£EPA
Un,:ec States
Env rcnmental Protection
Agency
Development
Wasni.ngton DC 20460
June 1994
Methods for
Assessing the
Toxicity of
Sediment-associated
Contaminants with
Estuarine and
Marine Amphipods
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Errata
Pages 80-82, Sections 11.4.5-11.4.5.3, Effects of Sediment-
associated Ammonia
These sections describe a procedure that can be used to reduce
ammonia concentrations in field-collected sediments prior to
conducting laboratory toxicity tests. For dredged material
testing under the Clean Water Act or the Marine Protection,
Research, and Sanctuaries Act, the following alternative
procedure should be used. This procedure was described in a
December 21, 1993 guidance memorandum issued by the U.S. EPA
Office of Wetlands, Oceans and Watersheds, U.S. EPA Office of
Science and Technology, and U.S. Army Corps of Engineers
Operations, Construction, and Readiness Division.
For dredged material testing the following procedure should be
used if it is necessary to reduce interstitial water ammonia
levels. Whenever chemical evidence of ammonia is present at
toxicologically important levels, and ammonia is not a
contaminant of concern, the laboratory analyst should reduce
ammonia in the sediment interstitial water to species-specific
no-effect concentrations (see table 11.4 on page 81). Ammonia
levels in the interstitial water can be reduced by sufficiently
aerating the sample and replacing two volumes of water per day.
The analyst should measure interstitial ammonia each day until it
reaches the appropriate species-specific no-effect concentration.
After placing the test organism in the sediment, the analyst
should ensure that ammonia concentrations remain within an
acceptable range by conducting the toxicity test with continuous
flow or volume replacement not to exceed two volumes per day.
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EPA 600/R-94/025
June 1994
Methods for Assessing the Toxicity
of Sediment-associated Contaminants with
Estuarine and Marine Amphipods
Office of Research and Development
U.S. Environmental Protection Agency
Narragansett, Rhode Island 02882
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Disclaimer
This document has been reviewed in accordance with I'.S. Environmental Protection
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 potentially
pose a threat to a variety of aquatic ecosystems. Sediment functions as a reservoir for
common contaminants such as pesticides, herbicides, polychlorinated biphenyls (P('Bs).
polycyclic aromatic hydrocarbons, and metals such as lead, mercury, and arsenic. In-
place contaminated sediment can result in de pan pa rate benthic 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 characterizations. The United States Environmental Protection
Agency (LSEPA) 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
contaminants 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 I'SEPA'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 LSEPA program offices to address the problem of contaminated sediment. The
sediment test methods in this manual will be used by LSEPA to make decisions under a
range of statutory authorities concerning such issues as: dredged material disposal.
registration of pesticides and toxic substances, Superfund site assessment, and
assessment and cleanup of hazardous waste treatment, storage, and disposal facilities.
The use of uniform sediment testing procedures by LSEPA 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
Rick Swartz, LSEPA, Newport, OR.
m
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Abstract
A laboratory method is described for determining the short-term toxicity of
contaminated whole sediments using marine and estuarine amphipod crustaceans.
Sediments may be collected from estuarine or marine environments or spiked with
compounds in the laboratory. A test method is outlined that may be used with any of
four amphipod species, including Ampelisca abdita, Eohaustorius estuarius, Leptocheirus
plumulosus, and Rhepoxynius abronius. The toxicity test is conducted for 10 d in 1 L
glass chambers containing 175 inL of sediment and 800 mL of overlying water.
Overlying water is not renewed, and test organisms are not fed during the toxicity tests.
Temperature and salinity of overlying water, and choice of negative control sediment.
are species-specific. The choice of reference sediment may also be species-specific under
certain applications. The endpoint in the toxicity test is survivaJ, and reburial of
surviving amphipods is an additional measurement that can be used as an endpoint.
Procedures are described for use with sediments from oligohaline to fully marine
environments.
IV
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Acknowledgements
This document is a general purpose testing manual for estuarine and marine sediments.
The approaches have also been described in Swartz et al. (1985), Scott and Redmond
(1989), DeWitt et al. (1>>89), Schlekat et al. (1992), ASTM (1992) and ASTM (1994a).
The manual incorporates general guidelines that reflect the consensus of the Freshwater
Sediment Toxicity Assessment Committee and the U.S. Environmental Protection Agency
(I SEPA) Program Offices. Members of the Freshwater Sediment Toxicity Assessment
Committee 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, MI; 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.E. Schlekat and K.J. Scott, SAIC,
Narragansett, Rl and the document was prepared under contract No. 68-CO-004.
Contributors to specific sections of this manual are listed alphabetically 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, RI
K.J. Scott, SAIC, Narragansett, RI
2. Sections 10-11; Culture and Test Methods
C.E. Schlekat, SAIC, Narragansett, RI
K.J. Scott, SAIC, Narragansett, RI
R. Swartz, USEPA, Newport, OR
T. Dewitt, USEPA, Newport, OR
3. Section 12; Statistical Analysis
J. Heltshe, SAIC, Narragansett, RI
R.A. Hoke, SAIC, Hackensack, NJ
H. Uee. USEPA, Newport, OR
T.J. Norberg-King, USEPA, Duluth, MN
C.E. Schlekat, SAIC, Narragansett, RI
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•4. Section 13; Precision and Accuracy
G.A. Burton, Wright State University, Dayton, OH;
T.J. Norberg-King, L'SEPA, Duluth, MN
C.E. Schlekat, SAIC, Narragansett, RI
K.J. Scott, SAIC, Narragansett, RI
Review comments from the following individuals are gratefully acknowledged: B.L.
McGee, University of Maryland, Queenstown, MD and M.S. Redmond. Northwest
Aquatic Science, Newport, OR; 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.; D. Klemm, EMSU and L. Cast, TA1,
Newtown, OH; G. Hanson, OSW, Washington, D.C; R. Swartz, S. Ferraro, ERL-N,
Newport, OR; J. Arthur, R. Spehar, and C. Stephan, ERL-D, Duluth, MN.
Participation by the following laboratories in the round-robin testing is greatly
appreciated: Aqua Survey, Incorporated, Flemington, NJ; Battelle Marine Science
Laboratory, Sequim, WA; Environment Canada, Atlantic Regional Laboratory,
Dartmouth, NS; EVS Environmental Consultants, North Vancouver, BC; Science
Applications International Corporation, Environmental Testing Center, Narragansett,
RI; USACE, Waterways Experiment Station, Vicksburg, MS; USEPA, Edison, NJ;
USEPA, Gulf Breeze, FL: USEPA, Narragansett, RI; USEPA, Newport, OR.
USEPA's Office of Science and Technology provided support for the development of this
manual.
VI
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Contents
Disclaimer ij
Foreword iii
Abstract iv
Acknowledgements v
Figures \ii
Tables , viii
1. Introduction 1
1.1 Significance of Use 1
1.2 Program Applicability 3
1.3 Scope and Application 7
1.4 Performance-based Criteria 15
2. Summary of Method 16
2.1 Method Description and Experimental Design 16
2.2 Types of Tests 18
2.3 Test Endpoints 18
3. Definitions 19
4. Interferences 22
4.1 General Introduction * 22
4.2 Non-Contaminant Factors 24
4.3 Changes in Unavailability 25
4.4 Presence of Indigenous Organisms 26
5. Health, Safety, and Waste Management 27
5.1 General Precautions 27
5.2 Safety Equipment 27
5.3 General Laboratory and Field Operations 28
5.4 Disease Prevention 29
5.5 Safety Manuals 29
5.6 Pollution Prevention, Waste Management, and Sample Disposal 29
6. Facilities, Equipment, and Supplies 30
6.1 General 30
6.2 Facilities . 30
6.3 Equipment and Supplies 31
7. Water, Reagents, and Standards 36
7.1 Water 36
7.2 Reagents 38
7.3 Standards 38
VII
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Contents (continued)
8. Sample Collection, Storage, Manipulation, and Characterization 39
8.1 Collection 39
8.2 Storage 39
8.3 Manipulation 40
8.4 Characterization 43
9. Quality Assurance and Quality Control 45
9.1 Introduction 45
9.2 Performance-based Criteria 45
9.3 Facilities, Equipment, and Test Chambers 46
9.4 Test Organisms 46
9.5 Water 46
9.6 Sample Collection and Storage 46
9.7 Test Conditions 46
9.8 Quality of Test Organisms 47
9.9 Quality of Food 47
9.10 Test Acceptability 47
9.11 Analytical Methods 48
9.12 Calibration and Standardization 48
9.13 Replication and Test Sensitivity 48
9.14 Demonstrating Acceptable Performance 48
9.15 Documenting Ongoing Laboratory Performance 49
9.16 Reference Toxicants 49
9.17 Record Keeping 50
10. Collection, Culture, and Maintaining of Test Organisms 53
10.1 Life History 53
10.2 Species Selection 55
10.3 Field Collection 55
10.4 Holding and Acclimation 62
10.5 Culture Procedure for Leptocheirus plumulosus 64
11. Test Method 100.4 Ampelisca abdita, Eohaustorius estuarius, Leptocheirus
plumulosus, or Rhepoxynius abronius 10-d Survival Test for Sediments 68
11.1 Introduction 68
11.2 Recommended Test Method for Conducting a 10-d Sediment Toxicity
Test with Ampelisca abdita, Eohaustorius estuarius, Leptocheirus
plumulosus, or Rhepoxynius abronius 68
11.3 General Procedures 72
11.4 Interpretation of Results 78
vni
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Contents (continued)
12. Data Recording, Data Analysis and Calculations, and Reporting 83
12.1 Data Recording 83
12.2 Data Analysis 83
12.3 Data Interpretation 109
12.4 Reporting 110
13. Precision and Accuracy 112
13.1 Determining Precision and Accuracy 112
13.2 Accuracy 113
13.3 Replication and Test Sensitivity 113
13.4 Demonstrating Acceptable Laboratory Performance 113
13.5 Precision of Sediment Toxicity Test Methods 115
References 122
Appendix
A. Sample data sheets 135
IX
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Figures
Figure 12.1 Treatment response for a Type I and Type II error.
89
Figure 12.2 Power of the test vs percent reduction in treatment response relative to
the control mean at various CV's (8 replicates, alpha = 0.05 (one-
tailed)) 90
Figure 12.3 Power of the test vs percent reduction in treatment response relative to
the control mean at various CV's (5 replicates, alpha = 0.05 (one-
tailed)) 91
Figure 12.4 Power of the test vs percent reduction in treatment response relative to
the control mean at various CV's (8 replicates, alpha = 0.10 (one-
tailed)) 92
Figure 12.5 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)} 94
Figure 12.6 Fffect of alpha and beta on the number of replicates at various CV's
(assuming combined alpha + beta = 0.25) 95
Figure 12.7 Decision tree for analysis survival and growth data subjected to
hypothesis testing 104
Figure 12.8 Decision tree for analysis of point estimate data 106
Figure A.I Field collection and laboratory holding data sheet 136
Figure A.2 Data sheet for % h reference toxicant test 137
Figure A.3 Data sheet for daily observations during the 10-d solid phase test 138
Figure A.4 Data sheet for 10-d solid phase test 139
Figure A.5 Data sheet for test summary 140
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Tables
Table 1.1 Sediment quality assessment procedures 4
Table 1.2 Statutory needs for sediment quality assessment 6
Table 1.3 Rating of selection criteria for estuarine and marine sediment toxicity
testing organisms 11
Table 4.1 Advantages and disadvantages for use of sediment tests 23
Table 6.1 Equipment and supplies for culturing and testing estuarine and marine
amphipods. Supplies are for all species unless specified 34
Table 9.1 Recommended test conditions for conducting reference-toxicity tests . 51
Table 10.1 Comparison of habitat characteristics and other life history parameters
of four estuarine and marine amphipod species used in sediment
toxicity tests 57
Table 11.1 Test conditions for conducting a 10-d sediment toxicity test with Ampelisca
abdita, Eohaustorius estuarius, Leptocheirus plumulosus, or Rhepoxynius
abronius 69
Table 11.2 General activity schedule for conducting a sediment toxicity test with
Ampelisca abdita, Eohaustorius estuarius, Leptocheirus plumulosus, or
Rhepoxynius abronius 71
Table 11.3 Test acceptability requirements for a 10-d sediment toxicity test with
Ampelisca abdita, Eohaustorius estuarius, Leptocheirus plumulosus, or
Rhepoxynius abronius 73
Table 11.4 Application limits for 10-d sediment toxicity tests with Ampelisca
abdita, Eohaustorius estuarius, Leptocheirus plumulosus, or Rhepoxynius
abronius 81
Table 12.1 Suggested a levels to use for tests of assumptions 87
Table 13.1 Inter-laboratory precision for survival of Rhepoxynius abronius in 10-d
whole sediment toxicity tests using seven sediments 117
Table 13.2 Inter-laboratory precision for survival of Ampelisca abdita in
10-d whole sediment toxicity tests using four sediments 118
Table 13.3 Interlaboratory precision for survival of Eohaustorius estuarius in 10-d
whole sediment toxicity tests using four sediments 120
XI
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Tables (continued)
Table 13.4 Inter-laboratory precision for survival of Leptocheirus plumulosus in
10-d whole sediment toxicity tests using four sediments 121
XII
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Section 1
Introduction
1.1 Significance of Use
1.1.1 Sediment provides habitat for many estuarine and marine organisms 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
sediment. Mounting evidence exists of environmental degradation in areas where
USEPA Water Quality Criteria (WQC) 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 contaminants in sediment may be several orders of magnitude higher
than in the overlying water; however, bulk sediment concentrations have not been
strongly correlated to bioavailability (Burton, 1991). Partitioning or sorption of a
compound between water and sediment may depend on many factors including: aqueous
solubility, pH, redox, affinity for sediment organic carbon and dissolved organic carbon.
grain size of the sediment, sediment mineral constituents (oxides of iron, manganese, and
aluminum), and the quantity of acid volatile sulfides in sediment (Di Toro et al., 1990;
Di Toro et al., 1991). Although certain chemicals are highly sorbed to sediment, these
compounds may still be available to the biota. Contaminated sediments may be directly
toxic to aquatic life or can be a source of contaminants for bioaccumulation in the food
chain.
1.1.2 Assessments of sediment quality have commonly included sediment chemical
analyses and surveys of benthic community structure. Determination of sediment
contaminant concentrations on a dry weight basis alone offers little insight into
predicting adverse biological effects because bioavailability may be limited by the
intricate 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 fluctuations, physical parameters, and biotic interactions. In order to
obtain a direct measure of sediment toxicity, 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 contamination that can only be inferred
from chemical or benthic community analyses. The L'SEPA 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
methods will provide high quality data needed for the national inventory and for
regulatory programs to prevent, remediate, and manage contaminated sediment
(Southerland et al., 1991).
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1.1.3 The objective of a sediment test is to determine whether contaminants in sediment
are harmful to or are hioaccumulated by benthic organisms. The tests can be used to
measure interactive toxic effects of complex contaminant mixtures in sediment.
Furthermore, knowledge of specific pathways of interactions among sediments 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
hioavailability. (2) investigate interactions among contaminants, (3) compare the
sensitivities of different organisms, (4) determine spatial and temporal distribution of
contamination, (5) evaluate hazards of dredged material, (6) measure 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 chemicals
and biological responses. Results of toxicity tests with test materials spiked into
sediments at different concentrations may be reported in terms of an LC50 < median
lethal concentration), an EC50 (median effect concentration), an IC50 (inhibition
concentration), or as a NOEC (no observed effect concentration) or LOEC (lowest
observed effect concentration). However, spiked sediment may not be representative of
contaminated sediment 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 responses.
1.1.5 Evaluating effect concentrations for chemicals in sediment requires knowledge of
factors controlling their bioavailability. Similar concentrations of a chemical in units of
mass of chemical per mass of sediment dry weight often exhibit a range in toxicity in
different sediments (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 bioavailability of non-ionic organic compounds in sediment is
often inversely correlated with the organic carbon concentration. Whatever the route of
exposure, these corn Nations of effect concentrations to interstitial water concentrations
indicate that predicted or measured concentrations in interstitial water can be used to
quantify the exposure concentration to an organism. Therefore, information on
partitioning of chemicals between solid and liquid phases of sediment is useful for
establishing effect concentrations (Di Toro et al., 1991).
1.1.6 Field surveys can be designed to provide either a qualitative reconnaissance of the
distribution of sediment contamination or a quantitative statistical comparison of
contamination among sites. Surveys of sediment toxicity are usually part of more
comprehensive analyses of biological, chemical, geological, and hydrographic data.
Statistical correlations may be improved and sampling costs may be reduced if
subsampies are taken simultaneously for sediment tests, chemical analyses, and benthic
community structure.
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1.1.8 Table 1.1 lists several approaches the USEPA has considered for the assessment of
sediment quality (L'SEPA, 1992c). These approaches include: (1) equilibrium
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 Range Effects median (see Chapman, 1989: L'SEPA, 1989a: LSEPA, 199<)a:
I'SEPA, 1990b; and LSEPA, 1992b for a critique of these methods). The sediment
assessment approaches listed in Table 1.1 can be classified as numeric (e.g., equilibrium
partitioning), descriptive (e.g.. whole sediment toxicity tests), or a combination of
numeric and descriptive approaches (e.g., Apparent Effects Threshold: I'SEPA, 1992c).
Numeric methods can be used to derive chemical-specific sediment 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
evidence is the most desirable approach for assessing the effects of contaminants
associated with sediment (Long and Morgan. 1990). Hazard evaluatioas integrating data
from laboratory exposures, chemical analyses, and benthic community assessments
provide strong complementary evidence of the degree of pollution-induced degradation
in aquatic communities (Chapman 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 recently, 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 a L'SEPA management strategy for
contaminated sediments: (1) in-place sediment should be protected from contamination
to ensure beneficial uses of surface waters. (2) protection of in-place sediment should be
achieved through pollution prevention 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 surface waters in the United States. The goal of the CWA is to
restore and maintain physical, chemical, and biological integrity of the nation's waters
(Southerland et al., 1992). Federal and state monitoring programs traditionally have
focused on evaluating water column problems caused by point-source dischargers.
During the next few years, 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 methods will provide
high quality data needed for the national inventory and for regulatory program to
prevent, remediate, and manage contaminated sediment (Southerland et al., 1992).
1.2.3 The Office of Water (OW), the Office of Pesticide Programs (OPP), the Office of
Pollution Prevention and Toxic Substances (OPPT). the Office of Solid Waste (OSW),
and the Office of Emergency and Remedial Response (OERR) are all committed to the
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Table l.l
Sediment quality assessment procedures
Method
lype
Numeric Descriptive Combination Approach
Equilibrium Partitioning
Tissue Residues
Interstitial Water Toxicitv
Benthic Community Structure
A sediment quality value fur a
given contaminant is determined
by calculating the sediment
concentration of the contaminant
that corresponds to an interstitial
water concentration equivalent to
the USEPA water quality
criterion for the contaminant.
Safe sediment concentrations of
specific chemicals are established
by determining the sediment
chemical concentration that
results in acceptable tissue
residues.
Toxicity of interstitial water is
quantified and identification
evaluation procedures are
applied to identify and quantify
chemical components responsible
for sediment toxicity.
Environmental degradation is
measured by evaluating
alterations in benthic community
structure.
Note: Modified from USEPA (1992c).
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Table 1.1
Sediment quality assessment procedures (continued)
Method
Type
Numeric Descriptive Combination Approach
Whole sediment Toxicity
and Sediment Spiking
Sediment Quality Triad *
Apparent Effects Threshold
(AET)
Test organisms are exposed tn
sediments that may contain
known or unknown quantities nf
potentially toxic chemicals. At
the end of a specified time
period, the response of the test
organisms is examined in relation
to a specified endpoint. Dose-
response relationships can he
established by exposing lest
organisms to sediments that have
been spiked with known amounts
of chemicals or mixtures of
chemicals.
Sediment chemical
contamination, sediment toxicit).
and benthic community structure
are measured on the same
sediment sample.
Correspondence between
sediment chemistr>. toxicitv. and
field effects is used to determine
sediment concentrations that
discriminate conditions of
minimal, uncertain, and major
biological effects.
The sediment concentration of a
contaminant above which
statistically significant biological
effects (e.g., sediment tuxii.it> i
are always expected. AFT
are empirically derived from
paired field data for sediment
chemistry and a range of
biological effects indicators.
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Table 1.2
Statutory needs for sediment quality assessment
Law1
Area of Need
CERCLA • Assess need for remedial action with contaminated sediments; assess degree
of clean-up required, disposition of sediments
OVA • NPDES permitting, especially under Best Available Technology (BATi in
water-quality-limited water
• Section 403(c) criteria for ocean discharges; mandatory additional
requirements to protect marine environment
• Section 301 (g) waivers for publicly owned treatment works (POTVVS)
discharging to marine waters
• Section 404 permits for dredge and fill activities (administered by the
Corps of Engineers)
FIFRA • Review uses of new and existing chemicals
• Pesticide labeling and registration
MPRSA • Permits for ocean dumping
NEPA • Preparation of environmental impact statements for projects with surface
water discharges
TSCA • Section 5: Pre-manufacture notice reviews for new chemicals
• Sections 4, 5 and 6: Reviews for existing chemicals
RCRA • Assess suitability (and permit) on-land disposal or beneficial use of
contaminated sediments considered "hazardous"
CERCLA Comprehensive Environmental Response, Compensation and Liability Act
(Superfund)
OV A 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 Recoverv Act
Note:
Modified from Dickson et al. (1984) and Southerland et al. (1992).
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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 identified risks would
trigger regulatory actions.
1.2.4 Tiered testing should include a hierarchy of tests with the tests in each successive
tier becoming progressively more rigorous, complex, and costly (Southerland et al. 1992i.
Guidance needs to be developed to explain how information within each tier would
trigger regulatory action. The guidance could be program specific, describing decisions
based on a weight of evidence approach, a pass-fail approach, or comparison to a
reference site depending on statutory and regulatory requirements. There are now two
approaches for tiered testing used by USEPA: (1) the Office of Water-U.S. Armv C'orps
of Engineers dredged material testing framework and (2) the OPP ecological risk
assessment tiered testing framework. Tier 1 of the dredged material testing framework
consists of a review of existing chemical and biological data or an inventory of nearhv
.sources. In Tier 2, chemical data are compared to water and sediment quality criteria.
Tier 3 evaluations consist of acute toxicity and bioaccumulation testing, and a
comparison of the results to a reference area. Tier 4 studies consist of site-specific field
studies. The OPP testing framework consists of acute toxicity testing in Tier 1. followed
by chronic (early life stage) toxicity testing in Tier 2 and further chronic toxicity testing
(full life cycle) in Tier 3. A tiered testing framework has not yet been chosen for
Agency-wide use, but some of the components have been identified to be standardized.
These components are toxicity tests, bioaccumulation tests, chemical criteria, and other
measurements that may have ecological significance including benthic communitv
structure evaluation, colonization rate, and in situ sediment testing within a mesocosm
U SEPA, 1992a).
1.3 Scope and Application
1.3.1 Procedures are described for testing estuarine and marine amphipod crustaceans
in the laboratory to evaluate the toxicity of contaminants associated with whole
sediments. Sediments may be collected from the field or spiked with compounds in the
laboratory. A toxicity method is outlined for four species of estuarine and marine
sediment-burrowing amphipods found within United States coastal waters. The species
are Ampelisca abdita, a marine species that inhabits marine and mesohaline portions of
the Atlantic coast, the Gulf of Mexico, and San Francisco Bay; Eohaustorius estuarius. a
Pacific coast estuarine species; Leptocheirus plumulosus, an Atlantic coast estuarine
species; and Rhepoxynius abronius, a Pacific coast marine species. Generally, the
method described may be applied to all four species, although acclimation procedures
and some test conditions (i.e., temperature and salinity) will be species-specific (Sections
10 and 11). The toxicity test is conducted for 10 d in 1 L glass chambers containing 1"5
mL of sediment and 8(10 mL of overlying seawater. Exposure is static (i.e., water is not
renewed), and the animals are not fed over the 10 d exposure period. The endpoint in
the toxicity test is survival, and reburial of surviving amphipods is an additional
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measurement that can he used as an endpoint. Procedures are described for use with
sediments with pore water salinity ranging from >0 c'rr to fully marine.
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 Leptocheirus
plumulosusi. (2) develop standard sediment bioaccumulation tests (i.e.. 28-d exposures
with the bivalve Macuma nasuta and the polychaete Nereis virens) (Lee et al., 1989), (3)
refine sediment spiking procedures, (4) refine sediment dilution procedures, (5) refine
sediment Toxicity Identification Evaluation (TIE) procedures, and (6) produce additional
data on confirmation of responses in laboratory tests with natural populations of benthic
organisms. This information will be described in future editions of the manual.
1.3.3 This methods manual serves as a companion to the freshwater sediment testing
method manual (LSEPA. 1994a).
1.3.4 Procedures described in this manual are based on the following documents:
Swartz et al. (1985). DeVVitt et al. (1989), Scott and Redmond (1989). Schlekat et aJ.
(1992). ASTM (1992), and Environment Canada (1992). This L'SEPA manual outlines
specific test methods for evaluating the toxicity of sediments with A. abdita, E. estuarius,
L. plutnulosus. and R. abrontus. While standard procedures are described in the
manual, further investigation of certain issues could aid in the interpretation of test
results. Some of these issues include the effect of shipping on organism sensitivity,
additional performance criteria for organism health, and confirmation of responses in
laboratory tests with natural benthos populations.
1.3.5 General procedures described in this manual might be useful for conducting tests
with other estuarine or marine organisms (e.g.. Corophium spp., Grandidieretla japonica,
Lepidactylus dytiscus, Streblospio benedicti), although modifications may be necessary.
Altering the procedures described in this manual may alter bioavaiiability and produce
results that are not directly comparable with results of acceptable procedures.
Comparison of results obtained using modified versions of these procedures might
provide useful information concerning new concepts and procedures for conducting
sediment tests with aquatic organisms. If tests are conducted with methods different
from those described 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 L'SEPA allows the use of indigenous species only where State regulations
require their use or prohibit importation of the recommended species. Where state
regulations prohibit the importation or use of the recommended test species, permission
should be requested from the appropriate regulatory agency before their using
indigenous species.
1.3.7 Where States have developed culturing and testing methods for indigenous species
other than those recommended in the manual, data comparing the sensitivity of the
substitute species and one or more of the recommended species must be obtained with
8
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sediments or reference toxicants, to ensure that the species selected are at least as
sensiti\e 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 (DeVVitt et al.. 1989: Swartz. 1989).
Ideally, a test organism should: (I) have a lexicological database demonstrating relati\e
sensitivity 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 from culture or through field collection,
(5) be easily maintained in the laboratory, (6) be easily identified. (7) be ecological!} 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 physico-chemical characteristics
(e.g.. grain size), and (10) be compatible with selected exposure methods and endpoints
(ASTM. 1993a). Methods utilizing selected organisms should also be ill) peer reviewed
(e.g., journal articles, ASTM guides) and (12) confirmed with responses with natural
populations of benthic organisms.
1.3.8.2 Of these criteria (Table 1.3), a database demonstrating relative sensitivity to
contaminants, contact with sediment, ease of culture in the laboratory or availabilitv for
field-collection, ease of handling in the laboratory, tolerance to varying sediment
physico-chemical characteristics, and confirmation with responses with natural benthic
populations were the primary criteria used for selecting A. abdita, E. estuarius,
L. plumulosus, and R. abronius for the current edition of the manual. The species
chosen for this method are intimately associated with sediment, due to their tube-
dwelling or free-burrowing, and sediment ingesting nature. Amphipods have been used
extensively to test the toxicity of marine, estuarine, and freshwater sediments (Swart/ et
at., 1985: DeVVitt et al., 1989; Scott and Redmond, 1989; DeWitt et al.. 1992a: Schlekat
et al.. 1992; ASTM, 1992). The selection of test species for this manual followed the
consensus of experts in the field of sediment toxicology who participated in a workshop
entitled "Testing Issues for Freshwater and Marine Sediments". The workshop was
sponsored by LSEPA Office of Water. Office of Science and Technology, and Office of
Research and Development, and was held in Washington. D.C. from 16-18 September
1992
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available, Crandidierella japonica (ASTM, 1992), was not selected because workshop
participants felt that the use of this species was not sufficiently broad to warrant
standardization of the method. Environment Canada (1992) has recommended the use
of the following amphipod species for sediment toxicity testing: Amphiporeia virginiana.
C Ontphium volutator. Eohaustorius washingtonianus, Foxiphalus xiximeus, and
l.eptocheirus pinguis. A database similar to those available for A. abdita, E. estuarius,
L. plumulosus, and R. abronius must be developed in order for these and other
organisms to be included in future editions of this manual.
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. Several studies have
evaluated the sensitivities of A. abdita, E. estuarius, L. plumulosus, or R. abronius. either
relative to one another, or to other commonly tested estuarine or marine species. For
example, the sensitivity of marine amphipods was compared to other species that were
used in generating saltwater Water Quality Criteria. Seven amphipod genera, including
Ampelisca abdita and Rhepoxynius abronius, were among the test species used to generate
saltwater \\ater Quality Criteria for 12 chemicals. Acute amphipod toxicity data from
•4-d water-only tests for each of the 12 chemicals was compared to data for (1) all other
species. (2) other benthic species, and (3) other in fan rial species. Amphipods were
generally of median sensitivity for each comparison. The average percentile rank of
amphipods
10
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Table 1.3 Rating of selection criteria for estuarine and marine sediment toxicity testing organisms
Criterion
Ampelixca abdita
Kohaustorius estuarius l^ptocheirus plumulosus Rhepoxyniua abronius
Relative sensitivity
toxicity database
Round-robin studies
conducted
Contact with sediment
Laboratory culture
Maintain in laboratory
Taxonomic
identification
Kcological importance
Geographical
distribution1
Sediment physico-
chemical tolerance
Lield-validated
Peer-reviewed
Kndpoinls monitored
All,, PAC,
PA(
AIL
PAC
Survival
Survival, reburial
Survival
Survival, reburial
Note:
All, = Atlantic Coast, PAC = Pacific Coast,
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among all species tested was 57.2%; among all benthic species. 55.5*7; and, among all
infuunal species, 54.3%. Thus, amphipods are not uniquely sensitive relative to all
species, benthic species, or even infaunal species (D. Hansen, USEPA. Narragansett, RI,
personal communication). Additional research may be warranted to develop tests using
species that are consistently more sensitive than amphipods. thereby offering protection
to k-ss sensitive groups.
