ test or Bartlett's test. The
option of using nonparametric statistics on the entire set
of data ia also an alternative.
14.2.4.9.2 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
treatment 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 re-
quires data analysis with the Wilcoxon Rank Sum test
with Bonferroni's adjustment. Steel's Many-One Rank
test is a nonparametric test for comparing treatments
with a control. This test is an alternative to the Dunnett's
Procedure, and may be applied to data when the nor-
mality assumption has not been met. Steel's test re-
quires equal variances across treatments and the con-
trol but is thought to be fairly insensitive to deviations
from this condition (USEPA, 1993a). Wilcoxon's Rank
Sum Test is a nonparametric test to be used as an
alternative to the Steel's test when the number of repli-
cates are not the same within each treatment. A
Bonferroni's adjustment of the pairwise error rate for
74
-------
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, 1993a).
14.2.4.9.3 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 indi-
vidual comparison is performed independently at a
smaller a (than used in an experiment-wise comparison)
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 par-
ticular 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 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.
14.2.4.9.4 The recommended procedure for pairwise
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 com-
parison test should give results similar to the LSD. If the
treatments do not contain equal numbers of replicates,
the appropriate analysis is the t-test with Bonferroni's
adjustment. For comparisons that maintain an
experiment-wise a error rate Dunnett's test is recom-
mended for comparisons with the control.
14.2.4.9.5 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. Dunnett's procedure can only be used
when the same number of replicate test chambers have
been used at each treatment and the control.
14.2.4.9.6 To perform the individual comparisons, cal-
culate the 1 statistic for each treatment and control
combination, as follows:
where Y! = mean for each treatment
Y, = mean for the control
Sw = square root of the within mean square
n, = number of replicates in the control
n = number of replicates for treatment "i"
To quantify the sensitivity of the Dunnett's test, the
minimum significant difference (MSD=MDD) may be
calculated with the following formula:
where d = Critical value for the Dunnett's Proce-
dure
S =
n =
The square root of the within mean
square
The number of replicates per treatment.
assuming an equal number of replicates
at all treatment concentrations
n, = Number of replicates in the control
14.2.5 Methods for Calculating LCSOs,
ECSOs, and ICps
14.2.5.1 Figure 14.8 outlines a decision tree for analysis
of point estimate data. USEPA manuals (USEPA, 1985;
USEPA, 1989b; USEPA, 1993b; USEPA, 1993c) dis-
cuss in detail the mechanics of calculating LC50 (or
EC50) or ICp values using the most current methods.
Data Survival Point Estimates
I
Two or More Partial Mortalities
Yes J. No
Significant Chi Square Test One Partial Mortality
Yes | No
Yes
No
Graphical
| Linear Interpolation]
Trimmed Spearman-Karber I
LC50 and 95% Confidence Intervals
Figure 14.8 Decision tree for analysis of point estimate data.
75
-------
The most commonly used methods are the Graphical,
Probit, Trimmed Spearman-Karber and the Linear Inter-
polation Methods. In general, results from these meth-
ods should yield similar estimates. Each method is
outlined below and recommendations are presented for
me use of each method.
14.2.5.2 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 concen-
trations. 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.
14.2.5.3 Due to the intensive nature of the calculations
for the 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 with the methods discussed below (except for
the Graphical Method) were developed by USEPA and
can be obtained by sending a diskette with a written
request to USEPA, Environmental Monitoring Systems
Laboratory (EMSL), 26 W. Martin Luther King Drive,
Cincinnati, OH 45268 or call 513/569-7076.
14.2.5.4 Graphical Method. This procedure estimates
an LC50 (or EC50) by linearly interpolating between
points of a plot of observed percentage mortality versus
the base 10 logarithm (Iog10) of treatment concentration.
The only requirement for its use is that treatment mor-
talities bracket 50%.
14 2.5.4.1 For an analysis using the Graphical Method
the data should first be smoothed and adjusted for
mortality in the control replicates. The procedure for
smoothing and adjusting the data is detailed in the
following steps: Let p0, pv ..., pk denote the observed
proportion mortalities for the control and the k treat-
ments. 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 p, , then
where pf = the smoothed observed proportion mor-
tality for concentration i.
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:
Pf = (Pf-p5)/(i-P§)
where pf = the smoothed observed proportion mor-
tality for the control
pf = the smoothed observed proportion mor-
tality for concentration i.
14.2.5.5 The Probit Method. This method is a para-
metric statistical procedure for estimating the LC50 (or
EC50) and the associated 95% confidence interval
(Finney, 1971). The analysis consists of transforming
the observed proportion mortalities with a Probit trans-
formation, and transforming the treatment concentra-
tions to Iog10. Given ihe assumption of normality for the
tog10 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 correspond-
ing to 70% mortality is z70 or =.52. The LC50 is calcu-
lated from the regression and is the concentration asso-
ciated 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 associated 95%
confidence interval, two or more of the observed propor-
tion mortalities must be between zero and one. The
original percentage mortalities should be corrected for
control mortality using Abbott's formula (Section
14.2.5.4.1; Finney, 1971) before the Probit transforma-
tion is applied to the data.
14.2.5.5.1 A goodness-of-fit procedure with the
chi-square statistic is used to determine if 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. Most computer programs
that generate Probit estimates also generate confidence
interval estimates for the LC50. These confidence inter-
val estimates on the LC50 may not be correct if replicate
mortalities are pooled to obtain a mean treatment re-
sponse (USEPA-USCOE, 1994). This can be avoided
by entering the Probit-transformed replicate responses
and doing a least squares regression on the trans-
formed data.
14.2.5.6 The Trimmed Spearman-Karber Method.
The Trimmed Spearman-Karber Method is a modifica-
tion of the Spearman-Karber, nonparametric statistical
procedure for estimating the LC50 and the associated
95% confidence interval (Hamilton et al., 1977). This
procedure estimates the trimmed mean of the distribu-
76
-------
tion of the Iog10 of the tolerance. If the log tolerance
distribution is symmetric, this estimate of the trimmed
mean is equivalent to an estimate of the median of the
log tolerance distribution. Use of the Trimmed
Spearman-Karber Method is only appropriate when the
requirements for the Probit Method are not met (USEPA,
1993b; USEPA, 1993c). This method is only appropriate
for lethality data sets.
14.2.5.6.1 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 for the LC50 estimate,
one or more of the smoothed, adjusted, observed pro-
portion mortalities must be between zero and one.
14.2.5.6.2 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 14.2.5.4.1). Calculate the amount
of trim to use in the estimation of the LC50 as follows:
Trim = max(pf, 1-p£)
where pf = the smoothed, adjusted proportion mor-
tality for the lowest treatment concen-
tration, exclusive of the control.
p£ = the smoothed, adjusted proportion mor-
tality for the highest treatment concen-
tration.
k = the number of treatment concentrations,
exclusive of the control.
14.2.5.7 Linear Interpolation Method. This method
calculates a toxicant concentration that causes a given
percent reduction (e.g., 25%, 50%, etc.) in the endppint
of interest and is reported as an ICp value (1C = Inhibi-
tion Concentration; where p = the percent effect). The
procedure was designed for general applicability in the
analysis of data from chronic toxicity tests, and the
generation of an endpoint from a continuous model that
allows a traditional quantitative assessment of the preci-
sion of the endpoint, such as confidence limits for the
endpoint of a single test, and a mean and coefficient of
variation for the endpoints of multiple tests.
14.2.5.7.1 As described in USEPA (1993b; 1993c), 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 re-
sponse for the previous concentration, (2) follow a piece-
wise linear response function, and (3) are from a ran-
dom, 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 ana
identically distributed.
1 4.2.5.7.2 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 re-
sponse of interest, the (p) percent reduction from the
control.
14.2.5.7.3 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 concentra-
tion are considered in order of increasing concentration.
starting 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 ( Y,), it is used
as the response. If it is larger than the control mean, il 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 com-
pared to the mean observed _response for the next
higher toxicant concentration (Y3). Again, if the mean
observed response for the next higher toxicant concen-
tration 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 first two, and the mean is used as the response
for the control and two lowest concentrations of toxicant.
This process is continued for data from the remaining
toxicant concentrations. Unusual patterns in the devia-
tions from monotonicity may require an additional step
of smoothing. Where Y, decrease monotonically. the Y,
become M; without smoothing.
14.2.5.7.4 To obtain the ICp estimate, determine the
concentrations C , and CJf, which bracket the response
M, (1 - p/100), where M, 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
calculated as follows:
where C
tested concentration whose observed
mean response >s greater than M,(1 -
p/100).
tested concentration whose observed
mean response is less than M,(1 - p/
100).
77
-------
M. = smoothed mean response for the con-
trol.
Mj = smoothed mean response for con-
centration J.
Mjf,= smoothed mean response for con-
centration J -i- 1.
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.
14.2.5.7.5 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 for the true mean.
In the bootstrap method, the test data Yj; 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* esti-
mate. The distribution 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 estimated by the standard deviation of the
individual ICp* estimates. Empirical confidence intervals
are derived from the quantiles of the ICp* empirical
distribution. For example, 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 undetec-
ted 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.
14.2.6 Analysis of Bioaccumulation Data. In some
cases, body burdens will not approach steady-state
body burdens in a 28-d test (Lee et al., 1994). Organic
compounds exhibiting these kinetics will probably 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 metals
may gradually increase over time so that 28 d is inad-
equate 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.
14.2.6.1 Biotic Sampling. In the long-term studies, the
exposure should continue until steady-state body bur-
dens are attained. ASTM (1988b) recommends a mini-
mum of five sampling periods (plus t0) when conducting
water exposures to generate bioconcentration factors
(BCFs). Sampling in a geometric progression is also
recommended with sampling 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.
14.2.6.1.1 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 contaminant 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 greater than 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 versus Day 42) are not critical, though for
comparative purposes, samples should be taken at t28.
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.
14.2.6.1.2 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).
14.2.6.2 Abiotic Samples. 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 docu-
ment whether such depletions have occurred, replicate
sediment samples should be collected for physical and
chemical analysis from each sediment type at the begin-
ning and the end of the exposure. Archiving sediment
samples from every biological sampling period also is
recommended.
14.2.6.3 Short-Term Uptake Tests. 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.
14.2.6.4 Estimating Steady-State. In tests where
steady-state cannot be documented, it may be possible
to estimate steady-state concentrations. Several meth-
ods have been published that can be used to predict
78
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steady-state contaminant levels from uptake and depu-
ration 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
contaminants 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 at., 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 (l-e-"2"')
where Ct
Cs
Ks
K2
t
= contaminant concentration in tissue
at time t
bioconcentration studies in fish, see Davies and Dobbs
(1984), Spacie and Hamelink (1982), and ASTM (1988b).
