United States
Environmental Protection
Agency
Office of Research and
Development
Washington DC 20460
EPA/600/R-93/183
September 1993
oEPA Guidance Manual
Bedded Sediment
Bioaccumulation Tests
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EPA/600/R-93/183
September 1993
GUIDANCE MANUAL: BEDDED SEDIMENT BIOACCUMULATION TESTS
Henry Lee II1, Bruce L. Boese1, Judy Pelletier2,
Martha Winsor2, David T. Specht1, and Robert C. Randall1
Bioaccumulation Team
Pacific Ecosystem Branch
Environmental Research Laboratory - Narragansett
U.S. Environmental Protection Agency
Hatfield Marine Science Center
Newport, Oregon 97365
2AScI Corporation
Hatfield Marine Science Center
Newport, Oregon 97365
September, 1989
ERL-N Contribution No. Nlll
OS Printed on Recycled Paper
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PREFACE
The recommendations in this document are based on our
experience and best scientific judgment. It is our hope that the
procedures suggested here will aid scientists in conducting
"routine" and experimental sediment bioaccumulation tests, as
well as aid regulators in determining when bioaccumulation tests
are needed, in evaluating the QA/QC procedures of such tests, and
in interpreting the results. The recommendations made here,
however, do not constitute an official policy or standard
procedure by the U.S. Environmental Protection Agency. Mention
of trade names or commercial products does not constitute
endorsement or recommendation of use by the Environmental
Protection Agency.
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1993 UPDATE TO "BIOACCUMULATION GUIDANCE MANUAL"
In the four years since we wrote this document, the "Guidance
Manual" has become the standard for assessing dredge materials
proposed for ocean disposal (U.S. EPA/ACE, 1991), dredge materials
proposed for inland disposal (U.S. EPA/ACE, 1993 draft), and
several scientific studies. Fortunately, the procedures in this
document appear to have stood the test of time relatively well.
Nonetheless, there are several modifications. These modifications,
as well as procedures for freshwater organisms, are being
incorporated into an ASTM guidance document that should be out in
1994 or 1995.
I. VERSIONS OF GUIDANCE MANUAL
The 1989 version of the Guidance Manual was not published and
was distributed only as photocopies. In the 1989 version, Table
IV-3A in the Appendix IV was in error. In 1991, the Army Corps of
Engineers agreed to photocopy and distribute the Manual in support
of the new "Green Book" (U.S. EPA/ACE, 1991). This version was
single spaced and contained a corrected Table IV-3A. The 1993
version contains the corrected Table IV-3A, corrects an error in
Figure IV-1, and makes a few editorial corrections.
II. NON-INDIGENOUS TEST SPECIES
Over the last few years, there has been a growing awareness of
the ecological and economic damage caused by introduced species.
Because both east and west coast species are often used in
bioaccumulation tests, there is a real potential of introducing
bioaccumulation test species or associated fauna and flora (e.g.,
pathogens, algae used in transporting the worms) . Any user of this
document needs to understand that it is their responsibility to
assure that non-indigenous species are not released into the
environment.
The general procedure to contain non-indigenous species is to
collect and then poison all water, sediment, organisms, and
associated packing materials (e.g., algae, sediment) before
disposal. Chlorine bleach can be used as a poison. Double
containment system is used to keep any spillage from going down the
drain. Guidance on procedures used in toxicity tests can be found
in Appendix B of DeWitt et al. (1992). Permits to import, hold,
and use non-indigenous species may be required by state Fish and
Game departments or other state agencies. Flow-through tests can
generate large quantities of water, and the researcher needs to
plan on having sufficient storage facilities.
III. SEDIMENT RENEWAL
We no longer recommend the addition of supplemental sediment
i i i
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during 28-day exposures (Chapter X) . In tests longer than 28 days,
sediment addition is still an option to maintain pollutant
concentrations and food quality/quantity as is periodic renewal of
all the test sediment.
IV. DURATION OF TEST
Our recent work on field sediments contaminated with DDT, DDT
metabolites, and dieldrin indicates that 28 days may be
insufficient to obtain 80% of steady-state tissue residues for a
number of higher Kow compounds (see Boese and Lee, 1992 for summary
of unpublished data and an expanded review of the percent of
steady-state obtained in 28 days). These results emphasize the
importance of conducting tests of at least 28 days duration.
In tests where it is critical to measure the residue within
_>80% of steady-state, it may be necessary to conduct longer term
tests or at least to expose a few individuals for a longer duration
to test whether steady-state was achieved. In screening tests
where is important not to underestimate the residues, the residues
obtained from 28-day tests could be multiplied by a "steady-state
correction factor". The "steady-state correction factor" is the
reciprocal of the decimal fraction of the amount of steady-state
tissue residue obtained after 28 days. For example, if the tissue
residue after 28 days is 0.33 of the residue at steady state, the
correction factor would be I/.33 = 3. These correction values
would be obtained from previously conducted lab studies (e.g.,
Table IV-1 of Guidance Manual; Boese and Lee, 1992).
V. GUT PURGING
As far as we are aware, the effects of gut purging have not
yet been assessed quantitatively. Therefore, the recommendations
in Chapter X and Appendix X-l remain the best available guidance.
VI. TERMINOLOGY AND COMPARISON OF BIOACCUMULATION MODELS
Unfortunately, the terminology used in bioaccumulation is not
standardized and has continued to evolve, or at least change, since
the Guidance Manual was written. The "thermodynamic-based
bioaccumulation" model referred to in Appendix 1-1 of the Guidance
Manual is better termed the "equilibrium partitioning
bioaccumulation" (EqP) bioaccumulation model. Since the term
"accumulation factor" (AF) of the EqP bioaccumulation model first
appeared in the peer-reviewed literature, the term "biota sediment
accumulation factor" (BSAF) has been used by some authors. It
appears that the two terms are interchangeable, though the reader
is cautioned to check the units. Though it was not emphasized in
Appendix 1-1 of the Guidance Manual, AFs have units of gC/gL (see
Lee, 1992) and it is critical that both the tissue pollutant and
lipid concentrations be measured in or converted to the same units
IV
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(i.e., both wet or both dry) . Additionally, the sediment pollutant
concentrations and the total organic carbon (TOO concentration of
the sediment need to be in consistent units.
The guidance manual for open water disposal of dredge
materials (U.S. EPA/ACE, 1991) uses the "theoretical
bioaccumulation potential" (TBP). The TBP is the application of
the EqP model using an accumulation factor of 4.0. Again the
reader is cautioned to use consistent units for lipid and tissue
residue and for sediment and lipid concentrations.
We now recommend the use of "ks" rather than "k^" for the
first-order uptake rate coefficient from sediment to help
distinguish it from uptake from water (see Lee, 1991). It is also
recommended to consider ks a coefficient rather than a constant.
More thorough comparisons of bioaccumulation models than given
in Appendix 1-1 can be found in Boese and Lee (1992), Landrum et
al. (1992), and Lee (1992).
VII. FIELD VALIDATION
The bioaccumulation test methodology is presently being
evaluated by comparing residues in field-captured Macoma nasuta
with residues in laboratory-exposed M. nasuta and Nereis virens.
The initial data for sum DDT with M. nasuta indicates that only 34%
of the steady-state residue was obtained within 28 days at the one
station where a long-term (90 day) test was conducted. However,
when the 28-day residues from other stations were multiplied by the
steady-state correction factor, the lab residues were within 3-fold
of residues in field-captured M. nasuta in 7 of 8 cases. This
comparison included 6 sites that varied by almost 3 orders-of-
magnitude in sum DDT sediment concentrations, indicating that the
procedures work over a wide range of sediment contamination.
VIII. REFERENCES
Boese, B. and H. Lee II. 1992. Synthesis of Methods to Predict
Bioaccumulation of Sediment Pollutants. U.S. EPA Report.
ERL-Narragansett No. N232. 87 pp.
DeWitt, T., M. Redmond, J. Sewall, and R. Swartz. 1992. Develop-
ment of a Chronic Sediment Toxicity Test for Marine Benthic
Amphipods. Chesapeake Bay Program/TRS 89/93.
Landrum, P., H. Lee II, M. Lydy. 1992. Toxicokinetics in aquatic
systems: Model comparisons and use in hazard assessment. Environ.
Toxicol. Chem. 11:1709-1725.
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Lee II, H. 1991. A clam's eye view of the bioavailability of
sediment-associated pollutants. in Organic Substances and
Sediments in Water. Vol. Ill: Biological. R. Baker (ed.), Lewis
Publ., N.Y., pp 73-93.
Lee II, H. 1992. Models, muddles, and mud: Predicting
bioaccumulation of sediment-associated pollutants. in Sediment
Toxicity Assessment. G. Allen Burton (ed.), Lewis Publ., N.Y. pp
267-291.
U.S. EPA/ACE. 1991. Evaluation of Dredged Material for Ocean
Disposal (Testing Manual). U.S. EPA Report No. 503/8/91/001,
Office of Marine and Estuarine Protection, Washington, DC.
U.S. EPA/ACE. 1993. Evaluation of Dredged Material Proposed for
Discharge in Inland and Near Coastal Waters - Testing Manual
(Draft).
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ACKNOWLEDGEMENTS
We greatly appreciate the time and effort Jack H. Gentile,
Peter F. Landrum, Norman Rubinstein, and Richard K. Peddicord
spent in reviewing a document of this magnitude. We also
appreciate the comments of James Heltshe and Donald J. Reish on
specific chapters. All reviewers offered important comments and
suggestions, and the document benefited from their insights. We
also appreciate Richard Latimer's initial prodding for us to
initiate this task. George Ditsworth, and Donald Schults allowed
us to use their unpublished data. Karl Rukavina, Richard Lapan,
and Tricia Lawson offered numerous editorial comments and helped
assure there were no typoos left in the document. The data on
the sediment pollutant concentrations in Yaquina Bay were
originally collected from research partially funded by the Puget
Sound Program of Region X of the U. S. Environmental Protection
Agency. Environmental Research Laboratory-Narragansett
Contribution No. Nlll.
VI 1
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TABLE OF CONTENTS
PREFACE i i
1993 UPDATE i i i
ACKNOWLEDGEMENTS vii
LIST OF TABLES xi i
LIST OF FIGURES Xl'ii
ABSTRACT xi V
CHAPTER I: INTRODUCTION 1
CHAPTER II: CONTROL VERSUS REFERENCE SEDIMENT 6
A. DEFINITION OF CONTROL AND REFERENCE SEDIMENT 6
B. CRITERIA FOR CONTROL AND REFERENCE SEDIMENTS 10
C. STANDARD REFERENCE SEDIMENT 13
CHAPTER III. PRINCIPLES OF EXPERIMENTAL DESIGN 16
A. OBJECTIVES AND DEFINITIONS 16
B. HYPOTHESES TESTING 18
C. REPLICATION 20
D. RANDOMIZATION 27
E. PSEUDOREPLICATION 27
F. AVOIDING OR REDUCING PSEUDOREPLICATION 30
G. COMPOSITING SAMPLES 33
CHAPTER IV: TEST DURATION AND SAMPLING SCHEDULES 36
A. STANDARD 28-DAY BIOACCUMULATION TEST 36
B. LONG-TERM UPTAKE TESTS 41
C. ESTIMATING STEADY-STATE TISSUE RESIDUES FROM UPTAKE
AND DEPURATION RATES 44
CHAPTER V: SEDIMENT COLLECTION, HOMOGENIZATION, MANIPULATION,
AND STORAGE 48
A. SEDIMENT COLLECTION AND TRANSPORT 48
B. SEDIMENT SPIKING AND MANIPULATION 54
C. LABORATORY SEDIMENT STORAGE 54
D. SEDIMENT PREPARATION AND HOMOGENIZATION 55
CHAPTER VI: SEDIMENT CHARACTERIZATION 58
A. GRAIN SIZE 58
B. TOTAL SOLIDS CONTENT 60
C. ORGANIC CARBON 61
D. ADDITIONAL SEDIMENT CHARACTERISTICS 63
E. INTERSTITIAL WATER 65
IX
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CHAPTER VII: ORGANISM SELECTION 69
A. SELECTION CRITERIA 69
B. RECOMMENDED SPECIES 70
C. NUMBER OF SPECIES TESTED AND MULTIPLE
SPECIES TESTS 72
CHAPTER VIII: ORGANISM COLLECTION, MAINTENANCE, TRANSPORT,
AND ACCEPTABILITY 74
A. COLLECTION AND TRANSPORT 74
B. CULTURING AND PURCHASING TEST ORGANISMS 77
C. PRE-EXPERIMENTAL MAINTENANCE 78
D. ORGANISM ACCEPTABILITY AND BACKGROUND
CONTAMINANT LEVELS 82
CHAPTER IX: SEDIMENT EXPOSURE SYSTEMS 85
A. SYSTEM REQUIREMENTS 85
B. EXPOSURE SYSTEM DESIGN 91
C. MULTIPLE SPECIES EXPOSURE CHAMBERS 95
CHAPTER X: EXPERIMENTAL INITIATION, MAINTENANCE,
AND SAMPLING 97
A. EXPERIMENTAL INITIATION AND MAINTENANCE 97
B. SCHEDULE FOR ABIOTIC AND BIOTIC POLLUTANT SAMPLES .... 102
C. METHODS OF BIOTIC SAMPLING 105
D. GUT PURGING 106
E. ACCEPTABLE LEVELS OF MORTALITY Ill
F. CHAIN OF CUSTODY 112
CHAPTER XI: POLLUTANT AND LIPID ANALYSIS 114
A. POLLUTANT ANALYSIS 114
B. LIPID ANALYSIS 118
C. SAMPLE STORAGE 121
D. REPORTING OF RESULTS 122
CHAPTER XII: STATISTICAL ANALYSES 123
A. TESTS FOR NORMALITY AND HOMOGENEITY OF VARIANCES 124
B. PAIRWISE COMPARISONS 126
C. MULTIPLE COMPARISONS 131
D. INTERPRETATION OF COMPARISONS OF TISSUE RESIDUES 133
E. ADDITIONAL ANALYSES 134
CHAPTER XIII: REGULATORY STRATEGIES FOR USE OF
BIOACCUMULATION DATA 137
A. NO FURTHER DEGRADATION 138
B. TISSUE RESIDUE EFFECTS 139
C. WATER QUALITY CRITERION TISSUE LEVEL APPROACH 140
D. FDA ACTION LIMITS 142
E. HUMAN HEALTH RISK ASSOCIATED WITH SHELLFISH 143
F. TROPHIC TRANSFER OF POLLUTANTS INTO PELAGIC
FOOD WEBS „.......„„„...„ 144
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APPENDICES
(Numbered sequentially by Chapter)
APPENDIX 1-1: ADDITIONAL METHODS TO PREDICT BIOACCUMULATION..147
A. FIELD COLLECTION 147
B. BIOACCUMULATION FACTORS AND ACCUMULATION FACTORS 149
C. TOXICOKINETIC BIOACCUMULATION MODELS 153
APPENDIX 1-2: SAFETY AND WASTE DISPOSAL 157
A. PERSONNEL SAFETY 157
B. HAZARDOUS WASTE DISPOSAL 157
APPENDIX III-l: DETERMINING NUMBER OF REPLICATES 159
APPENDIX IV-1: ADEQUACY OF 10-DAY AND 28-DAY EXPOSURES 164
APPENDIX IV-2: ALTERNATE TEST DESIGNS 170
A. SHORT-TERM TESTS 170
B. ESTIMATING STEADY-STATE FROM UPTAKE RATES 170
C. GROWTH DILUTION 171
APPENDIX IV-3: CALCULATION OF TIME TO STEADY-STATE 174
APPENDIX V-l: TECHNIQUES FOR SEDIMENT MANIPULATION 178
A. SEDIMENT SPIKING 178
B. INCREASING SEDIMENT TOC 182
C. DECREASING SEDIMENT POLLUTANT AND/OR
TOC CONCENTRATIONS 185
APPENDIX VII-1: SELECTION CRITERIA FOR TEST SPECIES 187
A. INDIGENOUS VERSUS SURROGATE SPECIES 187
B. REQUIRED CRITERIA 188
C. DESIRABLE CHARACTERISTICS 190
D. RECOMMENDED AND SECONDARY SPECIES 196
APPENDIX VIII-1: SOURCES FOR TEST ORGANISMS 198
APPENDIX IX-1: SPECIAL PURPOSE EXPOSURE CHAMBERS 199
A. CLAMBOX 199
B. WORMTUBES 199
C. SEDIMENT RESUSPENSION SYSTEMS 201
APPENDIX X-l: ADDITIONAL TECHNIQUES FOR CORRECTING FOR
GUT SEDIMENT 204
A. MODIFICATIONS TO 24-HOUR PURGE AND DISSECTION 204
B. CALCULATING POLLUTANT MASS OF GUT SEDIMENT 205
C. USE OF CONSERVATIVE TRACE ELEMENTS 206
GLOSSARY: 208
BIBLIOGRAPHY 213
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LIST OF TABLES
(Numbered sequentially by Chapter or Appendix)
TABLE II-1: REPRESENTATIVE CONTROL SEDIMENT
CONCENTRATIONS 11
TABLE III-l: RANGES OF COEFFICIENTS OF VARIATIONS (CV) FOR
VARIOUS ORGANISMS AND POLLUTANTS 24
TABLE IV-1: PERCENT OF STEADY-STATE TISSUE RESIDUE
OBTAINED AFTER 10-DAY AND 28-DAY EXPOSURES 38
TABLE VI-1: WENTWORTH GRADE CLASSIFICATION OF SEDIMENTS .... 59
TABLE VI-2: RATIOS FOR CONVERTING LOSS ON IGNITION (LOI)
TO TOTAL ORGANIC CARBON (TOC) 64
TABLE VII-1: PERTINENT CHARACTERISTICS OF TEST SPECIES 71
TABLE VIII-1 REPRESENTATIVE CONTROL ORGANISM TISSUE
RESIDUES 84
TABLE X-l: ERRORS ASSOCIATED WITH GUT SEDIMENT/PURGING 107
TABLE X-2: DEPURATION LOSS OF POLLUTANTS DURING 24 AND
72 HOUR GUT PURGES 110
TABLE XI-1: U.S. EPA CONTRACT LABORATORY PROGRAM
QUANTITATION LIMITS FOR WATER AND SEDIMENT WITH
ESTIMATES FOR TISSUE MATRICES 116
TABLE XI-2: PSDDA LOW LIMITS OF DETECTION FOR WATER,
SEDIMENT AND TISSUE MATRICES 117
TABLE XII-1: SUMMARY OF STATISTICAL ANALYSES 129
TABLE XII-2: EXAMPLES OF ANALYSES AND INTERPRETATION
OF RESULTS 135
TABLES IN APPENDICES
TABLE IV-1A: INFORMATION GAINED AND REQUIREMENTS OF DIFFERENT
APPROACHES TO ESTIMATING BENTHIC TISSUE
RESIDUES 167
TABLE IV-3A: ESTIMATED TIME TO OBTAIN 95% OF
STEADY-STATE TISSUE RESIDUE 176
XI 1
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LIST OF FIGURES
(Numbered sequentially by Chapter or Appendix)
FIGURE 1-1: STEPS IN CONDUCTING SEDIMENT BIOACCUMULATION
TESTS 4
FIGURE II-1: OVERLAP OF CONFIDENCE INTERVALS IN ORGANISMS
EXPOSED TO ACCEPTABLE AND UNACCEPTABLE
REFERENCE SEDIMENTS 9
FIGURE III-l: COEFFICIENT OF VARIATION VS. SAMPLE SIZE FOR
VARIOUS MINIMUM DETECTABLE DIFFERENCES 23
FIGURE III-2: MINIMUM DIFFERENCE DETECTABLE BETWEEN
TREATMENTS FOR A SPECIFIC COEFFICIENT OF
VARIATION AND SAMPLE SIZE 26
FIGURE III-3: RANDOM AND PSEUDORANDOM REPLICATION SCHEMES .. 29
FIGURE IV-1: TYPICAL UPTAKE-DEPURATION CURVE 46
FIGURE IX-1: REPRESENTATIVE SEDIMENT EXPOSURE SYSTEM 93
FIGURE XII-1: SAMPLING SCHEMES FOR COMPARISON-WISE VS.