1.3.8.4 Williams et al. (1986) compared the sensitivity of the K. abronius 10-d whole
sediment test, the oyster embryo (Crassostrea gigas) 48-h abnormality test, and the
bacterium (Photobacterium phosphoreum) 1-h luminescence inhibition test (i.e., the
Microtox4 test) to sediments collected from 46 contaminated sites in Commencement
Bav, \\ A. Rhepoxynius abronius were exposed to whole sediment, while the oyster and
bacterium tests were conducted with sediment elutriates and extracts, respectfully.
Microio\" was the most sensitive test with 63% of the sites eliciting significant
inhibition of luminescence. Significant mortality of/?, abronius was observed in 40% of
test sediments, and oyster abnormality occurred in 35% of sediment elutriates.
Complete concordance (i.e., sediments that were either toxic or not-toxic in all three
tt'sLs) uas observed in 41% of the sediments. Possible sources for the lack of
concordance at other sites include interspecific differences in sensitivity among test
organisms, heterogeneity in contaminant types associated with test sediments, and
differences in routes of exposure inherent in each bioassay. These results highlight the
importance of using multiple assays when performing sediment assessments.
1.3.8.5 Several studies have compared the sensitivity of combinations of the four
amphipods to sediment contaminants. For example, there are several comparisons
between A. abdita and R. abronius, between E. estuarius and R. abronius, and between
A. abdita and L. plumulosus. There are fewer examples of direct comparisons between
E. estuarius and L plumulosus, and no examples comparing L. plumulosus and
R. abronius. There is some overlap in relative sensitivity from comparison to
comparison within each species combination, which appears to indicate that all four
species are within the same range of relative sensitivity to contaminated sediments.
1.3.8.5.1 Word et al. (1989) compared the sensitivity of A. abdita and R. abronius to
contaminated sediments in a series of experiments. The experiments followed protocols
developed specifically for each species; thus, .4. abdita was tested at 20°C. whereas
R. abronius was tested at 15°C. Experiments were designed to compare the sensitivity of
the protocols rather than to provide a comparison of the response of the organism.
Sediments collected from Oakland Harbor, CA, were used for the comparisons. Twenty-
six sediments were tested in one comparison, while 5 were tested in the other. Analysis
of results using Kruskal Wallace rank sum test for both experiments demonstrated that
R. abronius exhibited greater sensitivity to the sediments than A. abdita. Long and
Buchman (1989) also compared the sensitivity of .4. abdita and R. abronius to sediments
from Oakland Harbor, CA. They also determined that A. abdita showed less sensitivity
than R. abronius, but they also showed that A. abdita was less sensitive to sediment grain
size factors than R. abronius.
12
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1.3.8.5.2 DeVVitt et al. (1989) compared the sensitivity of E. estuarius and R. abnmius to
sediment spiked with flouranthene and field-collected sediment from industrial
waterways in Puget Sound, WA, in 10-d tests, and to aqueous cadmium (CdCk in a 4-d
water-only test. The sensitivity of E. estuarius was from two (to flouranthene-spiked
sediment) to seven (to one Puget Sound, WA, sediment) times less sensitive than
R. abronius in sediment tests, and ten times less sensitive to CdCl, in the water-only test.
These results are supported by the findings of Pastorak and Becker (1990) who found
the acute sensitivity of E. estuarius and R. abronius to be generally comparable to each
other, and both were more sensitive than Neanthes (survival and biomass endpoints).
Panope (survival), and Dendraster (survival).
1.3.8.5.3 Leptocheirus plumulosus was as sensitive as the freshwater amphipod Hyalelta
azteca to an artificially created gradient of sediment contamination when the latter uus
acclimated to oligohaline salinity (i.e., 6 ftr) (McGee et al., 1993). DeVVitt et al. t!992bi
compared the sensitivity of L. plumulosus with three other amphipod species, two
molluscs, and one polychaete to highly contaminated sediment collected from Baltimore
Harbor, MD. that was serially diluted with clean sediment. Leptocheirus plumulosus was
more sensitive than the amphipods Hyalella azteca and Lepidactylus dytiscus and
exhibited equal sensitivity with E. estuarius. Comparisons using dilutions of sediment
collected from Black Rock Harbor. CT. show that ,4. abdita shows greater sensitivity
than L plumulosus when the latter is tested at 20CC (SAIC. 1993a). However.
L. plumulosus is more sensitive at 25°C, the temperature at which chronic test methods
with this species are being developed (DeYVitt, 1992a). than A. abdita at 20CC (SAIC.
1993a).
1.3.8.6 Limited comparative data is available for concurrent water-only exposures of all
four species in single-chemical tests. Studies that do exist generally show that no one
species is consistently the most sensitive.
1.3.8.6.1 The relative sensitivity of the four amphipod species to ammonia was
determined in ten-d water only toxicity tests in order to aid interpretation of results of
tests on sediments where this toxicant is present (SAIC, 1993c). These tests were static-
exposures that were generally conducted under conditions (e.g., salinity, photoperiod)
similar to those used for standard 10-d sediment tests. Departures from standard
conditions included the absence of sediment and a test temperature of 20 C for
L. plumulosus, rather than 25°C as dictated in this manual. Sensitivity to total ammonia
increased with increasing pH for all four species. The rank sensitivity was R. abronius •-•
,4. abdita > E. estuarius > L. plumulosus.
1.3.8.6.2 Cadmium chloride has been a common reference toxicant for all four species
in 4-d exposures. DeVVitt et al. (1992a) reports the rank sensitivity as R. abronius >
A. abdita > L. plumulosus > E. estuarius at a common temperature and salinity of 15 C
and 28 9rr. A series of 4-d exposures to cadmium that were conducted at species-specific
temperatures and salinities showed the following rank sensitivity: .4. abdita =
L. plumulosus = R. abronius > E. estuarius (SAIC, 1993a: SAIC, 1993b: and SAIC.
19930.
13
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1.3.8.6.3 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). For example. Reish (1988)
reported the relative toxicity of six metals (arsenic, cadmium, chromium, copper,
mercury, and zinc) to crustaceans, polychaetes, pelecypods, and fishes and concluded
that no one species or group of test organisms was the most sensitive to all of the metals.
1.3.8.7 The sensitivity of an organism is related to route of exposure and biochemical
response to contaminants. Sediment-dwelling organisms can receive exposure via from
three primary sources: interstitial water, sediment particles, and overlying water. Food
type, feeding rate, assimilation efficiency, and clearance rate will control the dose of
contaminants from sediment. Benthic invertebrates often selectively consume different
particle sizes (Harkey et al.. 1994) or particles with higher organic carbon
concentrations which 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 some amphipods (Landrum, 1989) and clams (Boese et al., 1990) uptake through the
gut can exceed uptake across the gills for certain hydrophobic compounds. Organisms
in direct contact with sediment may also accumulate contaminants by direct adsorption
to the body wall or by absorption through the integument (Knezovich et al., 1987).
1.3.8.8 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 , flouranthene. organochlorines, and metals have been correlated with either
the concentration of these chemicals in interstitial water or in the case of non-ionic
organic chemicals, concentrations in sediment on an organic carbon normalized basis (Di
Toro et al., 1990; Di Toro et al., 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 important. Therefore,
behavior and feeding habits of a test organism can influence its ability to accumulate
contaminants from sediment and should be considered when selecting test organisms for
sediment testing.
1.3.8.9 The use of A. abdita, E. estuarius* and R. abronius in laboratory toxicity studies
has been field validated with natural populations of benthic organisms (Swartz et al..
1994 for E estuarius: Swartz et al., 1982 for R. abronius). While no laboratory
information is available, a review of the distribution of L. plumulosus in Chesapeake Bay
indicates that its distribution is negatively correlated with the degree of sediment
contamination (Pfitzenmeyer, 1975: Reinharz, 1981).
1.3.8.9.1 Data from I'SEPA Office of Research and Development's Environmental
Monitoring and Assessment program were examined to evaluate the relationship
between survival of Ampelisca abdita in sediment toxicity tests and the presence of
amphipods. particularly ampeliscids, in field samples. Over 200 sediment samples from
two years of sampling in the Virginian Province (Cape Cod, MA, to Cape Henry, VA)
14
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were available for comparing synchronous measurements of A. abdita survival in to\icit>
tests to henthic community enumeration. Although species of this genus were union^ tlie
more frequently occurring taxa in these samples, ampeliscids were totally absent from
stations that exhibited A. abdita test survival <60*~r of that in control samples.
Additionally, ampeliscids were found in very low densities at stations with amphipod test
survival between 60 and 80"5- i.,J. Scott. SAIC, Narraganseit. Ri, personal
communication). These data indicate that tests with this species are predictive of
contaminant effects on sensitive species under natural conditions.
1.3.8.9.2 Swartz et al.(1982) compared sensitivity of R. abronius to sediment collected
from sites in Commencement Bay, W A, to benthic community structure at each site.
Mortality of R. abronius was negatively correlated with amphipod density, and
phovocephalid amphipods were ubiquitously absent from the most contaminated areas.
Schlekat et al., (1994) reported general good agreement between sediment tests with H
azteca and benthic community responses in the Anacostia River. Washington. DC.
1.3.8.9.3 Sediment toxicity to amphipods in 10-d toxicity tests, field contamination, and
field abundance of benthic amphipods were examined along a sediment contamination
gradient of DDT (Swartz et al.. 1994). Survival of £. estuarius and R. abronius in
laboratory toxicity tests was positively correlated 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 ug DDT (-t-metabolites)/g organic carbon. The threshold for abundance of
amphipods in the field was about 100 ug DDT (+metabolites)/g organic carbon.
Therefore, correlations between toxicity, contamination, and biology indicate that acute
sediment toxicity tests can provide reliable evidence of biologically adverse sediment
contamination in the field.
1.4 Performance-based Criteria
1.4.1 ISEPA's Environmental Monitoring Management Council (EMMC) recommended
the use of performance-based methods in developing chemical analytical standards
(Williams. 1993). Performance-based methods were defined by EMMC as a monitoring
approach which permits the use of appropriate methods that meet pre-established
demonstrated performance standards (Section 9.2).
1.4.2 The key consideration for methods used to obtain test organisms, whether the
they are field-collected or obtained from culture, is having healthy organisms of known
quality. A performance-based criteria approach was selected as the preferred method
through which individual laboratories should evaluate culture methods or the quality of
field-collected organisms rather than by control-based criteria. This method was chosen
to allow each laboratory to optimize culture methods, determine the quality field-
collected organisms, and minimize effects of test organism health on the reliability and
comparability of test results. See Table 11.3 for a listing of performance criteria used to
assess the quality of cultured (i.e., L, plumulosus) and field-collected amphipods. and to
determine the acceptability of 10-d sediment toxicity tests.
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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 a laboratory method for determining the short-term
toxicity of contaminated whole sediments using marine and estuarine amphipod
crustaceans. Test sediments may be collected from estuarine or marine environments
or spiked with compounds in the laboratory. A single test method is outlined that ma>
be used with any of four amphipod species, including Ampelisca abdita, Eohaustorius
estuarius, Leptocheirus plumulosus, and Rhepoxynius abronius. The toxicity test is
conducted for 1(1 d in 1 L glass chambers containing 175 mL of sediment and 800 mL of
overlying water. Overlying water is not renewed, and test organisms are not fed during
the tovicity tests. Temperature and salinity of overlying water, and choice of control
sediment li.e., negative control), are species-specific. The choice of reference sediment
may be species-specific under certain applications. The endpoint in the toxicity test is
survival, and reburial of surviving am phi pods is an additional measurement that can be
used as an endpoint. Procedures are described for use with sediments from oligohaline
to fully marine environments.
2.1.2 Experimental Design. The following section is a general summary of experimental
design. See Section 12 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 sediment is a sediment that is essentially free of contaminants 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 non-
point 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 physico-chemical characteristics such as
sediment texture may influence the response of test organisms (DeWitt et al., 1988). The
physico-chemical characteristics of test sediment 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
evaluated if the limits are exceeded in a test sediment. See Section 10.1 for tolerance
limits of each species for physico-chemical characteristics. If the physico-chemical
charucteristic(s) of a test sediment exceed the tolerance limits of the test organism, it
16
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may be desirable to include a control sediment that encompasses those characteristics, or
to choose a test organism with tolerance limits that are not exceeded b> the phvsico-
chemical characteristics in question. The effects of some sediment characteristics on the
results of sediment tests may be able to be addressed with regression equations i l)e\\itt
et al.. 1988: Ankley et al., 1994).
2.1.2.2 The experimental design depends on the purpose of the study. Variables that
need to be considered include the number and type of control sediments, the number of
treatments and replicates, and water quality characteristics. For instance, the purpose
of the study might be to determine a specific endpoint such as an LC50 and mav include
a control sediment, a positive control, and several concentrations of sediment spiked
with a chemical. A useful summary of field sampling design is presented b\ Green
(1979). See Section 12 for additional guidance on experimental design and statistics.
2.1.2.3 If the purpose of the study is to conduct a reconnaissance field survev to identifv
contaminated sites for further investigation, the experimental design might include onlv
one sample from each site to allow for maximum spatial coverage. The lack of
replication at a site usually precludes statistical comparisons (e.g., ANOYA), but these
surveys can be used to identify contaminated sites for further study or mav be evaluated
using regression techniques (Sokal and Rohlf, 1981; Steel and Torrie. 1980).
2.1.2.4 In other instances, the purpose of the study might be to conduct a quantitative
sediment survey to determine statistically significant differences between effects among
control and test sediments from several sites. The number of replicates per site should
be based on the need for sensitivity or power (Section 12). In a quantitative survev,
replicates (separate samples from different grabs collected at the same site) would need
to be taken at each site. Chemical and physical characteristics of each of these grabs
would be required for sediment testing. Separate subsamples might be used to
determine within-sample variability or to compare test procedures (e.g., comparative
sensitivity among test organisms), but these subsamples cannot be considered to be true
field replicates for statistical comparisons among sites (ASTM. 1993b).
2.1.2.5 Sediments often exhibit high spatial and temporal variability (Stemmer et al..
1990a). Therefore, replicate samples may need to be collected to determine variance in
sediment characteristics. Sediment should be collected with as little disruption as
possible; however, subsampling, compositing, or homogenization of sediment samples
may be necessary for some experimental designs.
2.1.2.6 Site locations might be distributed along a known pollution gradient, in relation
to the boundary of a disposal site, or at sites identified as being contaminated in a
reconnaissance survey. Both spatial and temporal comparisons can be made. In pre-
dredging studies, a sampling design can be prepared to assess the contamination 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 experimental test design, and statistical
analysis of the data, is the experimental unit. The experimental unit is defined as the
17
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smallest physical entity to which treatments can be independently assigned (Steei and
Torrie. 1980) and to which air exchange between test chambers are kept to a minimum.
\s the number of test chambers per 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, decreases, and the power of a significance test increases
(Section 12). Because of factors that might affect results within test chambers and
results of a test, all test chambers should be treated as similarly as possible. Treatments
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 A toxiciry method is outlined for four species of estuarine and marine amphipod,
including Ampelisca abdita, Eohaustorius estuarius, Leptocheirus plumulosus, and
Rhepoxynius abronius (Section 11). The manual describes procedures for testing
sediments from oligohaline to fully marine environments.
2.3 Test Endpoints
2.3.1 The primary endpoint measured in the toxicity test is survival. Reburial of
surviving amphipods in control sediment is an additional measurement that can be used
as an endpoint. Behavior of test organisms should be qualitatively observed daily in all
tests (e.g., avoidance of sediment).
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Section 3
Definitions
3.1 Terms
The following terms were defined in Lee (1980), NRC (1989). I SEPA (1989h>. I'SEPA-
L'SCOE (1991), I SEPA-LSCOE (1994), Lee et al. (1994), ASTM <1994a). or ASTM.
i!993b).
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 which 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 essentially free of contaminants and is used
routinely to assess the acceptability of a test. Any contaminants in control sediment ma>
originate from the global spread of pollutants and does not reflect any substantial input
from local or non-point 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 sediment 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 uould
represent the background conditions 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 evaluations.
3.1.1.6 Interstitial water or pore water. Water occupying space between sediment or soil
particles.
3.1.1.7 Spiked sediment. A sediment to which a material has been added for
experimental purposes.
3.1.1.8 Reference-toxicity test. A test conducted in conjunction with sediment 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.
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3.1.1.M 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 sediment 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 AY) observable Effect Concentration (NOEC). The highest concentration of a
toxicant to which organisms 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
\alue 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 which causes an adverse effect on the
test organisms (i.e.. where the value for the observed response is statistically significant
different from the controls).
3.1.1.14 Lethal concentration (LC). The toxicant concentration 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 concentration that would cause an
effect in a given percent of the test population. Identical to LC when the observable
adverse effect is death. For example, the EC50 is the concentration of toxicant that
would cause death in 50% of the test population.
3.1.1.1 ft Inhibition concentration (1C), The toxicant concentration that would cause a
given percent reduction in a non-quantal measurement for the test population. 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.
3.1.2 Grammatical Terms
3.1.2.1 The words "must", "should", "may", "can", and "might" have very specific
meanings in this manual.
3.1.2.2 "Must" is used to express an absolute requirement, that is, to state that a test
ought to be designed to satisfy the specified conditions, unless the purpose of the test
requires a different design. "Must" is only used in connection with the factors that
directly relate to the acceptability of a test.
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3.1.2.3 "Should" is used to state that the specified condition is recommended and ought
to be met if possible. Although a violation of one "should" is rarely a serious matter,
violation of several will often render the results questionable.
3.1.2.4 Terms such as "is desirable." "is often desirable," and "might be desirable" are
used in connection with less important factors.
3.1.2.5 "May" is used to mean "is (are) allowed to," "can" is used to mean "is (are)
able to," and "might" is used to mean "could possibly." Thus, the classic distinction
between "may" and "can" is preserved, and "might" is never used as a synonym for
either "mav" or "can."
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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 is observed when contaminants
are present at elevated concentrations, or there is less toxicity than expected.
4.1.2 There are three categories of interfering factors: those characteristics of
sediments affecting survival independent of chemical concentration (i.e., non-
contaminant factors); changes in chemical bioavailability 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 sediments, 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 physical, chemical, or biological characteristics of the sediment. Maintaining the
integrity of a field-collected sediment during removal, transport, mixing, storage, and
testing is extremely difficult and may complicate the interpretation of effects. Direct
comparisons of organisms exposed in the laboratory and in the field would be useful to
verify laboratory results. However, spiked sediment may not be representative of
contaminated sediment 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 responses of organisms.
4.1.3.1 Laboratory sediment testing with field-collected sediments may be useful in
estimating cumulative effects and interactions of multiple contaminants in a sample.
Tests with field samples usually cannot discriminate between effects of individual
chemicals. Most sediment samples contain a complex matrix of inorganic and organic
contaminants with many unidentified compounds. 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). Sediment spiking can also be used to
investigate additive, antagonistic, or synergistic effects of specific contaminant mixtures
in a sediment sample (Swartz et al.. 1988).
4.1.4 Methods which 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., <1() d: I'SEPA-
22
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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 bioavailability.
• Spiked sediment may not be representative of field contaminated 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 ecological effects.
Note: Modified from Swartz (1989).
23
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ISCOE. 1977; ISEPA-LSCOE, 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 procedures and culturing of
infaunal organisms with a variety of feeding habits including suspension and deposit
feeders is needed.
4.1.5 Despite the interferences discussed in this section, existing sediment testing
methods can be used to provide a rapid and direct measure of effects of contaminants
on benthic communities. Laboratory tests with field-collected sediment can also be used
to determine temporal, horizontal, or vertical distribution of contaminants 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 regulatory decision-making (e.g.,
I'SEPA, 1986a). Furthermore, sediment tests with complex contaminant mixtures are
important tools for making decisions about the extent of remedial action for
contaminated aquatic sites and for evaluating the success of remediation 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
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4.3 Changes in Bioavailability
4.3.1 Sediment toxicity tests are meant to serve as an indicator of contaminant-related
toxicity that might be expected under field or natural conditions. Although the tests are
not designed to simulate natural conditions, there is concern that contaminant
availability in laboratory toxicity test is different from \vhat 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 characteristics 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 per particle system. Similarly, oxidation of anaerobic sediments increases
the availability of certain metals (Di Toro et al.. 1990). Because the availability of
contaminants may be a function of the degree of manipulation, this manual recommends
that handling, storage, and preparation of the sediment for actual testings be as
consistent as possible. Although very disruptive of natural sediment physical features,
all test sediments should be press-sieved sometime before testing and re-homogenized
immediately before introduction to the test chambers if warranted (See Section 8.3.1).
Press-sieving is performed primarily to remove predatory organisms, large debris, or
organisms taxonomically similar to the test species. Certain I'SEPA program offices
may recommend that sediments should not be press-sieved. Also, it may not be
necessary to press-sieve sediments if previous experience has demonstrated the absence
of potential interferences, including predatory or competitive organisms or large debris,
or if large debris or predators can be removed with forceps or other suitable tools. The
presence of an abundance of amphipods that are tavonomically similar to the test species
should prompt press-sieving. This is particularly true if endemic Ampeliscidae are
present and A. abdita is the test species because it may be difficult to remove all of the
resident amphipods from their tubes. If sediments must be sieved, it may be desirable to
perform select analyses (e.g., pore-water metals or DOC, AYS, TOO on samples before
and after sieving to document the influence of sieving on sediment chemistry.
4.3.3 Testing sediments at temperatures different from that in the field might affect
contaminant solubility, partitioning coefficients, or other physical and chemical
characteristics. Interaction between sediment and overlying water and the ratio of
sediment to overlying water may influence bioavailability (Stemmer et al.. 1990bi.
4.3.4 Depletion of aqueous and sediment-sorbed contaminants 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 organism, 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). Inter-
particle diffusion or advection and the quality and quantity of sediment organic carbon
can also affect sorption kinetics.
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4.3.5 The route of exposure may be uncertain and data from sediment tests may be
difficult to interpret if factors controlling the bioavailability of contaminants in sediment
are unknown. Bulk-sediment chemical concentrations may be normalized to factors
other than dry weight. For example, concentrations of non-ionic organic compounds
might be normalized to sediment organic-carbon content (LSEPA, 1992c) and certain
metals normalized to acid volatile sulfides (Di Toro et al., 1990). Even with the
appropriate normalizing factors, determination of toxic effects from ingestion of
sediment or from dissolved chemicals in the interstitial water can still be difficult
(Lamberson and Swartz, 1988).
4.3.6 Salinity of the overlying water is an additional factor that can affect the
bioavailability of metals. Some metals (e.g., cadmium) are more bioavailable at lower
salinities. Therefore, if a sediment sample from a low salinity location is tested with
overlying waters of high salinity, there is the potential that metal toxicity may be
reduced. The suite of species provided in this manual allow these tests to be conducted
over the range of pore water salinities routinely encountered in field-collected sediments
from North American estuarine and marine environments.
4.4 Presence of Indigenous Organisms
4.4.1 Indigenous organisms may be present in field-collected sediments. An abundance
in the sediment sample of the test organism, or organisms taxonomically similar to the
test organism, may make interpretation of treatment effects difficult. The presence of
predatory organisms can also adversely affect test organism survival. For example,
Redmond and Scott (1989) showed that the polychaete Nephtys incisa will consume
Ampelisca abdita under toxicity test conditions. Previous investigators have inhibited the
biological activity of sediment with sieving, heat, mercuric chloride, antibiotics, or
gamma irradiation (Day et al., 1992). Although further research is needed to determine
effects on contaminant bioavailability from treating sediment to remove or destroy
indigenous organisms, estuarine and marine sediments must be press-sieved before the
start of a sediment toxicity test if the presence of predatory organisms is suspected (See
Section 8.3.1.1).
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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 preparation of a formal
written, health and safety plan, which is provided to each laboratory staff member, (3)
an ongoing training program on laboratory safety, and (4) regular safety inspections.
5.1.2 This manual addresses procedures which 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 responsibility 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 the manual, it is beyond
the scope of the manual to encompass all safety requirements necessary to conduct
sediment tests.
5.1.3 Collection and use of sediments may involve substantial risks to personal safety
and health. Contaminants in field-collected sediment may include carcinogens.
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
respirators as appropriate, (2) manipulating sediments under a ventilated hood or in an
enclosed glove box, and (3) enclosing and ventilating the exposure system. Personnel
collecting sediment samples and conducting tests should take all safety precautions
necessary for the prevention of bodily injury and illness which might result from
ingestion or invasion of infectious agents, inhalation or absorption of corrosive or toxic
substances through skin contact, and asphyxiation because of lack of oxygen or presence
of noxious gases.
5.1.4 Before sample collection and laboratory work, personnel should determine that all
required safety equipment and materials have been obtained and are in good condition.
5.2 Safety Equipment
5.2.1 Personal Safety Gear
5.2.1.1 Personnel should use safety equipment, such as rubber aprons, laboratory coats.
respirators, gloves, safety glasses, face shields, hard hats, and safety shoes as
appropriate. The degree of protection should vary according to the level contamination
associated with the test sediments. Generally, a higher degree of coverage should be
adopted in all aspects of testing sediments that may harbor hazardous levels of
27
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compounds. Coverage for testing control or moderately contaminated sediment does not
have to be as stringent.
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 All 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 Laboratory personnel should be trained in proper practices for handling and using
chemicals that are encountered during the procedures described in this manual.
Routinely encountered chemicals include acids and organic solvents. Special handling
and precautionary guidance in Material Safety Data Sheets should be followed for
reagents and other chemicals purchased from supply houses. All containers should be
adequately labeled to indicate their contents.
5.3.2 Work with some sediments may require compliance 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 bactericidal 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 fumes 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.
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.
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5.4 Disease Prevention
5.4.1 Personnel handling samples which are known or suspected to contain human
wastes should be given the opportunity to he immunized against hepatitis B. tetanus.
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 LSEPA (I986b) 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 identification
rules and land disposal restrictions, and to protect the air, water and land by
minimizing and controlling all releases from fume hods and bench operations. Also,
compliance is require with any sewage discharge permits and regulations. For further
information on waste management, consult "The Waste Management Manual for
Laboratory Personnel" available from the American Chemical Society's Department of
Government Relations and Science Policy, 1155 16th Street N.W., Washington, D.C.
20036.
5.6.2 Guidelines for the handling and disposal of hazardous materials should be strictly
followed. The Federal Government has published regulations for the management 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.
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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-tovicant" test with each potential test species, in which all test chambers contain a
control sediment (sometimes called the negative control), and clean overlying water for
each amphipod species to be tested. Survival of the test organism will demonstrate
whether facilities, water, control sediment, and handling techniques are adequate to
achieve acceptable species-specific control survival. Evaluations may also be conducted
of the magnitude of the within- and between-chamber variance in a test.
6.2 Facilities
6.2.1 The facility should include separate areas for culturing and testing to reduce the
possibility of contamination by test materials and other substances, especially volatile
compounds. Holding, acclimation, and culture chambers should not be in a room in
which sediment tests are conducted, stock solutions or where sediments are prepared, or
equipment is cleaned. Test chambers may be placed in a temperature controlled
recirculating water bath, environmental chamber, or equivalent facility with
temperature control. Enclosure of the test systems is desirable to provide ventilation
during tests to limit exposure of laboratory personnel to volatile substances.
6.2.2 Light of the quality and illuminance normally obtained in the laboratory is
adequate (about 5
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filters (e.g., BALSTOV, C-l filter), or equivalent. Laboratory ventilation systems
should be checked to ensure that return air from chemistry laboratories or sample
handling areas is not circulated to culture or testing rooms, or that air from testing
rooms does not contaminate culture rooms. Air pressure differentials between rooms
should not result in a net flow of potentially contaminated air to sensitive areas through
open or loosely-fitting doors.
6.3 Equipment and Supplies
6.3.1 Equipment and supplies that contact stock solutions, 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 fluorocarbon
plastics should be used whenever possible to minimize leaching, dissolution, and
sorption. High-density plastic containers are recommended for holding, acclimation, and
culture chambers. These materials should be washed in detergent, acid rinsed, and
soaked in flowing water for a week or more before use. 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 mentioned
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.
6.33 General Equipment
6.3.3.1 Environmental chamber or equivalent facility with photoperiod and temperature
control (5 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 weighing 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 which are at the
upper and lower ends of the range of the weight values 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
non-toxic plastic labware. 10 to 1000 mL for making test solutions.
6.3.3.6 Volumetric pipets -- Class A. 1 to 100 mL.
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6.3.3.7 Serological pipets -- 1 to 10 mL, graduated.
6.3.3.8 Pipet bulbs and fillers -- PROPIPET\ or equhalent.
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 measuring water temperature.
6.3.3.12 National Bureau of Standards Certified thermometer (see L'SEPA Method
170.1; L'SEPA. 1979b).
6.3.3.13 Dissolved oxygen, pH/selective ion, and salinity meters for routine physical and
chemical measurements. Unless a test is being conducted to specifically measure the
effect of one of these measurements, a portable field-grade instrument is acceptable. A
temperature compensated salinity refractometer is useful for measuring salinity of water
overlying field collected sediment.
6.3.3.14 Equipment for measuring ammonia (i.e.. ammonia-specific probe) is also
necessary.
6.3.3.15 See Table 6.1 for a list of additional equipment and supplies.
6.3.4 Test Chambers
6.3.4.1 The test chambers to be used in sediment toxicity tests are 1 liter glass
containers (beakers or wide-mouthed jars) with an internal diameter of 10 cm. Each
test chamber should have a cover. Acceptable test chamber covers include watch-
glasses, plastic lids, and 9 cm diameter glass culture dishes. It may be necessary to drill
a hole in each cover to allow for the insertion of a pipette for aeration.
6.3.5 Cleaning
6.3.5.1 All non-disposable sample containers, test chambers, tanks, and other equipment
that has 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. CarefulU rinse once with fresh, dilute (10'Y, V :V) hydrochloric or nitric acid
to remove scale, metals, and bases. To prepare a lO^r solution of acid, add 10
mL of concentrated acid to 90 mL of deionized water.
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4. Rinse twice with deionized water.
5. Rinse once with full-strength, pesticide-grade acetone to remove organic
compounds (use a fume hood or canopy).
6. Rinse three times with deionized water.
6.3.5.2 All test chambers and equipment should be thoroughly rinsed or soaked with the
toxicity test diluent water immediately before use in a test.
6.3.5.3 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, 1988), 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.
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Table 6.1 Equipment and supplier >r culturing and testing estuarine and
marine amphipods. Su| ?s are for all species unless specified.