For application of this procedure for sediment, see Lee
et al. (1994). Recent studies of the accumulation of
sediment-associated contaminants by benthos suggest
that the kinetics for freshly dosed sediments may require
a more complex formulation to estimate the uptake
clearance constant than that presented above (Landrum,
1989).
14.2.6.4.3 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
= contaminant concentration in sediment where S = time to apparent steady-state (days)
uptake rate coefficient in tissue (g sed
g-1 day1)
depuration constant (day1)
time (days)
As time approaches infinity, the maximum or equilibrium
contaminant concentration within the organism (Ctmax)
becomes
Ctmax =CsxKs/K2
Correspondingly, the bioaccumulatton factor (BAF) for a
compound may be estimated from
BAF= Ks/K2
14.2.6.4.1 This model assumes that the sediment con-
centration and the kinetic coefficients are invariant. Deple-
tion 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 physiol-
ogy such as reproduction will also invalidate the model.
Despite these potential limitation, the model can provide
estimates of steady-state tissue residues.
14.2.6.4.2 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
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
relationship between Kow and k2 for fish species (Spacie
and Hamelink, 1982):
K2 = antilog[1.47-0.414 x log(Kow)]
The relationship between S and k2 and between k2 and
Kow is summarized in Table 14.2. Estimated time (days)
to reach 95% of contaminant 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 depu-
rated (decimal fraction of tissue residue lost per day).
Table 14.2 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 available, then the table may be used
Table 14.2 Estimated Time to Obtain 95 Percent of Steady-State
Tissue Residue
S (days)
Log Kow
1
2
3
4
5
6
7
8
9
K2
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
79
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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 contaminants is un-
known. 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.
14.2.6.4.4 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 ex-
tremely low exchange rates with the environment (Bryan,
1976). Thus, the induction of metal binding proteins may
result in decreased depuration rate constants in organ-
isms exposed to the most polluted sediments. Addition-
ally, structure-activity relationships that exist for organic
contaminants (e.g., relationship between Kow and BCFs)
are not well developed for metals.
14.3 Data Interpretation
14.3.1 Sediments spiked with known concentrations of
contaminants can be used to establish cause and effect
relationships between chemicals and biological re-
sponses. Results of toxicity tests with test materials
spiked into sediments at different concentrations may be
reported in terms of an LC50 (median lethal concentra-
tion), an EC50 (median effect concentration), an IC50
(inhibition concentration), or as an NOEC (no observed
effect concentration) or LOEC (lowest observed effect
concentration; Section 3). Consistent spiking procedures
should be followed in order to make interlaboratory
comparisons (Section 8.3).
14.3.2 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
correlated to interstitial water concentrations, and effect
concentrations in interstitial water are often similar to
effect concentrations in water-only exposures. The bio-
availability of nomonic organic compounds are often
inversely correlated with the organic carbon concentra-
tion of the sediment. Whatever the route of exposure,
the correlations of effect concentrations to interstitial
water concentrations indicate predicted or measured
concentrations in interstitial water can be useful for
quantifying the exposure concentration to an organism.
Therefore, information on partitioning of chemicals be-
tween solid and liquid phases of sediment may be useful
for establishing effect concentrations.
14.3.3 Toxic units can be used to help interpret the
response of organisms to multiple contaminants in sedi-
ment. 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 mea-
sured concentration of a chemical in pore water by the
water-only LC5Q for the same chemical (Ankley et al.,
1991 a). Toxicity expressed as toxic units may be summed
and this may provide information on the toxicity of
chemical mixtures (Ankley et al., 1991 a).
14.3.4 Field surveys can be designed to provide either a
qualitative reconnaissance of the distribution of sedi-
ment contamination or a quantitative statistical compari-
son of contamination among sites (Burton and Ingersoll,
1994). Surveys of sediment toxicity are usually part of
more comprehensive analyses of biological, chemical,
geological, and hydrographic data. Statistical correlation
can be improved and costs reduced if subsamples are
taken simultaneously for sediment toxicity or bioaccu-
mulation tests, chemical analyses, and benthic commu-
nity structure.
14.3.5 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. Hazard evaluations integrating data from labora-
tory exposures, chemical analyses, and benthic commu-
nity assessments provide strong complementary evi-
dence of the degree of pollution-induced degradation in
aquatic communities {Chapman et al., 1992; Burton,
1991).
14.3.6 Toxicity Identification Evaluation (TIE) proce-
dures can be used to help provide insights as to specific
contaminants responsible for toxicity in sediment
(USEPA, 1991 a; Ankley and Thomas, 1992). For ex-
ample, the toxicity of contaminants such as metals,
ammonia, hydrogen sulfide, and nonionic organic com-
pounds can be identified using TIE procedures.
14.3.7 In terpretation of Comparisons of Tissue Resi-
dues. If the mean control tissue residues at Day 28 are
not significantly greater than 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 values are high, the data should be discarded
and the experiment conducted again after determining
the source of contamination.
80
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14.3.7.1 Comparisons of the 28-d control (or reference)
tissue residues and 28-d treatment tissue residues de-
termines whether there was statistically significant bio-
accumulation due to exposure to test sediments. Com-
parisons between control and reference tissue residues
at Day 28 determine whether there was a statistically
significant bioaccumulation due to exposure to the refer-
ence sediment. If no significant difference is detected
when treatment tissue residues are compared to a set
criterion value (e.g., FDA Action Limit) with a one-tailed
test, the residues must be considered equivalent to the
value even though numerically the mean treatment tis-
sue residue may be smaller.
14.3.7.2 BAFs and BSAFs. 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 necessary 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. See
Lee et al. (1994) provide examples of this approach.
14.3.7.3 Comparing Tissue Residues of Different
Compounds. In some cases, it is of interest to compare
the tissue residues of different compounds. For ex-
ample, Rubinstein et al. (1987) compared the uptake of
thirteen different PCB congeners to test for differences
in bioavailability. 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 t-tests and ANOVAs are inappropriate. A re-
peated 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 compounds 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.
14.4 Reporting
14.4.1 The record of the results of an acceptable
sediment test should include the following information
either directly or by referencing available documents:
14.4.1.1 Name of test and investigator(s), name and
location of laboratory, and dates of start and end of test.
14.4.1.2 Source of control or test sediment, method for
collection, handling, shipping, storage and disposal of
sediment.
14.4.1.3 Source of test material, lot number if appli-
cable, composition (identities and concentrations of ma-
jor ingredients and impurities if known), known chemical
and physical properties, and the identity and
concentration(s) of any solvent used.
14.4.1.4 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.
14.4.1.5 Source, history, and age of test organisms;
source, history, and age of brood stock, culture proce-
dures; and source and date of collection of the test
organisms, scientific name, name of person who identi-
fied the organisms and the taxonomic key used, age or
life stage, means and ranges of weight or length, ob-
served diseases or unusual appearance, treatments,
holding procedures.
14.4.1.6 Source and composition of food, concentra-
tions of test material and other contaminants, procedure
used to prepare food, feeding methods, frequency and
ration.
14.4.1.7 Description of the experimental design and test
chambers, the depth and volume of sediment and over-
lying water in the chambers, lighting, number of test
chambers and number of test organisms/treatment, date
and time test starts and ends, temperature measure-
ments, dissolved oxygen concentration (as percent satu-
ration) and any aeration used before starting a test and
during the conduct of a test.
14.4.1.8 Methods used for physical and chemical char-
acterization of sediment.
14.4.1.9 Definition(s) of the effects used to calculate
LC50 or EC50s, biological endpoints for tests, and a
summary of general observations of other effects.
14.4.1.10 A table of the biological data for each test
chamber for each treatment including the control(s) in
sufficient detail to allow independent statistical analysis.
14.4.1.11 Methods used for statistical analyses of data.
14.4.1.12 Summary of general observations on other
effects or symptoms.
14.4.1.13 Anything unusual about the test, any devia-
tion from these procedures, and any other relevant
information.
14.4.2 Published reports should contain enough infor-
mation to clearly identify the methodology used and the
quality of the results.
81
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Section 15
Precision and Accuracy
15.1 Determining Precision and
Accuracy
15.1.1 Precision is a term that describes the degree to
which data generated from replicate measurements dif-
fer and reflects the closeness of agreement between
randomly selected test results. Accuracy is the differ-
ence 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.
Quantitative determination of precision and accuracy in
sediment 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 15.2).
15.1.2 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 15.5.1) preci-
sion or multi-laboratory (interlaboratory or reproducibil-
ity; Section 15.5.2) precision. Intralaboratory precision
reflects the ability of trained laboratory personnel to
obtain consistent results repeatedly when performing
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 how reproduc-
ible a method is when conducted by a large number of
laboratories using the same method, organism, and
samples. Generally, intralaboratory results are less vari-
able than interlaboratory results (USEPA, 1991b; USEPA,
1993a; USEPA, 1994b; USEPA, 1994c; Hall et al.,
1989; Grothe and Kimerle, 1985).
15.1.3 A measure of precision can be calculated using
the mean and relative standard deviation (percent coef-
ficient of variation, or CV% = standard deviation/mean x
100) of Ihe calculated endpoints from the replicated
endpoints 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
effect levels derived from statistical analyses of hypoth-
esis testing. The CVs may be very high when testing
extremely toxic samples. For example, if there are mul-
tiple replicates with no survival and one with low sur-
vival, the CV may exceed 100%, yet the range of
response is actually quite consistent. Therefore, addi-
tional estimates of precision should be used, such as
range of responses, and minimum detectable differ-
ences (MOD) compared to control survival or growth.
Several factors can affect the precision of the test,
including test organism age, condition, sensitivity, han-
dling and feeding of the test organisms, overlying water
quality, and the experience of the investigators in con-
ducting tests. For these reasons, it is recommended that
trained laboratory personnel conduct the tests in accor-
dance with the procedures outlined in Section 9. Quality
assurance practices should include (1) single laboratory
precision determinations using reference toxicants for
each of the test organisms that are used to determine
the ability of the laboratory personnel to obtain precise
results. These determinations should be made before
conducting a sediment test and should be routinely
performed as long as whole sediment tests are being
conducted; (2) control charts (Section 15.3) should be
prepared for each reference toxicant and test organism
to determine if the test results are within prescribed
limits; and (3) tests must meet the minimum criteria of
test acceptability specific for each test organism (Tables
11.3, 12.3, 13.4; USEPA, 1991b).
15.1.4 Intralaboratory precision data are routinely cal-
culated for test organisms using water-only 96-h expo-
sures to a reference toxicant, such as KCI. Intralabora-
tory precision data should be tracked using a control
chart. Each laboratory's reference toxicant data will
reflect conditions unique to that facility, including dilution
water, culturing, and other variables (Section 9). How-
ever, each laboratory's reference loxicant CVs should
reflect good repeatability.