EXPERIMENT-WISE ERROR RATES 132
FIGURE XIII-1: POSSIBLE REGULATORY STRATEGY FOR HUMAN
HEALTH CRITERIA IN ASSESSING SEDIMENT
CONTAMINATION 146
FIGURES IN APPENDICES
FIGURE V-1A: EFFECT OF SEDIMENT SELECTION ON INGESTED
DOSE IN NATURAL AND ORGANICALLY ENRICHED
SEDIMENT 183
FIGURE IX-1A: CLAMBOX EXPOSURE CHAMBER 200
FIGURE IX-IB: WORMTUBE EXPOSURE SYSTEMS 203
XI
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ABSTRACT
GUIDANCE MANUAL:
BEDDED SEDIMENT BIOACCUMULATION TESTS
Henry Lee II, Bruce L. Boese, Judy Pelletier,
Martha Winsor, David T. Specht, and Robert C. Randall
Bioaccumulation tests with bedded sediments are the most
direct method of deriving tissue residue data required for
evaluation of dredge materials and for quantitative ecological
and human risk assessments. Bioaccumulation tests are also an
important experimental tool for identifying the factors
regulating the bioavailability of sediment-associated pollutants
and to test various Sediment Quality Criteria approaches.
However, the procedures for conducting such tests have not been
standardized, making it difficult to compare studies. This
manual gives detailed guidance on how to conduct "routine" bedded
sediment bioaccumulation tests with marine or estuarine deposit-
feeding organisms. All phases of the process are covered, from
formation of the experimental design, through the actual
exposures, to statistical analysis and interpretation of the
results.
Because the interpretation of tissue residue data is often
relative to "control" and "reference" sites, the acceptability of
such sites is considered. The importance of an appropriate
experimental design, including sufficient statistical power and
replication, is stressed. Based on recommendations for
statistical power and the use of one-tailed tests, a minimum of
eight replicates is recommended. Methods to avoid or reduce
"pseudoreplication, a common statistical problem in toxicity
tests, are also discussed.
For the data generated from the bioaccumulation tests to be
useful in quantitative risk assessments, tissue residues should
be within 80% of the steady-state tissue residue. Based on a
review of both 10-day and 28-day tests, we conclude that the 10-
day test will not meet this criterion for many environmentally
important pollutants, such as PCBs and DDT. Therefore, we
recommend a 28-day exposure as the routine duration for
bioaccumulation tests. Techniques for conducting long-term
exposures (>28 days) and kinetic approaches based on uptake and
depuration rates are also presented for cases when more accurate
estimates of steady-state tissue residues are required.
Sediment collection and preparation, including spiking
techniques, are discussed as are techniques for collecting and
maintaining test species in the laboratory. Based on a number of
criteria, including a required criterion for sediment-ingestion,
five species are recommended as suitable for routine testing.
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Another eight species are identified as potential "secondary"
species. The water quality and sediment requirements for
exposure chambers are discussed, and in most cases, these
requirements can be achieved with relatively simple static or
flow-through systems. Specific sampling schedules and techniques
are given for the routine 28-day exposures. To allow comparisons
among studies, we recommend the Bligh-Dyer method as the standard
lipid technique, or, if another lipid method is used, to
intercompare with Bligh-Dyer.
The statistical analysis of the data is discussed, and the
use of one-tailed tests is recommended when comparing a test
tissue residue(s) to reference or control tissue residue(s), as
would commonly be the case when testing for "no further
degradation". Besides the "no further degradation approach",
other regulatory strategies for using tissue residue data are
presented.
xv
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CHAPTER I. INTRODUCTION
Sediments are the ultimate sink for many of the pollutants
entering the marine/estuarine environment through industrial and
municipal discharges, dredge materials, and non-point runoffs.
Sediments are an especially important repository for compounds
that sorb strongly to particles, such as organic pollutants with
high octanol-water partitioning coefficients (Kow) (e.g., PCBs,
DDT) and many of the heavy metals. As a general rule, these are
the same compounds that bioaccumulate to high levels.
Accumulation of pollutants in bedded sediments (i.e., deposited
rather than suspended sediments) reduces their direct
bioavailability to pelagic organisms but increases their exposure
to benthic organisms.
Bioaccumulation of these sediment-associated pollutants by
benthic organisms can result in a number of ecological and human
health impacts. Bioaccumulation of pollutants can result in
acute and chronic effects in individual benthic organisms, which
ultimately can translate into alterations in benthic community
structure and function. Because the benthos are the primary food
for demersal predators, predation on contaminated benthos is an
important pollutant uptake route for many ecologically and
economically important fishes and invertebrates. Once introduced
into the pelagic foodweb, trophic transfer can spread the
pollutants to higher trophic levels, including sea birds, marine
mammals, and human consumers. If tissue concentrations in
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shellfish or fishes exceed state or Federal regulatory criteria,
such as the FDA Action Levels, areas may be closed to commercial
and recreational fishing with the resultant economic loss.
Given the importance of bioaccumulation by benthic
organisms, regulatory agencies need scientifically credible,
cost-effective methods to measure benthic tissue residues
resulting from exposure to existing sediments, as well as methods
to predict tissue residues resulting from projected levels of
sediment contamination. Additionally, development of
standardized protocols to assess the bioavailability of sediment -
associated pollutants are required to assist in the development
of Sediment Quality Criteria. This need to assess or predict
tissue residues in benthic organisms has long been recognized,
and a variety of field, laboratory, and theoretical approaches
have been developed, as discussed in Appendix 1-1.
Of these approaches, the laboratory test offers great
promise both in assessing existing field sediments or dredge
materials and as an experimental technique to gain insights into
the factors regulating bioavailability. Unfortunately, the
techniques for conducting sediment bioaccumulation bioassays have
varied considerably depending on the specific regulatory or
experimental goals, making it difficult to compare results.
Probably the most frequently used procedure has been the 10-day
test to assess "bioaccumulation potential" of dredge materials
(U.S.EPA/U.S.ACE, 1977), though recent evidence indicates the 10-
day exposure is not adequate (see Chapter IV and Appendix IV-1).
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The purpose of this manual is to develop a standardized
approach to conducting sediment bioaccumulation tests with
sediment-ingesting organisms exposed to bedded marine/estuarine
sediment. The manual covers all aspects of the bioassay
procedure, from experimental design to interpretation of the
results (see Figure 1-1), though detailed guidance in certain
areas (e.g., analytical methods) is beyond the scope of this
manual. These guidelines are designed for the "routine" testing
of sediments and are not tailored toward any specific regulation
or geographical location. The data obtained from the recommended
28-day test should, in most cases, generate the type of
information required for quantitative ecological and human health
risk assessments. Users of this manual must recognize, however,
that the proposed standard 28-day test may underestimate actual
steady-state field tissue residues under certain conditions.
These possible sources of underestimation and alternate test
methods are discussed.
The procedures presented here are based on our own
experiences, results from numerous published sediment
bioaccumulation tests, and to the extent appropriate, on the
standard bioconcentration tests for water uptake (ASTM, 1984),
the draft ASTM guidelines for sediment toxicity (ASTM, 1988a) and
sediment storage, characterization, and manipulation (ASTM,
1988b), and the draft Ecological Evaluation of Proposed Discharge
of Dredged Material into Ocean Waters (U.S. EPA/U.S. ACE, 1988)
in preparation by Battelle. The reader is cautioned that the
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FIGURE 1-1
Steps in Conducting Sediment Bioaccumulation Tests
Define Control and Reference Sediment
Chap. II
Formulate hypotheses and experimental design
Chap. Ill
Set duration and sampling schedule
Chap. IV
i
Collect Sediment
Chap. V
Choose test species
Chap. VII
1
Characterize sediment
Chap. VI
1
Collect/maintain test
species — Chap. VIII
r
I
Construct exposure
system — Chap. IX
Initiate experiment — Chap. X
Pollutant analysis — Chap. XI
Statistical analysis - Chap. XII
Interpretation - Chap. XIII
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final versions of these draft documents may differ in their
recommendations, and the final documents should be consulted.
The reader is also cautioned that the area of sediment
bioavailability/bioassays is highly dynamic and there are diverse
opinions among these various sources. When there is no
consensus, our rationale for recommending a technique is
discussed in the text or one of the appendices. A manual such as
this must be considered a "living" document, as modifications
will inevitably be needed both as a result of future research and
as we gain a better understanding of the information required for
quantitative risk assessments.
Though the methods in the manual have been directed toward
"routine" 28-day tests with marine/estuarine sediment, many of
the laboratory procedures are applicable to experimental
exposures of different durations and to brackish/freshwater
sediments. The standard 28-day test recommended here is not
exclusive of other methods to assess or predict bioaccumulation
(see Appendix 1-1). The specific technique(s) used will depend
upon the accuracy and precision required and the specific goals.
In many cases, the various approaches complement each other and
could be used sequentially or concurrently.
When working with contaminated field sediments or
experimentally spiked sediments, adherence to safe laboratory
practices is critical at all stages, as is disposal of all wastes
in an environmentally proper and legal manner. The specifics of
laboratory safety and waste disposal are beyond the scope of this
manual, but some guidelines are given in Appendix 1-2.
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CHAPTER II: CONTROL VERSUS REFERENCE SEDIMENT
Tissue residues occurring in organisms exposed to the test
sediment(s) are statistically compared to those occurring in
organisms exposed either to "control" or "reference" sediments.
Thus, the difference between control and reference sediments is
critical to the interpretation of the results. Before initiating
a test, determine if a particular sediment constitutes an
acceptable control and whether to use both a control and a
reference sediment or just a control sediment.
A. DEFINITIONS OF CONTROL AND REFERENCE SEDIMENTS
A "control" sediment is a pristine sediment or, more
practically, a sediment with very low levels of pollutants.
Essentially any contaminants in control sediments originate from
the global spread of pollutants and do not reflect any
substantial input from local point or non-point sources. The
comparison of the test sediment versus the control is a measure
(within the statistical limits of the test) of any
bioaccumulation from the test sediment beyond the inevitable
global background contamination. The use of control sediment
also provides information on any contamination from the seawater
or the exposure system. To the extent possible, grain size, TOC,
and other key physical characteristics of the control sediment
should closely resemble those of the test sediment.
-------
In comparison, a "reference" site may contain low to
moderate levels of pollutants. There are two slightly different
ways in which to use a reference sediment. In the first case,
the reference sediment is used as an indicator of the localized
sediment conditions exclusive of the specific pollutant input
being studied. Such sediment would be collected near the site of
concern, and would represent the background conditions resulting
from any localized pollutant inputs as well as the global input.
This is the manner in which reference sediment is used in the
dredge material evaluations (U.S. EPA/U.S. ACE, 1988). This
document states that reference sediment should be collected "near
the disposal site but should not have been influenced by previous
disposal of dredged materials." As the purpose is to compare the
reference sediments to the dredge material (i.e., test.sediment),
the reference sediments should be similar to the dredge material
in grain size, TOC, and other physical-chemical characteristics.
A second, though less common, use of reference sediments is
as a measure of the tissue residues at a particular site before a
specific pollutant input. This differs from the previous
application in that the reference sediment is used as a measure
of background conditions at the specific site rather than as an
indicator of background conditions at another location. For
example, if a sewage outfall pipe was being relocated in an
urbanized near-shore environment, tissue residues in organisms
exposed to sediment collected at the present discharge site (test
sediment) would be compared to those in organisms exposed to
-------
sediment from the new site (reference sediment). The difference
in the tissue residues is an estimate of how much the relocation
of the sewage discharge will increase the tissue residues at the
new site. As the purpose of this comparison is to predict what
will happen at a specific location, it may be impossible to
closely match the physical-chemical characteristics of the
reference and test sediments.
Understanding the type of information generated by reference
sediments is critical for correct interpretation of the tests.
Reference sediments usually contain measurable concentrations of
a number of pollutants. Uncontaminated organisms exposed to
reference sediments will eventually bioaccumulate pollutants to
the level of the organisms living at the reference site.
Comparing tissue residues from test sediments to those from a
reference sediment determines whether the test sediment results
in a significant incremental increase in tissue residues, not
whether there is any bioaccumulation (i.e., bioaccumulation
potential).