A. Biological Supplies
Brood stock of test organisms
TetraMin® (LP)
dried wheat leaves (LP)
dried alfalfa leaves (LP)
Neo-Novum® (LP)
Algae (e.g., Pseudoisochrysis paradoxa and Phaeodactylum tricornutum
[optional]) (LP)
B. Glassware
Culture chambers (30 cm x 45 cm x 15 cm plastic wash bin)
Test chambers (1 L glass jar or beaker)
Glass bowls
Wide-bore pi pets (4 to 6 mm ID)
Glass disposable pipets
Graduated cylinders (assorted sizes, 10 mL to 4 L)
C. Instruments and Equipment
Dissecting microscope
Stainless-steel sieves (e.g., U.S. Standard No. 25, 30, 35, 40, 50 mesh)
Photoperiod timers
Light meter
Temperature controllers
Thermometer
Continuous recording thermometer
Photoperiod tinier
Dissolved oxygen meter
pH meter
Selective ion meter
Ammonia electrode (or ammonia kit)
Salinity meter/temperature compensating salinity refractometer
Drying oven
Desiccator
Balance (0.01 mg sensitivity)
Refrigerator
Freezer
Note: LP = Leptocheirus plumulosus
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Table 6.1 Equipment and supplies for culturing and testing estuarine and
marine amphipods. Supplies are for aJl species unless specified
(continued)
Light box
Hemacy to meter
Mortar and pestle or blender (LP)
D. Miscellaneous
Ventilation system for test chambers
Air supply and air stones (oil free and regulated)
Glass hole-cutting bits
Glass glue
Aluminum weighing pans
Fluorescent light bulbs
Deionized water
Air line tubing
White plastic dish pan
Water squirt bottles
Shallow pans (plastic (light-colored), glass, stainless steel)
Sieve cups (mesh size <0.5 mm)
Dissecting probes
E. Chemicals
Detergent (non-phosphate)
Acetone (reagent grade)
Hexane (reagent grade)
Hydrochloric acid (reagent grade)
Reagents for preparing synthetic seawater (reagent grade Cad, »2 H2O, KBr. KCI.
MgCl, • 6 H2O, Na2B4O7»10 H:O, NaCI, Na HCO3. Na,SO4, SrCI, ^HX))
Formalin
Ethanol
Rose bengal
Cadmium chloride
Sodium dodecyl sulfate
Copper sulfate
Note: LP = Leptocheirus plumulosus.
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Section 7
Water, Reagents, and Standards
7.1 Water
7.1.1 Requirements
7.1.1.1 Sea water used to test and culture organisms should be uniform in quality.
Acceptable sea 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 I SEPA
(1993a) and ASTM (1994a) for a recommended list of chemical analyses of the water
supply.
7.1.2 Source
7.1.2.1 The source of natural water will depend to some extent on the objective of the
test and the test organism that is being used. All natural waters should be obtained
from an uncontaminated surface-water source upstream from or beyond the influence of
known discharges. Water should be collected at slack high tide, or within one h after
high tide. Suitable surface water sources should have intakes that are positioned to: (1)
minimize fluctuations in quality and contamination, (2) maximize the concentration of
dissolved oxygen, and (3) ensure low concentrations of sulfide and iron. Full strength
sea water should be obtained from areas where the salinity does not fall below 28 r«.
For estuarine tests, water having a salinity as near as possible to the desired test salinity
should be collected from an uncontaminated area. Alternatively, it may be desirable to
dilute full strength sea water with an appropriate fresh water source. Sources of fresh
water (i.e., 0 "r(] for dilution include deionized water, uncontaminated well or spring
water, or an uncontaminated surface-water source. Municipal-water supplies may be
variable and may contain unacceptably high concentrations of materials such as copper.
lead, zinc, fluoride, chlorine, or chlorainines. Chlorinated water should not be used to
dilute water utilized for culturing or testing because residual chlorine and chlorine-
produced oxidants are toxic to many aquatic organisms. Dechlorinated water should
only be used as a last resort for diluting sea water to the desired salinity since
dechlorination is often incomplete (ASTM, 1994a; I SEPA, 1993a).
7.1.2.2 For site-specific investigations, it is desirable to have the water-quality
characteristics of the overlying water (i.e.. salinity) as similar as possible to the site
water. For certain applications the experimental design might require use of water from
the site where sediment is collected. In estuarine systems, however, the pore water
salinity of sediments may not be the same as the overlying water at the time of collection
(Sanders et al., 1965).
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7.1.2.3 Water that might be contaminated with facultative pathogens may be passed
through a properly maintained ultraviolet sterilizer equipped with an intensity meter
and flow controls or passed through a filter with a pore size of 0.45 urn or less.
7.1.2.4 Natural sea water might need aeration using air stones, surface aerators, or
column aerators. Adequate aeration will stabilize pH, bring concentrations of dissolved
oxygen and other gases into equilibrium with air, and minimize oxygen demand and
concentrations of volatiles. The concentration of dissolved oxygen in source water
should be between 90 to 100% saturation to help ensure that dissolved oxygen
concentrations are acceptable in test chambers. Natural sea water used for
holding/acclimating, culturing, and testing amphipods should be filtered <<5 um) shortly
before use to remove suspended particles and organisms.
7.1.2.5 Water that is prepared from natural sea water should be stored in clean.
covered containers at 4°C and used within 2 d.
7.13 Reconstituted/Synthetic Seawater
7.1.3.1 Although reconstituted water is acceptable, natural seawater is preferable,
especially for tests involving chemicals whose bioavailability is affected by seawater
chemistry. Reconstituted water is prepared by adding specified amounts of reagent-
grade chemicals to high-purity distilled or deionized water (ASTM, 1988: LSEPA,
1993a). Acceptable high-purity water can be prepared using deionization, distillation, or
reverse-osmosis units (Section 6.3.3.2; LSEPA, 1993a). Test water can also 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 water. If large quantities of high quality deionized water are needed, it
may be advisable to supply the laboratory grade water deionizer with preconditioned
water from a mixed-bed water treatment system.
7.1.3.3 Reconstituted sea water is prepared by adding specified amounts of a suitable
salt reagent to high-purity distilled or deionized water (ASTM, 1988: LSEPA. 199la).
Suitable salt reagents can be reagent grade chemicals, commercial sea salts, such as
Forty Fathoms®, Instant Ocean®, or HW Marinemix£. Pre-formulated brine (e.g., 60 to
90 %r), prepared with dry ocean salts or heat-concentrated natural sea water, can also
be used.
7.1.3.4 A synthetic sea formulation called GP2 is prepared with reagent grade chemicals
that can be diluted with a suitable high-quality water to the desired salinity (I SEPA.
1994c).
7.1.3.5 To obtain the desired holding or acclimation salinity, sea salts or brine can be
added to a suitable freshwater or distilled water, or the laboratory's sea water suppl>
mav be diluted with a suitable freshwater or distilled water.
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7.1.3.6 The suitability and consistency of a particular salt formulation for use in holding
and acclimation should be verified by laboratory tests because some formulations can
produce unwanted toxic effects or sequester contaminants (Environment Canada. 1992).
Salinity and pH should be measured on each batch of reconstituted water.
7.1.3.7 Salinity, pH, and dissolved oxygen 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). Reconstituted sea
water should be filtered <<5 urn) shortly before use to remove suspended particles and
should be used within 24 h of filtration. USEPA (1993a) recommends using a batch of
reconstituted water within a two week period.
7.2 Reagents
7.2.1 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.
7.3 Standards
7.3.1 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.
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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 A benthic grab (i.e., PONAR, Smith-Maclntyre, Van Veen) or core sampler are
preferred sediment samplers because disturbance of sediment samples with these devices
is minimized relative to dredge samplers. Although selective sub-sampling, compositing.
and homogenization of sediment samples are necessary for most routine applications
addressed by this manual, collection and handling in the field should involve as little
disruption as possible. Disruption of sediment samples will cause the loss of sediment
integrity, and may cause changes in chemical speciation and chemical equilibrium
(ASTM, 1994a). Sediments are spatially and temporally variable (Stemmer et al..
1990a). Replicate samples should be collected to determine variance in sediment
characteristics. Sediments should be collected to a depth appropriate for the study
objectives. For example, 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 photolytic compounds. Removal of sediment from the sampling
device and subsequent allocation to storage containers or homogenization should be
accomplished using spoons, trowls, etc. made of, or coated in, inert materials (e.g..
Teflon®, kynar). Sediment samples should be cooled to 4CC in the field before return to
the laboratory or 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
System, Sensitech, Inc., Beverly, MA).
8.1.4 For additional information on sediment collection and shipment see ASTM
(1994a).
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 substantial!)
affect toxicity. Storage of sediment samples for several months at 4CC did not result in
significant changes in chemistry or toxicity (T. Dillon and H. Tatem. ISCOE. Vicksburg.
MS, personal communication: G.T. Ankley and D. Foe. I SEPA, Duluth, MM,
unpublished data); however, others have demonstrated changes in spiked sediment
within days to weeks (e.g.. Burton, 1991; Stemmer et al., 1990a). Sediments primarilv
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contaminated with non-ionic, nonvolatile organic compounds will probably change little
during storage because of their relative resistance to biodegradation and sorption to
solids. However, metals and metalloids may be affected by changing redox, oxidation, or
microbial metabolism (such as with arsenic, selenium, mercury, lead, and tin: all of
«hich are methylated by a number of bacteria and fungi). Metal contaminated
sediments may need to be tested relatively soon after collection with as little
manipulation 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 it'SEPA-
I'SCOE, 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 contaminant partitioning and should be
avoided (ASTM, 1994a; 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 headspace) at 4°C before the start of a test (Shuba et al.,
1978: ASTM, 1994a). 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.
8.3 Manipulation
8.3.1 Homogenization
8.3.1.1 Sediment samples tend to settle during shipment. As a result water above the
sediment should not be discarded, but should be mixed back into the sediment during
homogenization. If warranted, sediment samples should be press-sieved through a 1 or
2 mm mesh stainless steel screen to remove indigenous organisms. Press-sieving is
performed primarily to remove predatory organisms, large debris, or organisms
taxonomically similar to the test species. Certain I SEPA program offices may
recommend that sediments should not be press-sieved. Also, it may not be necessary to
press-sieve sediments if previous experience has demonstrated the absence of potential
interferences, including predatory or competitive organisms or large debris, or if large
debris or predators can be removed with forceps or other suitable tools. The presence
of an abundance of amphipods that are taxonomically similar to the test species should
prompt press-sieving. This is particularly true if endemic Ampeliscidae are present and
A. abdita is the test species because it may be difficult to remove all of the resident
amphipods from their tubes. If sediments must be sieved, it may be desirable to perform
select analyses (e.g.. pore-water metals or DOC, AVS. TOO on samples before and after
sieving to document the influence of sieving on sediment chemistry.
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8.3.1.2 If sediment is collected from multiple field samples, the sediment can he pooled
and mixed using stirring or a rolling mill, feed mixer, or other suitable apparatus (see
ASTM, 1994a). It is preferable to homogenize sediments by gentle hand mixing.
Although potentially disruptive, large numbers of sediments may demand the use of a
mechanical aid. Mechanical homogenization of sediment 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.-1 cm; Augers Unlimited, Exton, PA: Kemble et al.. 1994). These
procedures could also be used to mix test 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. 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 sediment. This short time period may not be long enough for sediments to
equilibrate with the spiked chemicals. Consistent 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 magnitude of interactive effects of
contaminants can be estimated by spiking a sediment with chemicals or complex waste
mixtures (Lamberson and Swartz, 1992). Sediments spiked with a range of
concentrations can be used to generate either point estimates (e.g.. LC50) or a minimum
concentration at which effects are observed (lowest observable effect concentration:
LOEC). The influence of sediment physico-chemical characteristics on chemical toxicity
can also be determined with sediment-spiking studies (Adams et al., 1985).
8.3.2.2 The test material(s) should be at least reagent grade, unless a test 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 concentration 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
disposal procedures.
8.3.2.2.1 Organic compounds have been added in the dry form or coated on the inside
walls of the mixing container (Ditsworth et al., 1990). Metals are general!) added in an
aqueous solution (ASTM. 1994a: Carlson et al., 1991; Di Toro et al.. 1990). If an
organic solvent is used, the solvent in the sediment should be at a concentration that
does not affect the test organisms. Concentrations of the chemical in the pore water and
in whole sediment should be monitored at the beginning and end of a test.
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8.3.2.3 I se 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 in the sediment should be at a concentration
that does not affect the test organism. The solvent control must contain the highest
concentration of solvent present and must be from the same batch used to make the
stock solution (see ASTM, 1988). 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 sediment 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 statistically
significant difference is detected between the two controls, only the solvent control may
be used for meeting the acceptability of the test and as the basis for calculating results.
The negative control might provide additional information on the general health of the
organisms tested. If no statistically significant difference is detected, the data from both
controls should be used for meeting the acceptability of the test and as the basis for
calculating the results (ASTM, 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 predicted 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 prediction 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 concentrations 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 non-polar organic compounds might be
normalized to sediment organic-carbon content and simultaneously extracted metals
might be normalized to acid volatile sulfides (Di Toro et al. 1990; Di Toro et al. 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.
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8.3.2.6 Addition of test materials) to sediment may be accomplished using various
methods, such as a: (1) rolling mill (preferred), (2) feed mixer, or (3) hand mixing
(ASTM, 1994a). 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. 1994a). 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 subsarnples is advisable to determine the degree of mixing homogeneity
(Ditsworth et al., 1990). Moreover, results from sediment-spiking studies should be
compared with the response of test organisms to chemical concentrations in natural
sediments (Lamberson and Swartz, 1988).
8.4 Characterization
8.4.1 All sediments should be characterized and at least the following determined:
salinity, pH. and ammonia of the pore water, organic carbon content (total organic
carbon, TOO, particle size distribution (percent sand, silt, clay), and percent water
content (ASTM, 1994a; Plumb, 1981). Salinity of sediment pore water should be
measured on the supernatant of an aliquot of the sediment using a refractometer or
conductivity meter. See Section 8.4.4.7 for methods to isolate pore water.
8.4.2 Other analyses on sediments might include: biological oxygen demand, chemical
oxygen demand, cation exchange capacity. Eh, total inorganic carbon, total volatile
solids, acid volatile sulfides, metals, synthetic organic compounds, oil and grease.
petroleum hydrocarbons, as well as interstitial water analyses for varous physico-
chemical parameters.
8.4.3 Macrobenthos may be quantified 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 descriptions of the sediment may include color, texture, presence of hydrogen
sulfide, and presence of indigenous organisms. Monitoring the odor of sediment samples
should be avoided because of potential hazardous volatile contaminants. It may be
desirable to describe color and texture gradients that occur with sediment depth.
8.4.4 Analytical Methodology
8.4.4.1 Chemical and physical data should be obtained using appropriate standard
methods whenever possible. For those measurements for which standard methods do
not exist or are not sensitive enough, methods should be obtained from other reliable
sources.
8.4.4.2 The precision, accuracy, and bias of each analytical method used should be
determined in the appropriate matrix: that is. sediment, water, tissue. Reagent blanks
and analytical standards should be analyzed and recoveries should be calculated.
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8.4.4.3 Concentration of spiked test material(s) in sediment, 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 desirable.
8.4.4.4 Separate chambers should be set up at the start of a test and destructively
sampled during and at the end of the test to monitor sediment chemistry. Test
organisms might be added to these extra chambers depending 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 contain any surface debris, any material
from the sides of the test chamber, or any sediment
8.4.4.6 Measurement of test material(s) concentration in sediment at the end of a test
can be taken by siphoning most of the overlying water without disturbing the surface of
the sediment, then removing appropriate aliquots of the sediment for chemical analysis.
8.4.4.7 A variety of procedures have been used to isolate interstitial water including
centrifugation, filtration, pressure, or by using an interstitial water sampler; however,
centrifugation without filtration is the recommended procedure (Ankley and Schubauer-
Berigan, 1994). Filtration may reduce concentrations of materials in interstitial water
(Schults et al., 1992). Care should be taken to ensure that contaminants do not
transform, degrade, or volatilize during isolation or storage of the 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 commitment by laboratory management and also includes the
following: (1) appointment of a laboratory quality assurance officer with the
responsibility and authority to develop and maintain a QA program. (2) preparation of a
Quality Assurance Project Plan with Data Quality Objectives, (3) preparation of written
descriptions of laboratory 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 LSEPA (1989c).
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. 14) test
conditions, (5) instrument calibration, (6) replication, (7) use of reference toxicants, 18)
record keeping, and (9) data evaluation.
9.1.3 Quality Control (QC) practices, on the other hand, consist of the more focused,
routine, day-to-day activities carried out within the scope of the overall QA program.
For more detailed discussion of quality assurance, and general guidance on good
laboratory practices related to testing see FDA (1978), LSEPA (1979a), LSEPA < 1980a).
LSEPA (1980b), L'SEPA (1993a). L'SEPA (1994b). I SEPA (1994c). DeVVoskin (1984).
and Taylor (1987).
9.2 Performance-based Criteria
9.2.1 LSEPA 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 which permits the use of appropriate 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 performance
standards.
9.2.2 Therefore, a performance-based criteria approach was selected as the preferred
method through which individual laboratories should evaluate culture methods or the
quality of field-collected organisms rather than by control-based criteria. This method
was chosen to allow each laboratory to optimize culture methods, determine the qualit)
of field-collected organisms, and minimize effects of test organism health on the
reliability and comparability of test results. See Table 11.3 for a listing of performance
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criteria for culturing L. plumulosus, determining the quality of field-collected organisms,
and evaluating the outcome of sediment tests.
9.3 Facilities, Equipment, and Test Chambers
9.3.1 Separate areaas for test organism culturing and testing areas must he provided to
avoid loss of cultures due to cross-contamination. Ventilation systems should be
designed and operated to prevent recirculation or leakage of air from chemical analysis
laboratories or sample storage and preparation areas into test organism culturing or
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 temperatures. Recommended materials should be used in the fabricating of
the test equipment which 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 overlying
water. This information is used to demonstrate that the facility, control sediment, water.
and handling procedures provide acceptable responses of test organisms (Section 9.14).
9.4 Test Organisms
9.4.1 The organisms should appear healthy, behave normally, feed well, and have low
mortality (e.g.. <15fr) in cultures, during holding, and in test controls. The species of
test organisms should be positively identified. Test organisms should not show signs of
disease or apparent stress (e.g., discoloration, unusual behavior).
9.5 Water
9.5.1 The quality of water used for organism culturing and testing is extremely
important. Overlying water used in culturing, holding, acclimation, and testing
organisms should be uniform in quality. Acceptable water should allow satisfactory
survival, growth or reproduction of the test organisms. See Section 7 for guidance on
selection and preparation of high quality test water.
9.6 Sample Collection and Storage
9.6.1 Sample holding times and temperatures should conform to conditions described in
Section 8.
9.7 Test Conditions
9.7.1 It is desirable to measure temperature continuously in at least one chamber
during the each test. Temperatures should be maintained within the limits specified for
each test. Dissolved oxygen, salinity, ammonia, and pH should be checked as prescribed
in Section 11.3.
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9.8 Quality of Test Organisms
9.8.1 If test organisms are obtained from culture, monthly reference-toxicity tests should
be conducted on all test organisms using procedures outlined in Section 9.16. 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.
9.8.2 The quality of test organisms obtained from an outside source, regardless of
whether they are from culture or collected from the field, must be verified b>
conducting a reference-toxicity test concurrently with the sediment test. For cultured
organisms, the supplier should provide data with the shipment describing the historv of
the sensitivity of organisms from the same source culture. For field-collected organisms.
the supplier should provide data with the shipment describing the collection location, the
time and date of collection, the water salinity and temperature at the rime of collection.
and collection site sediment for holding and acclimation purposes. If the supplier has
not conducted five reference toxicity tests with the test organism, it is the responsibilitv
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 identification of the test organisms.
and provide the taxonomic 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 L. plumulosus in cultures (see Section 10.5.8).
Additionally, survival in sediment tests conducted with A. abdita and L. plumulosus ina>
be affected by the nutritional suitability of food provided during holding and
acclimation.
9.10 Test Acceptability
9.10.1 For the test results to be acceptable, survival at 10 d must equal or exceed 90<'<
for all four amphipod species in the control sediment. See Table 11.3 for additional
requirements for acceptability of the tests.
9.10.2 An individual test may be conditionally acceptable if temperature, dissolved
oxygen, and other specified conditions fall outside specifications, depending on the
degree of the departure and the objectives of the tests (see Table 11.3). 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.
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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 I SEPA
methods manuals (I SEPA, 1979a: USEPA, 1979b; LSEPA, 1993d).
9.11.2 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.
9.12 Calibration and Standardization
9.12.1 Instruments used for routine measurements of chemical and physical
characteristics such as pH, dissolved oxygen, temperature, and salinity should be
calibrated before use each day according to the instrument manufacturer's procedures
as indicated in the general section on quality assurance (see L'SEPA Methods 150.1,
360.1. 170.1. and 120.1. ISEPA. 1979b). Calibration data should be recorded in a
permanent log.
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
per treatment, the significance level selected, and the type of statistical analysis. If the
variability remains constant, the sensitivity of a test will increase as the number of
replicates is increased. The minimum recommended number of replicates varies with
the objectives of the test and the statistical method used for analysis of the data
(Section 12).
9.14 Demonstrating Acceptable Performance
9.14.1 It is the responsibility of a laboratory to demonstrate 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 response 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) and same
data analysis methods. This should be done to gain experience for the toxicity tests and
as a point of reference for future testing. 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 12).
9.14.2 Before conducting tests with contaminated sediment, the laboratory should
demonstrate its abilitv to conduct tests by conducting five exposures in control sediment
48
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as outlined in Table 11.1. It is recommended that these five exposures with control
sediment be conducted concurrently with the five reference toxicity tests described in
Section 9.14.1.
9.14.3 Laboratories should demonstrate that their personnel are able to recover an
average of at least 90 ^ of the organisms from whole sediment. For example, test
organisms could be added to control 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 continuing basis is demonstrated by
conducting monthly 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, generating LCSOs using the same data
analysis method (Section 13).
9.15.2 Outliers, which are data falling outside the control limits, and trends of
increasing or decreasing sensitivity 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 credibility of the test results are suspect. In this case, the test
procedure should be examined for defects and should be repeated with a different batch
of test organisms.
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 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 Table 11.3 must be
considered when determining the acceptability of a sediment test. The acceptability of
the sediment test would depend on the experience and judgement 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 S% of the time, regardless of how well a laboratory
performs. For this reason, good laboratories which develop very narrow control limits
may be penalized if a test result which falls just outside the control limits is rejected dc
facto. The width of the control limits should be considered in decisions regarding
rejection of data (Section 13).
9.16 Reference Toxicants
9.16.1 Reference-toxicity tests should be conducted in conjunction with sediment tests to
determine possible changes in condition of a test organism (Lee, 1980). Water-only
reference-toxicity tests on cultured organisms should be conducted monthly, and should
be performed on each batch of field-collected organisms used for testing. Deviations
49
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outside an established normal 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 cadmium (available as cadmium chloride (CdCl:)),
copper (available as copper sulfate (CuSO4)), and sodium dodecyl sulflde (SDS) are
suitable for use. No one reference toxicant can be used to measure the condition of test
organisms in respect to another toxicant with a different mode of action (Lee, 1980).
Howe\er, it may be unrealistic to test more than one or two reference toxicants
routinely.
9. 16.3 Test conditions for conducting reference-toxicity tests with A. abdila, E. estuarius,
L. plumulosus, and R. abronius are outlined in Table 9.1.
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 toxicants;
detailed records of the test organisms used in the test(s), such as species, source, age,
date of receipt, and other pertinent information relating to their history and health;
information on the calibration of equipment and instruments; test conditions used; and
results of reference toxicant tests. Laboratory data should be recorded immediately to
prevent the loss of information or inadvertent introduction of errors into the record.
Original data sheets should be signed and dated by the laboratory personnel performing
the tests. For additional detail see Section 12.
50
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Table 9.1
Recommended test conditions for conducting reference-toxicitv tests
Parameter
Conditions
1. Test type:
2. Dilution series:
3. Toxicant:
4. Temperature:
5. Salinity:
6. Light quality:
7. Photoperiod:
8. Renewal of water:
9. Age and size of
test organisms:
10. Test chamber:
11. Volume of water:
12. Number of
organisms/chamber:
Water-only test
Control and at least 5 test concentrations (0.5
dilution factor)
Cd. Cu. Sodium dodecyl sulfate (SDS)
15°C for E. estuarius and R. abronius
20 C for A. abdita
25°C for L. ptumulosus
28 7d for ,4. abdita and R. abronius
20 7fi for E. estuarius and L. plumulosus
Chambers should be kept in dark or covered
with opaque material
24 h D
None
A. abdita: 3 - 5 mm (no mature males or
females)
E. estuarius: 3 - 5 mm
L. plumulosus: 2 - 4 mm (no mature males or
females)
R. abronius: 3 - 5 mm
1 L glass beaker or jar
800 niL (minimum)
n = 20 if 1 per replicate: n = 10 (minimum) if
>1 per replicate
51
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Table 9.1
Recom! nded test conditions for conducting reference-toxicity tests
(contin t)
Parameter
Conditions
13. Number of replicate
chambers/treatment:
1 minimum; 2 recommended
14. Aeration:
Recommended; but not necessary if
dissolved oxygen saturation can be achieved
without aeration
15. Dilution water:
16. Water quality:
17. Test duration:
Culture water, surface water, site water, or
reconstituted water.
Salinity, pH, and dissolved oxygen at beginning
and end of test. Temperature daily
96 h
18. Endpoint:
Survival (LC50); Reburial (EC50) optional for
E. estuarius and R. abronius
19. Test acceptability:
90% control survival
52
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Section 10
Collection, Culture, and Maintaining of Test Organisms
10.1 Life History
10.1.1 Ampelisca abdita: A. abdita is a tube-building amphipod in the family
Ampeliscidae. It occurs on the Atlantic coast from central Maine to central Florida,
although it is also found in the eastern portion of the Gulf of Mexico (Bousfield, 1973).
On the Pacific coast, it is present in San Francisco Bay, CA (Nichols et al., 1985:
Hopkins, 1986). They are small (adult length 4 to 8 mm), laterally compressed
amphipods. Healthy animals are opalescent pink and will remain tightly curled.
whereas unhealthy animals tend to be translucent white, and may uncurl (T. Thompson.
SAIC, Bothell, VVA, personal communication). Often dominant members of the henthic
community, A. abdita forms thick mats of tubes with amphipod densities up to
110,000/nr, and are often a dominant food source for bottom-feeding fish (Richards,
1963). The tubes are narrow and approximately 2 to 3 cm in length. A filter feeder,
A. abdita feeds on both particles in suspension and those from surflcial sediment
surrounding the tube. Ampelisca abdita is euryhaline, and has been reported in waters
that range in salinity from fully marine to 10 %r (Hyland, 1981). Laboratory tests have
shown the salinity application range of A. abdita in sediments is from 0 to 34 7« when
the salinity of overlying water is >28 9rr (Weisberg et al., 1992). This species generally
inhabits sediments from fine sand to mud and silt without shell fragments, although it
can also be found in relatively coarser sediments with a sizeable fine component. It is
often abundant in sediments with a high organic content. Analysis of historical data
shows little effect of sediment grain size on survival of A. abdita during 10 d sediment
toxicity tests (Long and Buchman, 1989; Weisberg et al., 1992). There is evidence that
sediments with >95% sand may elicit excessive mortality (J. Scott, SAIC. Narragansett,
Rl, personal communication). Ampelisca abdita have been collected at water
temperatures ranging from -2 to 27°C (J. Scott, SAIC, Narragansett. RI, and M.S.
Redmond, Northwest Aquatic Sciences, Newport, OR, unpublished data). Reproduction
patterns of .4. abdita vary geographically. In the colder waters of its range, .4. abdita
produces two generations per year, an over-wintering population that broods in the
spring, and a second that breeds in mid- to late-summer (Mills, 1963). In warmer
waters south of Cape Hatteras, NC, breeding might be continuous throughout the year
(Nelson, 1980). Juveniles are released after approximately two weeks in the brood
pouch. Juveniles take approximately 40 to 80 d for newly released juveniles to become
breeding adults under laboratory conditions at 20°C (Scott and Redmond, 1989).
10.1.2 Eohaustorius estuarius: Eohaustorius estuarius is a free-burrowing amphipod in
the family Haustoriidae. It is found on protected and semi-protected beaches from the
lower intertidal to shallow subridal waters exclusively on the Pacific coast from British
Columbia south to central California (Environment Canada, 1992). They are stout
(adult size range 3 to >5 mm) cup- or bell-shaped, dorsally compressed amphipods that
are grayish-brown or yellowish-brown in color (Environment Canada, 1992: ASTM.
1992). Eohaustorius estuarius are thought to be deposit feeders. It is an estuarine
species and has been reported in areas where pore water salinity ranges from 1 to 25 7rr
53
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iEnvironment Canada, 1992; R. Caldweli. Northwest Aquatic Sciences. Newport, OR.
personal communication). Laboratory studies have shown a salinity application range in
control sediments for E. estuarius from 0 to 34 c'(r. Eohaustorius estuarius inhabits
clean, medium-fine sand with some organic content. The species has exhibited
acceptable (i.e., >9()c"f) survival when exposed to clean sediments with a wide range of
grain sizes, with generally little affect on survival whether coarse-grained or fine-grained
(i.e., predominantly silt and clay) clean sediments are used (Environment Canada, 1992).
However, some correlation between survival and grain size exists (DeVVitt et al., 1989).
Fohaustorius estuarius has been collected from water temperatures from 0 to 23 C
lASTM. 1992: R. Caldweli. personal communication). Eohaustorius estuarius apparently
has an annual life cycle (Environment Canada, 1992; DeVVitt et al., 1989). Gravid
females are abundant in intertidal sediments from February through July (DeWitt et al..
1989; ASTM. 1992). However, reproduction might occur year-round because juveniles
are found throughout most of the year (DeVVitt et al., 1989).