15.1.5 To date, interlaboratory precision (round-robin)
tests have been completed with both Hyalella azteca
and Chironomus tentans using 4-d water-only and 10-d
whole sediment tests. The results of these round-robin
studies are described below.
15.2 Accuracy
15.2.1 The relative accuracy of toxicity tests cannot be
determined since there is no acceptable reference ma-
terial. The relative accuracy of the reference toxicity
82
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tests can only be evaluated by comparing test responses
to control charts.
15.3 Replication and Test Sensitivity
15.3.1 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 given level of variability
remains constant, the sensitivity of the test will increase
as the number of replicates is increased. The minimum
recommended number of replicates varies with the ob-
jectives of the test and the statistical method used for
analysis of the data (Section 14).
15.4 Demonstrating Acceptable
Laboratory Performance
15.4.1 It is the responsibility of a laboratory to demon-
strate its ability to obtain precise results with reference
toxicants before it performs sediment tests (Section
9.16). Intralaboratory precision, expressed as a coeffi-
cient of variation (CV), of the range for each type of test
to be used in a laboratory should be determined by
performing five or more tests with different batches of
test organisms, using the same reference toxicant, at
the same concentrations, with the same test conditions
(e.g., the same test duration, type of water, age of test
organisms, feeding), and same data analysis methods.
A reference toxicant concentration series (0.5 or higher)
should be selected that will consistently provide partial
mortalities at two or more concentrations of the test
chemical (Section 9.14, Table 9.1, 9.2).
15.4.2 The quality of test organisms obtained from an
outside source must be verified by conducting a
reference-toxicity test concurrently with the sediment
test. The supplier should provide data with the shipment
describing the history of the sensitivity of organisms
from the same source culture. If the supplier has not
conducted five reference toxicity tests with the test
organism, it is the responsibility of the testing laboratory
to conduct five reference toxicity tests before starting a
sediment test (Section 9.14.1).
15.4.3 Before conducting tests with contaminated sedi-
ment, the laboratory should demonstrate its ability to
conduct tests by conducting five exposures in control
sediment as outlined in Table 11.1, 12.1, or 13.1. It is
recommended that these five exposures with control
sediment be conducted concurrently with the five refer-
ence toxicity tests described in Section 15.4.2.
15.4.4 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 15.1), and the end-
points (LC50, NOEC, ICp) are examined to determine if
o
LU
O
Upper Control Limit
Central Tendency
Lower Control Limit
1 1 1 1 1 1 1 II 1 1 1 1
1 II 1 1 fc
10
15
20
O
o
O
Upper Control Limit (X + 2 S)
Central Tendency
Lower Control Limit (X - 2 S)
I I I I I I I I I I I I I I I I 1 I I I
10
15
20
Toxicity Test with Reference Toxicants
where
Figure 15.1
n-\
Successive toxicity values of toxicity tests.
Number of tests.
Mean toxicity value.
Standard deviation.
Control (cusum) charts: (A) hypothesis testing
and (B) point estimates (LC, EC, or 1C).
they are within prescribed limits. Control charts as de-
scribed in USEPA (1993a) 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 succes-
sive test result. After two years of data collection, or a
minimum of 20 data points, the control (cusum) chart
should be maintained using only the 20 most recent data
points.
15.4.5 The outliers, which are values falling outside the
upper and lower control limits, and trends of increasing
or decreasing sensitivity, are readily identified using
83
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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.
15.4.5.1 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 automatically 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%. All the performance
criteria outlined in Tables 11.3, 12.3, and 13.4 must be
considered when determining the acceptability of a sedi-
ment test. The acceptability of the sediment test would
depend on the experience and judgment of the investi-
gator and the regulatory authority.
15.4.6 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 are
suspect (LJSEPA, 1993a). In this case, the test proce-
dure should be examined for defects and should be
repeated with a different batch of test organisms.
15.4.7 Performance should improve with experience,
and the control limits tor point estimates should gradu-
ally narrow. However, control limits of ±2 SD, by defini-
tion, 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
penalized if a test that falls just outside the control limits
is rejected de facto. For this 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.
15.4.8 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 con-
centration 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, or Probit Method,
Graphical Method, or the Linear Interpolation Method
(Section 14).
15.4.9 The point estimation analysis methods recom-
mended in this manual have been chosen primarily
because 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.
15.5 Precision of Sediment Toxicity
Test Methods
75.5.1 Intralaboratory Precision
15.5.1.1 Intralaboratory precision of the Hyalella azteca
and Chironomus tentans 10-d tests (as described in
Tables 11.1 and 12.1) was evaluated at ERL-Duluth
using 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_ rather than 1.5 mL for C. tentans. The results of
the study are presented in Table 15.1. The mean sur-
vival 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).
15.5.2 Interlaboratory Precision
15.5.2.1 Interlaboratory precision using reference toxic-
ity tests and 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 labo-
ratories, contract laboratories, and academic laborato-
ries that had demonstrated experience in sediment tox-
icity testing (Table 15.2). The only exception to the
methods outlined in Table 9.1 and 9.2 was that 80%
rather than the current recommendation of 90% survival
was used to judge the acceptability of the reference
toxicity tests. The round robin study was conducted in
two phases for each test organism. The experimental
design for the round robin study required each labora-
tory 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. Criteria for selection of participants in the
Table 15.1 Intralaboratory Precision for Survival ol Hyalella
azteca and Chironomus tentans in 10-d Whole-
Sediment Toxicity Tests, June 1993*
Percent Survival
Individual
A
B
C
D
E
N
Mean
CV
H. azteca
85
93
90
84
100
5
90.4
7.2%
C. tentans
85
93
93
94
100
5
93.0
5.7%
Test sample was from a control sediment (T.J. Norberg-King,
USEPA, Duluth, MM, personal communication.) The test was
conducted at the same time by five individuals at ERL-Duluth. The
source of overlying water was from Lake Superior.
84
-------
round-robin study were that the laboratories had (1)
existing cultures of the test organisms, (2) experience
conducting tests with the organisms, and (3) would
participate voluntarily. The test methods for the refer-
ence 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.
15.5.2.2 In Phase 1, water-only reference toxicity {KCI)
tests were conducted with H. azteca for 96-h and LC50s
were calculated. In these tests, H. azteca were placed in
reconstituted hard water in 250-mL beakers containing a
small piece of plastic mesh substrate. Ten organisms
were randomly added to each of four replicates at five
concentrations of KCI and a control. The organisms
were fed 0.5 mL of a 1800 mg/L stock solution of YCT on
Day 0 and Day 2. Mortality was monitored at 24 h
intervals and the test was ended at 96 h (Table 9.2). In
Phase 2, the variability of the 10-d whole sediment test
procedure for H. azteca was evaluated using an auto-
mated water renewal exposure system (Table 11.1 and
Section B.3). This system consisted of eight replicate
300-mL beakers per treatment with each containing 10
organisms each. Each beaker contained a 100-mL ali-
quot of sediment and the overlying water was replaced
twice a day (Table 11.1). The test sediments that were
previously tested at ERL-Duluth to ascertain their toxic-
ity included a control sediment (RR 3), a moderately
Table 15.2 Participants in Round Robin Studies'
Chironomus tentans
Hyalella azteca
Laboratory
Lab A
LabB
LabC
LabD
LabE
LabF
LabG
LabH
Labi
LabJ
LabK
LabL
N
96 h
KCI
Test
Dec 92
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
3
4
9
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
a
96 h
KCI
Test
Oct92
Y
Y
Y
N
Y
Y
Y
Y
-*
Y
Y
Y
10
10-d
Sediment
Test
Mar 93
N
Y
Y
N
Y
Y
Y
N
Y
Y
Y
Y
9
contaminated sediment (RR 2), and a heavily contami-
nated test sediment (RR 1). Sediments RR 2 and RR 3
were contaminated primarily with copper. An additional
sediment heavily contaminated with polycyclic aromatic
hydrocarbons (RR 4) was tested by five laboratories. At
the end of a test, the sediment from each replicate was
sieved and surviving organisms were counted.
15.5.2.3 Ten laboratories participated in the H. azteca
reference toxicity test (Table 15.2). The results from the
tests with KCI are summarized in Table 15.3. The test
performance criteria of >80% control survival 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 LCSOs ranged from 232 to
372 mg/L KCI.
15.5.2.4 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 (Table 15.4).
All laboratories completed the tests; however, laboratory
C had 75% survival, which was below the acceptable
test criteria for survival (Table 11.3). For these tests, the
CV was calculated using the mean percent survival for
Table 15.3 Intel-laboratory Precision (or Hyalella azteca 96-h
LCSOs from Water-Only Static Acute Toxicity Tests
Using a Reference Toxicant (KCI) (October 1992)
Laboratory
Lab A
LabB
LabC
LabD
LabE
LabF
LabG
LabH
Labi
LabJ
LabL
N
Mean
CV
N
Mean
CV
KCI
LC50
(mg/L)
372
321
232
— '
325
276
297
336
142J
337
250
10
289.03
23.0%3
9
305.04
14.2%'
Confidence Intervals
Lower
352
294
205
1
282
240
267
317
101
286
222
Upper
395
350
262
1
374
316
331
356
200
398
282
Percent
Control
Survival
100
98
100
1
100
98
73
100
93
100
100
10
96.2%
8.3%
9
98.8
2.1%
' Y = Laboratory participated in testing sediment samples.
2 Test in January 1993.
3 Participated using C. ripar/us only.
' Did not intend to participate with C. tentans.
1 Laboratory did not participate in H. azteca test in October.
2Results are from a retest in January using three concentrations only;
results excluded from analysis.
3Mean 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 >80%.
85
-------
the eight laboratories that met the performance criteria
for the test. The CV for the control sediment (RR 3) was
5.8% with a mean survival of 94.5% with survival rang-
ing from 86% to 100%. For sediments RR 2 and RR 4,
the mean survival was 3.3% and 4.3%, respectively
(Table 15.4). 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 con-
taminated 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 between the test sediments and the control
sediment ranged from 5 to 24% with a mean of 11% (SD
= 6).
15.5.2.5 The Phase 1 C. tentans reference toxicity test
was conducted with KCI on two occasions (Tables 15.5
and 15.6). Both tests were conducted in 20 ml of test
solution in 30-mL beakers using 10 replicates per treat-
ment with 1 organism per beaker. Animals were fed 0.25
mL of a 4 g/L solution of Tetrafin® on Day 0 and Day 2
(Table 9.1). For the first reference toxicity test compari-
son, 10 laboratories participated, and eight laboratories
met the survival criteria of the round robin, which was
80% survival (Table 15.5). 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 correlated (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 15.5) as determined by head cap-
sule width.