Use of a reference site is appropriate when a "no further
degradation" approach is used to determine the suitability of an
industrial or municipal discharge or a disposal operation (see
Chapter XIII). Even in cases when the use of a reference
sediment is appropriate, the sediment should not contain more
than low to, at most, moderate levels of pollutants. If the
reference sediment contains too high a level of a contaminant,
the tissue residues in organisms exposed to reference sediment
-------
FIGURE II-1
Overlap of Confidence Intervals in Organisms
Exposed to Acceptable and Unacceptable Reference Sediments
Unacceptable
tissue residue
level
Tissue
residue
Acceptable Reference
Test
Reference
Unacceptable References
Test
Control
Control
Reference (b)
Reference (a)
-------
may not differ significantly from those in the test sediment even
though the organisms exposed to the test sediments accumulated an
unacceptable tissue residue (see Figure II-1). The situation in
Figure II-1 is an extreme example, but it does illustrate that
the results of a comparison of reference and test sediments
depends upon the absolute level and the variation in the
pollutant concentrations at the reference site.
B. CRITERIA FOR CONTROL AND REFERENCE SEDIMENTS
There are no simple criteria available to judge the
acceptability of a sediment as a control or reference sediment.
Ideally, the concentration of every anthropogenic pollutant
(e.g., PCBs, DDT) in a control sediment should be significantly
indistinguishable from zero, and the concentrations of naturally
occurring compounds (e.g., metals) should be within natural
levels. In practice, it will often be difficult to meet these
criteria. One alternative is to use the pollutant levels in
Table II-1, which gives a range of pollutant concentrations in
control sites on the West coast, as a guide. Sediment with
pollutant concentrations falling within the ranges in Table II-1
should represent adequate control values for the compounds
measured. Alternatively, the concentrations at a putative
control site can be compared to the sediment concentrations
(normalized by the silt-clay fraction) given in NOAA (1988) .
This document presents raw data for both organics and metals for
about 200 near-coastal sites throughout the United States with
the concentrations for the highest and lowest 10 stations in
10
-------
TABLE II-l: Representative Control Sediment Concentrations
COMPOUND
S. CALIF.1
BaPa
BFb
DDT
NAPHC
PAHd
PCB
Ag
As
Cd
Cr
Cu
Hg
Ni
Pb
Zn
(15-150)*
(<5.0-18)*
0.06-2.0
3-15
0.001-2
6.5-40
2.8-30
<1.0
<20.0
<10.0
<70.0
LOCATION
PUGET SOUND2 YAQUINA BAY, OR 3
7-30
7-80
0.03-0.6
3-30e
2-60
<0. 02-1.0
1.2
3-15
3.1-18.3
20.9
10-50
0.02-0.12
13.0
8.0
~
29-66
26.2
<0.01
<0.01f
<0.01
<0.01
0.559
0.47
19.3
6.3
14.5
5.5
26.3
Organics are in ppb dry wt. Metals are in ppm dry wt.
Not Considered Control Values
1 = Southern California (Bascom, 1984; Brown et al.,
Thompson et al.; 1984)
2 = Puget Sound, Washington (Konasewich et al., 1982)
3 = Yaquina Bay, Newport, OR (unpublished data)
f^Benzo (a) pyrene
"Benzofi,b,k)fluoranthene
^Napthalene
dPolyaromatic Hydrocarbons
®(Brown et al., 1984)
£(Schults, D.W., unpublished data.
U.S.EPA, Mar. Sci. Ctr, Newport, OR)
g(Swartz et al., 1984)
1984;
11
-------
tabular form. Sediment concentrations falling within or near the
lowest 10 stations would be acceptable as controls.
Concentrations substantially above those in Table II-1 or the
normalized values for the lowest 10 stations in NOAA (1988)
should not be considered controls, with the possible exception of
sediments containing natural high levels of certain metals.
Because the acceptability of a reference sediment depends in
part on the local background pollutant levels and how the
reference sediment will be used, no specific criteria will be
suggested here. However, the appropriateness of the proposed
reference site should be examined carefully if the silt-clay
normalized concentrations fall in the upper half of the
concentrations presented in NOAA (1988).
Because the regulatory interpretation of bioaccumulation
tests is often based on a comparative approach, having
scientifically defensible definitions and criteria for control
and reference sediments are critical. The suggestions presented
here represent a preliminary attempt at such criteria. To
develop more rigorous criteria, a regional statistical analysis
of the existing sediment data and any ecological effects data is
required. Defining control and reference concentrations by their
frequency of occurrence and by their correlation with adverse
biological effects would then be possible.
It is important to emphasize that the comparison of a test
sediment versus a reference sediment tests for an incremental
12
-------
increase in bioaccumulation and not whether any bioaccumulation
would result. Therefore, a control sediment treatment should
always be used in addition to any reference sediments.
C. STANDARD REFERENCE SEDIMENTS
Variation in organism behavior and physiology can
substantially affect pollutant uptake. For example, uptake in a
test species could vary seasonally in response to changes in
lipid content or temperature, or vary non-seasonally in response
to organism health or site of collection. The extent of this
variation should be assessed, especially if results will be
compared from tests conducted at different seasons or from tests
using organisms collected at different sites.
Organism variation can be assessed by using a "standard
reference sediment," a well characterized sediment containing a
known and constant pollutant concentration. This standard
reference sediment treatment is a positive control and would be
conducted in addition to the normal control, which is a negative
control. Differences among studies in tissue residues in the
standard reference sediment would measure the inherent variation
associated with a test species. Use of a standard reference
sediment would also help in standardizing results from different
laboratories and/or different species.
Although positive controls have been suggested for sediment
toxicity tests (e.g., Johns et al., 1989), they have not been
adequately considered for use in sediment bioaccumulation tests.
Part of the problem is the lack of any standardized sediment
13
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suitable for bioaccumulation tests. In the absence of a national
standard sediment, an interim solution is for each laboratory to
make its own standardized sediment.
Because of potential spatial and temporal variations in
pollutant concentrations in field sediments, the use of a
laboratory spiked sediment is recommended as the standard.
Spiking methods are discussed in Appendix V-l. The sediment used
for the reference should be collected at the site where the test
organisms are collected or, if that is impractical, the physical
characteristics (e.g., grain size, TOO should closely match
those at the collection site. The unspiked sediment can be
stored for long periods either by freezing or by drying.
Although both of these processes can affect the physical
integrity of a sediment, the purpose of the standard reference is
to provide a constant exposure regime, not necessarily a natural
one. Before either of these storage techniques are used,
survival and/or behavioral bioassays should be conducted on
previously frozen or dried uncontaminated sediment to assure that
the technique does not adversely affect the test species. The
sediment would be spiked just before its use.
Ideally, the standard reference sediment would be spiked
with a suite of compounds ranging in chemical properties.
Alternatively, a single neutral organic and/or a single metal
could be chosen as a representative compound(s). A specific PCB
congener, not an Aroclor, is a good candidate for the organics
because of the wealth of information on PCBs, their high
14
-------
bioaccumulation potential, and their resistance to metabolism.
We suggest using 2,2',4,4',5,5' hexachlorobiphenyl (IUPAC #153),
which is the most frequently occurring PCB congener in
environmental samples (McFarland and Clarke, 1989) and is readily
bioaccumulated by marine worms and clams (Rubinstein et al.,
1987; McElroy and Means, 1988; Lake et al., in review;
unpublished data). DDT is another possibility as an organic
reference toxicant. The use of radiolabeled PCB or DDT is
acceptable and would reduce the analytical load, though waste
disposal of a mixed waste could be a problem. Cadmium is
suggested as a general reference metal. Bioaccumulation of
sediment-associated cadmium has been studied in a number of
organisms (e.g., Ahsanullah et al., 1984) and has been suggested
as the reference toxicant for Neanthes growth tests (Johns et
al., 1989)
15
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CHAPTER III. PRINCIPLES OF EXPERIMENTAL DESIGN
Care in experimental design is necessary to ensure the data
generated are relevant to the problem under investigation, as
well as to maximize the information gained relative to the cost.
This chapter will discuss some basic principles of experimental
design and provide examples as they relate to bioaccumulation
tests. The following chapter presents specifics of the
experimental design such as test duration and sampling schedule.
A. OBJECTIVES AND DEFINITIONS
The objectives of these experiments are to quantify the
amount of pollutants bioaccumulated by organisms exposed to
sediments or dredge materials and to test whether this
accumulation is statistically greater than that 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) or dredge
material(s). A control sediment is always required to ensure
there is no contamination from the experimental set-up, but some
designs will also require a reference sediment (discussed in
Chapter II). Uptake from the control sediment or from the
reference sediment (when appropriate) is used to provide baseline
values to compare with uptake from the test sediment. Thus, the
reference sediment functions as the "control" treatment during
comparisons with test sediment but also functions as a test
16
-------
treatment during comparisons with the control sediment. Since
the statistical term "control" treatment could be confused with
the control sediment, in this chapter we will use the combined
descriptor control/reference when referring to the sediment used
as the "control" treatment.
The organism(s) to which a single application of treatment
is applied is the experimental unit. This will be either a
single organism or group of organisms (i.e., composite, see
Section G) placed in an aliquot of a particular type of sediment
in an exposure chamber. The specific type of sediment
constitutes the treatment, for example, the dredge material is
the test treatment. If a clam is placed in a beaker containing
sediment, the clam is the experimental unit and the beaker is the
exposure chamber. If several worms have to be composited to
supply sufficient biomass for chemical analysis, the group of
worms would constitute the experimental unit and the beaker or
aquarium containing them would constitute the exposure chamber.
If an aquarium is physically subdivided, such as containing
several beakers each with an aliquot of sediment, then the
organism(s) placed in each beaker is the experimental unit. The
important concept is that the treatment (sediment) is applied to
the experimental unit as a discrete unit.
Experimental units must be independent and not differ
systematically. This chapter will discuss the procedures
required to assure independence and randomization of the
17
-------
experimental units, as well as the importance of replicating the
experimental units to assure a sufficiently powerful statistical
test.
B. HYPOTHESIS TESTING
Statistical testing requires the establishment of the null
(Ho) and alternative (Ha) hypotheses prior to conducting an
experiment. In most cases, the tests for the bioaccumulation
bioassays will be one-tailed rather than two-tailed. One-tailed
tests are used because the purpose of the experiment is to
determine whether uptake from test sediment is significantly
higher than uptake from control/reference sediment. If uptake is
lower in the test sediment than in the control/reference
sediment(s), presumably no testing is required (see Chapter XII).
For these experiments, a one-tailed test will be performed where
Ho is that the mean tissue residue of organisms in a test
treatment is equal to the mean tissue residue of organisms in the
control treatment. Ha is that the mean tissue residue of
organisms in a test treatment is greater than the mean tissue
residue of organisms in the control treatment. Each test
treatment is compared to the control treatment separately.
Levels of statistical significance are stated by setting
values for Type I and II errors. A Type I error occurs when Ho
is rejected falsely (i.e., Ho is correct but rejected) and the
probability of a Type I error is usually termed "alpha" and given
a significance level of 0.05. In other words, if the tissue
residues are equal (Ho is true) and the experiment were to be
18
-------
repeated many times, an incorrect conclusion (i.e., Ho rejected
with the conclusion that tissue residues are not equal) would
occur 5% of the time. Type I error can be considered the
"discharger's " risk as it is the probability of incorrectly
ascribing bioaccumulation to a sediment or dredge material.
A Type II error occurs when Ho is falsely accepted (i.e., Ha
is true) and is termed "beta". The converse of a Type II error
(1-beta) is the statistical "power" of the test, which is the
probability of correctly rejecting Ho (i.e., Ha is correct).
We recommend a value of 0.05 for beta (power = 0.95) as the
standard for the bioaccumulation tests. This means that if there
were a true difference between test and control/reference tissue
residues and the experiment were to be repeated many times, an
incorrect conclusion (i.e., tissue residues equal when actually
there is bioaccumulation) would occur 5% of the time due to
chance. Type II error can be considered tne "environmental" risk
as it is the probability of incorrectly concluding that a
sediment or dredge material will not result in bioaccumulation.
Using a one-tailed test, as recommended here, instead of a two-
tailed test increases the power of the test given a set Type I
error and reduces the number of replicates required.
By using the same value (0.05) for both Type I and Type II
errors, an equal probability of error is assigned to both the
"discharger" risk and the "environmental" risk. An implicit
assumption of assigning equal risks is that the "cost" of making
either type of error is equal. It is difficult, and often
19
-------
subjective, to compare the monetary costs of pollution
treatment/dredging to the environmental/human health costs of
degradation of marine ecosystems. Therefore, in the absence of
other data, we believe the use of equal risk is the most
defensible procedure. In cases where it can be demonstrated that
the "dischargers" risk is substantially greater than the
"environmental" risk, the beta could be increased to 0.20, which
would reduce the probability (power) of correctly detecting
bioaccumulation to 80%.
Each pollutant must be considered and tested separately.
Different pollutant tissue residues determined from the same
experimental units are not independent and so can not be compared
using the standard statistical tests. The appropriate
statistical procedure for comparisons between different
pollutants (e.g., comparisons of PCB congeners from the same
tissue samples) is repeated measures ANOVA (see Chapter XII).
C. REPLICATION
An important principle in experimental design is the
replication of experimental units. Replication is the assignment
of a treatment to more than one experimental unit, which in the
bioaccumulation experiment is the organism (or composite of
organisms) to which a single treatment (e.g., test or
control/reference sediment) is applied. The variation among
replicates is a measure of the within-treatment variation which
includes random variation among individuals as well as sampling
20
-------
and analytical errors. This variation provides an estimate of
within-treatment error used for assessing the significance of
observed differences between treatments. In experiments without
replication, inferential statistical testing is not possible.
A minimum number of replicates is needed for sufficient
statistical power to determine whether the tissue residues of the
test organisms are greater than those of the control. The number
of replicates required can be calculated from an estimate of the
variance or coefficient of variation of tissue residue values and
a predefined minimum detectable difference between the two means.
The minimum detectable difference is the smallest absolute
difference between two means that is statistically
distinguishable. For example, if the tissue residues in the
organisms exposed to the test sediment must be at least twice as
great as those in the control sediment to be statistically
distinguishable, the minimum detectable difference of the means
is 2.0. Besides the absolute difference between two means, the
minimum detectable difference can be expressed as a proportion of
the mean or as a proportion of the variance (see Appendix III-l).
The smaller the minimum detectable difference, the greater
the number of replicates required for a given significance level
and power. Although there is no consensus on what constitutes an
acceptable minimum difference, we suggest the bioaccumulation
experiment be designed to detect a 2-fold difference between
tissue residues in the test and control sediments or the test and
reference sediments. In most cases, a 2-fold difference should
21
-------
provide a sufficiently precise result to address the ecological
and human health concerns. A smaller minimum detectable
difference would be required if the 2-fold range around the
control/reference sediment overlapped a human health or
environmental criterion (e.g., FDA action limit), though this
should be rare if the control/reference sediments are chosen
correctly.
In some cases, it may not be necessary to distinguish a 2-
fold difference in tissue residues among the treatments, such as
when there are large differences between the treatments. For
example, if the test sediment is suspected of being relatively
contaminated (e.g. pollutant concentrations greater than 50 times
control values), then a 5-fold minimum detectable difference
would be sufficient to find significant differences between the
treatments. In other cases, it may not be possible to achieve
the 2-fold minimum detectable difference with a reasonable
experimental design, such as with compounds with high analytical
variation. When it is impractical to achieve the 2-fold minimum
detectable difference, we recommend using sufficient replicates
to distinguish a 5-fold difference in tissue residues between
treatments, with the caveat that the 5-fold range around the
control/reference does not overlap a sediment or tissue residue
criterion or end-point.
Appendix III-l provides instructions on computing the number
of replicates (n). Figure III-l may be used to determine n from
a coefficient of variation and a minimum detectable difference
22
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to
U)
400-
o 300
i—
•<
-c
200
o 100
o
FIGURE III-1
STATISTICAL SIGNIFICANCE = 0.05
POWER OF TEST =0.95
ONE-TAILED TEST
I
10
I
11
12 13
6=5
6=4
6=3
6=2
6=1
i
H
SAMPLE SIZE
COEFFICIENT OF VARIATION VS SAMPLE SIZE FOR
VARIOUS MINIMUM DETECTABLE DIFFERENCES (5)
EXPRESSED AS A PROPORTION OF THE MEAN
-------
TABLE III-l: Ranges of Coefficient of Variations (CV) for Tissue
Residues Reported for Benthic Organisms
POLLUTANT
ORGANISM
CV
REFERENCE
Cadmium Modiolus demissus
Mytilus edulis
Mya arenaria
Mulinia lateralis
Callianassa australiensis
Mercury Modiolus demissus
Mytilus edulis
Copper Neanthes arenaceodentata
Zinc Nereis divers icolor
Octolasion tyrtaeum
Corbicula fluminea
Kepone Crassostrea virainica
PCB Octolasion tyrtaeum
Corbicula fluminea
Nereis virens
Uca spp.