10.1.3 Leptocheirus plumulosus: Leptocheirus plumulosus is a tube-building member of
the family Aoridae. It is found infaunally in subtidal portions of Atlantic Coast brackish
estuaries from Cape Cod, MA, to northern Florida (Bousfield. 1973; DeWitt et al.,
1992a). It is common in protected embay ments but has been collected in channels of
estuarine rivers up to depths of 10 m (Holland et al., 1988: Schlekat et al., 1992). and
has been reported to occur in depths up to 13 m (Shoemaker, 1932). They are relatively
large amphipods (adult length up to 1.3 cm) with cylindrically shaped bodies that are
brownish-grey in color. A distinguishing feature is a series of dark bands or stripes that
cross the dorsal surface of the pareons and pleons. In Chesapeake Bay, densities of
L. plumulosus can reach 28,987/m: and 24.133/m: in sandy and muddy sediments.
respectively (Holland et al., 1988). It feeds on particles that are in suspension and on
the sediment surface (DeWitt et al., 1992a). Leptocheirus plumulosus is found in both
oligohaline and mesohaline regions of east coast estuaries; ambient water salinity at
collection sites has ranged from 0 to 15 %< (Holland et el.. 1988; DeWitt et al., 1992a;
Schlekat et al., 1992; McGee et al., 1994). Laboratory studies have demonstrated a
salinity application range in control sediments of 0 to 32 "« ( Schlekat et al., 1992;
SA1C, 1993b). It is most often found in fine-grained sediment with a high proportion of
particulate organic material, although it has been collected in fine sand with some
organic content (Jordan and Sutton, 1984; Holland et al., 1988; Marsh and Tenore,
1990; DeWitt et al., 1992a; Schlekat et al.. 1992; McGee et al.. 1994). Analysis of
historical data for L. plumulosus reveals no effect of sediment grain size on survival in
control sediment Populations of L. plumulosus may be seasonally ephemeral, with
major population growth occurring in spring and large population declines occurring in
summer due to actions of predatory fish (Hines et al.. 1986) or absence of essential
micronutrients (Marsh and Tenore, 1990).
10.1.4 Rhepoxynius abronius is a free-burrowing amphipod in the family
Phoxocephalidae. It occurs on the Pacific Coast from Puget Sound, WA, to central
California in lower intertidal and nearshore subtidal zones to depths of 274 m offshore
(ASTM. 1992: Environment Canada, 1992; Lamberson and Swartz. 1988; Kemp et al.,
1985; Barnard and Barnard, 1982). Densities in the field are reported to range from
150 to 2200/nr (Lamberson and Swartz. 1988: Swartz et al.. 1985). It is a medium-sized
54
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(adult length from 3 to >5 mm) amphipod with a stout, somewhat rounded body shape.
Color may range from salmon pink to yellowish, grayish-brown to white with a pinkish-
brown hue (Environment Canada. 1992). Rhepoxynius abranius is a meiofaunal
predator, but it also ingests sedimentary organic material (Oakden, 1984). In the field.
R. abronius is found where pore water salinity is no lower than 20 7n (Environment
Canada, 1992). Laboratory tests have indicated that salinities below 25 V« ma> be toxic
to R. abronius (SA1C, 1993b; Swartz et al., 1985). Rhepoxynius abronius should
therefore not be chosen as the test species when the sediment pore water is <25 <7cr
(Swartz et al., 1985). Rhepoxynius abronius naturally inhabits clean, fine, sandy
sediments (ASTM, 1992). A number of studies have shown some reduction in survival
when this species is held in very fine-grained (predominantly silt and clay) sediment
(DeWitt et al., 1988; Long et al., 1990; McLeay et al., 1991: SAIC, 1993a; SAIC, 1993b).
Normally collected at temperatures ranging from 8 to 16°C, R. abronius has survived at
temperatures ranging from 0 to 20°C under laboratory conditions (ASTM, 1992).
Reproduction of R. abronius is annual, with peak production occurring from late winter
through spring (Kemp et al., 1985).
10.2 Species Selection
10.2.1 All four species have been routinely used to test sediments with a range of grain
size characteristics and pore water salinities. Selection of one or more of the four
species for a particular test/investigation should take into consideration the geographic
location of the testing facility and study area, the pore water salinity regime of the study
area, and the grain size characteristics of the sediment being tested. The species that is
used must exhibit tolerance to the physicochemical properties of every sediment included
in a particular study. Pore water ammonia concentrations may also enter into selection
of one species over others because the four species exhibit differential sensitivity to
aqueous ammonia. Most often it will not be necessary to discriminate among the four
species, and the decision to test one species above the rest may be driven by practical or
logistical concerns. For example, a testing facility may choose to primarily test one
species with a suitable local population in order to prevent potential complications
associated with shipping. However, sediments may be encountered with characteristics
that are outside of the tolerance range of one or more of the species. For example, grain
size limitations for A. abdita and R. abronius are <10% and :>90% fines, respectively. If
these species are exposed to sediments that exhibit textural characteristics outside of
these extremes, any mortality that is observed could be due to effects of grain size
independent of contaminants associated with the sediment. Ambiguity in interpretation
may be avoided by careful consideration of the test species given the sediment to be
tested. Comparative information is available for the four species on sediment grain size
sensitivity (Section 11.4.3), salinity application ranges (Section 11.4.4), and sensitivity to
aqueous ammonia (Section 11.4.5).
10.3 Field Collection
10.3.1 Field collection is presently the most common method for obtaining estuarine and
marine amphipods for sediment testing. All four species are commonly collected,
shipped, and held in the laboratory. Commercial vendors are available for all four
55
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species. The availability of the appropriate size class for each species may vary
seasonally. The collection site chosen should he one for which the presence of abundant
organisms of the correct size and age has been demonstrated previously, and
identification of the species has been confirmed taxonomically (e.g., Bousfield. 1973;
Barnard and Barnard, 1982). Collection areas should be relatively free of
contamination. All individuals in a test must be from the same source, because different
populations may exhibit different sensitivities to contaminants. The four species are
found in distinctly different habitats (Table 10.1).
10.3.2 Species-Specific Habitat Characteristics
10.3.2.1 Ampelisca abdita is found mainly in protected areas from the low intertidal zone
to depths of 60 m. On the Atlantic Coast, A. abdita ranges from central Maine to south-
central Florida and the eastern Gulf of Mexico. It can also be found on the Pacific
Coast in San Francisco Bay. Ampelisca abdita is euryhaline, and has been reported in
waters that range from fully marine to 10 7f(. This species generally inhabits sediments
from fine sand to mud and silt without shell fragments, although it can also be found in
relatively coarser sediments with a sizeable fine component. This species is often
abundant in sediments with a high organic content. Aggregations of .4. abdita are
indicated by an abundance of tubes on the sediment surface, location of which can be
facilitated by looking through a glass-bottom bucket. Although populations may be
seasonally ephemeral, .4. abdita is routinely collected year-round for toxicity testing
from subestuaries of N'arragansett Bay, RI. It is also routinely collected in San
Francisco Bay, CA.
10.3.2.2 Eohaustorius estuarius is found on protected and semi-protected beaches from
mid-water level to shallow subtidal, within the upper 10 cm of sediment along the Pacific
coast from British Columbia south to at least central California (ASTM 1992;
Environment Canada, 1992). Eohaustorius estuarius can be found on open coasts in beds
of freshwater streams flowing into the ocean, and in sand banks in estuaries, above the
level of other regional eohaustorids (£. sawyeri and E. washingtonianus) (Environment
Canada, 1992). It is an estuarine species, and has been reported in areas where pore
water salinity ranges from 1 to 25 %c (Environment Canada, 1992; R. Caldwell,
Northwest Aquatic Sciences, Newport, OR, personal communication). Eohaustorius
estuarius inhabits clean, medium-fine sand with some organic content It is routinely
collected for toxicity tests from Beaver Creek near Newport, OR, and on the west coast
of Vancouver Island. BC. Canada.
10.3.2.3 Leptocheirus plumulosus is found in subtidal portions of Atlantic Coast brackish
estuaries from Cape Cod, MA, to northern Florida. It is common in protected
embay merits, but has been collected in channels of estuarine rivers up to depths of 10 m.
Leptocheirus plumulosus is an estuarine species and has been reported in areas where
salinity at the sediment-water interface ranges from 1 to 15 ?« (DeVVitt et al.. 1992a:
Schlekat et al., 1992; McGee et al., 1994). It is most often found in fine-grained
sediment with a high proportion of particulate organic material, although it has been
collected in fine silty sand with some organic content. Populations of L. plumulosus
may be seasonally ephemeral, with major population growth occurring in spring, and
56
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Table 10.1
Comparison of habitat characteristics and other life history parameters of four estuarine and marine
amp hi pod species used in sediment toxicity tests
Characteristic
Ampelifca
abdita
Kohaustorius
estuarius
Ijeptocheirus
plumulosus
Rhepoxynius
abninius
Substrate Keiaiion
Zoogeography
Habitat
Life Cycle
Availability
Kcological Importance
Tube dwelling, closed1
Atiantic-Cuir- San
Francisco Bay4-5
Poly-upper mesohaline1
40 to 80 days'
Field- Potential culture1
High
Free burrowing2
Pacific"
Oligo-mesohaline2-'
Annual2
Field2
Tube dwelling, open1
Atlantic'
Oligo- mesohaline1
30 to 40 days'
Field-Culture'1112
High
Free burrowing1
Pacific'
Polyhaline'7
Annual10
Field1
Bousfield, 1973
DeVVitt et al.,
Barnard and Barnard, 1982
Nichols et al., 1985
Hopkins, 1986
Environment Canada, 1992
Swart/ et al., 1985
Scott and Redmond, 1989
DeWitt et al., 1992a
Kemp et al., 1985
Schlekat et al., 1992
McCee et al., 1993
57
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large population declines occurring in summer due to actions of predatory fish (Mines et
al., 1986) or absence of essential micronutrients (Marsh and Tenore, 1990). Leptocheirus
plumulosus has been collected for toxi< y tests from several areas in the Maryland
portion of Chesapeake Bay, including uie Magothy, Chester, Corsica, and Wye rivers.
Organisms have been collected for testing year-round from the Magothy River sub-
estuary of Chesapeake Bay (C. Schlekat. SAIC. Narragansett, Rl. and B. McGee,
University of Maryland. Queenstomi, MF>. unpublished data/personal communication).
10.3.2.4 Rhepoxynius abronius occurs in lower inter tidal and nearshore sub tidal zones
on the Pacific Coast from Puget Sound, W V to central California. Primary habitats of
R. abronius include nearshore subtidal zones on the Pacific Ocean coastline, and sub-
and intertidal zones within polyhaline portions of estuaries in the Pacific Northwest. It
is found where the pore water salinity is no lower than 20 %r. Rhepoxynius abronius
naturally inhabits clean, fine sand. It has been collected for use in toxicity tests from
Lower Yaquina Bay, OR (Swartz et al., 1985), and West Beach, Whidbey Island, W A
(Ramsdell et al., 1989; Word et al., 1989).
10.33 Collection Methods
10.3.3.1 Collection methods are species-dependent. Ampelisca abdita and L. plumulosus
are subtidal, and can be collected with a small dredge or grab (e.g., PONAR, Snuth-
Mclntyre, or Van Veen), or by skimming the sediment surface with a long-handled, fine-
mesh net. Eohaustorius estuarius and R. abronius occur both intertidally and subtidally.
Subtidal populations can be collected as above, and intertidal populations can be
collected using a shovel. At least one-third more amphipods should be collected than
are required for the test
10.3.3.2 All apparatus used for collecting, sieving, and transporting amphipods and
control-site sediment should be clean and made of non-toxic material. They should be
marked "live only" and must never be used for working with formalin or any other
toxic materials and should be stored separately from the aforementioned. The
containers and other collection apparatus should be cleaned and rinsed with distilled
water, deionized water, dechlorinated laboratory water, reconstituted seawater, or
natural seawater from the collection site or an uncontaminated source before use.
10.3.3.3 To minimize stress, amphipods should be handled carefully, gently, and quickly,
and only when necessary. Sieving should be performed by slow immersion in collection
site water. Once sieved, attempts should be made to keep amphipods submersed in
collection site sea water at the ambient collection temperature at all times. Amphipods
that are dropped, or injured should be discarded. Once separated from the sediment,
amphipods must not be exposed to direct sunlight.
10.3.3.4 Amphipods can be isolated from collection site sediment by gentle sieving. The
mesh size of the sieve will depend on the species collected. Sieves with 0.5 mm mesh
should be used for sediment containing ,4. abdita and L plumulosus. Larger 4. abdita,
which should not be used in the test, should be excluded by sieving first with a 1.0 mm
screen. When sieving A. abdita. only about half of the amphipods will be extracted from
58
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their tubes. The tube mat should be placed undisturbed for 20 to 30 min coax the
remaining animals out. Sieves with 1.0 mm mesh should be used for E. estuarius and
R. abronius.
10,3.3,5 Collection-site water should be used to sieve sediment in the field. A 2-cm
thick laver of sieved collection site sediment should be placed in transport containers,
and this sediment covered with collection-site water. Detritus and predators recovered
by sieving should be removed, and the collected amphipods should be gently washed into
the transport containers with collection site water.
10.3.3.6 The salinity and temperature of surface and bottom sea water at the collection
site should be measured and recorded. An adequate portion of collection site sediment
should be returned with the amphipods to serve as both laboratory holding sediment
and for use as control sediment in the toxicity test.
10.3.3.7 During transport to the laboratory, amphipods should be kept in sieved
collection-site sediment at or below the collection site temperature. Containers of
amphipods and sediment should be transported to the laboratory in coolers with ice-
packs, and the water in the containers of amphipods should be aerated if transport time
exceeds 1 h.
10.3.3.8 An alternate collection method for .4. abdita involves transporting intact field-
collected tubes to the laboratory for isolation of amphipods. This method is
advantageous because separation of A. abdita from its tubes may be time-consuming
when attempted in the field, a practice which may be impractical in cold winter months.
Amphipod tubes are collected as described in 10.3.1. and placed on a 0.5 mm sieve. The
sieve should be shaken vigorously to remove most of the sediment, leaving the intact
tubes. The tubes should be placed into a covered bucket that contains a sufficient
quantity of collection site water to cover the collected material, and transported to the
laboratory as described in Section 10.2.3.7. In the laboratory, the tubes should be
removed from the collection buckets and placed on a sieve series consisting of a 2 mm
mesh sieve over a 0.5 mm mesh sieve. Amphipods should be forced from their tubes by
spraying collection-temperature sea water on the material present on the 2 mm sieve.
When all the tube material has been sprayed, the 0.5 mm sieve should be shaken
vigorously to separate amphipods from any material that is present. The 0.5 mm sieve
should then be completely submersed, at which point the amphipods will float on the
water surface. The amphipods should then be skimmed from the surface with a small
aquarium net and transported to a container with sea water at the appropriate
temperature. The shaking process should be continued until only a few amphipods
remain in the sieve.
10.3.4 Life Stage and Size
10.3.4.1 The life stage for amphipods used in sediment toxicity tests will depend on the
species tested. For A. abdita and L. plumulosus, sub-adult individuals should always be
selected for testing. The life cycle of these species is relatively short, so the likelihood of
senescence and any effects that could be associated with reproductive
59
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development/maturation are minimized if young individuals are selected. Euhaustorius
estuarius and R. abronius are annual species with longer life spans than .4. abdita and
L. plumulosus. Mature individuals can be used providing they are within the
recommended size range.
10.3.4.2 The size range of test animals should be kept to a minimum regardless of the
chosen species. For all species, mature female amphipods, which are distinguishable by
the presence of embryos in the brood pouch or oviduct, should not be selected for
testing. Additionally, mature male A. abdita and L. plumulosus should not be used.
Recommended size ranges for the four species are as follows:
10.3.4.3 Ampelisca abdita: 3 to 5 mm; or those amphipods retained on a 0.71 mm sieve
after passing through a 1.0 mm sieve. Adult male animals must not be tested: they are
active swimmers and die shortly after mating.
10.3.4.4 Eohausturius estuarius: 3 to 5 mm; or those amphipods retained on a 1.0 mm
sieve. Large individuals (i.e., >5 mm) should not be tested because they might be
senescent
10.3.4.5 Leptocheirus plumulosus: 2 to 4 mm: or those amphipods retained on a 0.5
mm sieve after passing through a 0.71 mm sieve (P. Adolphson, Old Dominion
University, Norfolk, VA, personal communication).
10.3.4.6 Rhepoxynius abronius: 3 to 5 mm; or those amphipods retained on a 1.0 mm
sieve. Large individuals (i.e., >5 mm) should not be tested because they might be
senescent
10.3.5 Shipping Methods
10.3.5.1 All four species have been routinely shipped from the collection site to the
laboratory for sediment toxicity testing. Currently, shipping from the collection site is
necessary for many testing laboratories because culture methods are not available for all
four species. It is critical that standard, demonstrated shipping methods are utilized to
ensure that consistently healthy animals are used in successive toxicity tests.
Additionally, the amphipods that are received by a laboratory must meet the shipping
acceptance criteria recommended for each species. Shipping methods and acceptance
criteria will vary depending on the species used.
10.3.5.2 Ampelisca abdita: Collected amphipods should be shipped within 24 h of
collection. Acceptable methods are available for shipping A. abdita in sediment and in
water. For shipping in sediment, small plastic "sandwich" containers (approximately
500 mL) with scalable lids should be used. The containers are filled three-quarters full
with a minimum depth of 2 cm of sieved fine-grain collection-site sediment and then to
the top with well-aerated sea water. No more than 200 amphipods should be added to
each container. Amphipods should be allowed to burrow into the sediment and build
tubes before the containers are sealed. Containers should be sealed with lids under
water to eliminate any air pockets. For shipping in water-only, scalable plastic bags
60
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(approximately 1 L) should be used. Amphipods in their tubes should be placed in bags
and a sufficient amount of collection site water should be added to keep the tubes moist.
The bag should be filled with pure oxygen before sealing, and then placed into a second
bag. Bags should be placed in a container that has a layer of material (i.e.. styrofoam or
newspaper) sufficiently thick to prevent excessive movement over a layer of ice-packs.
The shipping container should be marked to prevent it from being inverted.
10.3.5.3 Eohaustorius estuarius and Rhepoxynius abronius: Shipping methods for these
organisms are essentially the same. Small plastic "sandwich" containers (approximately
500 mL) with scalable lids should be used. The containers are filled three-quarters full
with sieved collection site sediment (fine sand) and then with a 1 cm layer of collection
site sea water. Not mare than 100 amphipods should then be added and allowed to
burrow. After the animals have burrowed, the overlying water should be poured off,
but the sediment should be moist. The containers are then sealed and ready for
shipment.
10.3.5.4 Leptocheirus plumulosus: Several methods have successfully been used to ship
L. plumulosus, including the A. abdita sediment/overlying water method (10.2.5.2) and
the E. estuariuslR. abronius wet-sediment method (10.2.5.3) as described above.
Additionally, L. plumulosus have been successfully shipped in "sandwich" containers.
cubitainers, and thick plastic bags containing only well-aerated collection-site sea water.
10.3.6 Performance Criteria for Shipped Amphipods
10.3.6.1 The process of ensuring the availability of healthy amphipods on the day that
the test is set up begins when the animals arrive in the laboratory from the supplier.
Although the ultimate performance criterion for amphipods utilized in sediment toxicity
tests is achievement of >90% survival in control sediment, it would be desirable to
assess the quality and acceptability each batch of shipped amphipods using the criteria
that follow. For all four species, biological criteria should include an exhibition of active
swimming behavior upon placement in water, full digestive tracts, and an acceptable
color. Ampelisca abdita should be opalescent pink, E. estuarius should be grayish- or
yellowish white, L. plumulosus should be brownish or orangish-gray, and R. abronius
should be salmon pink, grayish- or yellowish-brown, or white with a pinkish-brown hue.
Mortality among the shipped animals should not exceed 5%. No sexually mature
animals should be included in shipments of -4. abdita or L. plumulosus. The shipping
containers should arrive intact, and the temperature of water or sediment in shipping
containers should be between 4 and 10°C. Information on physical parameters of the
collection site, including at least temperature and salinity, should be provided by the
supplier. Finally, a quantity of collection site sediment should be included as substratum
for amphipods during the acclimation period, and for use as control sediment in the test.
It may be desirable for the testing facility to stipulate these criteria to the supplier
when the animals are ordered. If these criteria are not met, the animals may have
experienced stress during shipment, and >90% survival in control sediment may not be
achieved.
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10.4 Holding and Acclimation
10.4.1 Density. Amphipods should be held and acclimated (if necessary) in containers
(4 to 8 L volume) that contain a 2 to 4 cm layer of collection site sediment that has been
sieved through a 0.5 mm mesh screen. Approximately 350 amphipods should be added
to each 8 L container. Amphipod density should not exceed 1 amphipod/cm:.
10.4.2 Duration. Depending on temperature and salinity at the collection site,
amphipods may have to be acclimated to standard test conditions. If necessary, changes
in temperature or salinity to bring amphipods from the collection site conditions to the
test conditions should be made gradually. Once test conditions are achieved, amphipods
should be maintained at these conditioas for at least two d before testing to allow for
acclimation. Amphipods held for more than ten d should not be used for testing because
they may not satisfy performance control criteria. Temperature and salinity should be
measured at least daily during the period when amphipods are being adjusted to the
conditions of the test water. Thereafter, temperature, salinity, pH, and dissolved oxygen
should be measured in the holding containers at least at the start and end of the
acclimation period, and preferably daily.
10.4.3 Temperature. Overlying water temperature must not be changed by more than
3 C per day during acclimation to the test temperature. Once the test temperature is
reached, amphipods must be maintained at that temperature for a minimum of 2 d. A
water bath, an incubator, or temperature-regulated room can be used for temperature
acclimation.
10.4.4 Salinity. It is unlikely that either A. abdita or R. abronius will require salinit\
acclimation because the collection site salinity for these two species will likely be within
3 arr of the test salinity of 28 7rr. Salinity of water used for temperature acclimation for
these species, if necessary, should be the test salinity, or 28 "rr. The target test salinity
for E. estuarius and L. plumulosus is 20 9rr. and it is likely that the collection site
salinity will be considerably lower than this for both species. Upon arrival in the
laboratory, the water used to hold E. estuarius and L. plumulosus should be adjusted to
20
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10.4.6 Water
10.4.6.1 Provided that it is acceptable to the test organisms, either an uncontaminated
supply of natural sea water or reconstituted sea water can be used for holding and
acclimation (Section 7). At a minimum, healthy amphipods must exhibit acceptable
survival in holding water, and must not exhibit signs of stress, such as unusual behavior
or changes in appearance.
10.4.6.2 If natural sea water is used, it should be obtained from an uncontaminated area
known to support a healthy, reproducing population of the test species or comparatively
sensitive species. Reconstituted sea water is prepared by adding commercially available
sea salts to water from a suitable source, in quantities sufficient to provide the desired
salinity. Pre-formulated brine (e.g., 60 to 90 %<•) prepared with dry ocean salts or heat-
concentrated natural sea water can also be used. To obtain the desired holding or
acclimation salinity, sea salts or brine can be added to a suitable fresh water, natural
estuarine water, or the laboratory's sea water supply. The suitability and consistency of
a particular salt formulation for use in holding and acclimation should be verified by
laboratory tests because some formulations can produce unwanted toxic effects or
sequester contaminants (Environment Canada, 1992). Reconstituted water should be
intensively aerated for two weeks before use (Environment Canada. 1992). Suitable
sources of water used for preparing reconstituted sea water include deionized water or
distilled water, or an uncontaminated natural surface water or ground water.
Chlorinated water must never be used because residual chlorine and chlorine-produced
oxidants are highly toxic to many aquatic animals. Dechlorinated municipal drinking
water should be used only as a last resort because dechlorination is often incomplete.
10.4.6.3 Assessments of the quality of the water used for holding and acclimation and
for preparing reconstituted sea water should be performed as frequently as required to
document acceptability. Analyses of variables including salinity, temperature, suspended
solids, pH, dissolved oxygen, total dissolved gasses, ammonia, nitrite, pesticides, and
metals are recommended. Sea water used for holding and acclimating amphipods
should be filtered (<5 urn) shortly before use to remove suspended particles and
organisms. Holding/acclimation water prepared from natural sea water should be used
within 2 d of filtration/sterilization whereas reconstituted sea water should be used
within 24 h of filtration/sterilization.
10.4.7 Feeding. Ampelisca abdita and L. plumulosus require supplemental feeding
during holding/acclimation. Ampelisca abdita should be fed daily, whereas L. plumulosus
should be fed every other day. Both species should be supplied with an algal ration
consisting of Pseudoisochrysis paradoxa or Phaeodactylum tricornutum that is provided
in conjunction with sea water renewal. See Stein (1973) for procedures to culture algae.
After 75% of the overlying water has been removed, each holding container should be
renewed with sea water at the appropriate salinity that contains algae at a concentration
of at least 1 x 106 cells/mL. Leptocheirus plumulosus should also be provided with dry
food ration, consisting at a minimum of finely powdered TetraMin®. It may be desirable
to grind the dry food in a blender. Each container should Deceive approximately 0.5 g
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dry food/350 arnphipods. Eohaustorius estuarius and R. abronius will utilize organic
material in the holding sediment as food and do not require supplemental feeding,
10.4.8 Acceptability of aniniais. Am phi pods counted into the holding/acclimation
chambers should be active and appear healthy. Any individuals that fail to burrow or
fail to make tubes (i.e., A. abdita) in holding sediment or that appear unhealthy during
the holding/acclimation period should be discarded. Apparently dead individuals should
also be discarded. If greater than 59r of the amphipods emerge or appear unhealthy
during the 48 h preceding the test, the entire group should be discarded and not used in
the test. Additionally, the group should be discarded if more than 10% of the
amphipods die or become inactive during the holding period before testing.
10.5 Culture Procedure for Leptocheirus plumulosus
10.5.1 The culturing method below is based on a method described in DeWitt et al.
(1992a). LSEPA sponsored a workshop was held from 12 to 13 January 1994 in
Newport, OR, on sediment testing (both acute and chronic) and culturing of Leptocheirus
plumulosus. Participants arrived at a consensus on recommending that laboratories with
little or no experience follow what has worked for other laboratories first. A consensus
among workshop participants was reached on a recommended diet for cultures, and this
will be stated. These procedures should not be considered definitive because many
issues contributing to optimal culture productivity have yet to be addressed. A periodic-
renewal culture system is used. It consists of culture bins that contain aerated sea water
over a thin (~1 cm) layer of clean, fine-grained sediment in which the amphipods
burrow. Culturing areas must be separate from testing areas to avoid loss of cultures
because of cross-contamination.
10.5.2 Starting a Culture. Amphipods for starting a laboratory culture of L. plumulosus
should be obtained from a source with a verified culture (Table 10.1). Alternatively,
L. plumulosus can be obtained from field populations. The taxonomy of the animals
must be confirmed before they are introduced into existing laboratory populations. In
addition, the ability of the wild population of sexually reproducing organisms to cross-
breed with existing laboratory populations must be determined. Sensitivity of the wild
population to select contaminants should also be documented. The temperature and
salinity of the shipped water containing amphipods should be gradually adjusted to 20 C
and 20 %r, respectively, at rates not exceeding 3°C per d and 5 7rr per d. Feeding and
regular maintenance should begin once the acclimation period is over. Cultures should
be started with approximately 300 mixed-age animals, of which only 100 should be
reproductively active adults (i.e., individuals >3 mm in length).
10.5.3 Culture Chambers. Culture chambers should be amenable to easy maintenance.
Plastic dishpans (30 cm x 40 cm x 15 cm) have been used successfully by several
laboratories (DeWitt et al., 1992a), and are recommended. Aeration, provided through
an aquarium air stone, should be vigorous and constant.
10.5.4 Culture Sediment. Cultures should be established with a thin layer (~1 cm) of
sediment that is spread on the bottom of the culture chamber. The sediment that is
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used for culture purposes should be the same as the control sediment used in sediment
toxicity tests. Suitable sources for culture sediment include the amphipod collection site
or another area that harbors sediment within the physico-chemical tolerance of
L. plumulosus that are listed in Table 4.1. Culture sediment should be press sieved
through a <0.5 mm screen before using to avoid the presence of indigenous organisms in
culture.
10.5.5 Culture Water. Culture water should be from the same source used for holding
and acclimating test organisms and for conducting toxicitv tests. See Section 7.1.2 for
acceptable sources of seawater. Cultures of L. plumulosus are maintained at a salinity of
20 ?«. To obtain this salinity, sea water should be diluted with deionized water.
Seawater used for culturing amphipods should be filtered (<5 um) shortly before use to
remove suspended particles and organisms. Water that may be contaminated with
pathogens should be treated shortly before use by filtration (<0.45 (am), either alone or
in combination with ultraviolet sterilization. Culture water prepared from natural sea
water should be used within 2 d of filtration/sterilization whereas reconstituted sea water
should be used within 24 h of filtration/sterilization. Culture water should be renewed
three times per week in conjunction with feeding.
10.5.6 Temperature and Photoperiod. Cultures should be maintained at 20~C. The
reproductive rate of L. plumulosus increases at temperatures greater than 2U°C,
necessitating more frequent culture thinning. Temperatures below 20°C may not foster
sufficiently prolific reproductive rates. Fluorescent lights should be on a 16L:8D
photoperiod at a light intensity of 500 to 1000 lux.
10.5.7 Food and Feeding
10.5.7.1 The following section is based on recommendations made at the I'SEPA-
sponsored "Leptocheirus plumulosus Workshop" held in Newport, OR. from 12 to 13
January 1994. The recommendations follow the consensus of experts who culture
L. plumulosus for use in sediment toxicity testing. It was concluded that laboratories
unfamiliar with this species should utilize the specific diet recommended below.
Modifications to the diet could then proceed by laboratories in order to optimize culture
practices as long as the modifications satisfied the performance criteria.
10.5.7.2 Culture chambers should be provided with food in conjunction with water
renewal. Three times a week, approximately 60<7r of culture water should be siphoned
from each culture chamber and replaced with the same volume of renewal water. The
renewal water should consist of seawater, cultured phytoplankton, and deionized water
combined to a salinity of 20 9rr and an algal density of approximately 10" cells mL '.
The proportions will vary depending upon the salinity of the seawater and the density of
the cultured phytoplankton. The algae used should be the chrysophyte Pseudoisochrysis
paradoxa and the diatom Phaeodactylum tricornutum mixed 1:1 v/v. Other algal species
can be used if it can be demonstrated that they foster amphipod growth and
reproductive rates equal to those of the aforementioned algal species.
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10.5.7.3 The cultures should also be provided with dry food just after the water change
is complete. The recommended dry food consists of a mixture of TetraMin*. dried
alfalfa, dried wheat leaves, and Neo-Novum*1. TetraMin* is a fish food flake, and is
available at most aquarium-supply stores. Dried alfalfa and dried wheat leaves are
available at health food stores. Neo-Novums is a maturation feed for use in shrimp
mariculture, and is available from Argent Chemical Laboratories, Redmond, \V A. The
mixture should be prepared at the following proportions: 48.5^r Tetra\1inE + 24f> dried
alfalfa + 24^ dried wheat leaves + 4.5% Neo-Novum®. The dry food mixture should be
made in advance in 400 g batches. The mixture should be thoroughly homogenized into
a fine powder. A blender will provide best results; alternatively, a mortar-and-pestle
can be used.