15.5.2.6 For the second Phase 1 KCI reference toxicity
tests with C. tentans, seven laboratories participated
(Table 15.6). 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.
15.5.2.7 In the 10-d whole sediment test with C. tentans
eight laboratories participated. The same three sedi-
ments used in the H. azteca whole sediment test were
used for this test (Table 15.7). All test conditions were
those as described in Table 12.1 with the exception of
the feeding rate of 1.0 mL rather than 1.5 mL for C.
tentans. Three laboratories did not meet the control
criteria for acceptable tests of >70% survival in the
control (RR 3) sediment; Table 12.3). For the five labo-
ratories 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%. For
Table 15.4 Interiaboratory Precision for Survival of Hyalella azteca in 10-d Whole Sediment Toxicity Tests Using Four Sediments
(March 1993)
Mean Percent Survival (SD) in Sediment Samples
Laboratory
Lab A
LabB
LabC
LabD
LabE
LabF
LabG
LabH
Labi
LabJ
LabK
LabL
N
Mean 1 3
CV 1
N
Mean 2"
CV2
RR 1
76.2
57. 522
i
46.2
72.5
50.0
— '
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)
RR2
2.5
1.22
1
0
23.7
0
1
0
0
0
0
9
3.0
256%
8
3.3
253%
1
(7.1)
(0)
(0)
(18.5)
(0)
(0)
(0)
(0)
(0)
RR3
1
97.5
75.0*
1
97.5
98.7
100
— '
86.2
96.2
95.0
86.2
9
93.0
9.0%
8
94.5
5.8%
(Control)
(4.6)
(17.7)
(7.1)
(3.5)
(0)
(10.6)
(5.2)
(5.3)
(18.5)
RR4
— '
11.2
1.22
1
—
0
3.3
— '
—
2.5
—
—
5
3.6
121%
4
4.3
114%
(13.6)
(0)
(0)
(5.2)
(7.1)
' Laboratory did not participate in H. azteca test in March.
;' 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 >80%.
86
-------
Table 15.5 Intel-laboratory Precision for CMronomus tentans 96-h LCSOs from Water-only Static Acute Toxicity Tests Using a
Reference Toxicant (KC1) (December 1992)
Labora-
tory
Lab A
Lab B
LabC
LabD
LabE
LabF
LabG
LabH
Labi
Lab J
N
Mean I5
CV 1
N
Mean 26
CV2
KCI
LC50
(9/L)
6.19
6.83
5.00
3.17
2.00s
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
{%)
75'
100
100
100
80
80
95
95
100
65'
10
89.0
14.5%
8
93.8
9.3%
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%
Instar
at
Start
of Test
3,4
3
3
3
3
1
3
3
4
2,3
Age at
Start
of Test
(day)
10
13
11
11
11
12
11
7
10
7
10
10.3
17.9%
8
10.75
15.2%
Control survival below minimum acceptable level.
Unable to calculate LC50 with trimmed Spearman Karber; no confidence interval could be calculated.
Confidence intervals cannot be calculated as no partial mortalities occurred.
No animals were measured.
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 >80%.
Table 15.6 Interlaboratory Precision for Chironomus tentans 96-h LCSOs from Water-only Static Acute Toxicity Tests Using a
Reference Toxicant (KCI) (May 1993)
Labora-
tory
Lab A
LabB
LabC
LabD
LabE
LabF
LabG
LabH
Labi
LabJ
n
Mean 1"
CV 1
n
Mean 2 5
CV2
KCI
LC50
(9/L)
1
6.65
1
5.30
5.11
3.61
5.36
— '
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
Control
Survival
(%)
—
90
—
553
100
90
93
—
95
100
7
89
17.5%
6
94.7
4.8%
Age at
Start
of Test
(day)
—
12
—
10
11
10
12
10-11
13
7
11.1
9.46%
6
11.2
9.13%
Did not participate in reference toxicity test in April.
Confidence intervals cannot be calculated as no partial mortalities occurred.
Control survival below minimum acceptable level.
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%.
87
-------
the 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 15.8). When the RR 1 data for each laboratory
were compared to the control for that laboratory, the
minimum detectable difference for survival among labo-
ratories ranged from 2.3 to 12.1 with a mean of 8% (SD
= 4).
15.5.2.8 For C. tentans, growth is a sensitive indicator
of sediment toxicity (Ankley et al., 1993) and growth was
also measured in the round-robin comparison (Table
15.8). Using the data from five laboratories with accept-
able control survival in the control sediment (RR 3), the
mean weight of C. tentans for the control sediment (RR
3) was 1.254 mg (CV = 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 measure-
ments were obtained for C. tentans in sediment RR 2
because of the high mortality. The mean minimum de-
tectable difference for growth among laboratories meet-
ing 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.
15.5.2.9 These tests exhibited similar or better preci-
sion than many chemical analyses and effluent toxicity
test methods (USEPA, 1991b; USEPA, 1993a). The
success rate for test initiation and completion of the
EPA's round-robin evaluations is a good indication that
a well equipped and trained staff will be able to success-
fully conduct these tests. This is an important consider-
ation for any test performed routinely in any regulatory
program.
Table 15.7 Interlaboratory Precision for Survival of Chtronomus tentans in 10-d Whole-Sediment Toxicity Tests Using Three
Sediments (May 1993)
Laboratory
RR1
Mean Percent Survival (SD) in Sediment Samples
RR2
RR 3 (Control)
Lab A
LabB
LabC
LabD
LabE
LabF
LabG
LabH
Labi
LabJ
N
Mean 13
CV 1
N
Mean 2*
CV2
1
67.5
15.02
60.0*
85.0
87.52
90.0
— '
97.5
93.8
8
74.5
36.7%
5
86.8
135%
(14.9)
(12.0)
(20.0)
(11.9)
(12.5)
(13.1)
(4.6)
(11.8)
1
2.5
O2
O2
0
O2
12.5
1
0
0
8
1.88
233%
5
3.0
181%
(7.1)
(0)
(0)
(0)
(0)
(3.5)
(0)
(0)
1
98.8
62.52
66.3J
93.8
43.82
87.5
— '
98.8
81.2
8
79.1
25.1%
5
92.0
8.3%
(3.5)
(26.0)
(27.7)
(9.2)
(30.2)
(10.3)
(3.5)
(8.3)
' Old not participate in C. tentans test 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%.
88
-------
Table 15.8 Interiaboratory Precision for Growth of Chlronomus tentans in 10-d Whole-Sediment Toxicity Tests Using Three
Sediments (May 1993)
Laboratory
Growth—Dry Weight in mg (SO) in Sediment Samples
RR 1
RR2
RR 3 (Control)
Lab A
LabB
LabC
LabD
LabE
LabF
LabG
LabH
Labi
Lab J
n
Mean 1 3
CV 1
n
Mean 24
CV2
i
0.370
0.8832
0.21 52
0.657
0.2 102
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
0
0
0
0
0
. ,1
0
0
8
—
—
5
—
—
(0)
(0)
(0)
(0)
(0)
(0)
(0)
(0)
1
1.300
0.504
1.070
0.778
0.610
1.710
i
1.300
1.180
8
1.056
38.3%
5
1.254
26.6%
(0.060)
(0.212)
(0.107)
(0.169)
(0.390)
(0.250)
(0.006)
(0.123)
' 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.
4 Mean 2 and CV 2 exclude data points for all sediment samples from laboratories that did not meet minimum control survival of >70%.
89
-------
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Appendix A
Summary USEPA Workshop on
Development of Standard Sediment Test Methods
A.1 The USEPA Office of Water, Office of Science and
Technology, and Office of Research and Development
held a workshop September 16-18, 1992, in Washing-
ton, DC, to provide an opportunity for experts in the field
of sediment toxicology and staff from USEPA regional
and Headquarters program offices to discuss the devel-
opment of standard freshwater and marine sediment
testing procedures (USEPA, 1992a). As part of USEPA's
Contaminated Sediment Strategy, the Agency's pro-
gram offices agreed to develop and use consistent tests
for the assessment of sediment contamination. USEPA
sponsored research to address uncertainties associated
with the use of sediment tests discussed at the work-
shop. The results of discussions held at the workshop
were used to identify research issues and develop manu-
als for conducting sediment toxicity and bioaccumula-
tion tests. The following test organisms were selected
for sediment test method development in 1993: (1)
Freshwater toxicity tests: Hyalella azteca and Chironomus
tentans, (2) Freshwater bioaccumulation tests:
Lumbriculus variegatus, (3) Marine toxicity tests:
Ampelisca abdita, Rhepoxynius abronius, Eohaustorius
estuarius, and Leptocheirus plumulosus, (4) Marine bio-
accumulation tests: Macoma nasuta and Nereis spp.
A.2 If funds are available in future years, additional work
will be started on developing chronic toxicity tests, toxic-
ity identification evaluation (TIE), and test development
for other organisms. USEPA plans to develop two other
methods documents: (1) one on sediment spiking and
(2) one on sediment collection, handling, and storage.
Parts of the document on collection, handling, and stor-
age methods are already under development for a Qual-
ity Assurance and Quality Control guidance document
that will supplement both the Inland and Ocean Testing
Manuals for disposal of dredged material
(USEPA-USCOE, 1991; 1994).
A.3 Before the workshop on July 13, 1992, a question-
naire was sent out to freshwater workshop invitees and
other selected researchers. Information was requested
on culture methods for and testing of H. azteca, C.
tentans, and L variegatus and any additional organisms
used for sediment tests. The discussion topics for cultur-
ing and testing of the three species were ranked in order
of importance for developing standard methods based
on the similarity of the issues across all tests. The
following section summarize results of the questionnaire
for each test organism.
1. Development of a Standard Testing Method for
Hyalella azteca. Twenty-one responses to the sur-
vey were received and eighteen laboratories re-
ported information on H. azteca (USEPA, 1992a
and Table A.1). The summary of the survey re-
sponses follow. The most common response is un-
derlined and when no item is underlined it indicates
no single most common response. Published proce-
dures for conducting sediment toxicity tests with H.
azteca are also listed in Table A.2.