HCB Ma coma nasuta
BaP Amphipods
Ma coma inquinata
Abarenicola pacif ica
Napthalene Ma coma inquinata
Phenanthrene Ma coma inquinata
Abarenicola pacif ica
Chrysene Ma coma inquinata
Abarenicola pacif ica
*Samples were composited resulting in (usually)
1. Breteler and Saksa, 1985
2. Jackim et al . , 1977
3. Ahsanullah et al . , 1984
4. Pesch and Morgan, 1978
5. Renfro and Benayoun, 1975
6. Mac et al. , 1984
7. Morales-Alamo and Haven, 1983
8. McFarland et al . , 1985
9. Rubinstein et al . , 1984
10. U.S. EPA, 1986b
11. unpublished data, 1989
12. Reichert et al . , 1985
13. Augenfeld et al . , 1982
14. Roesijadi et al . , 1978
4-54%
4-61%
18-22%
35-49%
5-67%
5-34%
5-53%*
8-60%
42%
12-30%
7-8%
8-80%
2-23%
10-74%
5-40%
31-75%
23-33%
4-22%
4-36%
9-24%
50-100%
17-56%
10-31%
11-46%
2-46%
lower CV's
1
1
2
2
3
1
1
4
5
6
6
7
6
6,8
9
10
11
12
13
13
14
13
13
13
13
24
-------
expressed as a proportion of the mean using a significance level
of 0.05 and a power of 0.95 (beta value of 0.05) for one-tailed
tests. If no other information is available, the coefficient of
variations for tissue residues in various benthic species given
in Table III-l can used as guides for Figure III-l. The values
in Table III-l should be used as lower estimates as many were
derived from composites (which will produce lower CVs than will
individual samples), used radiolabeled compounds, and are the
results from successful, published experiments.
ASTM (1984) recommends at least four replicates to determine
bioconcentration factors. Because of the likelihood of a greater
variation in sediment exposures compared to water exposures, we
recommend a minimum of eight replicates as the "default" number
of replicates to provide a statistical power of 95%. Figure III-
2 may be used to determine if eight replicates are adequate for a
specified coefficient of variation and minimum detectable
difference expressed as a proportion of the mean. In some cases,
when variability is low or less power is required, as few as five
replicates can be used, though five should be an absolute
minimum. In this discussion, the number of replicates refers to
the number analyzed for tissue residues and not the number
exposed. It is prudent to include an extra replicate or two for
each treatment in case of mortality or the loss of samples during
chemical analysis.
25
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to
400 H
300-
FIGURE III-2
STATISTICAL SIGNIFICANCE = 0.05
POWER OF TEST =0.95
ONE-TAILED TEST
MINIMUM DETECTABLE DIFFERENCE
THE MINIMUM DIFFERENCE DETECTABLE BETWEEN TREATMENTS
(EXPRESSED AS A PROPORTION OF THE MEAN)
FOR A SPECIFIC COEFFICIENT OF VARIATION AND SAMPLE SIZE (N)
-------
D. RANDOMIZATION
Randomization is the unbiased assignment of treatments to
the experimental units (i.e., organisms or composites of
organisms) ensuring that no treatment is favored and that
observations are independent. This is necessary for valid
statistical testing. Randomization is often performed by using
tables of random numbers. For these experiments, it is important
to randomly assign the organisms to the control and test
treatments, to 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 to randomize the location of exposure
units. For example, a bias in the results may occur if
assignments are not randomized and all the largest animals are
placed in the same treatment.
E. PSEUDOREPLICATION
The appropriate assignment of treatments to experimental
units is critical to avoid a common error in design and analysis
recently 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).
1. Lack of Replication
The simplest form of pseudoreplication is treating
subsamples as true replicates of the experimental unit (Figure
III-3a). For example, two aquaria are prepared, one with
27
-------
control sediment, the other with test sediment, and five
organisms are placed in each aquarium. Even if each organism is
analyzed individually, the five organisms are not true replicates
because the treatment (i.e., sediment type) is applied to the
aquarium as a whole and not to each individual organism
separately. In this case, the experimental unit is the five
organisms and each organism is a subsample, therefore, there is
no replication of experimental units in this particular design.
2. Segregation
A less obvious form of pseudoreplication is the physical
segregation of replicates by treatment, potentially resulting in
a systematic error (bias) and lack of independence. For
example, all the control experimental units are placed in one
area of a room and all the test experimental units are in another
(Figure III-3b). Spatial effects (e.g., different lighting,
temperature) could bias the results for one set of treatments
making it impossible to distinguish true effects of the treatment
from the effects due to the physical layout of the experiment.
Random physical intermixing of the experimental units is
necessary to avoid this type of pseudoreplication.
A more common form of segregating replicates is the use of
separate aquaria for each treatment. For example, segregation
would occur if all the control experimental chambers (e.g.,
beakers) are placed in one aquarium and all the test experimental
chambers in another aquarium (Figure III-3c). Any effects due to
28
-------
FIGURE III-3
Random and Pseudorandom
Replication Schemes
a. no replication
b. segregation
C. segregation
aquarium 1
d. randomized
with interdependent \ 1—
replication g| j—j |—j
aquarium 2
cm
e. completely
randomized
f. randomized
block
cm
cm
a-d
e-f
aquarium 1 aquarium 2 aquarium 3
control experimental unit
test experimental unit
pseudoreplication
strict replication
29
-------
temperatures or different lighting conditions could bias the
results for one of the treatments. Replicate aquaria are
necessary in this case. Section III-F gives suggestions on
addressing this type of pseudoreplication.
3. Randomization With Interdependent Replicates
Randomized spatial interspersion does not necessarily
preclude pseudoreplication. If the replicates are physically
interdependent, spurious effects can bias one treatment over
another. This can occur if all the aquaria replicates of the
control are serviced by the same water supply system while all
the treatment aquaria replicates are serviced by another water
supply system (Figure III-3d). Any differences between supply
systems may potentially bias one set of aquaria over another.
Thus, even if the aquaria replicates are physically interspersed,
the replicates are not independent. To avoid pseudoreplication,
each experimental unit should have its own water or air supply,
all branching off a common supply and there should be no flow of
water from one exposure system to another.
F. AVOIDING OR REDUCING PSEUDOREPLICATION
1. Avoiding Pseudoreplication
Pseudoreplication can be avoided 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 interspersion and independence.
The simplest design that avoids pseudoreplication is the
completely randomized design (Figure III-3e). In this design,
30
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treatments are randomly assigned to the experimental units
independent of location and each experimental unit is maintained
in a separate exposure chamber with a separate water and air
supply.
A randomized block design is also appropriate. A block is a
set of relatively homogeneous units to which treatments are to be
applied, such as all the beakers within an aquarium. In the
randomized block design, all the treatments are randomly assigned
to each block, and there are multiple blocks. For example, if
there are two treatments and eight beakers per aquarium, each
aquarium is randomly assigned four beakers with control sediment
and four beakers with test sediment (for another example see
Figure III-3f). One drawback of this design, however, is that
since both test and control organisms are in one aquarium
(block), the potential exists for contamination of controls by
test sediment. This is especially likely with organisms that
eject sediment into the water, such as Macoma during the
production of pseudofeces. If this design is used, the aquaria
and/or control exposure chambers need to be monitored to assure
that cross-contamination does not occur.
2. Reducing Pseudoreplication Effects
Totally avoiding pseudoreplication may be difficult or
impossible given resource constraints. For example, one common
experimental design segregates the experimental treatments in
separate aquaria. In this case, the beakers containing the test
sediment are placed in separate aquaria from beakers containing
31
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the control sediment (see Figure III-3C). Such a design avoids
the problem of cross-contamination between the test and control
sediment and does not require a separate aquarium for each
beaker. However, because the beakers are segregated by treatment
type, their distribution is not random. In such cases, the
experimental unit may be defined as the replicated unit
(organism(s) in the beaker with each beaker as a replicate), but
with the stated assumption that there is no effect due to the
physical segregation (aquaria effect in this example).
With this design, we recommend using replicate aquaria for
each treatment type to enable comparison of results between
aquaria within a given treatment using a nested ANOVA. If
aquaria effects are apparent, the data from one or more aquaria
may be considered invalid, or the differences due to the aquarium
effect may be deemed trivial compared to the treatment effects.
For example, if there is a significant difference among test
aquaria results, but that difference is much less than the
difference between test and control aquaria, the aquaria effects
may be considered unimportant to the results of the experiment.
However, moderate to large differences between aquaria of the
same treatment would suggest a local contamination problem or
other type of bias and the experiment should be repeated. If no
significant aquaria effects are detected, the organism(s) within
each beaker are properly considered the experimental unit and
each beaker a replicate. The analysis (see Chapter XII) is then
performed as if the beakers were not segregated into aquaria.
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G. COMPOSITING SAMPLES
Compositing consists of combining samples (e.g., organisms,
sediment) and chemically analyzing the mix rather than the
individual samples. The chemical analysis of the mix provides an
estimate of the average concentration of the individual samples
making up the composite. Compositing will be used in
bioaccumulation experiments primarily when the biomass of an
individual organism is insufficient for chemical analysis.
Several individuals can be composited into a single experimental
unit with sufficient biomass and the analysis performed on the
composite. Compositing is also used when the cost of analysis is
high. For example, sediment pollutant analysis can be based on
sediment composited from several exposure chambers of the same
treatment to reduce the analytical sample load. Replicate
sediment or tissue composites (i.e., experimental units) are
required if statistical testing is planned.
For the tissue composite to be unbiased, the individuals
must be randomly assigned to the various treatments. Each
organism or sediment sample added to the composite must be of
equal size (i.e., wet weight) and the composite must be
completely homogenized before taking a sample. If compositing is
performed in this manner, the value obtained from the analysis of
the composite is the same as an average obtained from analyzing
each individual sample (within any sampling and analytical
errors) . If replicate composites are made, the variance of the
replicates will be less than the variance of the individual
33
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samples, providing a more precise estimate of the mean value.
This increases the power of a test between means of composites
over a test between means of individuals or samples for a given
number of samples analyzed.
However, if composites are made of individuals or samples
varying in size (e.g., varying weights of sediment samples) or
quality (e.g., disproportionate number of gravid females in one
composite), the value of the composite and the mean of the
individual organisms or sediment samples are no longer
equivalent. The variance of the replicate composites will
increase, decreasing the power of a test between means. In
extreme cases, the variance of the composites can exceed the
population variance (Tetra Tech, 1986a). Therefore, it is
important to keep the individuals or sediment samples comprising
the composite equivalent in size and quality. If sample sizes
vary, consult the tables in Schaeffer and Janardan (1978) to
determine if replicate composite variances will be higher than
individual sample variances, which would make compositing
inappropriate.
It is not advisable to composite samples if an estimate of
the population variance is required or if information about the
range in values obtained for individuals is needed. For example,
tissue samples should not be composited if it is important to
know the percent of individuals exceeding the FDA Action Limits.
Compositing also requires more individuals (assuming individuals
34
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can be analyzed) so it is not advised when space or cost keeps
the number of individuals at a minimum. When there is extra
sediment or tissue, archive individual samples in case a measure
of the population variance or the concentration in a particular
exposure chamber is desired latter.
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CHAPTER IV: TEST DURATION AND SAMPLING SCHEDULES
Besides the statistical issues addressed in the previous
chapter, an environmentally-relevant bioaccumulation test
requires an appropriate exposure duration and sampling schedule
for organism, sediment, and water samples. This chapter
discusses these topics for the standard 28-day and long-term
bioaccumulation tests, as well as a kinetic approach. Additional
discussion of the 10-day versus 28-day tests are give in Appendix
IV-1. Alternative test designs, which may be applicable under
special circumstances or for research purposes, are presented in
Appendix IV-2.
A. STANDARD 28-DAY BIOACCUMULATION TEST
1. Steady-State and Duration
Ideally, the duration of a bioaccumulation test should be
sufficient for the organisms to reach steady-state tissue
residues, where steady-state is operationally defined as the lack
of any significant difference (ANOVA, alpha = 0.05) among tissue
residues taken at three consecutive sampling intervals (ASTM,
1984). The time to reach or approach steady-state varies
drastically among different compounds, but in general, the tests
should be designed to generate environmentally-relevant data on
high Kow organics (e.g., PCBs, DDT) and heavy metals. Therefore,
we recommend a 28-day exposure as the standard duration. As
discussed in Appendix IV-1, a 28-day exposure will result in
tissue residues within 80% of the steady-state tissue residues in
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most cases. When steady-state is not approached within 28 days,
tissue residues of organics usually appear to be within 2-4 fold
of steady-state concentrations (Table IV-1), which is considered
acceptable for the ASTM bioconcentration test (ASTM, 1984).
Although a 28-day sediment exposure will not assure attainment of
steady-state for all environmentally important compounds, it will
improve the predictive power of the test compared to the commonly
used 10-day exposure. For cases where more accurate estimates of
the steady-state tissue residues are needed, a long-term
bioaccumulation test (Section B) or a kinetic uptake approach
(Section C) is recommended.
2. Biotic Sampling Schedule
Biological samples are used to determine the amount of
pollutants accumulated from the test sediment and to
statistically compare these values to the amount of pollutants
accumulated from control and reference sediments. To set the
baseline conditions for these comparisons, bioassay organisms
should be analyzed for pollutant and lipid content immediately
before initiation of the experiment (tr, samples) . As discussed
in the previous chapter (Chapter III), eight replicates are
assumed as the number required to achieve sufficient statistical
power. Therefore, eight replicate organisms or composites (i.e.,
experimental units) should be analyzed at tQ. The organisms
sampled at tQ should be chosen randomly from the same set of
organisms used in the various sediment treatments. If
compositing of individuals is necessary to obtain sufficient
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TABLE IV-1: Percent of Steady-State Tissue Residue of Neutral
Organics Obtained After 10 and 28 Day Exposures to
Bedded Sediment.
Compound
% of Steady-State
Tissue Residue
10 -DAY 28 -DAY Species
Estimated
by Ref .
Phenanthrene 100 100
Benzo(a)pyrene 96 100
Benzo(a)pyrene 96 100
Phenanthrene 94 100
Hexachlorobiphenyl 88 100
Phenanthrene 67 95
Aroclor 1260 53 100
Benzo(a)pyrene 43 75
Chrysene 43 87
Hexachlorobenzene 35 70
Benzo(a)pyrene 32 66
Aroclor 1242 29 82
Aroclor 1260 27 100
Aroclor 1254 27 100
Aroclor 1242 18 87
Aroclor 1254 12 82
Aroclor 1254 9 25
Macoma inquinata G
Hexagenia limbata K
Mysis relicta K
Mysis relicta K
Hexagenia limbata K
Pontoporeia hoyi K
Macoma balthica G
Macoma inquinata K
Macoma inquinata G
Macoma nasuta K
Pontoporeia hoyi K
Cerastodema edule G
Cerastodema edule G
Cerastodema edule G
Nereis virens G
Macoma balthica G
Nereis virens K
1
3
3
3
3
3
4
1
1
2
3
4
4
4
4
4
5
K = Steady-state tissue residue estimated from kinetic uptake
model.
G = Steady-state tissue residues estimated from graphs of tissue
residues versus time. Often, the 10-day and 28-day values
had to be interpolated. "Steady-state" was defined as the
time period with the maximum tissue residue.
SOURCES:
1 = Augenfeld et al., 1982
2 = Boese et al., in press and unpublished data
3 = Landrum and Poore, 1988
4 = Langston, 1978
5 = McLeese et al., 1980
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biomass, the same compositing scheme should be used for all
sampling periods throughout the experiment. At the end of the
28-day test period (t28), eight replicate organisms or composites
should be taken from each of the treatments and analyzed for
pollutants and lipids. For comparing test and control sediments,
the simplest design results in 24 tissue samples (8 controls at
tQ, 8 controls at t28, and 8 test at t28) .
We recommend including an extra one or two replicates in
each treatment in case a sample is lost. Additionally, several
extra individuals or composites should be taken at the initiation
of the experiment. These extra samples should be frozen until
the tissue residue data has been analyzed and interpreted. The
method of physically sampling the organisms is discussed in
Chapter X.