10.5.7.4 The amount of dry food added will depend on the density of each culture bin.
Newly started culture bins (i.e., those with 300 mixed-age animals) should receive 0.25 to
0.3 g of the dry food mixture per feeding for the first two weeks. Thereafter, 0.5 g of
the dry food mixture should be provided per feeding. Less food is provided during the
first two weeks as excess food may result in microbial build-up on the sediment surface.
The appropriate amount of food should be measured out and sprinkled on the water
surface at the center of the culture bin, where it will disperse and settle evenly.
10.5.8 Culture Maintenance
10.5.8.1 Observations and Measurements. Cultures should be observed daily to ensure
that aeration is adequate in all culture chambers. Inspection for the presence of
oligochaete and polychaete worms and copepods should be conducted weekly. The
presence of excessive densities of these or other competing or predacious organisms
should prompt renewal of culture sediment after separating amphipods from other
organisms. Cultures should be inspected for the presence of microbial build-up on the
sediment surface in conjunction with water changes and feeding. This build-up appears
as a white or gray growth that may originate near uneaten food. Presence of microbial
build-up may indicate that the amount dry food is in excess required by the amphipods.
Addition of the dry food mixture to culture chambers with surficial microbial build-up
should temporarily cease until the build-up is no longer present.
10.5.8.2 Healthy cultures are characterized by an abundance of burrow-openings on the
sediment surface. Although amphipods may leave their burrows to search for food or
mates, they will ordinarily remain in their burrows under the daylight portion of the
photoperiod. Amphipod density may therefore only be estimated by examining the
number of burrow openings. An abundance of animals (e.g., >15 per culture bin) on the
sediment surface could indicate inadequate sediment quality, low dissolved oxygen
concentrations, or overcrowding. A culture chamber with an abundance of amphipods
or unhealthy individuals on the sediment surface should be examined closely, and the
dissolved oxygen concentration should be measured. If the dissolved oxygen
concentration is below 60% saturation the culture chamber should be sieved, and the
population and culture sediment examined. If the population is too dense (i.e., >1500
adults per culture chamber), it should be thinned as described below. If the sediment
becomes an unacceptable habitat, i.e., if it is black and sulfurous below the sediment
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surface, or contains an excess of competitive or predacious organisms, the health)
surviving amphipods should be placed in a new culture chamber with newly prepared
culture sediment.
10.5.8.3 Water temperature should be measured daily and dissolved oxygen should be
measured weekly. Cultures should be continuously aerated. Salinity should be
measured after water renewal, or 3 times per week. Ammonia and pH should be
measured with each new batch of sediment.
10.5.8.4 Culture Density. Leptocheirus plumulosus can be prolific, and care must be
taken to ensure that culture chambers do not get overcrowded. Amphipods in
overcrowded culture chambers may be stressed because of food and space limitations.
causing the fecundity of females to drop below 5 eggs per female. Culture density must
not exceed 1.5 amphipods cm'2 (i.e., 1500 animals per culture chamber) and should
ideally be maintained at approximately 0.4 amphipods cm2 (i.e., 400 adults per culture
chamber). To avoid overcrowding, cultures should be thinned every two months by
sieving through a 1 mm mesh screen, allowing young amphipods to pass through the
screen and remain in the sediment. Approximately 100 to 150 adult amphipods should
be selected from the sieved population and returned to the culture tub. The remainder
should be used to start new cultures or discarded.
10.5.9 Obtaining Amphipods for Starting a Test
10.5.9.1 Leptocheirus plumulosus used in tests should be started with pre-reproductive
animals that are 2 to 4 mm in length. To obtain animals in this size range, sediment
from culture chambers containing mixed-size amphipods should be poured over a sieve
series that consists of the following sequence of mesh sizes: 0.71 mm, 0.50 mm, and
0.25 mm. Animals retained on the 0.50 mm mesh screen should be washed into a
shallow glass pan. The smaller animals from this group should be selected for toxicity
testing. Gravid females should be avoided.
10.5.9.2 Alternatively, test animals within a narrow size range are obtained by isolating
the smallest amphipods which are allowed to grow until they reach a testable size. To
obtain the smallest amphipods, first transfer sediment from culture chambers containing
mixed-size amphipods over a sieve series that consists of the following sequence of mesh
sizes: 1.0 mm, 0.5 mm, and 0.25 mm. Animals retained on the 0.25 mm mesh screen
should be small juveniles that are 1.1 to 2.0 mm in length. They will take approximately
two weeks to reach testable size after isolation. The amphipods retained on the 0.25 mm
screen should be washed into a culture chamber that is set up as a normal culture
chamber, i.e., containing a thin (~1 cm) sediment layer and maintained under culture
conditions for the two week interim period. By the end of the two week grow-out
period, the animals should be of testable size (i.e., 2 to 4 mm), and be within a narrow
size and age range.
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Section 11
Test Method 100.4
Ampelisca abdita, Eohaustorius estuarius, Leptocheirus plumulosus,
or Rhepoxynius abronius 10-d Survival Test for Sediments
11.1 Introduction
11.1.1 Ampelisca abdita, Eohaustorius estuarius, Leptocheirus plumulosus, and
Rhepoxynius abronius have been used extensively to test the toxicity of estuarine and
marine sediments. The choice of these amphipod species as test organisms is based on
sensitivity to sediment-associated contaminants, availability and ease of collection,
tolerance of environmental conditions (e.g., temperature, salinity, grain-size), ecological
importance, and ease of handling in the laboratory. Additionally, the species chosen for
this method are intimately associated with sediment by nature of their burrowing or
tube-dwelling and feeding habits. Field validation studies have shown that amphipods
are absent or have reduced abundances at sites where toxicity in laboratory tests.
Amphipod sediment toxicity tests have been successfully performed for regulatory and
research purposes by numerous laboratories, including state and federal government
agencies, private corporations, and academic institutions. Test guidance for A. abdita,
E. estuarius, L. plumulosus, and R. abronius has previously been developed (ASTM,
1992). The four species chosen are representative of both estuarine and marine habitats
and sediments that span the spectrum of particle sizes from fine-to coarse-grained
sediment. Thus, either alone or in combination, they may be used to measure toxicity of
any commonly encountered estuarine or marine sediment
11.1.2 Specific test methods for conducting the 10-d sediment toxicity test for the
amphipods Ampelisca abdita, Eohaustorius estuarius, Leptocheirus plumulosus, and
Rhepoxynius abronius are described in Section 11.2. Test method 100.4 was developed
based on Swartz et al. (1985); DeVVitt et al..(1989); Scott and Redmond (1989); Schlekat
et al. (1992): ASTM (1992); and Environment Canada (1992). Results of tests using
procedures different from the procedures described in Section 11.2 may not be
comparable and these different procedures may alter bioavailability. Comparison of
results obtained using modified versions of these procedures might provide useful
information concerning new concepts and procedures for conducting sediment tests with
estuarine or marine organisms. If tests are conducted with procedures different from
the procedures described in the manual, additional tests are required to determine
comparability of results (Section 1.3).
11.2 Recommended Test Method for Conducting a 10-d Sediment Toxicity Test with
Ampelisca abdita, Eohaustorius estuarius, Leptocheirus plumulosus, or Rhepoxynius
abronius.
11.2.1 Recommended conditions for conducting a 10-d sediment toxicity test with
A. abdita, E. estuarius, L. plumulosus, and R. abronius 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
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Table 11.1 Test conditions for conducting a 10-d sediment toxicity test with
Ampelisca abdita, Eohaustorius estuarius, Leptocheirus plumulosus, or
Rhepoxynius abronius
Parameter
Conditions
1. Test type:
2. Temperature:
3. Salinity:
4. Light quality:
5. Illuminance:
6. Photoperiod:
7. Test chamber:
8. Sediment volume:
9. Overlying water volume:
10. Renewal of overlying water:
11. Size and life
stage of amphipods:
12. Number of organisms/
chamber:
13. Number of replicate
chambers/treatment:
14. Feeding:
Whole sediment toxicity test, static.
15°C: E. estuarius and R. abronius
202C: A. abdita
25°C: L plumulosus
20 9rr: E. estuarius and L. plumulosus
28 9rr: A. abdita and R. abronius
Wide-spectrum fluorescent lights
5(10 - 1000 lux
24L:OD
1-L glass beaker or jar with -10 cm I.D.
175 mL (2 cm)
800 mL
None
A. abdita: 3 - 5 mm (no mature males or
females)
E. estuarius: 3 • 5 mm
L. plumulosus: 2 - 4 mm (no mature males
or females)
R. abronius: 3 - 5 mm
20 per test chamber
Depends on objectives of test. At a
minimum, four replicates must be used.
None
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Table 11.1 Test conditions for conducting a 10-d sediment toxicity test with
Ampelisca abdita. Eohaustorius estuahus, Leptocheirus plumulosus, or
Rhepoxynius abronius (continued)
Parameter
Conditions
15. Aeration:
16. Overlying water:
17. Overlying water quality:
Test duration:
19. Endpoints:
20. Test acceptability:
Water in each test chamber should be aerated
overnight before start of test and throughout
the test: aeration at rate that maintains >90
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Table 11.2 General activity schedule for conducting a sediment toxicity test with
Ampelisca abdita, Eohaustorius estuarius, Leptocheirus plumulosus, or
Rhepoxynius abronius.
Dav Activity
-10 to -3 Collect or receive amphipods from supplier and place into collection site
sediment. Alternatively, separate 2 - 4 mm L. plumulosus from cultures.
-9 to -2 Acclimate and observe amphipods to species-specific test conditions.
Feed A. abdita and L. plumulosus. Monitor water quality (e.g.,
temperature, salinity, and dissolved oxygen).
-1 Observe amphipods, monitor water quality. Add sediment to each test
chamber, place chambers into exposure system, and start aeration.
0 Measure pore water total ammonia, salinity, and pH. Measure
temperature of overlying water in test chambers. Transfer 2(1 amphipods
into each test chamber. Archive 20 test organisms for length
determination.
1 Measure temperature. Observe behavior of test organisms and ensure
that each test chamber is receiving air. Measure dissolved oxygen in test
chambers to which aeration has been cut-off.
2 Measure total water quality (pH, temperature, dissolved oxygen, salinity.
total ammonia of overlying water). Observe behavior of test organisms
and ensure that each test chamber is receiving air.
3 to 7 and 9 Same as Day 1.
8 Same as Day 2.
10 Measure temperature. End the test by collecting the amphipods with a
sieve.
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chambers/treatment, and water quality characteristics should be based on the purpose of
the test and the methods of data analysis (Section 12). 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 A. abdita, E. estuarius.
L. plumulosus, and R. abronius must be conducted at the species-specific temperature
and salinity with a 24 h light photoperiod at a illuminance of about 500 to 1000 lux
(Table 11.1). Test chambers are 1 L glass chambers containing 175 mL of sediment and
800 mL of overlying seawater. Twenty amphipods are added to each test chamber at
the start of a test The size range of the amphipods will depend on species that is being
tested (see Section 10.3.4 for allowable size range for each species). The number of
replicates/treatment depends on the objective of the test Five replicates are
recommended for routine testing (see Section 12). Exposure is static (i.e., water is not
renewed), and the animals are not fed over the 10 d exposure period. Overlying water
can be culture 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. For all other applications, the characteristics of the
overlying water for each species should be chosen according to Table 11.1. Requirements
for test acceptability are summarized in Table 11.3.
11.3 General Procedures
11.3.1 Introduction of Sediment. On the day before the addition of amphipods (Day -1).
each test sediment (either field collected or laboratory spiked) should be homogenized by
stirring in the sediment storage container or by using a rolling mill, feed mixer, or
other suitable apparatus. Control and reference sediments are included. Sediment
should be visually inspected to judge the extent of homogeneity. Excess water on the
surface of the sediment can indicate separation of solid and liquid components. If a
quantitative measure of homogeneity is required, replicate subsamples should be taken
from the sediment batch and analyze for TOC. chemical concentrations, and particle
size. Spiked sediments should not be homogenized before introduction into test
chambers because the equilibrium between the spiked contaminant and the sediment
partitioning factors may be disrupted.
11.3.1.1 A 175-mL aliquot of thoroughly homogenized sediment is added to each test
chamber. It is important that an identical volume be added to each replicate test
chamber; at a minimum the volume added should equate to a depth of 2 cm in the test
chamber. The sediment added to the test chamber should be settled either by tapping
the side of the test chamber against the side of the hand or by smoothing the sediment
surface with a nylon, fluorocarbon. or polyethylene spatula. Highly contaminated
sediment should be added to test chambers in a certified laboratory fume hood.
11.3.2 Addition of Overlying Water. To minimize disruption of sediment as test
seawater is added, a turbulence reducer should be used. The turbulence reducer mav be
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Table 11.3 Test acceptability requirements for a 10-d sediment toxicity test with
Ampelisca abdita. Eohaustorius estuarius, Leptocheirus plumulosus. or
Rhepoxynius abronius.
A. It is recommended for conducting a 10-d test with A. abdita, E. estuarius. L,
plumulttsus, or R. abronius that the following performance criteria are met:
1. Size, life stage, and reproductive stage of amphipods must be within the
prescribed species-specific ranges at the end of the test (Section 10.3.4).
2. Average survival of amphipods in the control sediment must be greater than
or equal to 90<7r at the end of the test.
3. Salinity, pH, and ammonia in the overlying water and sediment grain size
are within tolerance limits of test species.
B. Performance-based criteria for culturing L. plumulttsus include:
1. Laboratories should perform monthly %-h water-only reference-toxicity
tests to assess the sensitivity of culture organisms. If reference-toxicit> 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. Records should be kept on the frequency of restarting cultures.
3. Laboratories should record the pH and ammonia of the cultures at least
quarterly. Dissolved oxygen and salinity should be measured weekh.
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.
C. Performance-based criteria for field-collected amphipods include:
1. Laboratories should perform reference-toxicant tests on each batch of field-
collected amphipods received used in a sediment test (Section 9.16).
2. Acclimation rates to test salinity and temperature should not exceed 3 ( and
5 <7rr per 24 h.
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Table 11.3 Test acceptability requirements for a 10-d sediment toxicity test with
Ampelisca abdita, Eohaustorius estuarius, Leptocheirus plumulosus. or
Rhepoxynius abronius (continued)
3. Amphipods received from commercial suppliers must exhibit active
swimming behavior upon placement in water, have full digestive tracts, and
display an acceptable color.
D. Additional requirements:
1. All organisms in a test must be from the same source.
2. It is desirable to start tests as soon as possible 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 must be included in a test.
5. The time-weigh ted average of daily temperature readings must be within
±1 :C of the desired temperature. The instantaneous temperature must
always be within ±3CC of the desired temperature.
6. Natural physico-chemical characteristics of test sediment collected from the
field should be within the tolerance limits of the test organisms.
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either a disk cut from polyethylene, nylon, or Teflon® sheeting (4 to 6 mil), or a glass
petri dish attached (open face up) to a glass pipette. If a disk is used as the turbulence
reducer, it should fit the inside diameter of the test chamber and have attached a length
of nylon monofilament (or nontoxic equivalent) line. The turbulence reducer is
positioned just above the sediment surface and raised as sea water is added to the 750-
mL mark on the side of the test chamber. The turbulence reducer is removed and
rinsed with test sea water between replicates of a treatment. A separate turbulence
reducer is used for each treatment. The test chambers should be covered, placed in a
temperature controlled water bath (or other acceptable equivalent) and gently aerated.
A test begins when the organisms are added to the test chambers (Day l)i.
11.3.3.1 Addition of Amphipods. On the following day (Day 0), amphipods are added to
the test chambers. Approximately one-third more amphipods than are needed for the
test should be sieved from the culture or control sediment in the holding container* s).
and transferred to a sorting tray. The additional animals allow for the selection of
healthy, active individuals. The sieve size for isolating amphipods from the culture or
control sediment will depend upon the selected species. Ampelisca abdita and
L. plumulosus should be isolated using a 0.5 mm sieve, whereas E. esiuarius and
R. abronius should be isolated using a 1.0 mm sieve. Sieving should be conducted with
sea water of the same temperature and salinity as the holding and test water. Once
isolated, active amphipods should be randomly selected using a transfer pipette or other
suitable tool (not forceps), and distributed among dishes or cups containing
approximately 150 mL of test sea water until each container has tvventv amphipods.
The number of amphipods in each dish should be verified by recounting before adding
to test chambers. To facilitate recounting, amphipods may be distributed to test
chambers in batches of 5 or 10 instead of the full complement of 20. The distribution of
amphipods to the test chambers must be executed in a randomized fashion.
11.3.3.2 Amphipods should be added to test chambers without disruption of the
sediment by placing a 6-mil polyethylene, nylon, or Teflon® disk on the water surface
and gently pouring the water and amphipods from the sorting container over the disk
into the test chamber. The disk should be removed once the amphipods have been
introduced. Alternatively, amphipods from the sorting container can be poured into a
sieve cup (mesh size <0.5 mm) and gently washed into the test chamber with test sea
water. Any amphipods remaining in the sorting container should be gently washed into
the test chamber using test sea water. The water level should be brought up to the 950-
mL mark, the test chamber covered, and aeration continued.
11.3.3.3 After the addition of the animals, the test chambers should be examined for
animals that may have been injured or stressed during the isolation, counting, or
addition processes. Injured or stressed animals will not burrow into sediments, and
should be removed. The period of time allowed for healthy amphipods to bury into test
sediments will depend upon the species used. Eohaustnrius estuarius, L. plumulosus, and
R. abronius should be allowed 5 to 10 min to bury into the test sediment. Ampelisca
abdita, which may take longer to build tubes, should be allowed 1 h. Amphipods that
have not burrowed within the prescribed time should be replaced with animals from the
same sieved population, unless they are repeated!) burrowing into the sediment and
75
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immediately emerging in an apparent avoidance response. In that case, the amphipods
are not replaced. The number of amphipods that are removed must be recorded.
11.3.4 Test Conditions
11.3.4.1 Aeration. The overlying sea water in each test chamber must be aerated
continuously after the water is added (i.e.. Days -1 through 10) except during
introduction of the test organisms. Compressed air, previously filtered and free of oil.
should be bubbled through a glass or plastic pipette and attached plastic tubing. The tip
of the pipette should be suspended 2 to 3 cm above the surface of the sediment layer so
as to not disturb the sediment surface. The concentration of dissolved oxygen (DO) in
the water overlying the sediment in the test chambers is maintained at or near
saturation by gently aerating the water. Air is bubbled through the test chamber at a
rate that maintains a >90% DO concentration, but does not cause turbulence or disturb
the sediment surface. If air flow to one or more test chambers is interrupted for more
than one h, DO must be measured in those test chambers to determine whether DO
concentrations have fallen below 6\)% of saturation. Results may be unacceptable for
test chambers in which aeration was interrupted and DO concentrations fell to below
60<~r saturation.
11.3.4.2 Lighting. Lights must be left on continuously at an intensity of 500 to 1000 lux
during the 10 d exposure period. The constant light increases the tendency of the
organisms to remain buried in the sediment, and thus to remain exposed to the test
material.
11.3.4.3 Feeding. The four species of amphipods used in this method must not be fed
during the 10-d exposure period.
11.3.4.4 Water Temperature. The test temperature will depend on the species that is
tested. Test temperatures were selected to be near the summertime thermal maximum
that each species would be expected to encounter in the environment. Eohaustorius
estuarius and R. abronius, the Pacific Coast amphipods. must be tested at 15=C.
Ampelisca abdita must be tested at 20°C and L. plumulosus at 253C.
11.3.4.5 Salinity. The salinity of the water overlying the test sediment will vary
depending on the selected test species. For routine testing, .4. abdita and R. abronius
should be tested at an overlying water salinity of 28 ^rr, whereas E. estuarius and
L. plumulosus should be tested at 20 7f(. Pore water salinity of each test sediment must
be measured prior to the initiation of a test. Sediment pore water should be obtained by
centrifugation. Alternatively, salinity can be measured before homogenization in the
water that comes to the surface in the sample container as the sediment settles. The
pore water salinity of the test sediment must be within the salinity application range of
the chosen amphipod species (Table 10.2). Rhepnxynius abronius cannot be tested when
sediment pore water salinities are <25 "vrr. Another species must be used for such
sediments. Ampelisca abdita, E. estuarius, and L. plumulosus can be tested over the
entire pore water salinity range (i.e.. 0 to 34 9M when the recommended species-specific
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overlying salinity is used. Depending on the objectives of the study, E. estuarius and
L. plumulosus may be tested with overlying water at salinities ranging from 1 to 32 '«.
11.3.5 Measurements and Observations
11.3.5.1 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 time-weighted average of daily temperature
readings must be within ±1°C of the desired temperature. The instantaneous
temperature must always be within ±3°C of the desired temperature.
11.3.5.2 Salinity, dissolved oxygen, and pH of the overlying water should be measured
daily in at least one test chamber per treatment, and at a minimum, they must he
measured in every test chamber at the beginning and the end of a test.
11.3.5.3 Ammonia must be measured in overlying water towards the beginning (e.g..
2) and towards the end of the test (e.g., day 8). Measurement of overlying water pH and
temperature should accompany each ammonia measurement. Simultaneous
measurements of ammonia, pH, and temperature in sediment pore water should be
measured at the beginning of the test. Pore water should be extracted after the
sediment has been press-sieved and homogenized. Samples of pore water should be
obtained by centrifugation.
11.3.5.4 Each test chamber must be examined at least daily during the 10 d test period
to ensure that airflow to the overlying sea water is acceptable. The number of
amphipods swimming in the water column and trapped in the air-water interface should
be noted. Amphipods caught in the air-water interface must be gently pushed down into
the water using a glass rod or pipette. The number of apparently dead animals should
be noted.
11.3.6 Ending a Test. Laboratories should demonstrate the ability of their personnel to
recover an average of at least 90% of the organisms from control sediment. For
example, test organisms could be added to control sediment and recovery could be
determined after 1 h (Tomasovic et al., 1994).
11.3.6.1 The contents of the test chambers must be sieved to isolate the test animals.
The mesh size for sieving the contents of the test chambers must be no larger than 0.5
mm. Test water should be used for sieving. Material retained on the sieve should be
washed into a sorting tray with clean test sea water. Ampelisca abdita are tube-builders.
and it will be necessary to make an effort to ensure that no tubes remain on the sieve.
The sieve should be slapped forcefully against the surface of the water to ensure that all
of the amphipods and tubes are dislodged from the screen. Eohaustorius estuarius,
L. plumulosus, and R. abnmius are easily removed from the sediment by the sieving
process.
1 1.3.6.2 Material that has been washed from the sieve into the sorting tray should be
carefully examined for the presence of amphipods. A small portion of the material
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should be sorted through at a time, removing arnphipods as they are found. Material
from tests conducted with .4. abdita will include tubes built by the arnphipods during the
test. The tubes must be carefully examined and teased apart under a dissecting
microscope or magnifying glass because .4. abdita will often remain in the tubes even
after vigorous sieving. Numbers of live, missing, and dead amphipods should be
determined and recorded for each test chamber. Missing animals are assumed to have
died and decomposed during the test and disintegrated: they should be included in the
number dead in calculations of the percent survival for each replicate treatment.
Amphipods that are inactive but not obviously dead must he observed using a low-power
dissecting microscope or a hand-held magnifying glass. Any animal that fails to exhibit
movement (i.e. neuromuscular twitch of pleopods or antennae) upon gentle prodding
with a probe should be considered dead.
11.3.7 Test Data. Survival is the primary endpoint recorded at the end of the 10-d
sediment toxicity test with .4. abdita, E. estuarius, L. plumulosus, and R. abronius. The
ability of surviving amphipods to rebury in clean control sediment can be used to
calculate effective mortality, that is, the sum of dead animals plus those survivors that
fail to rebun. This endpoint has been used for E. estuarius, L. plumulosus, and
R. abronius. If it is desired to determine reburial, surviving amphipods should be
transferred to containers holding a 2-cm layer of 0.5 mm sieved control sediment and an
overlying layer (>2 cm) of test sea water. Salinity of the test sea water for reburial
should be the same as that measured in the test chamber. The number of surviving
amphipods unable to rebury in control sediment after 1 h is recorded for each test
chamber and is used to calculate effective mortality.
11.4 Interpretation of Results
11.4.1 Section 12 describes general information for interpretation of test results. The
following sections describe species-specific information that is useful in helping to
interpret the results of sediment toxicity tests with .4. abdita. E. estuarius. L. plumulosus,
and R. abronius.
11.4.2 Influence of indigenous Organisms. Indigenous organisms may be present in
field-collected sediments. An abundance in the sediment sample of the test organism, or
organisms taxonomically similar to the test organism, may make interpretation of
treatment effects difficult. The presence of predatory organisms can also adversely
affect test organism survival. For example, Redmond and Scott (1989) showed that the
polychaete \ephtys incisa will consume Ampelisca abdita under toxicity test conditions.
11.4.3 Effect of Sediment Grain Size. All four species show tolerance to most sediment
types, with generally little effect on survival whether coarse-grained or fine-grained (i.e.,
predominantly silt and clay) clean sediments are used. However, adverse effects due to
the grain-size distribution of test sediment may occur when sediments that are either
extremely sandy or fine depending on the species of amphipod used. In order to
separate effects of sediment-associated contaminants from effects of particle size, an
appropriate clean control/reference sediment should be incorporated into the test when
test sediments are within the range of concern for each species. Alternatively, another
78
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species that is tolerant of the sediment extreme in question might be tested in
conjunction with the chosen species. Ranges of concern are outlined below.
11.4.3.1 Ampelisca abdita. Survival of Ampelisca abdita in sediment that is >951- sand
may elicit excess mortality, but this has not been quantified (J. Scott, SAIC,
Narragansett, RI, personal communication). Toxicity tests conducted with A. abdita on
sediments that are >95% sand should be conducted with a clean control sediment
characteristic of that test sediment.
11.4.3.2 Leptocheirus plumulosus. Leptocheirus plumulosus has exhibited >90% sur\i\al
in clean sediments ranging from - 100% sand to - 100% silt + clay (SAIC. 1993a:
SAIC. 1993b: Schlekat et al., 1992; J. Kavanaugh. University of West Florida. Gulf
Breeze, FL. personal communication).
11.4.3.3 Eohaustorius estuarius. Eohaustorius estuarius has exhibited acceptable (100'V i
survival when exposed to clean sediments ranging from 0.6 to 100% sand ilSEPA ERL-
Narragansett, Pacific Ecosystems Branch, Newport, OR, unpublished data). Howe\er.
E. estuarius naturally inhabits sandy sediments, and some correlation between survival
and grain size has been reported by DeVVitt et al. (1989) and SAIC <1993a; 1993b), with
increased mortality associated with increased proportions of fine-grained sediment.
Therefore, it may be desirable to include clean control sediments with a range of particle
sizes characteristic of those of the test sediment(s) in toxicity tests conducted with
E. estuarius.
11.4.3.4 Rhepoxynius abronius. Rhepoxynius abronius has been used to test sediments
with a wide range of sediment grain sizes. However, R. abronius naturally inhabits
clean, fine, sandy sediments, and a number of studies have shown some reduction in
survival when this species is held in very fine-grained (predominantly silt and clay)
sediment (DeVVitt et al., 1988; Long et al., 1990; McLeay et al., 1991; SAIC. 1993a:
SAIC, 1993b). Therefore, when test sediments are predominantly silts or clays, the
experimental design include a silt-clay control sediment with a range of particle sizes
characteristic of the test sediment(s). Alternatively, when the particle size of test
sediments are known, regression techniques can be used to evaluate potential effects of
fines on R.abronius survival (see DeWitt et al., 1988).
11.4.4 Effects of Pore Water Salinity. The four amphipod species exhibit variability in
their salinity tolerance ranges. There are two options available for laboratory sediment
testing regarding the choice of overlying water salinity for a given sediment. The
options are to either use the standard salinity for each test species, or to match the
salinity to that of the pore water. The range of pore water salinities in which a given
species can survive for ten days when using the species-specific overlying water salinity
is the salinity application range. The range of salinity in which a given species can
survive for ten days when the overlying water salinity is matched to that of the pore
water salinity is the salinity tolerance range. In either scenario, the potential for a toxic
response due to salinity alone exists if a species is exposed to conditions outside of its
range of tolerance. For estuarine sediments, it is very important to know the port1 water
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salinity of each sediment before testing is started, to choose a species that will not be
affected by the pore water salinity, and to use overlying water of an appropriate salinity.
11.4.4.1 Salinity tolerance ranges for each species are as follows: Ampelisca abdita:
20 to 32 Tc (SAIC, 1993b): Eohaustorius estuarius: 2 to 34 <7r(; Leptocheirus
plumulosus: 1.5 to 32 l~fr; Rhepoxynius abronius: 25 to 32 7rf. While there is some
evidence of salinity-related stress for E. estuarius and L. plumulosus at salinity extremes,
the breadth of salinity tolerance exhibited by these species (DeVVitt et a!., 1989; Schlekat
et al., 1992: SAIC. 1993b) is most likely sufficient for application to the majority of
sediments that may be encountered in an estuarine system (i.e., interstitial salinity
between 2 and 28 '/<• >. If it is desirable to have matching overlying and pore water
salinity from areas where pore water salinities are 0 to 2 7rr, an organism that has been
demonstrated to tolerate this salinity range should be used, either instead of or in
addition. The amphipod Hyalella azteca is one such species. Likewise, sediments
collected from areas of high salinity (i.e.. >32 7ff for L. plumulosus} should probably
utilize A. abdita, E. estuarius, or R. abronius.
11.4.4.2 Salinity application ranges for each species are as follows: Ampelisca abdita
with overlying water salinity of 28 to 32 7ff: 0 to 34 ?« (Weisberg et al., 1992; SAIC,
1993b); Eohaustorius estuarius with overlying water salinity of 20 7tr: <2 to 34 7((
(DeVVitt et al.. 1989; SAIC, 1993b); Leptocheirus plumulosus with overlying salinity of
20 Vc <1.5 to 32 ?rt (Schlekat et al., 1992; SAIC, 1993b) and Rhepoxynius abronius with
overlying water salinity of 28 to 32 7rr. 25 to 34 7f( (Swartz et al., 1985: Lamberson and
Swartz. 1988).