Table A.1 List of Laboratories Responding to the Survey
Laboratory
Hyalel/a
azteca
Chironomus Lumbriculus
tentans variegatus
Dept. of Fish. & Oceans, x
Canada
Environ. Canada, x
Burlington, ON
EPA-Duluth, Duluth, MN x
EPA Region 1, Lexington, x
MA
EPA Region 8, Denver, CO x
EPA-Newtown, Cincinnati, x
OH
EVS Consultants, x
Vancouver, BC
MD Dept. Environ., x
Baltimore, MD
Miami Univ., Oxford, OH x
Mich. State Univ., x
E. Lansing, Ml
NBS-Athens, Athens, GA x
NBS-Columbia, x
Columbia, MO
NOAA-Ann Arbor, x
Ann Arbor, Ml
Old Dominion, Norfolk. VA x
State of WA, x
Manchester, WA
Univ. of MS, University, x
MS
Univ. of Wl-Supenor, x
Superior, Wl
Wright State Univ., x
Dayton, OH
101
-------
Table A.2 Summary of Testing Procedures Used to Evaluate the Toxicity of Whole Sediments with Hyaletla azteca
Condition [1]
Temperature (CC) 20
Light 'intensity NR
•,'oot candies)
Priotopenod NR
Test c^arrber (rrL) 1000
Sediment volume (mL) 200
Overlying water 800
volume (mL)
Renewal rate of) 0
Overlying water
(additions/day)
Age of organisms juvenile
(days)
Size of organisms (mm) NR
Number of organisms/ 15
charrber
Number of replicate NR
chambers/treatment
Food RC
Aeration Yes
Overlying water Natural
Test duration (d) 10
Er.dpoints S
Test acceptability NR
(survival %)
Citations
[1] Nebeker etal. (1984a)
(2) Ingersoll and Nelson (1990)
[3] Ankley eta!. (1993a)
|4| Burton etal. (1989)
|5] Winger and Lasier (1993)
[6] Borgmann and Munawar (1989)
[2]
20
25-50
16-8
1000
200
100
1-4
juvenile
1-2
20
4
RC
None
Natural
10-28
S.G.rvl
80
[3]
22
NR
16-8
300
100
175
1-4
7-14
NR
10
3
YCT
None
Natural
10
S
80
Conditions
Food:
Endpoints:
Citation
[4]
20-25
50-100
16-8
300
40-50
160-200
variable
juvenile
NR
10
4
RC
DO<3
Natural
7
S
80
[5]
20-23
25-50
16-8
30-300
5-100
20-150
0-2
7-14
1-2
3-10
5-10
YCT, mL
None
Reconst.
10
S
80
YCT = yeast-cerophyll-trout enow, RC = Rabbit
TM = Tetramin®, ML =
maple leaves.
S = survival, G = growth (length or weight), M =
NR = Not reported
f6]
20-22
NR
16-8
2500
-150
-1350
0
0-7
NR
20
2
TM
Yes
Natural
28
S,G
NR
chow,
maturation,
102
-------
A. Survey Summary of Culture Methods for H. azteca
Flow: Static vs. renewal
Temperature: 19 to 25°C (23!O
Light: 16L8D photoperiod; about 500 to 1000 lux
Chamber: 1 L to 80 L
Age of organisms: Known age vs. mixed age
Frequency restart: Monthly, every two months
Water Quality: Natural vs. reconstituted
Source of Strains: ERL-Duluth, ERL-Corvallis. Burlington, Michigan State (most cultured in moderately-hard
or hard water)
Moderate
Leaves. Tetramin®. rabbit chow, diatoms, yeast, wheat grass, Chlorella, alfalfa, Nutrafin®,
YCT. paper towels, Selenastrum, Ankistrodesmus, brine shrimp, aquatic plants, sedi-
ment. Feed 2 to 3 times/week typical.
Aeration:
Feeding:
Substrate:
Leaves, nylon mesh, cotton gauze, 3-M web plastic, paper towels
Reference toxicants: Cd. Cu, KCI, Zn, NaCI, Cr (water-only exposures)
B. Survey Summary of Testing Procedures for H. azteca
Flow:
Aeration:
Temperature:
Light:
Chamber:
Sediment ratio:
Static vs. renewal
None or moderate
20 to 25°C (2Q!CJ
16L8D photoperiod; about 250 to 1000 lux
30 ml to 1 L (250 to 300 mL)
1:1 to 1:4 ratio sediment:water
Age of organisms: Known age (0 to 7 d, 7 to 14 d) vs. mixed age (size about 7 to 14 d) (sieved)
Number of organisms:5 to 20/chamber (10/chamber)
Number replicates: 2 to 10/treatment (3 to 5/treatment)
Duration: 2- to 28-d HO-d)
Feeding: None, Rabbit Chow, YCT, maple leaves, Tetramin®
Endpoints: Survival, length, weight, sexual maturation (males), young production, bioaccumulation
Acceptability: Survival (60%). length, weight
103
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2. Development of a Standard Testing Method for Chironomus
tentans. Twenty-one responses to the survey were received
and twelve laboratories reported information on C. tentans
(USEPA, I992a and Table A.1). The summary of the survey
responses follow. The most common response is underlined
and when no item is underlined it indicates no single most
common response. Published procedures for conducting sedi-
ment toxicity tests with C. tentans are also listed in Table A.3.
Table A.3 Summary of Testing Procedures Used to Evaluate the Toxicity of Whole Sediments with Chironomus tentans
Condition [1]
Temperature (°C) 20
Light intensity NR
(foot candles)
Photopenod NR
Test chamber (ml) 1000
Sediment volume (mL) 200
Overlying water 800
volume (rnL)
Renewal rate of 0
overlying water
(additions/day)
Age of organisms 2nd
(instar)
Size of organisms NR
Number of organisms/ 15
chamber
Number of replicate NR
chambers/treatment
Food TM.CP
Aeration Yes
Overlying water Natural
Test duration (d) 10
Endpomts S,G
Test acceptability NR
(survival %)
Citations;
|1] Nebekeretal. (I984a)
[2] Adams etal. (1985)
[3] Ankley etal. (1993a)
(4] Giesy etal. (1988)
J5] Wentsel etal. (1977)
[2]
22
-100
16-8
3000
-250
2000
0-5
2nd
0.15mg
25
2
TM
None
Natural
14
S,G
NR
Conditions:
Food:
Endpoints:
Citation
[3]
22
NR
16-8
300
100
175
1-4
2nd
NR
10
NR
TF
None
Natural
10
S,G
70
(4]
23
NR
NR
50
-7.5
47
0
2nd
0.5 g
1
15
TF
Yes
Natural
10
G
NR
[5]
22
NR
NR
2000
1500
-200
0
2nd
6-8 mm
20
NR
None
Yes
Natural
17
S,G
NR
CP - cerophyll, RC = Rabbit chow, TM = Tetramin®,
TF = Tetrafin®
S * survival, G = growth
NR - not reported
(length or weight),
M = maturation,
104
-------
A. Survey Summary of Culture Methods for C. tentans
Flow: Static vs. renewal
Temperature: 19 to 25°C f23°C)
Light: 16L8D photoperiod; about 500 to 1300 lux
Chamber: 1 L to 80 L
Age of organisms: Known age vs. mixed age
Frequency restart: 2x/week to every 6 months
Age restart: Egg masses to <24-h old larvae
Water Quality: Natural vs. reconstituted
Aeration: Moderate
Feeding: Tetramin®. Nutrafin®, YCT and algae, alfalfa and Tetrafin®, feed daily lo 3x/week
Substrate: Paper towels {bleached or unbleached): sand
Reference toxicants: Cu, NaCI, Cd, KCI (water-only exposures)
B. Survey Summary of Testing Procedures for C. tentans
Flow:
Aeration:
Temperature:
Light:
Chamber:
Sediment ratio:
Static vs. renewal
None or moderate
20 to 25°C (23°C)
16L:8D photoperiod; about 250 to 1300 lux
50 mL to 2 L
1:1 to 1:4 ratio sediment:water
Age of organisms: Known age (0 to 16 d; 10 to 14 d)
Number of organisms: 10 to 80/chamber MO to 15/chamber)
Number replicates: 2-15 (3 to 41
Duration: 2 to 14-d MO-d)
Feeding: Trout chow, Tetrafin®, YCT
Endpoints: Survival, weight
Acceptability: Survival (70%). weight (dry weight)
105
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3. Development of a Standard Testing Procedure for Lumbriculus
variegatus. Twenty-one responses to the survey were re-
ceived and five laboratories reported information on
L variegatus (USEPA, 1992a and Table A.1). The summary
of the survey responses follow. The most common response
is underlined and when no item is underlined it indicates no
single most common response. Published procedures for
conducting sediment bioaccumulation tests with L. variegatus
are also listed in Table A.4.
Table A.4 Summary of Testing Procedures Used to Conduct Whole Sediment Bioaccumulation Tests with Lumbriculus variegatus
Condition
[1]
Citation
I2J
' ' :50 g dry weight organisnv.sediment organic carbon
NR = not reported
Citations.
[1J Phippsetai (1993)
|2] Kukkonen and Landrum (1993)
[3] E L. Brunson, NBS, Columbia, MO, unpublished data
(4| Schuytema et al. (1988)
[3]
[4]
Temperature (°C)
Light intensity
(foot candles)
Photopenod
Test chamber (L)
Sediment volume (L)
Overlying water
volume (L)
Renewal rate of
overlying water
(additions/day)
Age of organisms
Loading (g/chamber)
Number of replicate
chambers/treatment
Food
Aeration
Overlying water
Test duration (d)
Test acceptability
20
NR
NR
3-5
1.5-2
1.5-3
2-6
Adult
1
NR
None
None
Natural/
Reconst.
10-60
NR
23
NR
Various
0.15-0.6
30-l80g
0.1-0.45
0.5-1
Adult
1:50*
3-4
None
Yes
Natural
10-60
Biomass
lipid
23
25-50
16-8
4
i
3
1
Adult
1
3-5
None
Yes
Natural
56
Biomass
20
NR
NR
3-3.8
0.3-0.35
2.7-3
0
Adult
0.1-0.39/L
3
Yes
Yes
Natural
28-44
NR
106
-------
A. Survey Summary of Culture Methods for L. variegatus
Flow: Static vs. renewal
Temperature: 22 to 24°C
Light: 16L8D photoperiod; illuminance unspecified
Chamber: 1 L to 80 L
Age of organisms: Mixed-age adults
Frequency restart: Monthly, every two months
Natural vs. reconstituted
Moderate
Water Quality:
Aeration:
Feeding:
Frozen silver cup trout chow, salmon starter, sediment, Tetramin®, yeast, wheat grass.
Chlorella, alfalfa, Nutrafin®, YCT, paper towels. Feed 2 to 3 times/week typical.
Substrate: Paper towels, sediment
Reference toxicants: No reference toxicants specified
B. Survey Summary of Testing Procedures for L. variegatus
Flow:
Aeration:
Temperature:
Light:
Chamber:
Sediment Ratio:
Static vs. renewal
None or moderate
10to23°C
16L:8D photoperiod; illuminance unspecified
1 to 6 L
1:1 to 1:4 ratio sediment: water (sediment volumes should be adequate to allow feeding
and burrowing)
Age of organisms: Adults, 3.8 cm.