Time-series samples may be taken during the 28-day exposure
to document uptake kinetics. This type of information can be
very helpful even if it is necessary to limit the analytical load
by taking only a single sample or, preferably, a single composite
at each sampling period. However, if the data will be
statistically compared to determine if steady-state has been
attained, replicates are required at each sampling period. The
sampling interval for these samples should approach a geometric
progression with sampling periods of no greater than one week
(e.g., day 0, 2, 4, 7, 14, 21, and 28). A sample at 10 days is
recommended if there are previous 10-day exposure data. Begin
the series on Monday to avoid weekend sampling.
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3. Abiotic Sampling: Frequency and Replicates
The physical and chemical properties of each test, control,
and reference sediment need to be characterized immediately after
collection. This includes, at a minimum, grain size
distribution, moisture content, pollutant concentrations, and TOG
(or LOI) (see Chapter VI). Depending upon the length of storage,
it may be necessary to remeasure these physical and chemical
parameters, with the possible exception of grain size
distribution, immediately prior to the start of the
bioaccumulation test (i.e., tQ). If these tQ samples will be
statistically compared to samples taken at the end of the test
period (t28), eight replicate samples are required.
At the end of the bioaccumulation test (t28), take sediment
samples from each exposure chambers for measurement of pollutant
concentrations, TOG, and moisture contents. It is usually not
necessary to remeasure grain size. Preferably, these analyses
should be conducted on the sediment from each beaker or aquarium
(i.e., experimental unit). Measurements on individual
experimental units may help explain any unexpected variation
among the replicates. If eight replicates are used per
treatment, this would result in a total of 24 sediment samples (8
controls at tQ, 8 controls at t28, and 8 test samples at t28).
If this is too large an analytical load, an alternative is
to analyze a composite sample from each treatment composed of
equal aliquots of sediment from each beaker or aquarium within
the treatment. Additionally, a sediment sample from each beaker
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or aquarium should be taken and archived. If the tissue residue
data are more variable than expected or if there are "unusual"
data points, these individual sediment samples should be
analyzed. Additionally, individual sediment samples should be
analyzed if the differences in pollutant concentrations in the tQ
and t2g sediment samples are greater than would be expected from
analytical variation alone. It would then be possible to
determine if significant changes in pollutant concentration or
TOG had occurred during the course of the experiment.
B. LONG-TERM UPTAKE TESTS
1. Criteria and Limitations
In some cases, body burdens will not approach within 80-90%
of the steady-state body burdens in a 28-day test (see Table IV-1
and Appendix IV-1). Organic compounds exhibiting these kinetics
will likely have a log Row > 5, be metabolically refractory
(e.g., highly chlorinated PCBs, dioxins), and exhibit low
depuration rates. Many of these same organic compounds biomagnify
in aquatic food webs and pose a human health risk. Additionally,
tissue residues of several heavy metals may gradually increase
over time so that 28 days is inadequate to approach steady-state.
Depending on the goals of the study, it may be necessary to
conduct an exposure longer than 28 days (or a kinetic study as
discussed in Section C) to obtain a sufficiently accurate
estimate of steady-state tissue residues of these compounds.
Although these longer term studies generate more accurate data
for these compounds, they require greater resources, increase the
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analytical load, and increase the likelihood of problems
involving the maintenance of the organisms and temporal changes
in sediment pollutant concentrations.
2. Biotic Sampling
In the long-term studies, the exposure should continue until
steady-state body burdens are attained. As mentioned, steady-
state is documented by the lack of any statistical difference in
tissue residues in three consecutive sampling intervals (ASTM,
1984). ASTM (1984) recommends a minimum of five sampling periods
(plus t0) when conducting water exposures to generate
bioconcentration factors (BCFs). For bioconcentration tests,
ASTM (1984) recommends sampling in a geometric progression with
sampling times reasonably close to S/16, S/8, S/4, S/2, and S,
where S is the time to steady-state. This sampling design
presupposes a fairly accurate estimate of time to steady-state,
which is often not the case with sediment exposures.
To document steady-state from sediment exposures, we
recommend placing a greater number of samples at and beyond the
predicted time to steady-state. With a pollutant expected to
reach steady-state within 28-50 days, samples should be taken at
days 0, 7, 14, 21, 28, 42, 56, and 70. If the time to steady-
state is much greater than 42 days, then additional sampling
periods at 2 week intervals should be added (e.g., day 84).
Slight deviations from this schedule (e.g., day 45 versus day 42)
are not critical, though for comparative purposes, samples should
be taken at t2g. An estimate of time to steady-state may be
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obtained from the literature or approximated from structure-
activity relationships (Appendix IV-3), though these values
should considered the minimum times to steady-state.
Compared to the ASTM bioconcentration sampling schedule,
this schedule increases the likelihood of statistically
documenting that steady-state has been obtained though it does
not document the initial uptake phase as well. If accurate
estimates of the first-order uptake coefficient (kl, see Section
C) are required, add sampling periods during the initial uptake
phase (e.g., days 0, 2, 4, 7, 10, 14, 17, etc.).
One problem with longer exposures is the greater probability
of the test organisms reproducing. Spawning can drastically
affect lipid content and possibly pollutant concentrations
(Niimi, 1983). Additionally, because many species die after
spawning, it is prudent to add extra replicates. Increasing the
total number of replicates by an additional 10-20% should suffice
in most cases. If not needed, archive these extra individuals at
the end of the test as replacement samples in case of analytical
failures or analyze them to increase the statistical power of the
final sampling period.
3. Abiotic Samples
The bioavailable fraction of the pollutants as well as the
nutritional quality of the sediment are more prone to depletion
in these extended tests than in the 28-day exposures. To
statistically document whether such depletions have occurred, at
least eight replicate sediment samples are required for physical
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and chemical analysis from each sediment type at the beginning
and the end of the exposure. Additionally, we recommend
archiving sediment samples from every biological sampling period.
To minimize the depletion of sediment pollutants or
nutrients, completely replace the sediment with stored sediment
or freshly spiked sediment on a regular basis (e.g., monthly).
Sediment must be renewed carefully to avoid damaging the test
organisms, especially polychaete worms. Another way to minimize
depletion of pollutants is to add fresh sediment periodically
(see Chapter X). Over a long experiment, however, the exposure
container may be entirely filled, necessitating the replacement
of the sediment anyway. Replenishment sediment should be sampled
and analyzed for the recommended parameters. Do not feed the
organisms a supplemental food (e.g., fish flakes) as this will
reduce exposure to ingested sediment and may result in an
underestimation of sediment bioavailability and steady-state
tissue residues.
C. ESTIMATING STEADY-STATE TISSUE RESIDUES FROM UPTAKE AND
DEPURATION RATES
Several methods have been published which can be used to
predict steady-state pollutant levels from uptake and depuration
kinetics (Spacie and Hamelink, 1982; Davies and Dobbs, 1984).
All of these methods were derived from fish exposures and most
use a linear uptake, first-order depuration model which may be
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modified for uptake of pollutants from sediment:
Ct(t) = kl*Cs/k2*(l-e~k2*t) (1)
Where: Ct = pollutant concentration in tissue at time t
Cs = pollutant concentration in sediment.
kl = uptake constant (day"1 )
k2 = depuration constant (day"1)
t = time (days)
As time approaches infinity, the maximum or equilibrium
pollutant concentration within the organism (Ctmax) becomes:
Ctmax = Cs*kl/k2 (2)
Correspondingly, the bioaccumulation factor (BAF) for a
compound may be estimated from:
BAF = kl/k2 (3)
The kinetic approach requires an estimate of the uptake rate
constant (kl) and the depuration rate constant (k2), which are
determined from the changes in tissue residues during the uptake
phase and depuration phase, respectively. The uptake experiment
should be short enough that an estimate of kl is made during the
linear portion of the uptake phase (Figure IV-1) to avoid an
unrealistically low uptake rate due to depuration. The
depuration phase should be of sufficient duration to smooth out
any loss from a rapidly depurated compartment (Figure IV-1) .
Unless there is reason to suspect that the route of exposure will
affect the depuration rate, it is acceptable to use a k2 derived
from a water exposure. The durations of the uptake and
depuration experiments will vary with animals species,
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FIG. IV-1: Idealized Uptake - Elimination Curve
"k M
Ct(t) =
Uptake Phase
k2/k *
( 1-e
Cs = k * Ct
Steady-state
Linear uptake phase
* Cs » k
Ct
Elimination Phase
Fast component
Slow component
(Contaminated Sediment) '(Clean Sediment)
TIME
tissue concentration (ug/g)
uptake constant (1/time)
sediment concentration (ug/g)
elimination constant (1/time)
t = time
Ct
k-,
Cs
2
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compound, pollutant concentration, analytical detection limits,
and test sediment. As a result, no specific guidelines will be
presented here. For a discussion of this method for
bioconcentration studies in fish, see Davies and Dobbs (1984),
Spacie and Hamelink (1982), and the ASTM standard practice for
conducting bioconcentration tests (ASTM, 1984). Effects of
growth on the estimation of kl and k2 and how to correct for
growth dilution effects on the estimate of steady-state tissue
residues are discussed in Appendix IV-2.
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CHAPTER V. SEDIMENT COLLECTION, HOMOGENIZATION,
MANIPULATION, AND STORAGE
Bioaccumulation tests use sediments collected in the field
and brought back to the laboratory or manipulated experimentally
in the laboratory. In both cases, the handling can result in
loss of fine sediments, interstitial water, and water soluble
compounds; oxidation of compounds; or contamination by metals and
organics. This disruption can change physicochemical properties
such as grain size distributions, pollutant concentrations,
sorption equilibria, speciation, and complexation, thereby
affecting pollutant bioavailability (Plumb, 1981; Jennett et al. ,
1980; Holme and Mclntyre, 1984). Although some changes are
unavoidable, they can be minimized with appropriate techniques.
In this chapter, we provide guidelines on handling sediments
during and after collection. We cannot, however, formulate a
standard operating procedure applicable in all cases because the
techniques used depend on the goal of the experiment and the
pollutants of concern. In particular, techniques optimally
suited to study metals may not be not be suitable for organic
compounds (Plumb, 1981; ASTM, 1988b).
A. SEDIMENT COLLECTION AND TRANSPORT
1. Depth of Collection and Sediment Collection Techniques
The depths from which sediments are collected can affect
bioaccumulation test results; therefore, a consistent depth
should be used in all collections. We recommend sampling the
upper 2-3 cm layer, a depth commonly used for toxicity and
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bioaccumulation tests and sediment chemistry (e.g., Jenne et al.,
1980; Plumb, 1981; Swartz et al., 1985a, 1986; NOAA, 1988;
Ferraro et al., 1990). Advantages in sampling the upper layer
include that the sediment is more recently deposited, more
consistent in pollutant concentrations, in contact with the
overlying water, and the most biologically active zone. However,
tests on dredge materials may require that representative samples
be collected from deeper layers, up to several meters deep, in
areas intended for dredging.
To collect intertidal sediment samples by hand, use
shovels, scoops, spatulas, or coring tubes. Hand skim or core
with one of the above mentioned tools the upper 2 cm sediment
layer. To maintain the sample layers intact, deposit the
sediment sample into an appropriate container or, if a corer is
used, plug the top and bottom of the tube. Core samples may be
sectioned later at specific depth-intervals for analytical and
bioaccumulation tests (Plumb, 1981; Holme and Mclntyre, 1984;
NOAA, 1988) .
Box corers and benthic grabs are commonly used to collect
sediments in subtidal waters. Sampler choice will vary according
to firmness of substrate, volume of sediment needed, and type of
ship available. Box corers are the preferred collection device
because they disturb sediment layers the least and retain fine
particles. A Smith-Mclntyre or modified Van Veen grab, though
more disruptive to sediment layers than a box corer, is
acceptable. Compared to the box corer, these grabs operate in
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sandier bottoms, are easier to handle, require fewer personnel,
and operate in heavier seas (Plumb, 1981; Holme and Mclntyre,
1984; Tetra Tech, 1986b; NOAA, 1988) . Scrape surficial sediment
from the grab or box corer samples and immediately store in
appropriate containers. Consider flocculent material part of the
sample (Lauenstein and Young, 1986) .
If depth profiles are of interest, the original sediment
layering must be preserved. Take core samples from the center of
the grab sample once on shipboard and section them vertically at
specific depth intervals (Plumb, 1981). To minimize oxidation
and changes in other chemical properties, place plastic or
Teflon bags or containers of appropriate composition and
diameter over the ends of core tubes and extrude samples to
specified depths.
Construct all collecting equipment with appropriate
materials and clean equipment to reduce the possibility of
contamination. When organic pollutants are the primary concern,
avoid contaminating sediment with various plastics, especially if
phthalate esters, used in flexible plastics, will be quantified.
When metals are the primary concern, avoid contaminating
sediments with any metal, including stainless steel. Subsampling
from the center of grabs as well as coating, covering, or lining
equipment with silicone rubber, TeflonR, plastic, polypropylene,
or polyethylene will eliminate direct contact of samples with the
equipment. For instance, TeflonR bags can be used to cover
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stainless steel scoops (Lauenstein and Young, 1986). When both
metal and organic contaminants are the interest, use glass when
possible.
To remove organics/metal contamination, initially wash the
equipment with a non-phosphate detergent, then consecutively
rinse with distilled water, a water-miscible organic solvent, 5-
10% hydrochloric or nitric acid, and finally deionized-distilled
water (Plumb, 1981; Lauenstein and Young, 1986; U.S. EPA/U.S.
ACE, 1988). Bake (>350°C) all glass equipment (e.g., jars,
corers, trays) before use. Glassware to be used in metal
analyses should be stored wrapped in TeflonR sheets or plastic
wrap, whereas glassware to used in organic analyses should be
stored wrapped in TeflonR or aluminum foil. Wash new plastic
containers and equipment as described and then leach them with
distilled water before use. Rinsing grabs or corers with
seawater between stations should suffice in most studies, though
it may be necessary to use a brush or a detergent to remove
highly cohesive sediments. When it is critical to remove all
contaminants, Lauenstein and Young (1986) recommended rinsing
grabs or corers with methylene chloride, followed by a seawater
rinse. However, methylene chloride generates a hazardous
substance, and its use on a ship would have to be carefully
controlled so as not to endanger the workers or release a
hazardous waste into the environment. Safer alternatives are
methanol and ethanol alcohol.
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Specifics of the field sampling design, such as the number
of sites and the number of samples per site, depend upon the
goals of the study and the type of spatial resolution required.
Guidance for designing field sampling programs can be found in
Green (1979), Elliott (1983), and Lauenstein and Young (1986).
2. Field Measurements
Upon collection, immediately determine sediment temperature
and salinity by inserting a thermometer and an electrode 1 cm
into the center of each sample (Plumb, 1981; Swartz et al.,
1986). When metals are of interest, immediately measure pH and
Eh, both of which require undisturbed samples (Holme and
Mclntyre, 1984). Important information recorded with each sample
should include the site (name, with latitude and longitude to
tenths of a minute), replicate number, depth, sampler
description, numbers and kinds of subsamples, sediment
characteristics, temperature, salinity, pH or Eh if measured,
odor-color, penetration depth, sieve size, vessel size, date and
time, weather conditions, names of chief scientist and team
members, and comments (Lauenstein and Young, 1986).
3. Field Storage and Sediment Transport
Physical, chemical, and biological changes in sediment
samples can occur rapidly, resulting in changes in sediment
quality and/or bioavailability during the transport of sediment.
Temperature, pH, and dissolved oxygen are often the rate
controlling factors for these changes (Jennett et al., 1980). To
diminish these effects, store the sediment sample in a bag or jar
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immediately after collection. TeflonR containers or brown
borosilicate glass jars with Teflon -lined lids are recommended
for both metal and organic samples, but regular glass jars with
TeflonR-lined lids are acceptable (Lauenstein and Young, 1986).
Containers need to be completely cleaned and stored in a covered
container to avoid contamination. Cleaning protocols used for
the sampling equipment also apply to storage containers.
Fill jars and bags completely with sediment to eliminate
airspace and retard oxidation of metals, but retain as much of
the interstitial water as possible (Lauenstein and Young, 1986;
U.S. EPA/U.S. ACE, 1988). Refrigerate sample containers in
insulated cartons or ice chests immediately after collection. To
maintain a temperature near 4°C, provide containers with
prefrozen, jelled refrigerant packs (e.g. Blue IceR) or ice.