11.4.5 Effects of Sediment-associated Ammonia. Field-collected sediments may contain
concentrations of ammonia that are toxic to amphipods. Water column no effect
concentrations for the four amphipod species are presented in Table 11.4. If ammonia
concentrations are above these values, mortality occurring after 10 d may be due in part
to effects of ammonia. Depending on test application, it may be desirable to lower the
ammonia concentration by manipulating the test system prior to introduction of test
organisms if measured ammonia in the overlying water is greater than the species-
specific no effect concentration. If sediment toxicity tests are conducted to evaluate the
acceptability of dredge material for disposal, the manipulations must be performed.
Manipulations involve flushing the test system by renewing a specified amount of
overlying water for up to two consecutive 24 h periods.
11.4.5.1 If ammonia is of concern to the regulatory application associated with the
sediment toxicity test, overlying water should be sampled approximately 1 cm above the
sediment surface prior to introduction of animals on Day 0. If overlying water ammonia
concentration are less than or equal to the species-specific no effect concentration listed
in Table 11.4, then the test may proceed normally. If overlying water ammonia
concentration is greater than the species-specific no effect concentration listed in
Table 11.4, then the test system must be flushed for 24 h at a rate of 6 volume
replacements/24 h.
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(600/R-94/025)
Errata
Pages 80-82, Sections 11.4.5-11.4.5.3, Effects of Sediment-
associated Ammonia
These sections describe a procedure that can be used to reduce
ammonia concentrations in field-collected sediments prior to
conducting laboratory toxicity tests. For dredged material
testing under the Clean Water Act or the Marine Protection,
Research, and Sanctuaries Act, the following alternative
procedure should be used. This procedure was described in a
December 21, 1993 guidance memorandum issued by the U.S. EPA
Office of Wetlands, Oceans and Watersheds, U.S. EPA Office of
Science and Technology, and U.S. Army Corps of Engineers
Operations, Construction, and Readiness Division.
For dredged material testing the following procedure should be
used if it is necessary to reduce interstitial water ammonia
levels. Whenever chemical evidence of ammonia is present at
toxicologically important levels, and ammonia is not a
contaminant of concern, the laboratory analyst should reduce
ammonia in the sediment interstitial water to species-specific
no-effect concentrations (see table 11.4 on page 81). Ammonia
levels in the interstitial water can be reduced by sufficiently
aerating the sample and replacing two volumes of water per day.
The analyst should measure interstitial ammonia each day until it
reaches the appropriate species-specific no-effect concentration.
After placing the test organism in the sediment, the analyst
should ensure that ammonia concentrations remain within an
acceptable range by conducting the toxicity test with continuous
flow or volume replacement not to exceed two volumes per day.
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Table 11.4 Application limits for 10-d sediment toxicity tests with Ampelisca abdita, Eohaustorius estuarius, Leptocheirus
plumulosus, or Rhepoxynius abronius
Parameter
Ampelisca
abdita
Eohaustorius
estuarius
Leptocheirus
plumulosus
Rhepoxynius
abronius
Temperature (°C)
Overlying Salinity (»*)
Grain Size (% silt/clay)
Ammonia (total mg/I, pH 7.7)
Ammonia (HI1 mg/I, pH 7.7)
Sulfides
20
> 10
> 10
<30
<0.4
NA
15
0-34
full range
<60
<0.8
NA
25
1.5 - 32
full range
<60
<0.8
NA
15
>25
<90
<30
<0,4
NA
HI = unionized ammonia
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i 1.4.5.2 After 24 h, the overlying water ammonia concentration must be measured
again. If it is less than or equal to the species-specific no effect concentration, testing
should be initiated by adding animals. The system must be flushed at a rate of 6 volume
replacements/24 h over the course of the test. Overlying water ammonia should be
measured again on Day 10 of the test.
11.4.5.3 If after the initial 24 h flushing period (i.e., that described in 11.4.5.1) the
overlying water ammonia concentration is still greater than the species-specific no effect
concentration, the system must be flushed for again 24 h at a rate of 6
volumereplacements/24 h. After the second flushing, ammonia concentrations in the
overlying water should be measured again, and if concentrations are less than or equal
to the species-specific no effect concentration listed in Table 11.4, then the test may
proceed as described in Section 11.4.5.2. If overlying water ammonia concentrations still
exceed the species-specific no effect concentration, it must be concluded that ammonia
cannot be reduced to no effect concentrations without concern for flushing other
contaminants from the sediment At this point, the test should still be conducted as
described in Section 11.4.5.2 After 24 h, the overlying water ammonia concentration
must be measured again. If it is less than or equal to the species-specific no effect
concentration, testing should be initiated by adding animals. The system must be
flushed at a rate of 6 volume replacements/24 h over the course of the test. Overlying
water ammonia should be measured again on Day 10 of the test.
11.4.5.3 If after the initial 24 h flushing period (i.e., that described in 11.4.5.1) the
overlying water ammonia concentration is still greater than the species-specific no effect
concentration, the system must be flushed for again 24 h at a rate of 6 volume
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Section 12
Data Recording, Data Analysis and Calculations, and Reporting
12.1 Data Recording
12.1.1 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.
12.1.2 A file should be maintained for each sediment test or group of tests on closely
related samples (Section 9). This file should contain a record of the sample chain-of-
custody; a copy of the sample log sheet; the original bench sheets for the test organism
responses during the sediment test(s); chemical analysis data on the sample(s): control
data sheets for reference toxicants; detailed records of the test organisms used in the
test(s), such as species, source, age, date of receipt, and other pertinent information
relating to their history and health; information on the calibration of equipment and
instruments; test conditions used; and results of reference toxicant tests. Original data
sheets should be signed and dated by the laboratory personnel performing the tests.
12.1.3 Example data sheets are included in Appendix A.
12.2 Data Analysis
12.2.1 Statistical methods are used to make inferences about populations, based on
samples from those populations. In most sediment toxicity tests, test organisms are
exposed to contaminated sediment 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. In any toxicity.
summary statistics such as means and standard errors for response variables (e.g.,
survival) should be provided for each treatment (e.g., pore-water concentration,
sediment).
12.2.1.1 Types of data. Two types of data can be obtained from sediment toxicity tests.
The most common endpoint in toxicity testing is mortality, which is a dichotomous or
categorical type of data.
12.2.1.2 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 studies designed to: (1) evaluate hazards of
dredged material, (2) assess site contamination in the environment (e.g., to rank areas
for clean-up), and (3) determine effects of specific contaminants, or combinations of
contaminants, through the use of sediment spiking techniques. Each of these broad
study designs has specific statistical design and analytical considerations, which are
detailed below.
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12.2.1.2.1 Dredged Material Hazard Evaluation. In these studies, n sites are compared
individually to a reference sediment. The statistical procedures appropriate for these
studies are generally pairwise comparisons. Additional information on toxicity testing of
dredged material and analysis of data from dredged material hazard evaluations is
available in USEPA-USCOE (1994).
12.2.1.2.2 Site Assessment of Field Contamination. Surveys of sediment toxicity often are
included in more comprehensive analyses of biological, chemical, geological, and
hydrographic data. Statistical correlation can be improved and costs may be reduced if
subsamples are taken simultaneously for sediment toxicity, chemical analyses, and
benthic community structure determinations. There are several statistical approaches to
field assessments, each with a specific purpose. If the objective is to compare the
response or residue level at all sites individually to a control sediment, then the pairwise
comparison approach described below is appropriate. If the objective is to compare
among all sites in the study area, then a multiple comparison procedure that employs an
experiment-wise error rate is appropriate. If the objective is to compare among groups
of sites, then orthogonal contrasts are a useful data analysis technique.
12.2.1.2.3 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 LCSO, EC50,
IC50, NOEC, or LOEC. 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.
12.2.2 The guidance outlined below on the analysis of sediment toxicity test data is
adapted from a variety of sources including Lee et al. (1994), USEPA ('993a), USEPA
(1993b), USEPA (1993c), and USEPA-USCOE (1994). The objectives of a sediment
toxicity test is to quantify contaminant effects on test organisms exposed to natural or
spiked sediments or dredged materials and to determine whether these effects are
statistically different from those occurring in a control or reference sediment Each
experiment consists of at least two 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 experimental set-up 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 sediment that represents an environmental condition or
potential treatment effect of interest.
12.2.2.1 Experimental Unit. During toxicity testing, each test chamber to which a single
application of treatment is applied is an experimental unit. 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.
12.2.2.2 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
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variation and provides an estimate of within-treatment error for assessing the
significance of observed differences between treatments.
12.2.2.3 Minimum Detectable Difference (MDD). 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 significance level and power increases. Because no
consensus currently exists on what constitutes a biologically acceptable MDD, 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 test results.
12.2.2.4 Minimum number of replicates. Four replicates per treatment or control are the
absolute minimum number of replicates for a sediment toxicity test However, USEPA
recommends five replicates for marine testing or eight replicates for freshwater testing
(USEPA, 1994a) for each control or experimental treatment. It is always prudent to
include as many replicates in the test design as are economically and logistically possible.
USEPA sediment toxicity testing methods recommend the use of 20 organisms per
replicate for marine testing or 10 organisms per replicate for freshwater testing
(USEPA, 1994a). An increase in the number of organisms per replicate in all
treatments, including the control, is allowable only if: (I) test performance criteria for
the recommended number of replicates are achieved and (2) it can be demonstrated that
no change occurs in contaminant availability due to the increased organism loading.
12.2.2.5 Randomization. Randomization is the UKbiased 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 the largest animals are placed in
the same treatment), (2) randomize the allocation of sediment (e.g., not take all the
sediment in the top of a jar for the control and the bottom for spiking), and (3)
randomize the location of exposure units.
12.2.2.6 Pseudoreplication. The appropriate assignment 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 treatments are not
replicated or the replicates are not statistically independent (Hurlbert, 1984). The
simplest form of pseudoreplication is the treatment of subsamples of the experimental
unit as true replicates. For example, two aquaria are prepared, one with control
sediment, 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
biological response and do not replicate the treatment (i.e., sediment type). In this case,
the experimental unit is the 10 organisms and each organism is a subsample. A less
obvious form of pseudoreplication is the potential systematic error due to the physical
segregation of exposure chambers by treatment. For example, if all the control exposure
chambers are placed in one area of a room and all the test exposure chambers are in
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another, spatial effects (e.g., different lighting, temperature) could bias the results for
one set of treatments. Random physical intermixing of the exposure chambers or
randomization of treatment location may be necessary to avoid this type of
pseudoreplication. 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.
12.2.3 The purpose of a toxicity test is to determine if the biological response to a
treatment sample differs from the response to a control sample. Table 12.1 presents the
possible outcomes and decisions that can be reached in a statistical test of such a
hypothesis. The null hypothesis is that no difference exists among the mean control and
treatment responses. The alternative hypothesis of greatest interest in sediment tests is
that the treatments are toxic, or contain concentrations of bioaccumulable compounds,
relative to the control or reference sediment.
12.2.3.1 Statistical tests of hypotheses can be designed to control for the chances of
making incorrect decisions. In Table 12.1, 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 making a Type I error. Historically, having chosen a, environmental
researchers have ignored P and the associated power of the test (1-fi).
12.2.3.2 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 analysis. The
consequences of a Type II statistical error in environmental studies should never be
ignored and may in fact be the most important criteria to consider in experimental
designs and data analyses which include statistical hypothesis testing. To paraphrase
Fairweather (1991), "The commitment of time, energy and people to a false positive (a
Type I error) will only continue until the mistake is discovered. In contrast, the cost of
a false negative (a Type II error) will have both short- and long-term costs (e.g., ensuing
environmental degradation and the eventual cost of its rectification)."
12.2.3.3 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, (3) the number of replicate units for the treatment and control
samples, (4) the number of animals exposed within a replicate exposure chamber, and
(5) the selected probabilities of Type I (a) and Type II (|3) errors.
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Table 12.1 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 Design Is
Balanced
0.10
0.05
0.01
0.10
0.05
Unbalanced2
0.25
0.10
0.05
0.25
0.10
1 N = total number of observations (replicates) in all treatments combined; n
number of observations (replicates) in an individual treatment
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12.2.3.4 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 (5. The MOD
should be established ahead of time based upon biological and program considerations.
The investigator has little control of the variance among replicate exposure chambers.
However, this variance component can be minimized by selecting test organisms that are
as biologically similar as possible and maintaining test conditions within prescribed
quality control (QC) limits.
12.2.3.5 The MDD 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 parametric analysis. If the desired MDD, 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 12.1 to determine how likely it is that a 20% reduction will be detected in the
treatment response relative to the control response. The CV is defined as 100% x
(standard deviation divided by the mean). In a test design with 8 replicates per
treatment and with an a level of 0.05, high power (i.e., >0.80) to detect a 20% reduction
from the control mean occurs only if the CV is 15% or less (Figure 12.1). The choice of
these variables also affects the power of the test. If 5 replicates are used per treatment
(Figure 12.2), 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%.
12.2.3.6 Relaxing the a level of a statistical test increases the power of the test
Figure 12.3 duplicates Figure 12.1 except that a is 0.10 instead of 0.05. Selection 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 12.1 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 12.3) and the CV remains at 15%, then there is a 90% probability (power) of
detecting a 20% reduction relative to the control mean. The latter example would be
preferable if an environmentally conservative analysis and interpretation of the data is
desirable.
12.2.3.7 Increasing the number of replicates per treatment will increase the power to
detect a 20% reduction in treatment response relative to the control mean (Figure 12.4).
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 treatment, 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 uniform organisms to reduce biological variability or increasing
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 response relative to the
control mean.
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TR = Control TR > Control
DECISION
TR = Control
TR > Control
Correct
1 - a
Type I
Error
a
Type II
Error
P
Correct
(Power)
NOTE: Treatment response (TR), alpha (a) represents the
probability of making a Type I statistical error (false positive); beta
((3) represents the probability of making a Type II statistical error
(false negative).
Figure 12.1 Treatment response for a Type I and Type II error.
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10
20 30 40
% Reduction of Control Mean
50
60
70
Figure 12.2 Power of the test vs percent reduction in treatment response relative to
the control mean at various CV's (8 replicates, alpha = 0.05 (one-tailed)).
90
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o
Q.
0.9 --
0.8 --
0.7 --
0.6 --
0.5 --
0.4 --
0.3 --
0.2 --
0.1 --
= 40%
10
20 30 40
% Reduction of Control Mean
50
60
70
Figure 12.3 Power of the test vs percent reduction in treatment response relative to
the control mean at various CV's (5 replicates, alpha = 0.05 (one-tailed)).
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o
Q.
0.9 --
0.8
0.7 --
0.6 --
0.5 --
0.4 --
0.3 --
0.2 --
0.1 T
0 t
-4-
10
20 30 40
% Reduction of Control Mean
50
60
70
Figure 12.4 Power of the test vs percent reduction in treatment response relative to
the control mean at various CV's (8 replicates, alpha = 0.10 (one-tailed)).
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12.2.3.8 The effect of the choice of a and (3 on number of replicates for various CV's is
illustrated in Figure 12.5 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 (3
of 0.15. In Figure 12.5, if a = (3 = 0.125, the number of replicates required to detect a
difference of 20% relative to the control is at a minimum. As a or p decrease, the
number of replicates required to detect the same 20% difference relative to the control
increases. However, the curves are relatively flat over the range of 0.05 to 0.20 and that
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
pre-selected percentage reduction in mean treatment response relative to the control
mean.
12.2 .4 Figure 12.6 outlines a decision tree for analysis of survival and growth data
subjected to hypothesis testing. In the tests described herein, samples or observations
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 non-parametric statistics text such as Conover (1980) may also be helpful.
12.2.4.1 Mean. The sample mean (x) is the average value, or Zx/n, where:
n = number of observations (replicates)
Xj = ith observation
ZXj = every x summed = Xj + x2 + x3 + . . . + xn
n
12.2.4.2 Standard Deviation. The sample standard deviation (s) is a measure of the
variation of the data around the mean and is equivalent to W. The sample variance, s2,
is given by the following "machine" or "calculation" formula:
2 _
d
n - 1
12.2.4.3 Standard Error of the Mean. The standard error of the mean (SE, or s/ifn)
estimates variation among sample means rather than among individual values. The SE
is an estimate of the SD among means that would be obtained from several samples of n
observations each. Most of the statistical tests in this manual compare means with other
means (e.g., dredged sediment mean with reference mean) or with a fixed standard (e.g.,
FDA action level; Lee et ah, 1994). Therefore, the "natural" or "random" variation of
sample means (estimated by SE), rather than the variation among individual obser-
vations (estimated by s), is required for the tests.
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I
o
Q.
0.2
8 10
No. of Replicates (n)
12
14
16
Figure 12.5 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)).
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Alpha (Betas 0.25-Alpha)
Figure 12.6 Effect of alpha and beta on the number of replicates at various CV's
(assuming combined alpha + beta = 0.25).
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12.2.4.4 Tests of Assumptions. In general, parametric statistical analyses such as t-tests
and analysis of variance are appropriate only if: (1) there are independent, replicate
experimental units for each treatment, (2) the observations within each treatment follow
a normal distribution, and (3) variances for both treatments are equal or similar. The
first assumption is an essential component of experimental design. The second and third
assumptions can be tested using the data obtained from the experiment. Therefore,
before conducting statistical analyses, tests for normality and equality of variances
should be performed.
12.2.4.4.1 Outliers (extreme values) and systematic departures from a normal
distribution (e.g., a log-normal distribution) are the most common causes of departures
from normality or equality of variances. An outlier is an inconsistent or questionable
data point that appears unrepresentative of the general trend exhibited by the majority
of the data. Outliers may be detected by tabulation of the data, plotting, or by analysis
of residuals. An explanation should be sought for any questionable data points.
Without an explanation, data points should only be discarded with extreme caution. If
there is no explanation, the analysis should be performed both with and without the
outlier, and the results of both analyses should be reported. An appropriate
transformation, such as the arcsine square root transformation, will normalize many
distributions (USERA, 1985). Problems with outliers can usually be solved only by using
non-parametric tests, but careful laboratory practices can reduce the frequency of
outliers.
12.2.4.4.2 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 observations so that the mean of residuals for each treatment and over all
treatments is zero. The Shapiro-Wilk's Test provides a test statistic W, which is
compared to values of W expected from a normal distribution. W will generally vary
between 0.3 and 1.0, with lower values indicating greater departure from normality.
Because normality is desired, one looks for a high value of W with an associated
probability greater than the pre-specified a level.
12.2.4.4.3 Table 12.2 provides a levels to determine whether departures from normality
are significant. Normality should be rejected when the probability associated with W
(or other normality test statistic) is less than a for the appropriate total number of
replicates (N) and design. A balanced design means that all treatments have an equal
number (n) of replicate exposure chambers. A design is considered unbalanced when
the treatment with the largest number of replicates (nmax) has at least twice as many
replicates as the treatment with the fewest replicates (nmin). Note that higher a levels are
used when the number of replicates is small, or when the design is unbalanced, because
these are the cases in which departures from normality have the greatest effects on t-
tests and other parametric comparisons. If data fail the test for normality, even after
transformation, nonparametric tests should be used for additional analyses.
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12.2.4.4.4 Tables of quantiles of W can be found in Shapiro and Wilk (1965), Gill
(1978), Conover (1980), USEPA (1989) and other statistical texts. These references also
provide methods of calculating W, although the calculations can be tedious. For that
reason, commonly available computer programs or statistical packages are preferred for
the calculation of W.
12.2.4.4.5 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 (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).
12.2.4.4.6 If no tests for equality of variances are included in the available statistical
software, Hartley's Fmax can easily be calculated:
Fmax = ( larger of sf , s\ )/( smaller of s\ , si )
When Fmax is large, the hypothesis of equal variances is more likely to be rejected. Fmax
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).
12.2.4.4.7 Levels of a for tests of equality of variances are provided in Table 12.2.
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.
12.2.4.4 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 arcsine-square root transformation. The
arcsine-square root transformation is commonly used on proportionality data to stabilize
the variance and satisfy the normality requirement. If the data do not meet the
assumption of normality and there are four or more replicates 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 test are adjusted.
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12.2.4.5.1 The arcsine-square root transformation consists 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 arcsine-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 arcsine, as follows.
a. For RPs greater than zero or less than one:
Angle (in radians) = arc sine^/(RP)
b. Modification of the arcsine when RP = 0.
Angle (in radians) = arcsine \ —
\ 4n
where n - number animals/treatment replicate.
c. Modification of the arcsine when RP = 1.0.
Angle = 1.5708 radians-(radians for RP = 0)
12.2.6.5 Two Sample Comparisons (N=2). The true population mean (u) and standard
deviation (a) are known only after sampling the entire population. In most cases
samples are taken randomly from the population, and the s calculated from those
samples is only an estimate of a. Student's t-values account for this uncertainty. The
degrees of freedom for the test, which are defined as the sample size minus one (n-1),
should be used to obtain the correct t-value. Student t-values decrease with increasing
sample size because larger samples provide a more precise estimate of u and 0.
12.2.4.6.1 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
hypothesis, 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 (HJ 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
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larger or smaller). A one-tailed test is used when there is an a priori reason to test for a
specific relationship between two means such as the alternative hypothesis that the
treatment mortality or tissue residue is greater than the control mortality or tissue
residue. In contrast, the two-tailed test is used when the direction of the difference is
not important or cannot be assumed before testing.
12.2.4.6.2 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
12.2.4.6.3 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.
12.2.4.6.4 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.
12.2.4.7 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 equality of
means x} and x2 from two independent samples with n, and n2 replicates and unequal
variances is:
where s] and si are the sample variances of the two groups. Although the equation
assumes that the variances 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 t distribution with degrees of freedom (df) given by Satterth wake's
(1946) approximation:
/ Oh - 1) + (slln^ I («2 - 1)
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
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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, the t-test with Bonferroni's adjustment can be used for data analysis
(USEPA, 1994b; USEPA, 1994c). When variances are equal;, an F test for equality is
unnecessary.
12.2.4.8 Nonparametric Tests. Tests such as the t-test, which analyze the original or
transformed data, and which rely on the properties of the normal distribution, are
referred to as parametric tests. Nonparametric tests, which do not require normally
distributed data, analyze the ranks of data and generally compare medians 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, nonparametric 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.
12.2.4.8.1 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 assumption is not met, data should be converted to normalized ranks
(rankits). Rankits are simply the z-scores expected for the rank in a normal
distribution. Thus, using rankits imposes a normal distribution 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 formula:
ranku =
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).
12.2.4.8.2 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 unequal variances using the rankits or ranks.
12.2.4.9 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 evaluating pair wise comparisons. After the applicable comparisons are
determined, the data must be tested for normality to determine if parametric statistics
are appropriate and whether the variances of the treatments are equal. If normality of
the data and equal variances are established, then an analysis of variance (ANOVA) may
be performed to address the hypothesis that all the treatments including the control are
equal. If normality or equality of variance are not established then transformations of
the data may be appropriate or nonparametric statistics can be used to test for equal
means. Tests for normality of the data should be performed on the treatment residuals.
A residual is defined as the observed value minus the treatment mean, that is, rik = oik -
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(kth treatment mean). Pooling residuals provides an adequate sample size to test the
data for normality.
12.2.4.9.1 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 which 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
A NOVA to get an estimate of the within treatment variance. After a variance estimate
is obtained, the means of the treatments with 0 variance may be tested against the other
treatment means using the appropriate mean comparison. Equality of variances among
the treatments can be evaluated with the Hartley FmM test or Bartlett's test. The option
of using nonparametric statistics on the entire set of data is also an alternative.
12.2.4.9.2 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 requires 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 normality assumption
has not been met. Steel's test requires equal variances across treatments and the
control, but is thought to be fairly insensitive to deviations from this condition (USEPA,
1993a). Wilcoxon's Rank Sum Tests is a nonparameteric test to be used as an
alternative to the Steel's test when the number of replicates are not the same within
each treatment A Bonferroni's adjustment of the pairwise error rate for comparison of
each treatment versus the control is used to set an upper bound of alpha on the overall
error rate. This is in contrast to the Steel's test with a fixed overall error rate for
alpha. Thus, Steel's tests is a more powerful test (USEPA, 1993a).
12.2.4.9.3 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 individual 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 particular sample to the control or reference.
However, if several samples were taken from the same area and the decision to accept or
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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.
12.2.4.9.4 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 comparison 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 recommended for comparisons with the control.
12.2.4.9.5 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.
12.2.4.9.6 To perform the individual comparisons, calculate the t statistic for each
treatment and control combination, as follows:
where Y( = Mean for each treatment
Yj = Mean for the control
Su = 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 Procedure
Sw = The square root of the within mean square
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n = The number of replicates per treatment, assuming an equal number of
replicates at all treatment concentrations
n, = Number of replicates in the control
12.2.5 Methods for Calculating LCSOs, ECSOs, and ICps.
12.2.5.1 Figure 12.8 outlines a decision tree for analysis of point estimate data. USEPA
(USEPA, 1985; USEPA, 1989b; USEPA, 1994b; USEPA, 1994c) discuss in detail the
mechanics of calculating LC50 (or EC50) or ICp values using the most current methods.
The most commonly used methods are the Graphical, Probit, Trimmed Spearman-
Karber and the Linear Interpolation Methods. In general, results from these methods
should yield similar estimates. Each method is outlined below and recommendations
presented for the use of each method.
12.2.5.2 Data for at least five test concentrations and the control should be available to
calculate an LC50 although each method can be used with fewer concentrations.
Survival in the lowest concentration must be at least 50% and an UC50 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.
12.2.5.3 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
recommended that the data be analyzed with the aid of computer software. A computer
program to estimate the LC50 values and associated 95% confidence intervals with the
methods discussed below (except for the Graphical Method) was 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.
12.2.5.4 The 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 (log,,,) of treatment concentration. The only requirement for its
use is that treatment mortalities bracket 50%.
12.2.5.4.1 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 p(l, p,, ..., pk denote the observed
proportion mortalities for the control and the k treatments. The first step is to smooth
the p; if they do not satisfy p,, < p, < ... < pk. The smoothing process replaces any
adjacent p:'s that do not conform to pn < p, < ... < pk with their average. For example, if
Pi is less than pM then:
where p) = the smoothed observed proportion mortality for concentration i.
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Data—Survival, Growth, Etc.
Test for Normality
Normal
Shapiro-Wilk's Test (N<50)
Non-Normal
Tests for Homogeneity of Variance
1 Transformation?
1 Barttett's |j Hartley's
Heterogenous Variances
1 Rankits
No
Homogenous Variances
Yes, N>2 +
No, N-2
>3 Replicates
Yes
[ Anova )
Equal Repl
1 No
! Bonterroni's 1
1
cation
Yes
1—^
Comparison- Wise Alpha
Fisher's LSD, Duncan's
hxpenment-Wise Alpha
Dunnett's
-^
*<
\
Equal Replication
jv«
Steel's
Many-One
Rank Test
i _
XNO
Wilcoxon
w/
Bonferroni
1
1 tndpoint
s
Figure 12.7 Decision tree for analysis survival and growth data subjected to hypothesis
testing.
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Adjust the smoothed observed proportion mortality in each treatment for mortality in
the control group using Abbott's formula (Finney, 1971). The adjustment takes the
form:
where p," = the smoothed observed proportion mortality for the control
p\ = the smoothed observed proportion mortality for concentration i.
12.2.5.5 The Probit Method. The Probit Method is a parametric statistical procedure
for estimating the LC50 (or EC50) and the associated 95% confidence interval (Finney,
1978). The analysis consists of transforming the observed proportion mortalities with a
probit transformation, and transforming the treatment concentrations to log,,,. Given
the assumption of normality for the log,,, of the tolerances, the relationship between the
transformed variables mentioned above is about linear. This relationship allows
estimation of linear regression parameters, using an iterative approach. A probit is the
same as a z-score: for example, the probit corresponding to 7095- mortality is z70 or =.52.
The LC50 is calculated from the regression and is the concentration associated with 50%
mortality or z=0. To obtain a reasonably precise estimate of the LC50 with the Probit
Method, the observed proportion mortalities must bracket 0.5 and the log,,, of the
tolerance should be normally distributed. To calculate the LC50 estimate and associated
95% confidence interval, two or more of the observed proportion mortalities must be
between zero and one. The original percentage mortalities should be corrected for
control mortality using Abbott's formula before the Probit transformation is applied to
the data.
12.2.5.5.1 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 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 interval estimates
on the LC50 may not be correct if replicate mortalities are pooled to obtain a mean
treatment response. This can be avoided by entering the probit-transformed replicate
responses and doing a least squares regression on the transformed data.
12.2.5.6 The Trimmed Spearman-Karber Method. The Trimmed Spearman-Karber
Method is a modification of the Spearman-Karber. non-parametric statistical procedure
for estimating the LC50 and the associated 95% confidence interval (Hamilton et al.,
1977). This procedure estimates the trimmed mean of the distribution of the log,,, 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 (I'SEPA. 1994b; liSEPA, 1994c). This method is only
appropriate for lethality data sets.
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Survival Point Estimates
f
Two or More Partial Mortalities
Yes J. No
Significant Chi Square Test
Yes | No
One Partial Mortality
Yes
Probit
No
Graphical
{linear Interpolation]
Trimmed Spearman-Karber 1
LC50 and 95% Confidence Intervals
Figure 12.8 Decision tree for analysis of point estimate data.
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12.2.5.6.1 To calculate the LC50 estimate with the Trimmed Spearman-Karber Method,
the smoothed, adjusted, observed proportion mortalities must bracket 0.5. To calculate
a confidence interval for the LC50 estimate, one or more of the smoothed, adjusted,
observed proportion mortalities must be between zero and one.
12.2.5.6.2 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). Calculate
the amount of trim to use in the estimation of the LC50 as follows:
Trim = max(p", 1 - p£)
where p, = the smoothed, adjusted proportion mortality for the lowest treatment
concentration, exclusive of the control.
p£ = the smoothed, adjusted proportion mortality for the highest treatment
concentration.
k = the number of treatment concentrations, exclusive of the control.
12.2.5.7 The Linear Interpolation Method. This method calculates a toxicant
concentration that causes a given percent reduction (e.g., 25%, 50%, etc.) in the
endpoint of interest and is reported as an ICp value (1C = Inhibition Concentration;
where p = the percent effect). The procedure was 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 precision 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.
12.2.5.7.1 As described in USEPA (USEPA, 1994b; USEPA, 1994c), the Linear
Interpolation Method of calculating an ICp assumes that the responses: (1) are
monotonically nonincreasing, where the mean response for each higher concentration is
less than or equal to the mean response for the previous concentration, (2) follow a
piecewise linear response function, and (3) are from a random, independent, and
representative sample of test data. If the data are not monotonically nonincreasing, they
are adjusted by smoothing (averaging). In cases where the responses at the low toxicant
concentrations are much higher than in the controls, the smoothing process may result
in a large upward adjustment in the control mean. In the Linear Interpolation Method,
the smoothed response means are used to obtain the ICp estimate reported for the test.