Number of organisms:Adequate number to provide tissue mass for analysis of residue of concern
Number replicates: 4 to 5/treatment
Duration: 10to28d
Endpoints: Bioaccumulation
Feeding: None
Acceptability: Adequate tissue mass for residue analysis
107
-------
A.4 Workgroup participants arrived at a consensus on
several culturing and testing procedures. Where it was
not possible to make a decision because of lack of
information, the group identified research items that
need further consideration before a specific decisions
could be made.
A.5 In developing guidance for culturing test organisms
to be included in the methods manual for sediment tests,
it was generally agreed that no one method had to be
used to culture organisms. The success of the tests
should rely on the health of the culture from which the
organisms were taken for testing. That is, having healthy
organisms of known quality and age for testing was
deemed to be the key consideration relative to culture
procedures. Therefore, performance-based criteria were
selected as the preferred approach laboratories should
use to evaluate cultures rather than using control-based
criteria. Performance criteria were chosen to allow each
laboratory to optimize culture techniques and meet qual-
ity control monitoring requirements.
A.6 The selection of test organisms for standardization
were based on (1) current and historical acceptance, (2)
logistical considerations, and (3) availability of testing
methods. Major differences between freshwater and
marine tests were discussed: (1) freshwater organisms
are cultured in the laboratory and marine organisms are
collected from the field, (2) freshwater organisms are
smaller and younger, (3) freshwater organisms are gen-
erally epibenthic and marine test organisms are gener-
ally infaunal, (4) freshwater test conditions are sensitive
to organic carbon, sediment oxygen demand, ammonia,
and the buffering capacity of the overlying water. The
overlying water is more stable in saltwater tests com-
pared to freshwater tests.
A.7 Performance-based culturing and testing criteria
were outlined for freshwater species at the workshop
(Table 11.3. 12.3, 13.4). Consensus was reached on
criteria that must be met and criteria that should be
considered. Factors that must be met include reference
toxicants with short-term water only exposures, survival
of control organisms, age or size of organisms at the
beginning or end of tests, and consistent water quality.
Factors that should be considered include parental sur-
vival, food quality, frequency of restarting cultures, and
time to emergence (midges). In addition, a
performance-based criterion for culture of L variegatus
to monitor is population doubling time (number of organ-
isms).
A.8 The following topics were discussed related to the
use of Hyatelta azteca and Chironomus spp. in sediment
toxicity tests: (1) Age of organisms used to start a test:
Hyalella azteca: Organisms of age 0- to 14-d old are
typically used to start a test. It may be best to test
organisms 7- to 14-d old, since organisms 0- to 7-d old
are difficult to recover from sediment. Chironomus
tentans: 9- to 11-d old organisms should be tested.
Chironomus riparius: 6- to 8-d old organisms should be
tested; (2) Length of test: The length of the test agreed
upon was 10 d with survival as the endpoint. Additional
research is ongoing to evaluate growth as an endpoint;
(3) Feeding: A minimal amount of food is required to
consistently achieve adequate control survival and
growth. Additional research is ongoing to evaluate the
influence of feeding on sediment toxicity; (4) Water
renewal: Limited renewal of overlying water was recom-
mended (about 1 to 2 volume/d): (5) Sediment volumes:
Sediment volume up to in the 200 mL have been rou-
tinely tested, but smaller volumes would be acceptable;
(6) Grain size: There does not seem to be a substantial
effect of sediment grain size in the 10-d exposures with
H. azteca. Additional research is ongoing to evaluate
the influence of grain size on the response of amphipods
and midges; (7) Strains of organisms: Different strains of
H. azteca have been used for testing. Reference toxi-
cant comparisons of the strains are needed.
A.9 The following topics were discussed related to the
use of L. variegatus sediment bioaccumulation tests: (1)
Age of test organisms: Adults should be tested; (2)
Length of test: 28 d. Additional research is ongoing to
evaluate duration of the exposure; (3) Feeding: No
feeding is required during the test; (4) Water renewal:
Limited renewal of overlying water was recommended
(about 1 to 2 volume/d); (5) Sediment volumes: Sedi-
ment volume up to in the 200 mL have been routinely
tested, but smaller volumes would be acceptable; (6)
Grain size: There does not seem to be a substantial
effect of sediment grain size in 10-d exposures with
L. variegatus; (7) Additional discussion topics: Standard
lipid content should be addressed in the document,
sediment avoidance may be important, and rigorous
techniques have been developed to purge the gut in
clean water. Research is needed to see if purging is
necessary.
108
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Appendix B
Exposure Systems
B.1 Renewal of overlying water is recommended during
sediment tests (Section 11.3,12.3,13.3). The overlying
water can be replaced manually (e.g., siphoning) or
automatically. Automated systems require more equip-
ment 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
suspension of sediment compared to manual renewal of
water.
B.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; Benoit et al., 1993; Zumwalt et al., 1994). The
water-delivery system should be calibrated before a test
is started to verify that the system is functioning prop-
erly. 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 B.3 and by Zumwalt et al. (1994) in
Section B.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).
B.3 Benoit et al. (1993) describe a sediment testing
intermittent-renewal (STIR) system (stationary or por-
table) for invertebrate toxicity testing with sediment.
Either stationary or portable systems enable the mainte-
nance of acceptable water quality (e.g., dissolved oxy-
gen) by automatically renewing overlying water in sedi-
ment tests at rates ranging from 1 to 21 volume renew-
als/d. The STIR system not only reduces the labor
associated with renewal of overlying water but also
affords a gentle exchange of water that results in virtu-
ally no sediment suspension. Both gravity-operated sys-
tems 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.
B.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 toxic-
ity tests. A STIR system (stationary or portable) solely
for sediment toxicity tests was designed, which offers a
simple, inexpensive approach for the automated re-
newal of variable amounts of overlying water (Figures
B.1 and B.2). This system is described below. The
system can be built as a two-unit system (Section B.3.2)
or with more exposure treatments (Section B.3.4). All
exposure systems consist of exposure holding tanks.
head tanks, head tank support stands, and a water bath
(Section B.3.2 and B.3.3). The automated delivery sys-
tem includes design descriptions for a support stand,
water renewal supply, and water delivery apparatus
(Section B.3.4).
B.3.2 Two Unit Portable STIR System Construction
(Figure B.1 and B.2)
B.3.2.1 Exposure Holding Tanks (2) (Figure B.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
bottom.
B.3.2.2 Head Tanks (2) (4-L capacity; Figure B.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: 13.9cm 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
109
-------
Figure 5 - Photo of portable mini-flow test system.
Figure B.1 Portable table top STIR system described in Benoit et al. (1993).
110
-------
Calibrated Volume Sight Tube
(1.3cm Clear Tube)
Head Box
(30.5 x 30.5 x 38cm High
Adjustable Float on
Threaded Rod
Toilet Tank Valve
•— Water Inlet
Timer Controlled Solenoid Valves
Optional
Automated
Water
Delivery
Apparatus
CD
;u
CO
Water Distribution
Manifold with Open Ends
(1.3cm plastic pipe)
' 6mm Pipe to Hose Adaptor
u ._ .Calibrated
Head Tank FlowTube
4L I
Water Bath
T
Circulator
Pump
• Water Bath Flow -
Holding
Tank
ft —
Self
Starting
Siphon
Outlet
1
ft
fl
Optional 1.2
or 3 Unit
"Add on"
Water Bath
_>
.o
/
Thermostat
All tanks and water bath drain to common 19L jug with air
vent and optional hose from jug to floor drain.
Figure B.2 Portable table top STIR system with several additional options as described in Benoit et al. (1993).
111
-------
Width (end)
Exposure Holding Tank
Width (end)
Head Tank
2.5cm
Qr1 Water pump inlet
Water pump inlet
2.5cm
..-' 3.2cm
Thermostat
Overflow
1.6cm
Length (side)
Basic Water Bath
Length (side)
Basic Water Bath with Optional Holes for Water Bath
Width (end)
Add on Water Bath for One Additional Unit
Figure B.3 Tanks for the STIR system in Benolt et al. (1993).
112
-------
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 mUmin.
B.3.2.3 Head Tank Support Stand (1) (Figure B.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:
2 Sides:
2 Ends:
16.7 cm x 33.7 cm
17.2 cm x 32.5 cm
17.2 cm x 16.7cm
3. Size is such that both head tanks fit into support
stand for storage and transport.
B.3.2.4 Water Bath (1) (Figure B.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
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 bot-
tom 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
downward 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
electrical 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.
B.3.2.5 Latex Rubber Mold: If you plan to construct a
substantial number of exposure test beakers, as de-
scribed 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. After the 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"
113
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clamp and physically pulling the beaker from the
mold.
B.3.3 Suggested options for more exposure treatments
(examples given are for a three unit treatment system)
B.3.3.1 Exposure Holding Tanks and Head Tanks:
Same dimensions as for two unit system except that
three (3) of each should be made.
B.3.3.2 Head Tank Support Stand (1) (Figure B.3)
1. Outer diameter: 16.7 cm wide x 49.5 cm long x 17.8
cm high
2. Cutting dimensions:{acrylic plastic, 6 mm)
1 Bottom: 16.7 cm x 49.5 cm
2 Sides: 17.2 cm x 48.3 cm
2 Ends: 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.
B.3.3.3 Water Bath (1) (Figure B.3)
1. Outer diameter: 33cm wide x 56.4 cm long x 7.4 cm
high
2. Cutting dimensions: (acrylic plastic, 6 mm)
1 Bottom: 33 cm x 71.7 cm
2 Ends: 33 cm x 6.8 cm
2 Sides: 55 cm x 6.8 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
from corner edges are shown in the drawing for
future additions of "add-on" water baths.
4. Motor mount and junction box installation are the
same as for two unit system.
B.3.3.4 "Add-On" Water Bath (example given is for one
additional unit treatment system; Figure B.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 con-
nection as shown in Figure B.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.
B.3.4 Optional Automated Water Delivery Apparatus
For Table Top STIR Systems (examples given are for a
three unit treatment system)
B.3.4.1 Support Stand: A stand to support the auto-
mated water delivery apparatus, shown in Figure B.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 B.2 sits on top of the stand, and the water distri-
bution manifold as shown in Figure B.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 B.2.
B.3.4.2 Water Renewal Supply: If tests will be con-
ducted in the local water supply, then the head box
water inlet shown in Figure B.2 is simply plumbed into
the supply line. However, if the tests are conducted with
transported water or with reconstituted 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 treatment system, re-
spectively. 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.
B.3.4.3 Operation of Water Delivery Apparatus: The
head box water inlet solenoid valve (Figure B.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 separate timer
control switch. With four test 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 B.2).
114
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B.3.4.3.1 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 L of 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.
B.3.4.3.2 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
ffow. One hour after they come on, the second timer will
shut off the solenoid valve inlet and water pump.