Make sure that samples are protected from the refrigerant to
avoid cross contamination and freezing of the sample.
Shipping containers must be durable and leak-proof or lined
with two heavy duty plastic bags. Add adequate absorbent
material to soak up any leaked liquid. Pack samples tightly,
using dividers between glass containers and fill all empty spaces
with packing material. Mark containers with "This End Up" and
"Fragile" labels. Ship samples by "overnight" or "24 Hour"
carrier to the laboratory immediately after completion of
sampling to protect sample quality. Refrigerate samples at 4C°
upon arrival. Guidance for shipping hazardous materials can be
found in CFR 49, Parts 100-177.
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B. SEDIMENT SPIKING AND MANIPULATION
Besides using field collected sediments, researchers can
experimentally manipulate sediments to test pollutant
bioavailability under defined conditions. Addition or spiking of
pollutants to sediments is the most frequent type of
manipulation. Other types of manipulations include the addition
of inert substances for producing a less polluted sediment and
the alteration of sediment organic content or particle size.
Sediment spiking and manipulation techniques have not been
standardized, and until standard methods are developed or the
techniques are intercalibrated, exercise caution when comparing
results from different techniques. Because manipulations can
alter properties of sediments, prepare and manipulate control
sediments in the same manner as test sediments. Several sediment
manipulation techniques are outlined in Appendix V-l.
C. LABORATORY SEDIMENT STORAGE
Keep the time between the collection and/or spiking of a
sediment and its use in bioassays to a minimum. If there is a
delay of more than about 24 hours, store the sediment in air-
tight containers at 4°C in the dark. Because freezing may
physically disrupt sediment, sediments for biological testing
should not be frozen or freeze-dried (Tetra Tech, I986b), with
the possible exception of sediment stored for use as a "standard
reference sediment" (Chapter II). If metals are the major source
of contamination in a field sediment, the sediment should be
54
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stored in the absence of air to minimize oxidation of reduced
forms. Nitrogen can be used to fill the headspace in the
container. Air-tight glass containers are recommended for
sediments polluted with either metals or organics (Plumb, 1981;
Lauenstein and Young, 1986). High density polyethylene and
TeflonR containers are also acceptable. Remove large organisms
and extraneous material, such as bivalves or twigs, from the
sediment before storing.
There is no consensus on maximum storage time other than
that it should be kept to a minimum. This lack of consensus
reflects the use of different end-points, sediment types, and
storage procedures. Little information exists storage effects,
though pollutants that are volatile, biodegrade rapidly, or
undergo rapid oxidation-reduction reactions should be the most
prone to changes in concentrations and/or bioavailability. Given
the present state-of-the-art, the maximum time recommended for
storage of dredge materials used for biological testing is 6
weeks at 4°C (U.S. EPA/U.S. ACE, 1988), which seems reasonable.
The storage of samples for analytical and physical analysis is
discussed in Chapter XI.
D. SEDIMENT PREPARATION AND HOMOGENIZATION
Before using a field sediment, remove any extraneous
materials (e.g., macroalgae, twigs, garbage, and rocks) and large
organisms (e.g., bivalves). Disturb the sediment as little as
possible during this process. The simplest technique is to
55
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gently spread the material out in a glass pan and remove any
large objects with forceps. However, if metals are of primary
importance, keep contact with air to a minimum and use plastic
tools.
If a field sediment is going to be experimentally
manipulated (e.g., spiked with a pollutant), sieve the sediment
through a 1-2 mm mesh sieve to remove the extraneous materials.
Using as small a volume of water as possible, sieve the sediment
over a large container (e.g., garbage pail). After letting the
suspended fines settle for 6 to 24 hours, carefully siphon off or
decant the overlying water and mix the settled fine particles
back into the sediment.
After settling or storing sediments, mix them well
immediately before taking aliquots for chemical analysis,
spiking, or using in bioaccumulation tests. This will assure
homogeneity as well as mix any separated interstitial water back
into the sediment. Stir with a spoon or rod made of an
appropriate material. If grab samples were divided into several
containers, mix the respective sediment samples together before
sampling or using in biological tests. Large masses of sediment
can be manually mixed in an appropriately cleaned glass tray or
plastic tub, or placed in jars and rotated on a rolling mill.
Homogenize control sediments in the same manner as test
sediments.
Visually inspect the sediment to judge the extent of
homogeneity. Excess water on the surface of the sediment can
56
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indicate separation of solid and liquid components. If a
quantitative measure of homogeneity is required, take replicate
(3-8) subsamples from the sediment batch and analyze for TOC,
chemical concentrations, or the percent fines.
Jenne et al. (1980) cautioned against prolonged stirring
which can abrade floes and change the sediment's physicochemical
properties, such as dissolved organic matter (DOM). However, all
changes to the sediment are probably impossible to avoid. Recent
results suggest that even stirring can increase interstitial
water DOM concentrations (DeWitt, T. pers. comm. OSU, Mar. Sci.
Ctr., Newport, OR).
57
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CHAPTER VI. SEDIMENT CHARACTERIZATION
Bioavailability of sediment-associated pollutants depends on
the physical and chemical composition of the sediment, which is
often specific to a particular site and may change due to
experimental manipulations (Gambrell et al., 1980; Jennett et
al., 1980). Therefore, always measure the parameters potentially
affecting bioavailability: pollutant concentrations, grain size
distributions, organic carbon, and total solids contents (Plumb,
1981; NOAA, 1988). A number of other sediment and interstitial
water measurements such as pH, Eh, cation exchange, and acid
volatile sulfides will aid interpretation of the results,
especially those of metals.
A. GRAIN SIZE ANALYSIS
Grain size analysis is the measure of the frequency and
distribution of the disaggregated mineral particles comprising
the sediment. Distributions are commonly reported on the
Wentworth scale, which classifies particles as coarse sand,
medium sand, fine sand, very fine sand, silt, and clay (Folk,
1980, Holme and Mclntyre, 1984) (Table VI-1). Particle sizes are
either expressed in millimeters or on a phi scale, where phi =
-Iog2 particle diameter in millimeters. Quantification of the
fine fraction (silt-clays <0.0625 mm or > 4 phi) is important
because pollutants predominantly associate with, and many
deposit-feeders ingest, this fraction.
58
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TABLE VI-1: Wentworth Grade Classification of Sediment
GRADE LIMITS
NAME
mm Phi
Coarse sand 1.000 - 0.500 +1.0
Medium sand 0.500 - 0.250 +2.0
Fine sand 0.250 - 0.125 +3.0
Very fine sand 0.125 - 0.062 +4.0
Silt 0.062 - 0.004 +8.0
Clay < 0.004 >8 . 0
59
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The sand fraction of a sediment is determined by first
drying the sediment (air dried or at <105°C) and then removing
the organic matter with 6% hydrogen peroxide (Buchanan, 1984) . A
known mass of dry sediment is then sieved through a standard
series of sieves graded on the Wentworth scale (e.g., <.500,
<.250, <.125, <.0625 mm, and pan) or on the phi scale (e.g., at
1/2 phi intervals) (Buchanan, 1984; Holme and Mclntyre, 1984).
Manually or mechanically shake about 25 grams of dry sediment for
about 20 minutes and then weigh the contents of each sieve. To
further characterize the silt-clay fraction (i.e, the sediment in
the bottom pan), an hydrometer and pipette analysis is used
(Jenne et al., 1980; Buchanan, 1984; Holme and Mclntyre, 1984).
Suspend the pan fraction in a graduated cylinder of
deionized/distilled water, take periodic water samples from a
known depth, and weigh the resultant suspended particles. The
weight of the silt-clay fractions can then be calculated from the
estimated settling velocities of particles of different sizes.
B. TOTAL SOLIDS CONTENT
Sediments are composed of both solids and interstitial water
(IW), with the relative proportion of the two phases varying due
to physical factors (e.g., percent sand) and biological factors
(e.g., intensity of bioturbation). The total solids content, the
percent of wet sediment comprised of particles, is used to
convert sediment pollutant concentrations from wet to dry weights
and to record changes in the ratio of water to sediment which can
cause desorption of pollutants from sediment (Plumb, 1981).
60
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Measure total solids content by drying a wet sediment sample at
105°C to a constant dry weight. The percent solids content is
the dry weight divided by the initial wet weight multiplied by
100 (APHA, 1985) . The percent moisture content of a sediment is
100% less the percent solids.
C. ORGANIC CARBON
Organic carbon content is one of the primary factors
regulating sediment bioavailability (e.g., Rubinstein et al.,
1983); therefore, measuring organic content on every sediment
tested is critical. One direct measure of organic matter is
total organic carbon (TOO, a measure of the total amount of
oxidizable organic carbon. In comparison to TOG, total carbon
(TC) is the measure of both the volatile organic and inorganic
nonvolatile materials, such as carbonates and bicarbonates (U.S.
EPA/U.S. ACE, 1988). Because of its relation to the binding of
organic pollutants, TOC is the more biologically relevant
measure.
Both TOC and TC can be determined by wet or dry (combustive)
oxidation techniques (Plumb, 1981; Holme and Mclntyre, 1984).
Oxidation of an untreated (no acidification) sediment measures
total carbon; whereas, oxidation of an acid treated sediment
measures TOC (Plumb, 1981). The common wet methods use a chromic
acid oxidation technique developed by Walkley and Black (1934;
also see Holme and Mclntyre, 1984) and el Wakell and Riley
(1956). Buchanan and Longbottom (1970) describe a technique to
61
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use on sediments containing coal. The differential combustion
method, used in commercially available carbon analyzers, oxidizes
the carbon with heat. The amount of C00 generated at different
£t
temperatures is a measure of the total and organic carbon.
Carbon analyzers are simpler to use than wet techniques and are
recommended. However, exercise care in using or purchasing
carbon analyzers because instruments designed for measuring TOC
in seawater may be too sensitive for sediments, requiring
extensive dilution of the samples.
A commonly used method to estimate organic matter is loss on
ignition (LOI) or total volatile solids (TVS), which is the
percent loss of weight after combustion. The organic matter is
combusted by heating dry sediment at 550°C for 1-4 hours. LOI is
calculated as the difference between the dried and combusted
sample weights divided by the dry weight, multiplied by 100 to
convert the number to a percent (Dean, 1974; Byers et al., 1978;
Plumb, 1981; APHA, 1985). Structurally bound water and
carbonates, found particularly in clays and in calcareous sands,
may be lost along with volatile solids during combustion,
distorting actual organic carbon values (Dean, 1974). Because
LOI includes these non-organic constituents, it is not a direct
measure of sediment organic carbon content.
When determining LOI, spread a small aliquot (a few grams)
of sediment out thinly in a porcelain ignition dish at least 75
mm in diameter. This exposes the sediment to an adequate oxygen
supply for oxidation of the organics. Clumps of sediment and/or
62
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large amounts of sediment have a tendency to block oxygen
transfer to the center of the clump, thereby yielding
significantly lower and/or inconsistent LOI results. We analyze
2g samples of sediments and ignite the dry total solids residue
for 1.5 hours at 550°C.
Of the two methods, LOI is simpler and cheaper than TOG.
However, most research and regulatory applications require a more
accurate measure than LOI (see Dean, 1974; Byers et al., 1978;
Mook and Hoskin, 1982). Because TOC is one of the parameters
used to calculate Accumulation Factors (see Appendix 1-1), we
recommend using LOI as a rapid survey method, but performing TOC
analyses on all sediments used in bioaccumulation tests. If
analyzing TOC on all sediments is impossible, determine the
conversion between LOI and TOC on a few samples or use .the values
in Table VI-2. However, use caution when using these conversion
factors because the relationship between LOI and TOC can vary
several fold among sediments (Dean, 1974; Byers et al., 1978;
Ditsworth, G., pers. comm., U.S. EPA, Mar. Sci. Ctr., Newport,
OR) .
D. ADDITIONAL SEDIMENT CHARACTERISTICS
Measure salinity in the overlying and interstitial water
during the initial characterization of sediments in the field. A
refractometer measures salinity with sufficient accuracy in most
cases and requires only a few drops of interstitial water. In
studies of metal bioavailability, measure pH and Eh, both of
63
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TABLE VI-2: Ratios For Converting Loss on Ignition (LOI) to Total
Organic Carbon (TOG) For Various Sediment Types.
Particle size
%>63um %<63um
36
med.
63.5
34.1
50.2
64
silt
36.0
65.9
49.8
TOG
1
10
1.21
2.29
2.10
LOI
2.
22
4.
7.
6.
13
.9
30
79
53
TOC/LOI
0
0
0
0
0
.47
.44
.28
.29
.30
Reference
Dean, 1974
Byers et al . , 1978
unpubl . data
unpubl . data
Ditsworth , pers . comm.
(Mean of 12 sediments)
19-81 19-81 0.53-3.50 2.43-10.10 .21-.37 Ditsworth1, pers. comm.
(Range of 12 sediments)
1 G. Ditworth, U.S.EPA, Mar. Sci. Ctr., Newport, OR
64
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which can affect pollutant solubility and mobilization (Engler,
1980). Measure these parameters during field collection,
repeating the measurements after handling and storage during
which values can change rapidly. Use a meter or probe, and a
platinum electrode, to measure pH and Eh, respectively. Insert
the probes 1-3 cm below the sediment surface and allow to
stabilize (Plumb, 1981). Discussion of pH and Eh can be found in
Fenchel (1969) and Pearson and Stanley (1979).
Additional characteristics potentially affecting metal
availability are amorphic oxides of iron and manganese, iron
sulfides, cation exchange sites, and selectively extractable
fractions using extractants such as hydrogen peroxide, acetic
acid, and organic chelates (Luoma and Jenne, 1976; Jennett et
al., 1980; Plumb, 1981). Consider measuring these parameters in
studies of metal bioavailability. Recent work suggests that acid
volatile sulfides (AVS) may be the primary factor regulating the
bioavailable fraction of many metals (DiToro et al., in review).
If further investigations support the initial findings, it may be
possible to use AVS as a "normalizer" for sediment metals much as
TOC is used for neutral organics. Morse et al. (1987) gives a
method to measure AVS.
E. INTERSTITIAL WATER
Interstitial water is an integral part of bedded sediments
and pollutants associated with the interstitial water may play a
major role in the uptake and toxicity of a number of compounds
(Adams et al., 1980; Swartz et al., 1988 ; Landrum 1989).
65
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Because of the potential role of pore water in controlling uptake
and toxicity, its characterization can be very informative.
"Dissolved" pollutant concentrations, complexed pollutant
concentrations, and dissolved organic matter (DOM) are the most
important parameters to measure.
A number of techniques have been used to collect
interstitial water for pollutant or nutrient analysis, including
squeezing of sediment cores (Presley et al., 1967; Robbins,
1977), centrifugation (Edmunds and Bath, 1976; Plumb, 1981;
Landrum et al., 1984), suctioning (Plumb,1981; Knezovitch and
Harrison, 1987), equilibration with dialysis membranes (Mayer,
1976) or porous TeflonR cups (Zimmermann et al., 1978), and
displacement with other liquids (Bately and Giles, 1980).
Manheim (1974) published a comparative study and Bately and Giles
(1980) reviewed several methods.
Centrifugation is a straight-forward technique suitable for
the routine collection of small to moderate amounts of
interstitial water for pollutant analysis. There is no standard
procedure, but centrifuging the sediment sample at 7000 - 9000
rpm for 5-10 minutes should suffice in most cases (Bately and
Giles, 1980; Plumb, 1981). Higher rpms are required if any
suspended particles remain, because particulate matter in the
supernatant will result in erroneously high "dissolved" pollutant
concentrations. If the speciation of metals will be examined,
the centrifuge tubes should be oxygen-free. Following
centrifugation, the supernatant is often vacuum filtered to
66
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remove particulate matter. There is no standard pore size, but a
0.45 urn NucleporeR membrane filter is frequently used (U.S.EPA,
1979; Plumb, 1981; Landrum et al., 1984). The filters should be
precleaned in 10% nitric acid for 24 hours and then soaked in
deionized water for another 24 hours (Lapan, R. pers. comm.
U.S.EPA, ERL-N, Narragansett, RI) A potential source of error is
sorption of dissolved pollutants onto the filter, resulting in an
underestimation of dissolved pollutant concentrations. Measure
the sorption of the pollutants onto the filter by determining the
loss of pollutants from standard solutions passed through the
filter.