No assumption is made about the distribution of the data except that the data within a
group being resampled are independent and identically distributed.
12.2.5.7.2 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 response of interest, the (p) percent
reduction from the control.
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12.2.5.7.3 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 concentration 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, it
is averaged with the control, and this average is used for both the control response (M,)
and the lowest toxicant concentration response (M2). This mean is then compared to the
mean observed response for the next higher toxicant concentration (Y3). Again, if the
mean observed response for the next higher toxicant concentration is smaller than the
mean of the control and the lowest toxicant concentration, it is used as the response. If
it is higher than the mean of the first two, it is averaged with the 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 deviations from monotonicity may require an additional step of
smoothing. Where Y( decrease monotonically, the Ys become M, without smoothing.
12.2.5.7.4 To obtain the ICp estimate, determine the concentrations Cj and CJ+I which
bracket the response Mt (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:
(C, +, - C,)
ICp = Cj + ( M1 (1 J + l J
where Cj = tested concentration whose observed mean response is greater than
M,(l - p/100).
Cj ^, = tested concentration whose observed mean response is less than Mj(l -
p/100).
M, = smoothed mean response for the control.
Mj = smoothed mean response for concentration J.
Mj t, = smoothed mean response for concentration J + 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.
12.2.5.7.5 Standard statistical methods for calculating confidence intervals are not
applicable for the ICp. The bootstrap method, as proposed by Efron (1982), is used to
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obtain the 95% confidence interval for the true mean. In the bootstrap method, the test
data YJJ is random!}' resampled with replacement to produce a new set of data Y^*, that
is statistically equivalent to the original data, but which produces a new and slightly
different estimate of the iCp (iCp*). This process is repeated at least 80 times (Marcus
and Holtzman, 1988) resulting in multiple "data" sets, each with an associated ICp*
estimate. The 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 quantiies of the ICp* empirical distribution.
For example, if the test data are resampled a minimum of 80 time, the empirical 2.5%
and the 97.5% confidence limits are about the second smallest and second largest ICp*
estimates (Marcus and Holtzman, 1988). The width of the confidence intervals
calculated by the bootstrap method is related to the variability of the data. When
confidence intervals are wide, the reliability of the 1C estimate is in question. However.
narrow intervals do not necessarily indicate that the estimate is highly reliable, because
of undetected violations of assumptions and the fact that the confidence limits based on
the empirical quantiies of a bootstrap distribution of 80 samples may be unstable.
123 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 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 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).
12.3.2 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 sediments (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 bioavailability of non-ionic organic compounds are often
inversely correlated with the organic carbon concentration 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 between solid and liquid phases of sediment
may be useful for establishing effect concentrations.
12.3.3 Toxic units can be used to help interpret the response of organisms to multiple
contaminants in sediment. A toxic unit is the concentration of a chemical divided by an
effect concentration. For example, a toxic unit of exposure can be calculated by dividing
the measured concentration of a chemical in pore water by the water-only LC50 for the
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same chemical (Ankley et al., 1991a). Toxicity expressed as toxic units may be summed
and this may provide information on the toxicity of chemical mixtures (Ankley et al.,
1991a).
12.3.4 Field surveys can be designed to provide either a qualitative reconnaissance of
the distribution of sediment contamination or a quantitative statistical comparison of
contamination among sites (Burton and Ingersoll, 1994). Surveys of sediment toxicity
are usually part of more comprehensive analyses of biological, chemical, geological, and
hydrographic data. Statistical correlation can be improved and costs reduced if
subsamples are taken simultaneously for sediment toxicity or bioaccumulation tests,
chemical analyses, and benthic community structure.
12.3.5 Descriptive methods such as toxicity tests with field-collected sediment should not
be used alone to evaluate sediment contamination. An integration of several methods
using the weight of evidence is needed to assess the effects of contaminants associated
with sediment. Hazard evaluations integrating data from laboratory exposures, chemical
analyses, and benthic community assessments provide strong complementary evidence of
the degree of pollution-induced degradation in aquatic communities (Chapman et al.,
1992; Burton, 1991).
12.3.6 Toxicity Identification Evaluation (TIE) procedures can be used to help provide
insights as to specific contaminants responsible for toxicity in sediment (USEPA, 1991a;
Ankley and Thomas, 1992). For example, the toxicity of contaminants such as metals,
ammonia, hydrogen sulfide, and non-ionic organic compounds can be identified using
TIE procedures.
12,4 Reporting
12.4.1 The record of the results of an acceptable sediment test should include the
following information either directly or by referencing available documents:
12.4.1.1 Name of test and investigator(s), name and location of laboratory, and dates of
start and end of test
12.4.1.2 Source of control or test sediment, method for collection, handling, shipping,
storage and disposal of sediment.
12.4.1.3 Source of test material, lot number if applicable, composition (identities and
concentrations of major ingredients and impurities if known), known chemical and
physical properties, and the identity and concentration^) of any solvent used.
12.4.1.4 Source and characteristics of overlying water, description of any pretreatment,
and results of any demonstration of the ability of an organism to survive or grow in the
water.
12.4.1.5 Source, history and age of test organisms; source, history and age of brood
stock, culture procedures; and source and date of collection of the test organisms,
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scientific name, name of person who identified the organisms and the taxonomic key
used, age or life stage, means and ranges of weight or length, observed diseases or
unusual appearance, treatments, holding procedures.
12.4.1.6 Source and composition of food, concentrations of test material and other
contaminants, procedure used to prepare food, feeding methods, frequency and ration.
12.4.1.7 Description of the experimental design and test chambers, the depth and
volume of sediment and overlying water in the chambers, lighting, number of test
chambers and number of test organisms/treatment, date and time test starts and ends.
temperature measurements, dissolved oxygen concentration (as percent saturation) and
any aeration used before starting a test and during the conduct of a test.
12.4.1.8 Methods used for physical and chemical characterization of sediment.
12.4.1.9 Definition(s) of the effects used to calculate LC50 or ECSOs, biological
endpoints for tests, and a summary of general observations of other effects.
12.4.1.10 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.
12.4.1.11 Methods used for statistical analyses of data.
12.4.1.12 Summary of general observations on other effects or symptoms.
12.4.1.13 Anything unusual about the test, any deviation from these procedures, and
any other relevant information.
12.4.2 Published reports should contain enough information to clearly identify the
methodology used and the quality of the results.
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Section 13
Precision and Accuracy
13.1 Determining Precision and Accuracy
13.1.1 Precision is a term that describes the degree to which data generated from
replicate measurements differ and reflects the closeness of agreement between randomly
selected test results. Accuracy is the difference between the value of the measured data
and the true value and is the closeness of agreement between an observed value and an
accepted reference value. 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 which 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, accuracy of the test methods has not been determined (Section 13.2).
13.1.2 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 13.5.1) precision or multi-laboratory
(interlaboratory or reproducibility: Section 13.5.2) precision. Intralahoratory 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 reproducible a method is when conducted by a large number of laboratories using
the same method, organism, and samples. Generally, intralaboratory results are less
variable than interlaboratory results (USEPA, 1991b; ISEPA. 1993a: USEPA, 1994b;
l.'SEPA, 1994c: Hall et al., 1989; Grothe and Kimerle, 1985).
13.1.3 A measure of precision can be calculated using the mean and relative standard
deviation (percent coefficient of variation, or CV% = standard deviation/mean x 100) of
the 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 hypothesis testing. The CVs may be very high when testing extremely toxic samples.
For example, if there are multiple replicates with no survival and one with low survival
the CV may exceed 100'7r. yet the range of response is actually quite consistent.
Therefore, additional estimates of precision should be used, such as range of responses
and minimum detectable differences (MDD) compared to control survival or growth.
Several factors can affect the precision of the test, including test organism age, condition,
sensitivity, handling of the test organisms, overlying water quality, and the experience of
the investigators in conducting tests. For these reasons, it is recommended that trained
laboratory personnel conduct the tests in accordance with the procedures outlined in
Section 9. Quality assurance practices should include: (1) single laboratory precision
determinations using reference toxicants for each of the test organisms which 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
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routinely performed as long as whole sediment tests are being conducted; (2) control
charts (Section 13.3) 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 (Table 11.3;
LSEPA, 1991 b).
13.1.4 Intralaboratory precision data are routinely calculated for test organisms using
water-only 96-h exposures to a reference toxicant, such as CdCl;. Intralaboratory
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). However, each laboratory's reference toxicant
CVs should reflect good repeatability.
13.1.5 To date, two interlaboratory precision (round-robin) tests have been completed
using 10-d whole sediment tests, one with Rhepoxynius abronius (Mearns et al., 1986).
and the other with Ampelisca abdita, Eohaustorius estuahus, and Leptocheirus plumulosus
(C. Schlekat, SAIC, Narragansett, RI, unpublished data). The results of these round-
robin study are described in Section 13.5.1.
13.2 Accuracy
13.2.1 The accuracy of toxicity tests cannot be determined since there is no acceptable
reference material. The relative accuracy of the reference toxicity tests can only be
evaluated by comparing test responses to control charts.
13.3 Replication and Test Sensitivity
13.3.1 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, the sensitivity of the test will increase
as the number of replicates is increased. The minimum recommended number of
replicates varies with the objectives of the test and the statistical method used for
analysis of the data (Section 12).
13.4 Demonstrating Acceptable Laboratory Performance
13.4.1 It is the responsibility of a laboratory to demonstrate its ability to obtain precise
results with reference toxicants before it performs sediment tests (Section 9.16).
Intralaboratory precision, expressed as a coefficient 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. This should
be done to gain experience for the toxicity tests and a point of reference for future
testing. 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).
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13.4.2 The quality of test organisms obtained from an outside source, regardless of
\vhether they are from culture or collected from the field, must be verified by
conducting a reference-toxicity test concurrently with the sediment test. For cultured
organisms, the supplier should provide data with the shipment describing the history of
the sensitivity of organisms from the same source culture. For field-collected organisms,
the supplier should provide data with the shipment describing the collection location, the
water salinity and temperature at the time of collection, and collection site sediment for
holding and acclimation purposes. 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 13.4.2).
13.4.3 Before conducting tests with contaminated sediment, the laboratory should
demonstrate its ability to conduct tests by conducting five exposures in control sediment
as outlined in Table 11.1. It is recommended that these five exposures with control
sediment be conducted concurrently with the five reference toxicity tests described in
Section 9.14.1.
13.4.4 A control chart should be prepared for each combination of reference toxicant
and test organism. End points 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 13.1), and the endpoint (LC50,
NOEC. ICp) are examined to determine if they are within prescribed limits. Control
charts as described in USEPA (1994a) and USEPA (1994b) 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 re-calculated with each successive test result. After two years
of data collection, or a minimum of 20 data points, the control chart should be
maintained using only the 20 most recent data points.
13.4.5 The outliers, which are values falling outside the upper and lower control limits,
and trends of increasing or decreasing sensitivity, are readily identified using control
charts. With an alpha of 0.05, one in 20 tests would be expected to fall outside of the
control limits by chance alone. During a 30 d period, if two reference toxicity tests out
of total previous 20 fall outside the control limits, the sediment toxicity tests conducted
during the that time in which the second reference toxicity test failed are suspect, and
should be considered as provisional and subject to careful review.
13.4.5.1 A sediment test may be acceptable if specified conditions of a reference toxicant
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 rage or if mortality in the control of the reference toxicity test
exceeds 10*7
-------
the overall credibility of the test system are suspect (USEPA, 1993a). In this case, the
test procedure should be examined for defects and should be repeated with a different
batch of test organisms.
13.4.7 Performance should improve with experience, and the control limits for point
estimates should gradually narrow. However, control limits of ±2 SD, by definition, will
be exceeded 5% of the time, regardless of how well a laboratory performs. Highly
proficient laboratories which develop a very narrow control limit may be unfairly
penalized if a test which 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.
13.4.8 The recommended reference toxicity test consists of a control and five or more
concentrations in which the endpoint is an estimate of the toxicant concentration which
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 12).
13.4.9 The point estimation analysis methods recommended 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 methods were considered in the selection process.
and it is recognized that the methods selected are not the only possible methods of
analysis for toxicity data.
13.5 Precision of Sediment Toxicity Test Methods
13.5.1 Intralaboratory Precision
13.5.1.1 Intralaboratory precision has not been evaluated for any of the four species.
13.5.2 Intel-laboratory Precision
13.5.2.1.1 Interlaboratory precision for R. abronius using 10-d whole sediment toxicity
tests using the methods described in this manual (Table 11.1) is described by Mearns et
al. (1986). Details of this study are described here. Five laboratories participated in the
study, including federal and state government laboratories, a contract laboratory, and an
academic laboratory. The laboratories were chosen because each had demonstrated
experience in sediment toxicity tests with R. abronius. The experimental design required
each laboratory to conduct 10-d whole sediment tests on a total of 7 sediment
treatments. One control sediment was tested. Three sediment treatments consisted of
control sediment that was amended with CdCl, to result in the following measured
concentrations: 4, 8, and 12 mg Cd/kg dry weight. Three field-collected sediments were
also used. They were collected from the following locations in Puget Sound, VYA:
Central Basin (Metro Seattle Station A600E), inner Sinclair Inlet, and Slip No. 1 in City
Waterway, Commencement Bay.
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13.5.2.1.2 Amphipods were collected from a depth of 6 m off West Beach, Whidbey
Island, WA, and distributed to each participating laboratory. Each laboratory used its
own source of clean seawater.
13.5.2.1.3 All five laboratories had >90% survival in control sediment, and thereby met
the performance criteria for the test Mean survival in control sediment was 96.4%, the
CV was 3.7%, and the range was from 92 to 100% (Table 13.1). Of the cadmium-
spiked sediments, survival was the least variable in the 4 mg/kg Cd treatment. Mean
survival was 96.2%, the CV was 4.2%, and the range was from 89 to 98%. The most
variable response was in the 12 mg/kg Cd sediment. Mean survival was 19%, the CV
was 79.1%, and the range was from 6 to 41%. City Waterway showed the least
variability among the field-collected sediments, with a mean survival of 83%, a CV of
6.4%, and a range from 74 to 87%. Sinclair Inlet showed the greatest variability among
the field-collected sediments, with a mean survival of 78.8%, a CV of 11.3%, and a
range from 67 to 88%.
13.5.2.2.1 Interlaboratory precision for A. abdita, E. estuarius, and L. plumules us using
10-d whole sediment toxicity tests is described by C. Schlekat (SAIC, Narragansett, Rl,
unpublished data). Details of this study are described below. The number of
participating laboratories varied with the test species: six for A. abdita, eight for
E. estuarius, and seven for L. plumulosus. Laboratories were chosen on the basis of
demonstrated experience with the particular test species. Each laboratory conducted 10-
d sediment toxicity tests on 4 sediment treatments. Sediment treatments were selected
for each species to include one negative control sediment and three contaminated
sediments. Highly contaminated sediment from Black Rock Harbor, CT, was diluted
with species-specific, non-contaminated control sediment, creating test sediments that
ranged in relative contamination from low to high.
13.5.2.2.2 Independent suppliers distributed amphipods to each laboratory. Ampelisca
abdita and Eohaustorius estuarius were field-collected from locations in Narragansett, Rl,
and Newport, OR, respectively. Leptocheirus plumulosus were obtained from cultures
located at the University of Maryland, Queenstown, MD. Each laboratory used its own
supply of clean seawater.
13.5.2.2.3 Mean survival of A. abdita in control sediment ranged from 85% to 100%
(Table 13.2). Five of the six laboratories achieved greater than 90% survival in control
sediment, which is the minimum survival that must be obtained in control sediment in
order for the test to be accepted. The grand mean was 94.5%, and the CV was 5.5. A
dose response was exhibited with decreasing survival with increasing proportions of
BRH sediment. Test sediments (i.e., 7%, 25, and 33% BRH dilutions) exhibited a higher
degree of variability than in control sediment. In 7% BRH sediment, mean survival
ranged from 20% in Laboratory 5 to 97% in Laboratory 6 (Table 13.2). Twenty-
percent BRH exhibited the greatest magnitude of variability, with a range of 1% to
90%. Thirty-three percent BRH also exhibited considerable variability. The overall
rank of sediment toxicity as measured by absolute mortality was consistent among
laboratories. One hundred percent of laboratories were in agreement for in
116
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Table 13.1 Inter-laboratory precision for survival of Rhepoxynius abronius in 10-d
whole sediment toxicity tests using seven sediments
Percent Survival
Lab
1
2
3
4
5
Mean
CV (%)
Control
92 (7)
96 (4)
100 (0)
94 (7)
100 (0)
96.4 (3.6)
3.7
4 mg/kjj
Cd
89(7)
98 (3)
97 (3)
99 (2)
98 (4.5)
96.2 (4.1)
4.2
8 mg/kK
Cd
87 (9)
90 (10)
78 (10.5)
50 (15)
77 (3)
76.4 (15.8)
20.7
(SD) in Sediment Samples
12 mn/kn
Cd
8 (3)
41 (11)
12 (7.5)
6 (5.5)
28(11.5)
19(15.5)
79.1
Central
Basin
83(11.5)
69 (7.5)
90 (8)
92 (5.5)
80 (3.5)
82.8 (9.1)
11.0
Sinclair
Inlet
78(13)
67(11)
87 (7.5)
88 (3)
74 (9)
78.8 (8.9)
11.3
Cit>
Waterway
74(11.5)
87(12)
83 (12.5)
84 (11)
87 (3)
83 (5.3)
6.4
Note: From Mearns et al., 1986
117
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Table 13.2 Inter-laboratory precision for survival of Ampelisca abdita in 10-d whole
sediment toxicity tests using four sediments
Percent Survival (SD) in Sediment Samples
Lab
1
2
3
4
5
6
Mean
CV (%)
Control
97.0 (4.5)
94.0 (8.9)
97.0 (4.5)
94.0 (8.9)
85.0 (7.1)
100.0 (0)
94.5 (5.2)
5.5
7% Black Rock
Harbor
63.0 (19.6)
75.0(6.1)
90.0 (3.5)
79.0 (17.8)
20.0(12.7)
97.0 (4.5)
70.7 (13.0)
38.9
20% Black Rock
Harbor
10.0 (7.9)
7.0 (4.5)
36.0 (9.6)
7.0 (AS)
1.0 (2.2)
90.0 (5.0)
25.2 (34.0)
135.1
33% Black Rock
Harbor
6.0 (4.2)
0.0 (0)
38.0(14.4)
3.0 (6.7)
1.0(2.2)
72.0(13.0)
20.0 (29.2)
146.2
Note: From: C. Schlekat et al., SAIC, Narragansett, RI, unpublished data.
118
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ranking control and 7% BRH sediments as the first and second least toxic sediments,
respectively (Table 13.2).
13.5.2.2.4 Every laboratory surpassed the minimum survival criteria of 90% survival in
control sediment with E. estuarius. The range was from 96 to 100%, with a Grand
Mean of 98.2% and a CV of 1.5 (Table 13.3). Grand Mean survival decreased with
increasing proportions of BRH. BRH sediment dilutions exhibited greater variability
than control sediment, with 25% BRH displaying the highest coefficient of variation. All
eight laboratories ranked survival of E. estuarius for control and 9% BRH as the least
and second least toxic, respectively (Table 13.2). With the exception of Laboratories 1
and 8, the rank for 25% and 42% BRH were appropriately third and fourth least toxic,
respectively.
13.5.2.2.5 Leptocheirus plumulosm exhibited a range of survival in control sediment
from 86% to 99% (Table 13.4). The Grand Mean was 91.8%, and the CV was 4.7.
Two laboratories, 3 and 5, failed to meet the minimum control sediment survival criteria
of 90%. Grand means displayed a dose response of decreasing survival with increasing
proportion of BRH sediment Coefficients of variation were uniformly higher in BRH
sediment dilutions as compared to control sediment, but did not vary greatly among
BRH sediments (Table 13.4). Laboratory 1 appeared to be an outlier with respect to
survival in BRH sediment dilutions, as survival of L. plumulosus was the lowest for all
three BRH sediments for any laboratory. The rank of sediments according to their
toxicity was generally consistent among laboratories. Agreement was 100% for control
and the highest BRH sediment; these were appropriately ranked 1 and 4, respectively
(Table 13.4). Laboratories 4 and 5 anomalously ranked 10% and 28% BRH as 3 and 2,
respectively, whereas the remaining laboratories ranked these sediments appropriately
according to the proportion of BRH.
13.5.2.3 These tests exhibited similar or better precision than many chemical analyses
and effluent toxicity test methods (USEPA, 1991b). The success rate for test initiation
and completion of this round-robin evaluation is a good indication that a well equipped
and trained staff will be able to successfully conduct this test. This is an important
consideration for any test performed routinely in any regulatory program.
119
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Table 13.3 Inter laboratory precision for survival of Eohaustorius estuarius in 10-d
whole sediment toxicity tests using four sediments
Lab
1
2
3
4
5
6
7
8
Mean
CV (%)
Percent Survival
(SD) in Sediment Samples
Control 9% Black
Rock Harbor
96.0 (6.5)
98.0 (2.7)
97.0 (2.7)
98.8 (2.7)
100.0 (0)
100.0 (0)
99.0 (2.2)
97.0 (6.7)
98.2 (1.5)
1.5
45.0 (19.7)
76.0(10.8)
89.0 (4.2)
59.0 (23.0)
75.0 (19.7)
69.0 (12.9)
79.0 (6.5)
53.0(14.4)
68.1 (14.7)
21.6
25% Black
Rock Harbor
6.0 (6.5)
46.0 (13,9)
59.0 (10.8)
47.2 (23.2)
36.0(12.4)
56.0 (18.8)
61.0 (10.8)
24.0 (14.7)
41.9(19.1)
45.5
42% Black
Rock Harbor
16.0 (9.6)
25.0(7.1)
45.0(10.0)
45.8 (27.0)
16.0 (9.6)
38.0(14.4)
50.0 (7.9)
29.0 (15.6)
33.1 (13.5)
40.9
Note: From: C. Schlekat et al., SAIC, Narragansett, Rl, unpublished data.
120
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Table 13.4 Inter-laboratory precision for survival of Leptocheirus plumulosus in 10-d
whole sediment toxicity tests using four sediments
Percent Survival (SD) in Sediment Samples
Lab
1
2
3
4
5
6
7
Mean
CV (%)
Control
91.3 (4.8)
91.0(8.9)
88.0 (8.4)
92.0 (7.6)
86.0 (10.2)
95.0(6.1)
99.0 (2.2)
91.8(4.3)
4.7
10% Black Rock
Harbor
6.0 (4.2)
62.0(11.0)
34.0 (15.2)
48.0 (23.9)
20.0 (9.4)
76.0 (10.2)
78.0 (13.0)
46.3 (27.7)
59.8
28% Black Rock
Harbor
5.0 (3.5)
51.0(15.6)
22.0(13.0)
59.0 (21.6)
28.0 (4.5)
65.0(14.6)
56.0 (4.2)
40.9 (22.6)
55.2
47% Black Rock
Harbor
2.5 (2.9)
33.0(11.5)
7.0 (5.7)
27.0(10.4)
12.0 (9.1)
38.0 (17.5)
26.0 (6.5)
20.8 (13.6)
65.5
Note: From: C. Schlekat et a!., SAIC, Narragansett, Rl, unpublished data.
121
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References
Adams, W.I'., R.A. Kimerle, and R.G. Mosher. 1985. An approach for assessing the
environmental safety of chemicals sorbed to sediments. In: Aquatic Toxicology and
Hazard Assessment: 7th Volume, ASTM STP 854, R.D. Cardwell, R. Purdy, and R.C.
Bahner, Eds., Philadelphia, PA: American Society for Testing and Materials, pp. 429-
453.
ASTM. 1988. Standard guide for conducting acute toxicity tests with fishes,
macroinvertebrates, and amphipods. ASTM 1993 Annual Book of Standards Vol. 11.04.
E729-88, Philadelphia, PA.
ASTM. 1992. Guide for conducting 10-day static sediment toxicity tests with marine
and estuarine amphipods. ASTM 1992 Annual Book of Standards Vol. 13.04, E1367-92,
Philadelphia, PA.
ASTM. 1993a. Standard guide for designing biological tests with sediments. ASTM
1994 Annual Book of Standards Vol. 11.04, E1525-93, Philadelphia, PA.
ASTM. 1993b. Standard terminology relating to biological effects and environmental
fate. ASTM 1994 Annual Book of Standards Vol. 11.04, E943-93, Philadelphia, PA.
ASTM. 1994a. Standard guide for collection, storage, characterization, and
manipulation of sediments for toxicological testing. ASTM 1993 Annual Book of
Standards Vol. 11.04. E1391-90, Philadelphia. PA.
ASTM. 1994b. Draft standard guide for the use of light in laboratory testing. Under
development by B.M. Greenburg, Department of Biology, University of Waterloo,
Waterloo. Ontario. ASTM Subcommittee E47.ll, January, 1994.
Ankley, G.T., M.K. Schubauer-Berigan, and R.A. Hoke. 1992. Use of toxicity
identification evaluation techniques to identify dredged material disposal options: A
proposed approach. Environ. Management. 16:1-6.
Ankley. G. and Thomas, N. 1992. Interstitial water toxicity identification evaluation
approach. In: Sediment classification methods compendium, pp. 5-1 to 5-14. EPA-823/R-
92-006, Washington, DC.
Ankley, G.T., D.A. Benoit, J.C. Balough, T.B. Reynoldson, K.E. Day, and R.A. Hoke.
1993. Development and evaluation of test methods for benthic invertebrates and
sediments: Effects of flow rate and feeding on water quality and exposure conditions.
Arch. Environ. Contain. Toxicol. 25:12-19.
Ankley. G.T. and M. K. Schubauer-Berigan. 1994. Comparison of techniques for the
isolation of pore water for sediment toxicity testing. Arch. Environ. Contam. Toxicol.: In
press.
122
-------
Ankley, G.T., D.A. Benoit, J.C. Balough, T.B. Reynoldson, K.E. Day, and R.A. Hoke.
Evaluation of potential confounding factors in sediment toxicity tests with three
freshwater benthic invertebrates. Environ. Toxicol. Chem. 13: 627-635, 1994a.
Ankley, G.T., Band, D.A., Balough, J.C., Reynoldson, T.B., Day, K.E., and Hoke, R.A.
1994. Evaluation of potential confounding factors in sediment toxicity tests with three
freshwater benthic invertebrates. Environ. Toxicol. Chem.:\n press.
Barnard, J.L., and C.M. Barnard. The genus Rhepoxynius (Crustacea: Amphipoda:
Phoxocephalidae) in American Seas. Smithsonian Contributions to Zoology, 357:1-49.
Bartlett, M.S. 1937. Some examples of statistical methods of research in agriculture
and applied biology. J. Royal Statist. Soc. Suppl. 4:137-183.
Boese B.L., Lee II, H., Specht, D.T., Randall, R.C., and Winsor, M.H. 1990.
Comparison of aqueous and solid-phase uptake for hexachlorobenzene in the tellinid
clam Macoma nasuta (Conrad): A mass balance approach. Environ. Toxicol. Chem.
9:221-231.
Bousfield, E.L. 1973. Shallow Water Gammeridean Amphipoda of New England.
Ithaca, NY: Cornell University Press, 312 pp.
Bureau of National Affairs, Inc. 1986. U.S. Environmental Protection Agency General
Regulation for Hazardous Waste Management. Washington, D.C.
Burton, G.A. 1991. Assessment of freshwater sediment toxicity. Environ. Toxicol.
Chem. 10: 1585-1627.
Burton, G.A., Jr. and C.G. Ingersoll. 1994. Evaluating the toxicity of sediments. In:
The Assessment of contaminated Great Lakes sediment. US Environmental Protection
Agency Report, Region V, Chicago, IL.: EPA-905-B94-002.
Carlson, A.R., G.L. Phipps, V.R. Mattson, P.A. Kosian, and A.M. Cotter. 1991. The
role of acid-volatile sulfide in determining cadmium bioavailability and toxicity in
freshwater sediments. Environ. Toxicol. Chem. 10:1309-1319.
Chapman, P.M., E.A. Power, and G. A. Burton, Jr. 1992. Integrative assessments in
aquatic ecosystems. In: Sediment toxicity assessment. G.A. Burton, Jr., Ed. Ann
Arbor, MI: Lewis Publishers, pp. 313-340.
Chapman, P.M. 1989. Current approaches to developing sediment quality criteria.
Environ. Toxicol. Chem. 8:589-599.
Chapman, P.M.. R.N. Dexter, and E.R. Long. 1987. Synoptic measurements of sediment
contamination, toxicity, and infaunal community composition (the Sediment Quality
Triad) in San Francisco Bay. Marine Ecology Progress Series 37:75-96.
123
-------
Conover, WJ. 1980. Practical Nonparametric Statistics. Second edition. John Wiley
and Sons, New York, NY, 493 p.
Craig, G.R. 1984. Bioassessment of sediments: Review of previous methods and
recommendations for future test protocols. IEC Beak consultants. Ltd. Mississauga,
Ontario.
Davenport, R. and A. Spacie. 1991. Acute phototoxicity of harbor and tributary
sediments from lower Lake Michigan. J. Great Lakes Res. 17:51-56.
Day, K.E., R.S. Kirby, and T.B. Reynoldson. 1992. The effects of sediment
manipulations on chronic sediment bioassays with three species of benthic invertebrates.
Abstract Presented at the 13'th Annual Meeting, Society of Environmental Toxicology
and Chemistry, Cincinnati, OH, Nov. 8-12.
DeVVitt, T.H., G.R. Ditsworth, and R.C. Swartz. 1988. Effects of natural sediment
features on survival of the Phoxocephalid amphipod, Rhepoxynius abronius. Marine
Envir. Res., 25:99-124.
DeVVitt, T.H., R.C. Swartz, and J.O. Lamberson. 1989. Measuring the acute toxicity of
estuarine sediments. Environ. Toxicol. Chem. 8:1035-1048.
DeVVitt, T.H., M.S. Redmond, J.E. Sewall, and R.C. Swartz. 1992a. Development of a
chronic sediment toxicity test for marine benthic amphipods. U.S. Environmental
Protection Agency. CBP/TRS/89/93.
DeVVitt, T.H., B.L. McGee, R. Alden, H. Phelps, and A. Pinkney. 1992b. A multi-
species, multi-lab comparison of the toxicity of a Chesapeake Bay sediment Presented
at the Thirteenth Annual Meeting of the Society for Environmental Toxicology and
Chemistry, Cincinnati, OH, November 8-12.