B.3.4.3.3 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 B.2.
B.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 unpublished data from ERL-Duluth (G.T.
Ankley, USEPA, Duluth, MM, personal communication).
B.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 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
completely separate tanks containing homogenized sedi-
ment from the same source as true replicates (of the
sediment "treatment") is inaccurate and is pseudorepli-
cation. Hence, because the same sediment is tested in
each beaker in a particular tank, and because the repli-
cation is focused on defining variability in the biology of
the organism (and the operator), this is essentially a
non-issue from a theoretical standpoint.
B.3.5.2 From a practical standpoint, it is important to
determine the potential influence of one beaker on an-
other 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 experi-
ment, four beakers of highly metal-contaminated sedi-
ment from the Keweenaw Waterway, Ml, were placed in
the same tank as four beakers containing clean sedi-
ment from West Bearskin Lake, MN. This was done in
two tanks; in one tank, 10 amphipods (Hyatelta azteca)
were added to each beaker, while in the other tank, 10
midges (Chironomus tentans) were placed in each bea-
ker. Controls for the experiment consisted of the West
Bearskin sediments assayed in separate "clean" tanks.
The four contaminated beakers were placed "upstream"
Table B.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) (ng/g)
WB2
WB
KW
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)
2 West Bearskin
3 Not determined
4 Keweenaw Waterway
of the four clean beakers to attempt to maximize pos-
sible exchange of contaminant. At the end of the test,
organism survival (and growth in for C. tentans) was
measured in two of the beakers from each site and
sediment Cu concentrations were determined in the
other two beakers from each site. The Keweenaw sedi-
ments contained concentrations of Cu in excess of
9,000 p.g/g (dry wt), and were toxic to both test species
(Table B.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 impor-
tant, there was no apparent increase in Cu concentra-
tions in the West Bearskin sediments held in the
Keweenaw tank (Table B.1).
B.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 concen-
trations were assayed: (1) in the midge test sediment
concentrations were about 150 jig dieldrin/g (dry weight)
and (2) in the amphipod test sediments contained in
excess of 450 \ig 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 organ-
isms 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 sediment assayed in the tank containing the spiked
sediment. This corresponded to the presence of mea-
surable dieldrin concentrations in unspiked Airport Pond
sediments in the tank with the mixed treatments
115
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Table 8.2 Sediment Dietdrin 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 Dkeldrin Concentrations and Organism Survival
and Growth in Airport Pond Sediment Tested in the
Same Tank as Dieldren-spiked Airport Pond
Sediment1
Sediment
AP2
AP
DAP"
AP
AP
DAP
Tank
1
2
2
3
4
4
Species
Amphipod
Amphipod
Amphipod
Midge
Midge
Midge
Survival
(%)
75
80
20
85
85
0
Dry wt
(mg/organism)
ND3
NO
ND
1.71
0.13
ND
Dieldrin
<0.01
0.07
446.4
<0.01
0.04
151.9
1 All values are the mean of duplicate observations (G.T. Ankley,
USEPA, Duluth, MN, unpublished data)
2 Airport Pond
3 Not determined
4 Dieldren-spiked Airport Pond
(Table B.2). The concentrations of dieldrin in the un-
spiked sediment, although detectable, were on the order
of 5,000-fold lower than the spiked sediments, indicating
relatively minimal transfer of pesticide.
B.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 bea-
kers containing clean Pequaywan Lake sediment, and
placed in a test tank with four beakers containing
Pequaywan Lake sediment without exogenous organic
carbon. Again, the control consisted of Pequaywan Lake
sediment held in a separate tank under otherwise identi-
cal test conditions. Assays were conducted, without
organisms, for 10 d. At this time, DO concentrations
were very low in the beakers containing trout chow-
amended sediment (ca., 1 mg/L, n = 4). However, over-
lying water DO concentration in the "untreated" vs. the
"treated" beakers in a separate tank, 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), interac-
tion between beakers within a tank is minimal.
B.3.5.5 One final observation germane to this issue is
worth noting. If indeed beakers of homogenized sedi-
ment within a test tank do not serve as suitable repli-
cates, 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 repli-
cates 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 common overlying water), this type of result would not
be expected.
B.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.
B.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).
B.4.1 Eight 35-mL polypropylene syringes equipped
with 18-gauge needles are suspended from a splitting
chamber (Figure B.4). The system is suspended above
eight beakers 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 B.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).
B.4.2 Water Splitting Chamber
B.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 x 30.5 cm glass bottom before assembly
(Figure B.4 and Table B.3). The glass bottom is made
from 4.8- (3/16 inch) or 6.4 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 B.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 adhe-
sive. Sharp glass edges should be sanded smooth using
a whetstone or a piece of carborundum wheel. After the
116
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Splitter Chamber
Top View
Stopper
Syringe
Overview
Leg Support
Splitter Chamber
Side View
Figure B.4 Water splitting chamber described in Zumwalt et al. (1994).
117
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Table B.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 (14.5 cm [5.75 inch])
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 silicone). These tubes are used
as sleeves for attaching the legs to the splitting cham-
ber. 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-thick x 2-cm-wide x 7-cm-long spacer is re-
quired so beakers and the optional waterbath can be slid
out the ends (Figure B.4). If the sleeves and legs are
attached to the ends of the splitting chamber, the bea-
kers and waterbath can be removed from the side. The
legs are inserted into the 12-mm tubes and secured
using nylon nuts or wingnuts. 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 under the splitting chamber.
B.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.
B.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.
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.
B.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.
B.4.3 Calibration and Delivery of Water to the Splitting
Chamber. 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 sy-
ringes 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 automatically 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 re-
cycled.
B.4.4 Waterbath and Exposure Beakers. The optional
waterbath surrounding the beakers is made from 3.2-mm
(1/8-inch) double-strength glass and is 15.8 cm wide x
29.5 cm long x 11.7 cm high (Figure B.4 [Figure B.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 ad-
hesive. The waterbath illustrated in Figure B.4 is op-
tional 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 tem-
perature control.
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B.4.4.5 Operation and Maintenance
B.4.4.5.1 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 toothbrush can be used to brush the outside of screens.
B.4.4.5.2 Overlying water can be aerated by suspend-
ing 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
suspended from the splitting chamber using stainless
steel hooks. Latex tubing (3.2-mm [1/8 inch] inner diam-
eter) 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.
B.4.4.5.3 The splitting chambers were used to deliver
water 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 sedi-
ment and 175 ml of overlying water at 23°C. Midges in
each beaker received a daily suspension of 4 mg
Tetrafin® flake food and survival and growth were mea-
sured 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 deliv-
ered with a single-cell diluler 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 between 24 beakers was less than
5%.
B.4.4.5.4 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 exposure compared with the beginning of the expo-
sure. 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 exposure and was about 2 to 3 mg/L
lower than inflowing water by the end of the exposure.
Survival and growth of midges were reduced with expo-
sure to metal-contaminated sediments. 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 similar toxic effects
on midges.
B.4.4.5.5 The system has been used to deliver 33 %o
salt water to exposure chambers for 10 d. Precipitation
of salts on the tips of the needles reduced flow from the
syringes. Use of a larger bore needle (16-gauge) re-
duced clogging problems; however, daily brushing of the
needle tips is required. Use of larger bore needles with
300-mL beakers containing 100mL 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.
119
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Appendix C
Food Preparation
C.1 Yeast, Cerophyl®, and Trout Chow (YCT) for
feeding the cultures and Hyalella azteca. Food should
be stored at 4°C and used within two weeks from
preparation; however, once prepared, YCT can be fro-
zen until use.
C.1.1 Digested trout chow is prepared as follows:
1. Preparation of trout chow requires one week.
Use starter or No. 1 pellets prepared according
to current U.S. Fish and Wildlife Service specifi-
cations. 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 (612/864-3181); 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 deion-
ized water. Mix well in a blender and pour into a
2-L separatory funnel or similar container. Di-
gest before use by aerating continuously from
the bottom of the vessel for one week at ambi-
ent laboratory temperature. Water lost due to
evaporation is replaced during digestion. Be-
cause of the offensive odor usually produced
during digestion, the vessel should be placed in
a ventilated area.
3. At the end of dige otion period allow material to
settle for a minimum of 1 h. Filter the 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.
C.1.2 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 ex-
cess material.
C.1.3 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,
alfalfa 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.
C.1.4 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 immedi-
ately, or it can be frozen until needed. Thawed
food is stored in the refrigerator between feed-
ings and is used for a maximum of two weeks.
Do not store YCT frozen over three months.
120
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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. Cul-
tures are fed 1 0 mUL on Monday and 5 mL/L on
Wednesday and Friday (USEPA, 1993).
C.2 Algal Food: Starter cultures of the green algae,
Selenastrum capricornutum and the diatom Navicula (or
Synedra) are available from the following sources: Ameri-
can Type Culture Collection (Culture No. ATCC 22662),
12301 Parklawn Drive, Rockville, MD 10852, or Culture
Collection of Algae, Botany Department, University of
Texas, Austin, TX 78712.
C.2.1 Algal Culture Medium for the green algae and
diatoms (Navicula or Synedra) prepared as follows
(USEPA, 1993a):
Table C.1 Nutrient Stock Solutions for Maintaining Algal Stock
Cultures
Stock Compound Amount dissolved in
solution 500 mL deionized water
1 . Macronutrients
A. MgCI2>6H2O 6.08 g
CaCI,-2H2O 2.20 g
NaN03 12.75g
B. MgSCv7H2O 7.35 g
C. K2HPO4 0.522 g
D. NaHCO3 7.50 g
2. Micron utrients
H3BO3 92.8 mg
MnCl2-4H2O 208.0 mg
ZnCI2 I.64 mg1
FeCI3-6H2O 79.9 mg
CoCI2-6H2O 0.71 4 mg2
Na,,MoCy2H2O 3.63 mg3
CuCI2-2H2O 0.006 mg"
NajEDTA-2H2O I50.0 mg
NajSeO4 1.196mg5
Table C.2 Final Concentration of Macronutrients and Micronu-
trlents in the Algal Culture Medium
Macronutrient Concentration Element Concentration
(mg/L) (mg/L)
NaNO3 25.5 N 4.20
MgCl2-6H2O 12.2 Mg 2.90
CaCI2-2H2O 4.41 Ca 1.20
MgSO4'7H2O 14.7 S 1.91
K2HPO4 1.04 P 0.186
NaHCO3 15.0 Na 11.0
K 0.469
C 2.14
Micronutrient Concentration Element Concentration
dig/L) (ng/L)
H3BO3 185 B 32.5
MnCI2-4H2O 416 Mn 115
ZnCI2 3.27 Zn 1.57
CoCI2-6H2O 1.43 Co 0.354
CuCI2-2H2O 0.012 Cu 0.004
Na,,MoO4'2H2O 7.26 Mo 2.88
FeCI3-6H2O 160 Fe 33.1
Na.,EDTA-2H2O 300 — —
Na2SeO4 2.39 Se 0.91
1 . Prepare stock nutrient solutions using reagent
grade chemicals as described in Table C.1 .