Pollutants in the interstitial water may either be truly
dissolved or complexed with DOM (Engler, 1980) . In general, the
complexed pollutants have a lower bioavailability than the free
forms (Luoma and Bryan, 1978; Jenne et al. 1980) . As DOM levels
may change due to handling (DeWitt, T., pers. comm., OSU, Mar.
Sci. Ctr., Newport, OR) or biotic effects (e.g., excretion of
organics), the bioavailability of interstitial water pollutants
may change over the course of an experiment or vary between field
and laboratory sediments. The concentration of DOM can be
determined either by using wet oxidation methods (Walkley and
Black, 1934; Holme and Mclntyre, 1984) or a carbon analyzer. As
mentioned, carbon analyzers that measure the relatively low
concentrations of DOM in interstitial water with good accuracy
are not as efficient at measuring the high concentrations of TOC
in sediment.
67
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Several techniques have been developed to measure the
concentrations of the dissolved and complexed pollutants in
interstitial water. A relatively simple technique has been
developed by Landrum et al. (1984) to separate and quantify
total, freely-dissolved, and colloidally-bound non-polar organic
pollutants in interstitial water. This technique uses a Sep Pak
C-18R cartridge to separate the bound and dissolved fractions.
The bound fraction of the pollutant will pass through, while the
dissolved fraction will be retained by the column.
Besides direct measurement, equilibrium interstitial water
pollutant concentrations can be calculated from the TOG, bulk
sediment concentration, and Koc for a compound (Karickhoff,
1984). Because all the aforementioned collection techniques may
alter interstitial water characteristics through adsorption/
desorption of pollutants or suspension of particulate matter
(Word et al., 1987), the calculated value for interstitial
equilibrium concentrations may be more accurate for very high Kow
compounds (Karickhoff, S., pers. comm., U.S. EPA, ERL-A, Athens,
GA). Calculated concentration values will not monitor temporal
changes in interstitial water but will serve as check on directly
measured concentrations. Measured concentrations substantially
higher than calculated could indicate suspended particles;
whereas, measured concentrations lower than expected could
indicate either that equilibrium had not been attained or that
the pollutant had adsorbed to the filter.
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CHAPTER VII. ORGANISM SELECTION
A. SELECTION CRITERIA
The choice of the test species can have a major influence on
the success, ecological significance, and interpretability of a
bioaccumulation test. Given the potential range in environmental
characteristics, no one species is best suited for all
conditions. There are, however, two characteristics required of
any bioaccumulation test species, as well as a number of other
desirable characteristics. These characteristics are summarized
in this chapter and Table VII-1 and discussed in more detail in
Appendix VII-1.
The first required criterion is that the test species ingest
sediment. This requirement is critical because recent work has
demonstrated that ingested sediment is the major uptake route for
higher Row compounds for some species (Landrum, 1989; Boese et
al., in press). Many benthic invertebrates are flexible in their
feeding mode, and this requirement does not preclude the use of
facultative filter-feeders (e.g., Macoma) as long as the only
exposure route during the experiment is from bedded sediment
(i.e, no resuspended particles or phytoplankton). The second
required attribute is that the test species be sufficiently
pollutant resistant to survive the duration of the exposure with
a minimum level of mortality. This requirement precludes the
species routinely used in sediment toxicity testing (e.g.,
Rheooxvnius).
69
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Besides the required criteria, there are a number of
desirable characteristics -- ease of collection, year-round
availability, ability to culture the species, adaptability to
laboratory conditions, suitable size, tolerance to a wide range
of sediment types and salinities, suitability for sublethal or
toxicokinetic tests, ecological or economic importance, having a
high bioaccumulation potential, compatibility with other species,
and a low capability of metabolizing PAHs and other contaminants.
The importance of these various criteria depends upon the
specific goals of the research and the sediment tested. However,
using an organism large enough to supply sufficient biomass for
chemical analysis is important in nearly all cases. Ideally, the
test species should be large enough to allow chemical analysis on
individuals. Even when individuals are composited, compositing a
smaller number of larger organisms is easier than dozens or
hundreds of smaller specimens.
B. RECOMMENDED SPECIES:
An evaluation of the suitability of potential test species
is summarized in Table VII-1. This evaluation is not based on
extensive comparative studies and should be considered a guide
rather than a definitive characterization of the species. Based
on this analysis, we identified five recommended bioaccumulation
test species and another eight "secondary" taxa. The recommended
species meet all or nearly all of the desired criteria and are
well established as bioaccumulation test species. The recommended
70
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TABLE VII-1
PERTINENT CHARACTERISITCS OF TEST SPECIES
SPECIES
Abarenicola spp.
Arenicola spp.
Callianassa spp.
Capitella spp.
*
Macoma balthica
*
Macoma nasuta
Nephtys incisa
*
Neanthes arenaceodentata
Nereis virens
Nereis diversicolor
Nucula spp.
Palaemonetes pugio
Yoldia limatula *
Feeding
Type
Fun
Fun
SSDF
SDF
SDF
SDF
SSDF
SDF/0
SDF/0
SDF/0
SSDF
SDF
SSDF
Biomass o Pollution Culture Commercial Bio.
S /«"> Tolerance Potential Availability lnfo
4-+ >is 4- - -
4-4- >is 4- -f
4-4- >i° -? - 4-
- >io ++ + +
4- >io 4- ~~ "~
+ + >io + - ~
+ >25 4. ~~ —
+ ? >28 4- 4- 4- +
4-4- >io ++ - +
4- + >10 4-4- - +
+ ? -4- — —
4-? >io - ? + 4-
4- >25 ? + - -
i
~r
4-4-
44-
4 +
4-
4-4
4-4-
44-
+
4-4-
4-
Fun = Funnel feeder
0 = Omnivore
SDF = Surface Deposit Feeder
SSDF = Subsurface Deposit Feeder
Recommended test species
+ = good, sufficient
++ = very good
- = poor, insufficient
Bio Info. = information on bioaccumulation toxicity
-------
species are the polychaetes Nereis diversicolor. Neanthes (Nereis)
virens, and the bivalves Macoma nasuta. Macoma balthica. and Yoldia
limatula. These species have been used in a substantial number of
experimental bioaccumulation studies and in regulatory monitoring.
Within their tolerance levels, these species should serve as
suitable test species, and we recommend using at least one of these
species in all tests, at least until the suitability of other
species has been demonstrated locally.
The secondary bioaccumulation species meet the required
characteristics but are to some extent deficient in one or more
of the important desired characteristics and/or there is
insufficient information to make a final evaluation. However,
some of these secondary taxa offer potential advantages such as
large size (arenicolid worms), additional phylogenetic groups
(i.e., crustaceans), adaptability to culturing (e.g., Neanthes
arenaceodentata). and high pollution tolerance (Capitella spp.).
The importance of these various advantages depend upon the site
specific situation (e.g., level of toxicity of sediment).
C. NUMBER OF SPECIES TESTED AND MULTIPLE SPECIES TESTS
Species as well as larger phylogenetic groups vary in their
tendencies to bioaccumulate pollutants both in response to their
modes of exposure and to their metabolic characteristics. The
extent of these interspecific variations are not well understood,
and both the magnitude and direction of species differences can
vary with pollutant (e.g., metals vs. organics) and perhaps with
72
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sediment type. Thus, utilization of two or more species from
different major taxa increases the probability of accurately
assessing the maximum field tissue residues.
The actual number of species and taxa used depends upon the
goals and scale of the project and the range of pollutants in the
sediment. In general, use of a single species should be adequate
for a general survey of an area or assessing a small discharge or
volume of dredge material. In interpreting the data from a
single species test, however, it should be recognized that no one
species is likely to maximize uptake from all pollutants. Two
species are recommended when assessing a moderate to large sized
discharge or dredging operation. The species should be of
different major taxa, and a polychaete and a bivalve are
recommended. It is especially important to include a bivalve if
PAH contamination is of concern, as bivalves have a reduced
capability to metabolize PAHs compared to amphipods or
polychaetes (Varanasi, et al. 1985). The addition of an
arthropod species or additional polychaete and/or bivalve species
may be justified when assessing a large discharge or dredging
operation, especially if there is a wide range of pollutants.
73
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CHAPTER VIII: ORGANISM COLLECTION, MAINTENANCE, TRANSPORT,
AND ACCEPTABILITY
To assure unbiased test results, bioassay organisms must be
in good health and have minimal background pollutant
contamination. Reasonable efforts must be taken to minimize
stress during collection and transport to holding facilities.
Holding facilities must provide high quality water and conditions
suitable for the maintenance of the test species. This chapter
describes techniques and facilities which will meet these general
requirements.
A. ORGANISM COLLECTION AND TRANSPORT
1. Field Collection of Test Organisms
The logistics of collecting intertidal species is usually
much simpler than those of collecting subtidal species, and
intertidal collection is recommended when possible. Infaunal
organisms can be collected by turning the sediment over with a
shovel and picking out larger species (e.g., clams) or by gently
sieving the sediment in the field. For most of the
bioaccumulation test species, a sieve size of 4-6 mm will collect
adequate numbers while minimizing damage and sorting time.
Collection equipment should not have been used in contaminated
sites or should have been adequately cleaned.
Subtidal organisms can be collected by grabs, dredges, or
suction samplers (see Holme and Mclntyre, 1984) . Dredges sample
a larger area than grabs and are usually more proficient at
collecting shallow-buried organisms, though there is a greater
74
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possibility of damaging specimens. Grabs are recommended for
collecting more deeply buried species. Suction lifts are also
useful for collecting larger, deeply buried bivalves, though they
require the use of SCUBA divers, and a greater likelihood of
damage exists. Electro-shocking, chemical poisons, and other
harsh methods of collection are not acceptable.
Remove organisms from the collection device as soon as
possible and submerge them in ambient seawater or sediment
contained in ice chests or uncontaminated plastic buckets. Avoid
overcrowding animals in collection containers. Discard organisms
with signs of disease or obvious defects (e.g., bivalves with
cracked shells).
State or local authorities may require collection permits or
ban collection from specified areas. Collection of regulated
species (e.g., bay clams) may require a local license, be limited
to a season, and preclude certain collection techniques.
Additional permits or precautions may be required when importing
non-indigenous species. Check with State authorities about the
local regulations before collecting or importing specimens.
2. Organism Transport
For organism maintenance, ASTM (1984) recommends not more
than a 3°C change in water temperature within a 12 hour period
and an oxygen concentration of between 60 and 100% of saturation.
If the time between collection and return to laboratory is short
(less than 1-2 hours) and ambient temperature is not extreme,
simple precautions should meet these requirements. If possible,
75
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collection buckets or ice-chests should be kept out of direct
sunlight and must not be left in closed vehicles. The water in the
containers should be changed periodically while collecting and
immediately before returning. If the time before returning to the
laboratory is several hours and/or air temperature is high, use a
portable aerator to maintain oxygen levels.
Successful long-distance transport of organisms, whether in
a vehicle or through the mail, requires packaging that retains
moisture and maintains an adequate supply of oxygen. This can be
accomplished by placing animals in a minimum amount of water (a
few milliliters) in a sealed container filled with air (e.g.,
Whirl-PakR bag). Alternatively, test animals may be placed
between wet nylon or seagrass (e. g., Zostera) and surrounded by
layers of wet paper towels, all contained in polyethylene
ZiplockR bags (Robinson, A., pers. comm. AScI, Mar. Sci. Ctr.,
Newport, OR; Gulf Specimen Co., pers. comm., Panacea, FL). Wet
sediments may also be used to retain moisture. These sediments
should have a low organic content (e.g., ashed sediment, beach
sand) as they are not as likely as natural sediment to turn
anoxic. Regardless of the moisture retaining agent, the
container should have a large air space to maintain aerobic
conditions. Air trapped within a plastic bag has the added
advantage in preventing animals from being crushed.
Containers with organisms should be placed in ice chests or
insulated shipping containers, with packets of jelled (e. g.,
Blue IceR) refrigerant placed at or taped to the inside of the top
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of the container. Jelled refrigerants are preferred to ice to
avoid melt water and a layer of insulating material should be
placed between the refrigerant and animals. Add sufficient
refrigerants to maintain the water temperature in the containers at
or a few degrees below the water temperature at the collection site
but not so much as to cold shock the organisms. Insulating
material should fill all extra space in the shipping container,
protecting and securing the bottles and bags in the carton. Pack
shipment containers to obtain a low center of gravity, and label
plainly to keep package upright. Every effort should be made to
provide overnight or 24-hour delivery. If the organisms are
transported by vehicle, periodically monitor the temperature and
drain any melt water and replace the ice as required.
B. CULTURING AND PURCHASING TEST ORGANISMS
A successful culture of an appropriate test species has the
advantages of providing a ready supply of specimens with a known
history. However, culturing of marine/estuarine organisms is not
a task to be undertaken lightly, and is usually justified only if
regular tests are planned. Although a few sediment ingesting
polychaetes can be cultured with relatively simple equipment (see
Appendix VII-1), the majority of recommended test species are not
routinely cultured.
Some test organisms can be purchased from biological supply
houses, local collectors, colleges, or bait shops. Purchasing
specimens can be cost-effective if the laboratory is not well
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equipped for collection (e.g., does not have a boat) or if tests
will be conducted only occasionally. There are several companies
that specialize in supplying bioassay organisms (see Appendix
VIII-1), although most do not presently supply appropriate
benthic bioaccumulation organisms on a routine basis. Check with
a supplier even if bioaccumulation test species are not currently
carried as availability of particular species may change or the
supplier may be able to fill special orders.
Maintain purchased organisms in the laboratory for at least
one week to acclimate them to local conditions and to monitor
their health. Before beginning bioaccumulation tests, analyze
the purchased organisms for background pollutant levels to
determine if they meet the criteria for control organisms (see
Table VIII-1).
C. PRE-EXPERIMENTAL MAINTENANCE
In general, the guidelines presented here are based on our
experience with Macoma nasuta or modified for deposit-feeders
from ASTM Standard Procedures (ASTM 1980, 1984) for maintaining
filter-feeding bivalves for bioconcentration tests. Most of the
bioaccumulation test species are adaptable to laboratory
conditions, so elaborate procedures are not usually required for
the maintenance of adults. Additional information on the
maintenance of benthic invertebrates can be found in King and
Spotte (1974), Dean and Mazurkiewicz (1975), Kinne (1976-1977), -
and National Research Council Committee on Marine Invertebrates
(1981).
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When maintaining a non-indigenous species, permits may be
required from state or local authorities. This may require fail-
safe precautions against accidental release of such organisms
into the local environment (i.e., double containment, diked
seawater drains, siphon breaks, etc.). Equipment, water, wastes,
and dead animals may require sterilization before disposal.
1. Water Quality
Constant water quality should be maintained in holding
aquaria, keeping dissolved oxygen between 60-100% saturation and
un-ionized ammonia concentrations <20 ug/L (ASTM, 1984). Flow-
through seawater with a minimum flow-rate of 1 L/hr per gram of
wet tissue is recommended as a means of maintaining water
quality. Filtering incoming seawater is generally inadvisable
with benthic invertebrates as the settling particles supply a
natural source of food. If flowing seawater is unavailable,
organisms can be maintained in static systems using collected
seawater. Store replacement seawater in covered containers in
the dark at 4°C to keep salinity and water quality constant. If
collection of natural seawater is impractical, artificial
seawater may be used, though it should be demonstrated that the
growth and behavior of the test species is not altered by using
artificial salts. Prepare artificial seawater with deionized
water or with distilled and charcoal-filtered water. Static
systems should always be aerated, and preferably equipped with a
recirculating aquarium filter with replaceable activated
charcoal.
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Regardless of whether flow-through or static systems are
used, the seawater should be analyzed for background levels of
pollutants, especially if it is collected from an urbanized area.
If a pollutant is detected in the water, its potential uptake can
be estimated by multiplying the water concentration by the
bioconcentration factor (BCF) for that compound. If the
calculated tissue residue is greater than that acceptable for a
control organism (Table VIII-1), a different water supply is
required. BCF values and methods to estimate BCFs can be found
in Bysshe et al. (1982).
Temperature should not vary more than 3°C in a 12 hour
period and salinity should not vary by more than 2 g/kg or 20% of
the average, whichever is larger (ASTM, 1984). In flow-through
systems, a storage tank within the laboratory will help
ameliorate natural fluctuations in temperature. In estuarine
areas, a storage tank may be necessary to supply high salinity
water during low salinity periods.