DeWoskin, R.S. 1984. Good laboratory practice regulations: A comparison. Research
Triangle Institute, Research Triangle Park, NC. 63 p.
Dickson, K.L., A.W. Maki, and W.A. Brungs, W.A. 1984. Fate and effects of sediment-
bound chemicals in aquatic systems. Pergamon Press, New York, NY.
Dillon, T.M. and A.B. Gibson. 1986. Bioassessment methodologies for the regulatory
testing of freshwater dredged material. Miscellaneous Paper EL-86-6, L'.S. Army
Engineer Waterways Experiment Station, Vicksburg, MS.
Di Toro, D.M., J.D. Mahony, D.J. Hansen, K.J. Scott, M.B. Hicks, S.M. Mays, and M.S.
Redmond. 1990. Toxicity of cadmium in sediments: The role of acid volatile sulfide.
Environ. Toxicoi Chem. 9:1489-1504.
Di Toro, D.M., C.S Zarba, D.J. Hansen, W.J. Berry, R.C. Swartz, C.E. Cowan, S.P.
Pavlou, H.E. Allen, N. A. Thomas, and P.R. Paquin. 1991. Technical basis for
124
-------
establishing sediment quality criteria for nonionic organic chemicals using equilibrium
partitioning. Environ. Toxicol. Chem. 10:1541-1583.
Ditsworth, G.R., D.W. Schults, and J.K.P. Jones. 1990. Preparation of benthic
substrates for sediment toxicity testing. Environ. Toxicol. Chem. 9:1523-1529.
Dixon, W.J. and F.J. Massey, Jr. 1983. Introduction to statistical analysis. Fourth
edition. McGraw-Hill Book Company, New York, NY. 678 p.
Efron, B. 1982. The Jackknife, the Bootstrap, and other resampling plans. CBMS 38,
Soc. Industr. Appl. Math., Philadelphia, PA.
Environment Canada. 1992. Biological test method: Acute test for sediment toxicity
using marine or estuarine amphipods. Environment Canada, Conservation and
Protection, Ottawa, Ontario. 83 p., EPS l/RM/26.
Fairweather, P.G. 1991. Statistical power and design requirements for environmental
monitoring. Aust. J. Mar. Freshwater Res. 42:555-567.
Finney, D.J. 1971. Probit analysis. Third edition, Cambridge, University Press, London,
333 p., 1971.
Food and Drug Administration. 1978. Good laboratory practices for non-clinical
laboratory studies. Part 58. Fed. Reg. 43(247):60013-60020 (December 22, 1978).
Gill, J.L. 1978. Design and analysis of experiments in the animal and medical sciences.
Vol. 3. Appendices. The Iowa State University Press, Ames, IA, 173 p.
Green, R.H. 1979. Sampling design and statistical methods for environmental
biologists. New York, NY: Wiley Interscience. 257 p.
Grothe, D.R. and R.A. Kimerle. 1985. Inter- and intra-laboratory variability in
Daphnia magna effluent toxicity test results. Environ. Toxicol. Chem. 4:189-192.
Hall, W.S., J.B. Patoczka, R.J. Mirenda, B.A. Porter, and E. Miller. 1989. Acute
toxicity of industrial surfactants to Mysidopsis bahia. Arch. Environ. Contam. Toxicol.
18:765-772.
Hamilton, M.A., R.C. Russo, and R.V. Thurston. 1977. Trimmed Spearman-Karber
method for estimating median lethal concentrations in toxicity bioassays. Environ. Sci.
Technol. 11:714-719. Correction 12:417 (1978).
Harkey, G.A., P.F. Landrum, and S.J. Klaine. 1994. Preliminary studies on the effect
of feeding during whole sediment bioassays using Chironomus riparius larvae.
Chemosphere, 28:597-606.
125
-------
Mines, A.H., A.M. Haddon, and L.A. Wiechert. 1990. Guild structure and foraging
impact of blue crabs and epibenthic fish in a subestuary of Chesapeake Bay. Mar. Ecol.
Prog. Ser.. 67:105-126.
Holland, A.F., A.T. Shaughnessy, L.C. Scott, V.A. Dickens, J.A. Ranasinghe, and J.K.
Summers. 1988. Progress report: Long-term benthic monitoring and assessment
program for the Maryland portion of Chesapeake Bay (July 1986-October 1987). PPRP-
LTB/EST-88-1. VERSAR, Columbia, MD.
Hopkins, D.R. 1986. Atlas of the distributions and abundances of common benthic
species in San Francisco Bay, California. Water Resources Investigations Report 86-
4003. Menlo Park, CA: L'.S. Geological Survey. 16 pp.
HylandJ.L. 1981. Comparative structure and response to (petroleum) disturbance in
two nearshore infaunal communities. PhD. Thesis, University of Rhode Island, 141 p.
Ingersoll C.G., F.J. Dwyer, and T.W. May. 1990. Toxicity of inorganic and organic
selenium to Daphnia magna (Cladocera) and Chironomus riparius (Diptera). Environ.
Toxicol. Chem. 9:1171-1181.
Ingersoll, C.G. and M.K. Nelson. 1990. Testing sediment toxicity with Hyalella azteca
(Amphipoda) and Chironomus riparius (Diptera). In: W.G. Landis and W.H. van der
Schalie (eds.), Aquatic toxicology and risk assessment, 13th volume. ASTM STP 1096.
Philadelphia, PA, pp. 93-109, 1990.
Jordan, R.A. and C.E. Sutton. 1984. Oligohaline benthic invertebrate communities at
two Chesapeake Bay power plants. Estuaries 7:192-212.
Karickhoff, S.W. and K.R. Morris. 1985. Sorption dynamics of hydrophobic pollutants
in sediment suspensions. Environ. Toxicol. Chem. 4:469-479.
Kemble, N.E., W.G. Brumbaugh, E.L. Brunson, FJ. Dwyer, C.G. Ingersoll, C.G., D.P.
Monda, and D.F. V oodward. 1994. Toxicity of metal-contaminated sediments from the
upper Clark Fork River, MT, to aquatic invertebrates in laboratory exposures. Environ.
Toxicol. Chem., in press.
Kemp, P.F., F.A. Code, and R.C. Swartz. 1985. Life history and production of the
Phoxocephalid amphipod Rhepoxynius abronius (Barnard). J. Crustacean Biology 5:449-
464.
Kemp, P.F., and R.C. Swartz. 1988. Acute toxicity of interstitial and particle-bound
cadmium to a marine infaunal amphipod. Mar. Environ. Res. 26:135-153.
Knezovich, J.P., F.L. Harrison, and R.G. Wilhelm. 1987. The bioavailability of
sediment-sorbed organic chemicals: A review. Water Air Soil Pollut. 32:233-245.
126
-------
Lamberson, J.O., T.H. DeWitt, and R.C. Swartz. 1992. Assessment of sediment toxicity
to marine benthos. In: Sediment toxicity assessment, G.A. Burton, Jr., Ed. Ann Arbor,
Ml: Lewis Publishers, pp. 183-211.
Lamberson, J.O., and R.C. Swartz. 1988. Use of bioassays in determining the toxicity
of sediment to benthic organisms. In: Toxic contaminants and ecosystem health: A
Great Lakes focus., M.S. Evans, Ed. New York, NY: John Wiley and Sons. pp. 257-279.
Landrum, P.F. 1989. Bioavailability and toxicokinetics of polycyclic aromatic
hydrocarbons sorbed to sediments for the amphipod, Pontoporeia hoyi. Environ. Sci.
Tech. 23:588-595.
Landrum, P.F., and W.R. Faust. 1992. Variation in the bioavaiiability of polycyclic
aromatic hydrocarbons sorbed to sediments for the amphipod Pontoporeia hoyi.
Environ. Toxicol. Chem. 11:1197-1208.
Lee, H., II, B.L. Boese, J. Pelletier, M. Winsor, D.T. Specht, and R.C. Randall. 1989.
Guidance manual: Bedded sediment bioaccumulation tests. USEPA Report No. 600/x-
89/302, ERL-Narragansett, 232 p.
Lee, D.R. 1980. Reference toxicants in quality control of aquatic bioassays. In: A.L.
Buikema and J. Cairns Jr. (eds.), Aquatic invertebrate bioassays. ASTM STP 715,
Philadelphia, PA. pp. 188-199.
Lee, II, H., B.L. Boese, and P.F. Landrum. 1994. Guide for determination of the
bioaccumulation of sediment-associated contaminants by benthic invertebrates (8/94
draft in ASTM review).
Long, E.R., M.F. Buchman, S.M. Bay, R.J. Breteler, R.S. Carr, P.M. Chapman, J.E.
Hose, A.L. Lissner, J. Scott, and D.A. Wolfe. 1990. Comparative evaluation of five
sediment toxicity tests with sediments from San Francisco Bay and Tomales Bay, CA.
Environ Toxicol. Chem. 9:1193-1214.
Long, E.R., and L.G. Morgan. 1990. The potential for biological effects of sediment-
sorbed contaminants in the NOAA National Status and Trends program. NOAA
Technical Memorandum NOS OMA 52, Seattle, WA. 175 pp + appendices.
Long, E.R. and M.F. Buchman. 1989. An evaluation of candidate measures of
biological effects for the National Status and Trends Program. NOAA Technical
Memorandum NOS OMA 45, Seattle, WA. 105 pp.
Marcus, A.H. and A.P. Holtzman. 1988. A robust statistical method for estimating
effects concentrations in short-term fathead minnow toxicity tests. Manuscript submitted
to the Criteria and Standards Division, U. S. Environmental Protection Agency, by
Battelle Washington Environmental Program Office, Washington, DC, June 1988, under
EPA Contract No. 69-03-3534. 39 p.
127
-------
Marsh, A.G., and K.R. Tenore. 1990. The role of nutrition in regulating the population
dynamics of opportunistic, surface deposit feeders in a mesohaline community. Limnol.
Oceanogr. 35(31:710-724.
McGee, B.L., C.E. Schlekat, D.M Boward, and T.L. Wade. 1994. Sediment
contamination and biological effects in a Chesapeake Bay marina. Ecotoxicology. In
Press.
McGee, B.L., C.E. Schlekat, and E. Reinharz. 1993. Assessing sublethal levels of
sediment contamination with the estuarine amphipod, Leptocheirus plumulosus. Environ
Tuxicol. Chem. 12:577-588.
McLeay, D.J., S. Yee, K. Doe, and S. Wade. 1991. Phase-II and Phase-Ill studies by
Environment Canada laboratories of 10-day tests for sediment toxicity using marine or
estuarine infaunal amphipods. Report prepared for Environment Canada (EP,
Conservation and Protection) and IGATG by McLeay Associates Ltd., West Vancouver,
B.C.
Mearns, A.J., R.C. Swartz, J.M. Cummins, P.A. Dinnel, P. Plesha, and P.M. Chapman.
1986. Inter-laboratory comparison of a sediment toxicity test using the marine
amphipod, Rhepoxynius abronius. Mar. Environ. Res. 19:13-37.
Mills, E.L. 1967. The biology of an amphipod crustacean sibling species pair. J.
Fisheries Res. Board Canada, 24:305-355.
National Research Council (NRC). 1989. Contaminated marine sediments—Assessment
and remediation, NRC, National Academy Press, Washington, DC.
Nelson, W.G. 1980. Reproductive patterns of gammaridean amphipods. Sarsia, 65:61-
71.
Nichols, F.H., and J.K. Thompson. 1985. Persistence of and introduced mudflat
community in south San Francisco Bay, California. Mar. Ecol. Prog. Ser., 24:83-97.
Nkedi-Kizza, P., P.S.C. Rao, and A.G. Hornsby. 1985. Influence of organic cosolvents
on sorption of hydrophobic organic chemicals by soils. Environ. Sci Tech., 19:975-979.
Oakden. J.M. 1984. Feeding and sediment selection in five species of Central
Californian phoxocephalid amphipods. J. Crustacean Biol. 4:233-247.
Oris, J.T. and J.P. Giesy. 1985. The photoenhanced toxicity of anthracene to juvenile
sunfish (Lepomis spp.). Aquat. Toxicol. 6:133-146.
Pastorak. R.A., and D.S. Becker. 1990. Comparative sensitivity of sediment bioassays at
three Superfund sites in Puget Sound. In: W.G. Landis and W.H. van der Schalie, Eds.,
Aquatic toxicology and risk assessment: Thirteenth volume, STP 1096. Philadelphia, PA:
American Society for Testing and Materials, pp. 123-139.
128
-------
Pfitezenmeyer, H. 1975. Benthos. In: A biological study of Baltimore Harbor, T. Koo,
Ed., Contribution No. 621. Center for Environmental and Estuarine Studies, University
of Maryland, Solomons, MD.
Plumb, R.H. 1981. Procedures for handling and chemical analyses of sediment and
water samples. U.S. Environmental Protection Agency/Corps of Engineers Technical
Committee on Criteria for Dredged and Fill Material. Contract EPA-48055720HI,
EPA/CE-81-1. 478 p.
Ramsdell, K.A., J.A. Strand, and V.I. Cullinan. 1989. Amphipod bioassay of selected
sediments from Sequim Bay, Washington. In: Oceans 1989, Volume 2: Ocean Pollution.
IEEE Publication No. 89CH2780-5, pp. 443-448.
Redmond, M.S., and K.J. Scott. 1989. Amphipod predation by the infaunal polychaete.
Nephtys incisa. Estuaries. 12(3):205-207.
Reinharz, E. 1981. Animal sediment relationships: A case study of the Patapsco River.
Open File No.6. Maryland Geological Survey, Baltimore, MD.
Reish, D.J. 1988. The use of toxicity testing in marine environmental research, Chapter
10. In: D.F. Soule and G.S. Kleppel (eds.), Marine organisms as indicators. Springer-
Verlag, New York. pp. 213-245.
Richards, S.W. The demersal fish populations of Long Island Sound: III Food for
juveniles from a mud locality. Bull. Bingham Oceanogr. Coll., 18:32-101, 1963.
Rohlf, F.J. and R.R. Sokal. 1981. Statistical Tables. W.H. Freeman and Company,
New York, NY.
SAIC. 1993a. Draft data report on comparative sensitivity of four amphipod species to
sediment-associated contaminants. Submitted to USEPA. EPA contract No. 68-CO-
0044. Prepared by Science Applications International Corporation, Narragansett, RI.
15 pp.
SAIC. 1993b. Draft data report on salinity tolerance and application for four
amphipod species. Submitted to USEPA. EPA contract No. 68-CO-0044. Prepared by
Science Applications International Corporation, Narragansett, RI. 22 pp.
SAIC. 1993c. Draft data report on comparative sensitivity of four amphipod species to
aqueous ammonia. Submitted to USEPA. EPA contract No. 68-CO-0044. Prepared by
Science Applicatioas International Corporation, Narragansett, RI. 30 pp.
Sanders, H.L., P.C. Mangelsdorf, Jr., and G.R. Hampson. 1965. Salinity and faunal
distribution in the Pocasset River, Massachusetts. Limnology and Oceanography
Supplement to Vol 10.-R216-R229.
129
-------
Satterthwaite, F.W. 1946. An approximate distribution of estimates of variance
components. Biom. Bull. 2:110-114.
Schaeffer, D.J. and K.G. Janardan. 1978. Theoretical comparison of grab and
composite sampling programs. Biom. J. 20:215-227.
Schlekat. C.E., B.L. McGee, and E. Reinharz. 1992. Testing sediment toxicity in
Chesapeake Bay with the amphipod Leptocheirus plumulosus: An evaluation. Environ
Toxicol. Chem.'\ 1:225-236.
Schlekat, C.E., B.L. McGee. D.M. Bowan, R. Reinharz, D.J. Velinsky and T.L. Wade.
1994. Tidal river sediments in the Washington, DC area. III. Biological effects
associated with sediment contamination. Estuaries 17: In press.
Schults, D.W., S.P. Ferraro, L.M. Smith. F.A. Roberts, and C.K. Poindexter. 1992. A
comparison of methods for collecting interstitial water for trace organic compounds and
metal analyses. Wat. Res. 26:989-995.
Schuytema, G.S., A.V. Nebeker, W.L. Griffis. and C.E. Miller. 1989. Effects of freezing
on sediments contaminated with DDT and endrin. Environ. Toxicol. Chem. 8:883-891.
Scott, K.J., and M.S. Redmond. 1989. The effects of a contaminated dredged material
on laboratory populations of the tubicolous amphipod, Ampelisca abdita. In: Aquatic
Toxicology and Hazard Assessment: 12'th Volume, ASTM STP 1027, I .M. Cowgill and
L.R. Williams. Eds. Philadelphia, PA: American Society for Testing and Materials.
pp. 289-303.
Scott. K.J. 1989. Effects of contaminated sediments on marine benthic biota and
communities. In: Contaminated marine sediments-Assessment and remediation.
National Research Council. National Academy Press, Washington, DC. pp. 132-154.
Shapiro, S.S. and M.B. Wilk. 1965. An analysis of variance test for normality (complete
samples). Biometrika 52:591-611.
Shoemaker, C.R. 1932. A new amphipod of the genus Leptocheirus from Chesapeake
Bay. Wash. Acad. Sci. 22: 548-551.
Shuba, P.J., H.E. Tatem, and J.H. Carroll. 1978. Biological assessment methods to
predict the impact of open-water disposal of dredged material. U.S. Army Corps of
Engineers Technical Report D-78-5Q, Washington, DC.
Snedecor. G.W. and G.C. Cochran. 1989. Statistical Methods. Eighth edition. The Iowa
State University Press, Ames, IA. 507 p.
Sokal, R.R. and F.J. Rohlf. 1981. Biometry, second edition. W.H. Freeman and
Company. New York.
130
-------
Southerland, E., M. Kravitz, and T. Wall. 1992. Management framework for
contaminated sediments (The USEPA Sediment Management Strategy). In: Sediment
Toxiciry Assessment, G.A. Burton. Ed. Ann Arbor, MI: Lewis Publishers, pp. 341-370.
Steel, R.G.D., and J.H. Torrie. I960. Principles and procedures of statistics. New
York, NY: McGraw Hill Book Co.
Stein, J.R. 1973. Handbook of phycological methods: Culture methods and growth
measurements. New York, NY: Cambridge 1,'niversity Press. 445 pp.
Stemmer, B.L., G.A. Burton Jr., and G. Sasson-Brickson. 1990a. Effect of sediment
spatial variance and collection method on cladoceran toxicity and indigenous microbial
activity determinations. Environ. Toxicol. Chem. 9:1035-1044,.
Stemmer, B. L., Burton, Jr.. G. A., and Leibfritz-Frederick, S. 1990b. Effect of
sediment test variables on selenium toxicity to Daphnia magna. Environ. Toxicol. Chem.
9:1035-1044.
Swartz, R.C., Code, F.A.. Lamberson, J.O, Ferraro, S.P., Schults. D.W., DeBen, W.A..
Lee II, H., and Ozretich, R.J. 1994. Sediment toxicity, contamination, and am phi pod
abundance at a DDT and dieldrin-contaminated site in San Francisco Bay. Environ.
Toxicol. Chem. 13:949-962.
Swartz, R.C., W.A. DeBen, K.A. Secru, and J.O. Lamberson. 1982. Sediment toxicity
and the distribution of amphipods in Commencement Bay, Washington, USA. Mar. Poll.
Bull. 13:359-364.
Swartz, R.C., W.A. DeBen, J.K.P. Jones, J.O. Lamberson, and F.A. Code. 1985.
Phoxocephalid amphipod bioassay for marine sediment toxicity. In: Aquatic Toxicolog>
and Hazard Assessment: 7th Volume, ASTM STP 854, R.D. Cardwell, R. Purdy. and
R.C. Bahner, Eds., Philadelphia, PA: American Society for Testing and Materials.
pp. 284-307.
Swartz, R.C., P.F. Kemp, D.W. Schultz, and J.O. Lamberson. 1988. Effects of mixtures
of sediment contaminants on the marine infaunal amphipod, Rhepoxynius abronius.
Environ. Toxicol. Chem. 7:1013-1020.
Swartz, R.C. 1989. Marine sediment toxicity tests. In: Contaminated Marine
Sediments - Assessments and Remediation, Washington, D.C.: National Academy Press.
pp. 115-129.
Taylor, J.K. 1987. Quality assurance of chemical measurements. Lewis PubL, Inc..
Chelsea, MI.
Tetra Tech, Inc. 1986. ODES statistical power analysis. Draft report, prepared for
Office of Marine and Estuarine Protection. I SEPA Contract NO. 68-01-6938, TC-3953-
03. Bellevue, WA.
131
-------
Tomasovic. M. et al. 1994. Recovery of Hyalella azteca from sediment. Environ. Toxicol.
Chem. 13:1311-1314.
I'SEPA. 1979a. Handbook for analytical quality assurance in water and wastewater
laboratories. EPA-600/4-79-019, Cincinnati, OH.
I'SEPA. 1979b. Methods for chemical analysis of water and wastes. EPA-600/4-79-020,
Cincinnati, OH.
I'SEPA. 1980a. Proposed good laboratory practice guidelines for toxicity testing.
Paragraph 163.60-6. Fed. Reg. 45:26377-26382 (April 18, 1980).
I'SEPA. 1980b. Physical, chemical, persistence, and ecological effects testing; good
laboratory practice standards (proposed rule). 40 CFR 772, Fed. Reg. 45:77353-77365
(November 21, 1980).
I'SEPA. 1985. Methods for measuring the acute toxicity of effluents to freshwater and
marine organisms. Third edition. EPA-600/4-85/013, Cincinnati, OH.
I'SEPA. 1986a. Quality criteria for water. EPA-440/5-86-001, Washington, DC.
I'SEPA. 1986b. Occupational health and safety manual. Office of Administration, EPA,
Washington, DC.
I'SEPA. 19S9a. Evaluation of the apparent effects threshold (AET) approach for
assessing sediment quality. Report of the sediment criteria subcommittee. SAB-EETFC-
89-027, Washington, DC.
I SEPA. 1989b. Short-term methods for estimating the chronic toxicity of effluents and
receiving waters to freshwater organisms. EPA-600/4-89/001, Cincinnati, OH.
I'SEPA. Toxic Substance Control Act (TSCA); Good laboratory practice standards,
Final Rule, Federal Register 54: 34034-34050. August 17, 1989c.
I SEPA. 1990a. Evaluation of the equilibrium partitioning (EqP) approach for assessing
sediment quality. Report of the sediment criteria subcommittee of the ecological
processes and effects committee. EPA-SAB-EPEC-90-006, Washington, DC.
I'SEPA. 1990b. Evaluation of the sediment classification methods compendium. Report
of the sediment criteria subcommittee of the ecological processes and effects committee.
KP \-SAB-EPEC-90-018. Washington, DC.
I'SEPA. 1990c. Analytical procedures and quality assurance plan for determination of
xenobiotic chemical contaminants in fish. EPA-600/3-90/023, Duluth, MM.
I'SEPA. I990d. Analytical procedures and quality assurance plan for determination of
PCDD/PCDF in fish. EPA-600/3-90/022, Duluth, MN.
132
-------
USEPA. 1991a. Sediment toxicity identification evaluation: Phase I (Characterization),
Phase II (Identification) and Phase III (Confirmation). Modifications of effluent
procedures. EPA 600/6-91/007, Duluth, MN.
USEPA. 1991b. Technical support document for water quality-based toxic control.
EPA-505/2-90/001, Washington, DC.
USEPA. 1992a. Proceedings from workshop on tiered testing issues for freshwater and
marine sediments. Washington, DC, September 16-18, 1992.
USEPA. 1992b. An SAB report: Review of sediment criteria development methodology
for non-ionic organic contaminants. Report of the sediment criteria subcommittee of the
ecological processes and effects committee. EPA-SAB-EPEC-93-002. Washington, DC.
USEPA. 1992c. Sediment classification methods compendium. EPA-823-R-92-006.
Washington, DC.
USEPA. 1993a. Methods for measuring the acute toxicity of effluents to freshwater and
marine organisms. Fourth edition. EPA 600/4-90/027F, Cincinnati, OH, 1993a.
USEPA. 1994a. Procedures for conducting sediment toxicity and bioaccumulation tests
using freshwater invertebrates. EPA-600/R-94/024. June 1994. I SEPA, Office of
Research and Development, Duluth, MN.
USEPA. 1994b. Short-term methods for measuring the chronic toxicity of effluents and
surface waters to freshwater organisms. Third edition. EPA-600/4-91/002. Cincinnati.
OH.
USEPA. 1994c. Short-term methods for measuring the chronic toxicity of effluents and
surface waters to marine and estuarine organisms. Second edition. EPA-600/4-91/002.
Cincinnati, OH.
USEPA-USCOE (U.S. Army Corps of Engineers). 1991. Evaluation of dredged material
proposed for ocean disposal (Testing manual). EPA-68-C8-0105. USEPA, Office of
Marine and Estuarine Protection, Washington, D.C.
USEPA-USCOE (U.S. Army Corps of Engineers). 1994. Evaluation of dredged material
proposed for discharge in inland and near coastal waters. EPA- 000/0-93/000.
Washington, DC.
USGS (U.S. Geological Survey). 1969. Techniques of water-resources investigations of
the USGS. Chapter Cl, Harold P. Guy, p. 58, Laboratory Theory and Methods for
Sediment Analysis; Book 5. Laboratory Analysis, I .S.G.S. Arlington, VA
Walters, D.B. and C.W. Jameson. 1984. Health and safety for toxicity testing.
Butterworth Publications, Woburn, MA.
133
-------
Weisberg. S.B., J.B. Frithsen, A.F. Holland, J.F. Paul, K.J. Scott, J.K Summers, H.T.
Wilson, R. Valente, D.G. Heimbuch, J. Gerritsen, S.C. Schimmel, and R.W. Latimer.
1992. EMAP-Estuaries Virginian Province 1990 Demonstration Project Report. EPA
600/R-92/100. I'SEPA, Environmental Research Laboratory, Narragansett, RI.
Wiederholm, T. , A.M. VViederholm, and VI. Goran. 1987. Fresh-water oligochaetes.
Water Air Soil Pollut. 36:131-154.
Williams. D.A. 1971. A test for differences between treatment means when several dose
levels are compared with a zero dose control. Biometrics. 27: 103-117
Williams, L.G., P.M. Chapman, and T.C. Ginn. 1986. A comparative evaluation of
marine sediment toxicity using bacterial luminescence, oyster embryo, and amphipod
bioassays. Marine Environmental Research, 19: 225-249.
Williams, L.R. 1993. Methods for the EPA's regulatory program. Environmental
Testing and Analysis. 2:36.
Winer, B.J. 1971. Statistical Principles in Experimental Design. Second edition.
McGraw-Hill Book Company, New York, NY. 907 p.
Word. J.Q., J.A. Ward, L.M. Franklin, V.I. Cullinan, and S.L. Kiesser. 1987.
Evaluation of the equilibrium partitioning theory for estimating the toxicity of the
nonpolar organic compound DDT to the sediment dwelling amphipod, Rhepoxynius
abronius. Battelle Marine Science Research Laboratory Report, Task 1, WA56, Sequim,
WA. 60 pp.
Word. J.Q., J.A. Ward, B. Brown, B.D. Walls, and S. Lemlich. 1989. Relative
sensitivity and cost of amphipod bioassays. In: Oceans '89 Volume 2: Ocean Pollution.
IEEE Publication No. 89CH2780-5. pp. 467-473.
Zar, J.H. 1984. Biostatistical Analysis. Second edition. Prentice-Hall, Inc., Englewood
Cliffs. NJ. 717 p.
134
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APPENDIX A
Sample Data Sheets
-------
FtelJ Collection and Laboratory Holding
Organism
Method of Field Collection: See SOP
Location:
Time:
Collection Date:
Used tor Experiments):
Temperature: Water:
Air:
S;Jimt\: o/oo
Holding Sediment:
Jar":
Collecuoti Date:
Tide Height:
C
c
Participants:
Weather:
Comments:
Number of Animus/Holding Jar:
Daie
Location
Temp
'C
Salinity
o.'oo
Renewal
Volume
iLi
Carboy
n
Date
Initials
Feedine
Volume
(L>
Species,
Initials
Jar
»
Animal
Sue
<
Date:
D.O.
pH
I
Initials
Date:
DO
PH
Initials
Number
Dead
Initials
Exper.
#
( nmments:
Figure A.I Fit-Id collection and laboratory holding data sheet.
136
-------
%hr Reference Toxicant Tesl-Dailv Data Sheet
Species:
Toxicant:
Experiment»
Text Sun Date.
Set-up Carboy
Set up and Daily Survival Check-.
Renewal <"arho\ f.
Test Level
mg/1.
Control
0 mg/1.
Rep
A
B
A
B
A
B
A
B
A
B
A
B
Temp °C
Observer
dale: 24hr
Total No.
(live+de-adi
Today*
No. l.i ve
dale: 4Shr
Toul No.
(live+dead)
Todavs
No. Live
date: 7Chr
Toul No
(live-fdeiuii
1 odays
No. Live
date' -i^hr
Toul No.
lli\e-fde4di
7txia>v
Vo. Liu-
Hmical DaLj
(\inU110DL1'
Test Level
mg/L
Control
Omg/1.
Rep
A
B
A
B
A
B
A
B
A
B
A
H
()h.si-r\e-
dale:
pH
DO
PTni
Salinity
PP<
dale.
PH
DO
PI1"'
Saljiut\
P)H
Figure A.2 Data sheet for 96 h reference toxicant test.
137
-------
10 Dav Solid Phase Tcst-Dajlv Dam Sheet
Date
hxpenmtnlff:
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tt
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Figure A.3 Data sheet for daily observations during the 10-d solid phase test.
138
-------
10 Dav Solid Phase Tesi-Phvsical Daui Sheet
Project
ExperiniL'ni *•:
Jar
Das
Da\
Da\
Dav
Figure A.4 Data sheet for 10-d solid phase test
139
-------
H; Da\ Solid Phase Test--BieokdrAMi OaLi Sheet
Project:
Species
hxpenment *
Date
.First Pick
bijlials
Time
Ja; «
1
2
3
4
S
ft
7
8
9
10
1 1
12
13
1-4
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1ft
17
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19
20
21
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24
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24
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[)eail
IN + the » live tnund In the picket
" It >|ii'' i'I I he animals j;e r/ussui^ . ic. >2 .it 2'ii. the -..unple :v.usl he (JA J
*"' F-.n.il * I ive = the tt live Irmn the I'lisl pick lecnunt - the » live linin the M
recount
Figure \.S Data sheet for test summary.
140
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