2. Add 1 mL of each stock solution, in the order
listed in Table C.1, to about 900 mL of deion-
ized water. Mix well after the addition of each
solution. Dilute to 1 L, mix well. The final con-
centration of macronutrients and micronutrients
in the culture medium is listed in Table C.2.
3. Immediately filter the medium through a 0.45
urn pore diameter membrane at a vacuum of not
more than 380mm (15 in.) mercury, or at a
pressure of not more than one-half atmosphere
(8 psi). Wash the filter with 500 mL deionized
water before use.
I2—Weigh out 164 mg and dilute to 100 mL. Add 1 mL of this
solution to micronutrient stock.
2CoCI2-6H2O—Weigh out 71.4 mgand dilute to 100 mL. Add 1 mLof this
solution to micronutrient stock.
3Na,,MoO4-2H2O—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.
,—Weigh out 119.6 mg and dilute to 100 mL. Add 1 mL of th is
solution to micronutrient stock.
4. If the filtration is carried out with sterile appara-
tus, filtered medium can be used immediately,
and no further sterilization steps are required
before the inoculation of the medium. The me-
dium 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 sub-
stantial loss of water by evaporation.
121
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C.2.2 Algal Cultures: Two types of algal cultures are
maintained: (1) stock cultures and (2) "food" cultures.
C.2.2.1 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
containing 100mL algal culture medium (pre-
pared 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 cul-
tures may be maintained at 25°C in environ-
mental chambers with cultures of other organ-
isms 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
provide 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 microor-
ganisms.
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 established outside sources of organisms
every four to six months.
C.2.2.2 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
cultures or cultures of other organisms if the
illumination is adequate (continuous "cool-white"
fluorescent 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 separately funnel, or are
manually mixed twice daily. If the cultures are
placed on a magnetic stir plate, heat generated
by the stirrer might elevate the culture tempera-
ture several degrees. Caution should be taken
to prevent the culture temperature from rising
more than 2 to 3°C.
C.2.2.3 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 al-
lowing the cultures to settle in a refrigerator for
at least three weeks and siphoning off the su-
pernatant.
2. The cell density (cells/mL) in the concentrate is
measured with an electronic particle counter,
microscope and hemocytometer, fluorometer,
or spectrophotometer and used to determine
the dilution (or further concentration) required to
achieve a final cell count of 3.0 to 3.5 X 107
cells/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
culture at 7 to 10 d will provide 4.5 X 109 algal
cells.
4. Algal concentrate can be stored in the refrigera-
tor for one month.
5. Cultures of Hyalella azteca are fed 10 mL/L on
Monday and 5 mUL on Wednesday and Friday
(ERL-Duluth, 1993).
122
-------
C.2.2.4 Establishing and Maintaining Stock Cultures of
Diatoms (Navicula sp. or Synedra sp.):
1. Upon receipt of the "diatom starter" culture (usu-
ally about 10 mL), a stock culture is started by
aseptically transferring 1 mL to each of several
250-mL culture flasks containing 100 mL algal
culture medium (prepared as described above).
The remainder of the starter culture can be held
in reserve for up to six months in a refrigerator
(in the dark) at 4°C.
2. The stock cultures are used as a source of
diatoms to initiate "diatom food" cultures. The
volume of stock culture maintained at any one
time will depend on the amount of 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 cul-
tures may be maintained at 23°C in environ-
mental chambers with cultures of other organ-
isms if the illumination is adequate (continuous
"cool-white" fluorescent lighting of about
1075 lux).
4. Cultures are mixed twice daily by hand.
5. Stock cultures can be held in the refrigerator
until used to start "diatom 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 provide an initial cell density of about
10,000 to 30,000 cells/mL in the new stock cul-
tures. Aseptic techniques should be used in
maintaining the stock algal cultures, and care
should be exercised to avoid contamination by
other microorganisms.
6. Stock cultures should be examined 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 established outside sources of organisms
every four to six months.
C.2.2.5 Establishing and Maintaining "Diatom food"
Cultures:
1. "Diatom food" cultures are started about 10 d
before use. About 20 mL of 7-d-old algal stock
culture (described in the previous paragraph)
are added to each liter of fresh culture medium
(e.g., 3 L of medium in a 4-L bottle). Aseptic
techniques should be used in preparing and
maintaining the cultures, and care should be
exercised to avoid contamination by other mi-
croorganisms. However, sterility of food cul-
tures is not as critical as in stock cultures be-
cause the food cultures are used in 7 to 10 d. A
one-month supply of diatom food can be grown
at one time and stored in the refrigerator.
2. Food cultures may be maintained at 23°C in
environmental chambers, but not with the algae
stock cultures. The illumination is continuous
"cool-white" fluorescent lighting of about
1075 lux). Higher temperatures can be prob-
lematic for diatom cultures.
3. Cultures are mixed continuously on a magnetic
stir plate (with a medium size stir bar) in a
moderately aerated separatory funnel, or are
manually mixed twice daily. Cultures become
very brown before harvesting. If the cultures are
placed on a magnetic stir plate, heat generated
by the stirrer might elevate the culture tempera-
ture several degrees. Caution should be taken
to prevent the culture temperature from rising
more than 2 to 3°C.
C.2.2.6 Preparing Concentrate of Diatoms for Use as
Food:
1. A diatom concentrate containing 1 X 109 cells/
mL is prepared from food cultures by centrifug-
ing the algae with a plankton or bucket-type
centrifuge, or by allowing the cultures to settle in
a refrigerator for at least three weeks and si-
phoning off the supernatant.
2. The cell density (cells/mL) in the concentrate is
measured with an electronic particle counter,
microscope and hemocytometer, fluorometer,
or spectrophotometer and used to determine
the dilution (or further concentration) required to
achieve a final cell count of 1 X 109 cells/mL.
3. Algal concentrate can be stored in the refrigera-
tor for one month.
4. Cultures of Hyaletla azteca are fed 10 mL/L on
Monday and 5 mL/L on Wednesday and Friday
(ERL-Duluth, 1993).
C.2.2.7 Cell counts:
1. Several types of automatic electronic and opti-
cal particle counters are available to rapidly
count cell number (cells/mL) and mean cell
volume (MCV; nrrvVcell). The Coulter Counter is
widely used and is discussed in detail in USEPA
(1978). When the Coulter Counter is used, an
aliquot (usually 1 mL) of the test culture is
diluted 10X to 20X with a 1% sodium chloride
electrolyte solution, such as Coulter ISOTON®,
to facilitate counting. The resulting dilution is
counted using an aperture tube with a 100-nm
123
-------
diameter aperture. 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 fol-
lowing 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
desired).
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
microscope 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
examination of the condition of the cells.
C.3 Tetrafin® food for culturing and testing C. tentans.
Food should be stored at 4°C and used within two
weeks from preparation or can be frozen until use.
1.
2.
Blend the Tetrafin® food in deionized water for
1 to 3 min or until very finely ground.
Filter slurry through an #110Nitex screen to
remove 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.
Tetrafin® food is added to each culture cham-
ber to provide about 0.04 mg dry solids/mL of
culture water. A stock suspension of the solids
is prepared 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 chamber volume is 8 L, 300 mg of food
would be added daily by adding 5 mL of a 56 g/
L stock suspension (USEPA, 1993).
4.
In a sediment test, Tetrafin® food (4 g/L)
added at 1.5 mL daily to each test chamber.
is
124
-------
Appendix D
Sample Data Sheets
125
-------
Culture
Acjar urn
A
B
C
D
E
F
Date o1 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
Egg
Masses
Produced
General
Comments
Initials of
Culturist
Figure D.1 Data sheet for the evaluation of a Chironomus tertians culture.
126
-------
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 (CuSO^ or KCI)
Reference Toxicant Supplier and Lot No.
Reference Toxicant Purity
Test Initiation Date
Toxicologist
Exposure Duration (H)
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 Toxicant 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.2 Data sheet for performing reference toxicant tests.
127
-------
S'.'d'rrert Sample Sojrce_
Date c' Test Initiation
To* col?, g:st Conducting Test.
Rechcate
Samo;ed
Temperature
TC)
Dissolved
Oxygen
(mg/L)
pH
Hardness
(mg/L)
Alkalinity
(mg/L)
Specific
Conductance
(umhos/cm)
Total
Ammonia
(mg/L)
Figure 0.3 Data sheet lor temperature and overlying water chemistry measurements.
128
-------
Study Code_
Study Name_
Building
Study Directory
Lead Technician
Daily Checklist for Sediment Tests
Diluter
Waterbath
Target temperature.
Acceptable Range .
Month
°C
°Cto
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
Figure D.4
Data sheet for daily checklist for sediment tests.
129
-------
Water Quality Data Sheet
Study Code
Study Name
Dissolved
Oxygen
(mg/L)
Date Test Day
_ , . Conductivity
Temp. Saliniy o/ H
'C (PPt) US)
Study Director Investigator
i"
Alkalinity 'z-
(ppm ^ p <8 1
c***c^c\ \ o. i -*. E ^
oauu3; ii ^ i ^
ml of Other
ISA
Volume of
Sample =
Sample Code
Meter #
Initials
Comments
Figure D.5
ml
ml
ml
ml
ml
ml
(ml Titrant x mult.
factor =)
ml
(ml Titrant x mult.
factor =)
Approved by_
ra
,o
I I
Date
ml
ml
Data sheet for water quality parameters.
130
-------
Chemistries
Test Typf
Organisrr
Test
Dates
I.D.
3 Sample Info Water Tvoe
Experimenter
Test
System
j Day 1 -1 0 1 2 3 4 5 6 7 8 9 10 Remarks
PH
DO (mg/L)
Temp (°C)
Hard/Alk
PH
DO (mg/L)
Temp (°C)
Hard/Alk
PH
DO (mg/L)
Temp (°C)
Hard/Alk
PH
DO (mg/L)
Temp (°C)
Hard/Alk
Figure D.6
Chemistry data sheet.
131
-------
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 0.7 Dally comment data sheet.
132
-------
Weight Data Form
Test Dates Species
Test Material Weighing Date Food
Location Oven Temp (°C) Age Organisms
Analyst Drying Time (h) Initial No/Rep
Wt of (mg) Wt of (mg)
oven dried pan + Mean dry (mg)
Sample Replicate pan organism Total (mg) Survival (#) wt./ Mean/Sample
Figure D.B Weight data sheet.
133
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