2. Sediment Quality
Maintain animals in a sufficient amount of clean sediment to
allow them to burrow naturally, which in nearly all cases will be
at least 3 cm. This sediment must be analyzed for pollutant
concentrations, which should not exceed the level acceptable for
a control sediment (Table II-1). Periodically add fresh sediment
of the same type to maintain an adequate food supply (i.e.,
detritus and associated microbes). One to three times a week,
add about two millimeters of fresh control sediment to the
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sediment surface. This sediment replenishment should be
sufficient if the organisms are not over-crowded. If the
sediments become heavily pelletized with fecal material, remove
the organisms and replace the sediment. The addition of other
types of food are not recommended except in special cases of
long-term maintenance. These foods include detritus or decaying
seaweeds, cultured marine phytoplankton and zooplankton,
microencapsulated diets, formulated feeds such as fish flakes
(e.g., TetraMinR), or small bits of clam or other tissues for
omnivores (Lee and Muller, 1972). Check the background pollutant
levels of all foods.
3. Organism Health And Acclimation
Field collected organisms should be held in the laboratory
for at least four days before commencing a bioassay, and
purchased organisms held for at least a week. Discard any
organism if injured or behaving abnormally. In general, animals
should not be held longer than two weeks before testing. If
longer maintenance periods are needed, the investigators should
have experience with the species and should monitor for any signs
of stress (e.g., reduced sediment processing rate, unusual tube
construction). A flow-through system is strongly advised if
long-term maintenance is planned.
To avoid the spread of diseases, organisms collected more
than a week apart should be maintained in separate aquaria, each
with an independent water supply. The organisms should be
checked every day or so, and any diseased, dying, and dead
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organisms promptly removed. Black spots on the surface of the
sediment can mark the location of dead organisms. Should a
question arise concerning the health of the animals, a behavioral
test such as time to rebury or analysis of lipid content is
recommended.
If holding and experimental conditions are different,
gradually acclimate the test organisms to experimental
conditions. This transition may be accomplished using serial
water dilutions until the proper temperature, salinity, and pH
are reached. Acclimation for temperature should proceed no
faster than 3°C in 72 hr (ASTM, 1984). Maintain animals at the
test temperature and salinity for at least two days before
commencement of an experiment. No more than 3% mortality is
permitted within 48 hr before the test (ASTM, 1984).
D. ORGANISM ACCEPTABILITY AND BACKGROUND CONTAMINANT LEVELS
Specimens selected for a test must tolerate the physical-
chemical conditions (e.g., TOC content, interstitial salinity) of
the test substrate and be free of disease or stress from capture
or handling. All specimens should be collected from the same
site, and preferably at the same time. All organisms used in a
given test should be as uniform in age and size as possible, and
bivalves should be of the same year class. For bioconcentration
tests, ASTM (1984) stipulates that the length (umbo to distal
valve) of the largest clam should be no greater than 1.5 times
larger than the smallest clam. For Macoma nasuta. this would be
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equivalent to a 3.7-fold difference in wet flesh weight. It is
important to correctly identify the test species, and voucher
specimens should be kept from each collection.
High pollutant background levels in the test specimens may
confound the results, making it difficult to detect differences
between treatments. Therefore, tissue residues in the test
organisms should be no greater than those expected in organisms
living in control sediment (see Chapter II). Approximate
acceptable background tissue concentrations for test species are
given in Table VIII-1. These values are from organisms collected
from sites which appeared to meet the criteria for a control
site. For compounds not listed in Table VIII-1, the ASTM (1984)
criterion of the background tissue residue not exceeding 10% of
the expected steady-state can be applied. First-order estimates
of steady-state tissue residues can be obtained from data on
other species or from the thermodynamic-based bioaccumulation
model for neutral organics (see Appendix 1-1).
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TABLE VIII-1: Representative Control Organism Tissue Residues
COMPOUND
Organics
(ppb wet wt.)
Various East
Coast Sitesa
Puget Sound*3 Yaquina Bay, ORC
CB
B(ibk)F
BaP
DDT
HCB
Naph
PAH
PCB
pesticides
<1.0-70
0.3-6.0*
<0.08-3.8
0.02-0.17
<1. 0-9.1
0.02-7.2
10-70
<0.03-0.6
<10
2.3-<10*
<1.0-<5.0
<130
<0.05
<2-17*
<2.0-10
1.9
3.9
Metals
(ppm wet wt.)
Ag
As
Cd
Cr
Cu
Hg
Ni
Pb
Zn
Various East
Coast Sitesa
0.2-2.6
1.5-3.9
<0.06-4.0
0.26-2.5
0.1-7.2
<0.05-1.2
<0.4-7.0
<0.6-2.6
2.4-30
Puget Sound" Yaquina Bay, ORC
<0.005
1.0
<2.0
a
b
c
*
Tetra Tech, 1985a
Konasewich et al., 1982
unpublished data
Tetra Tech, 1985a and Konasewich et al.,
1982
CB - Chlorinated benzenes
B(ibk)F - Benzo(i,b,k)fluoranthene
BaP - Benzo(a)pyrene
HCB - Hexachlorobenzene
Naph - Naphthalene
PAH - Polyaromatic hydrocarbons
PCB - Polychlorinated biphenyls
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CHAPTER IX. SEDIMENT EXPOSURE SYSTEMS
In designing the exposure systems, considerations for
maintaining an adequate environment for the test organisms must
be combined with considerations for pollutant behavior, costs of
construction and maintenance, and ease of operation. The
recommendations made here are based on the assumption that each
"exposure chamber" will hold a single experimental unit (e.g.,
individual organism or composite of a single species in a
beaker), but that these may be placed into a larger aquarium or
tank to maintain water quality. The "exposure system" is
composed of the replicate exposure chambers, any aquaria or tanks
which hold the exposure chambers, the water delivery system, and
any pollution abatement system. The recommendations are also
based on the standard 28-day exposure duration. Discussion of
specialized exposure chambers is limited to Appendix IX-1.
A. SYSTEM REQUIREMENTS
In designing the exposure system, two critical
considerations are the amount of sediment per individual and the
volume and turnover rate of water. Adequate amounts of sediment
and overlying water are required to assure that supplies of food
and pollutants are not substantially depleted, water column
oxygen concentration is not depressed, metabolites are diluted,
and the organism's feeding behavior is not impaired. The ASTM
criteria for bioconcentration (ASTM, 1984) and toxicity tests
(ASTM, 1980) offer general guidance to the design of a sediment
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exposure system, but several of their specific recommendations
had to be modified because of the different requirements for
deposit-feeders compared to fish and filter-feeders.
Additionally, the design criteria for maintaining a constant
pollutant concentration in the water are inapplicable as the
sediment instead of water is the exposure mode in these tests.
1. Sediment Requirements
Sediment serves as the habitat, source of food, and source
of pollutants for the test organisms. If insufficient sediment
is added, organisms may reingest the same particles.
Alternatively, if the fecal pellets are resistant to breakdown,
there may be a reduction in the appropriately sized particles,
especially with the more selective deposit-feeders. Both of
these processes could reduce the mass of pollutant available to
the test organisms. Although both reingestion and pelletization
of sediments occurs in field (see Lee and Swartz, 1980), the
rates may be exaggerated in laboratory systems. For this reason,
it is critical to supply a sufficient sediment mass during the
entire course of the laboratory exposure.
Assuming periodic sediment additions to the exposure
chambers (see Chapter X), we recommend initially adding at least
50 grams wet sediment for each 1 gram wet flesh tissue (excluding
shell) for surface deposit-feeding bivalves. For funnel-feeders
such as arenicolid worms, at least 200 grams of wet sediment to
each 1 gram of wet flesh tissue may be required for construction
of a normal feeding burrow. Besides the mass of sediment, the
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sediment must be deep enough to allow normal burying and feeding.
The initial depth for Macoma should be at least 2 cm and
preferably 3-5 cm, whereas a large lugworm may require 5-10 cm of
sediment. While these amounts should bracket the needs of most
organisms, the initial amount of sediment added should equal or
exceed consumption requirements for the 28-day exposure. For
example, if a particular species processes 2 grams of sediment
per gram of tissue per day, then at least 56 grams per gram of
tissue should be added initially. A more accurate estimate of
sediment requirements for selective deposit-feeders can be
generated by using the processing rates of the ingested size
fraction, though this information will .not usually be available.
If periodic sediment additions are not made, then the
initial amount added should exceed the total amount processed
over the duration of the experiment by at least 2-fold, and
preferably 5-fold. Thus, for the organism with a 2g/g-tissue/day
sediment processing rate, about 250-300 grams of sediment should
be added per gram of tissue. Compilations of sediment processing
rates (e.g.. Lee and Swartz, 1980) can be used to estimate these
requirements. It must be recognized, however, that in a
laboratory, an organism may deplete the food or pollutants within
its specific feeding zone regardless of the amount of sediment
added. This is especially likely with surface deposit-feeders.
For this reason, we strongly recommend periodic additions of the
treatment sediment, as discussed in Chapter X.
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2. Water Quality
The water quality requirements for conducting the tests are
similar to those for pre-experimental maintenance (Chapter VIII).
That is, dissolved oxygen should be between 60 and 100% of
saturation and the un-ionized ammonia concentration should not
exceed 20 ug/L (ASTM, 1984). These criteria can be met either by
using a flow-through system or a static batch-replacement mode.
For a flow-through system, ASTM (1984) recommends not more
than one filter-feeding bivalve (40-60 mm from umbo to edge of
distal valve) per liter per hour. This would be equivalent to
about a minimum flow of one L/hr/g wet tissue for an oyster.
However, this requirement is based on feeding, and does not
account for sediment oxygen demand. In addition to the flow rate
per gram tissue, flow-through systems should be designed to
achieve at least six turnovers a day.
In static systems, the water volume to loading ratio needs
to be sufficient to allow the maintenance of oxygen levels .>60%
of saturation. A gentle aeration helps maintain the oxygen level
as does changing the water two or three times a week. As an
example, 10 Macoma nasuta (mean wet flesh weight of about 1.3 g),
each in a 100 ml beaker with an initial 50 grams of sediment,
have been successfully maintained in a 10 L aquarium with 8 L of
filtered seawater (Ferraro et al., 1990). The aquarium was
gently aerated and the water changed three times a week.
In determining the oxygen demand for the system, it is
important to take into account the total sediment oxygen demand.
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In most cases, sediment microbial demand will be several fold
greater than the oxygen utilization by the test species.
Therefore, calculations based on sediment-free exposure systems
will underestimate the actual oxygen requirements. The total
Q
oxygen demand of sediments ranges from <1 to over 100 ml 02/mz/hr
(e.g., Hargrave, 1969; Smith et al., 1973; Smith, 1978; Davis and
Lee, 1983). In general, total oxygen demand will increase with
temperature and in organically enriched sediments, and the water
flow or volume should be increased accordingly.
Aeration will help ensure a proper oxygen concentration is
maintained, and is required in a static system. The air should
be filtered and free of oil and moisture. The volume should be
sufficient to turnover the water but not enough to resuspend the
sediment. This can be achieved with a supply of approximately
0.1 SCFH (standard cubic foot per hour) per 10 liter aquarium via
an air stone or pipette. Position the air stone or pipette
outside of beakers maintained in aquaria, or sufficiently far
above the surface to avoid resuspension in individual beakers or
aquaria. Check the air-stone or bubbler frequently and remove
any salt crystals forming at the orifices. If air is provided
from a compressed air tank, specify that the composition include
about 0.3-1.0% C02 to help control pH. If not specified, no C02
will be present.
Seawater is well buffered, but in static systems metabolites
and waste materials (i.e., ammonia) can build up, lowering pH.
Maintain pH between about 6.5 and 8.0 (ASTM, 1984). As mentioned,
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aeration will help maintain pH as will periodic replacement of
water. In static systems, the addition of clean, crushed oyster
shell to the bottom of the aquarium can provide a good buffering
system.
ASTM (1984) recommends that overlying water salinity should
vary less than 2g/kg or 20% of the average, whichever is higher.
In areas where salinity varies (as in water drawn from estuaries
with a seasonally high riverine contribution), store a quantity
of high salinity water sufficient for the expected period of low
salinity, or preferably, to maintain salinity over the duration
of the exposure.
Because phytoplankton and suspended material are a sink for
pollutants and a food for facultative filter-feeders, it is
important to filter the water to remove suspended particulates
(>5 um) during the test. Filtration can be accomplished with in-
line cartridge filters (commercially available with 2.5-5.0 um
pores) or in batch mode. The ASTM (1984) precautions concerning
the adequate concentration of phytoplankton necessary as a food
source are not relevant for deposit-feeders.
3. Temperature and Light
The tests should be conducted as close as possible to one of
the seven temperatures recommended by ASTM (1984) - 7, 12, 17,
22, 27, and 32°C. A temperature corresponding to the average
spring-summer temperature of the study site would simulate the
biologically most active season. Most commonly, this will be
12°C in the Pacific Northwest, 17°C in mid-latitudes, and 22 or
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27°C on the Gulf Coast. With flow-through systems, it can be
difficult to exactly adjust the temperature, though a large head
tank in a controlled-temperature room will temper the water. The
maximum difference between the minimum and maximum temperatures
must not exceed 10°C (ASTM, 1984).
Light should be provided by means of cool-white fluorescent
lights at an intensity of about 400 foot-candles. Other sources
(incandescent, fluorescent/incandescent, photosynthetically
active radiation augmented) may be required for special purposes.
Ultraviolet radiation, especially UV-B, is generally missing from
these artificially supplied spectra. Although UV-B radiation can
enhance the toxicity of certain pollutants (phototoxicity), this
probably will not greatly affect organisms buried in sediment.
ASTM (1984) recommends a 16h day, 8h night as a convenient
light/dark cycle. Schedules of 12/12 or 14/10 hrs day/night are
also acceptable, and may be useful in delaying maturation and
spawning of some species. We have routinely used a 12/12
schedule. These various day/night cycles can be controlled by
use of timing devices in the light circuits.
B. EXPOSURE SYSTEM DESIGN
1. Materials Compatibility
Materials used in the exposure system should not induce any
reaction by the organisms or affect pollutant concentration or
bioavailability. Borosilicate glass (Pyrex , Kimax , or
equivalent) and soft glass (soda-lime, window) have proved
generally non-reactive to metals and organics, and are the
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preferred materials where their fragility is not a major
limitation. Most rigid plastics (polyolefins, engineering resins
and fluoropolymers) are acceptable after conditioning, such as
soaking in deionized water for several days. Some plastics,
generally flexible types that contain mobile plasticizers
(phthalate esters), need to be tested for toxicity and pollutant
compatibility. These flexible plastics should not be used if the
uptake of phthalate esters will be studied. Because the alloy
components of many stainless steels may react with saltwater,
stainless steel should not be used in direct contact with
seawater. Choose another material if pollutant sorption to
internal surfaces of containers is a problem.
Any sealant used to construct chambers must be non-toxic,
such as Dow-CorningR #8641 clear, non-toxic silicone-rubber (i.e,
meets FDA Regulation 21 CFR 177.2600). Such materials are
usually specified for aquarium use and do not contain fungicides
(e.g., arsenic compounds). Exposed sealant at joints should be
minimal. Place sealant used for mechanical reinforcement on the
outside of the joint. Plastics and sealants must be chosen
carefully, as both may sorb pollutants. Product literature on
the material is helpful in determining the compatibility of a
particular plastic to a pollutant.
2. Exposure Chamber
The actual exposure chamber can consist of glass boxes,
beakers, aquaria, or other containers of appropriate material
(see Figure IX-1). For most species, beakers are an inexpensive
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FIGURE IX-1
Representative Sediment Exposure System
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exposure chamber. The diameter of the beaker and the sediment
depth in the beaker should be sufficient to allow the organism to
bury and/or construct normal tubes. With Macoma nasuta. about 50
mm diameter beakers (Corning #1000, 100 ml) with about 50 g of
sediment were sufficient for 25-32 mm clams (anterior to
posterior measurement) to bury normally. With 35-48 mm clams, 68
mm diameter beakers (Corning #1000, 250 ml) with about 100 g of
sediment were required. In both cases, the initial depth of the
sediment was about 3-5 cm depending on sediment type. If the
beakers are placed into an aquarium, the beaker height should be
several centimeters less than the water height to allow for
circulation into the beakers. For funnel-feeding arenicolid
worms, a long narrow glass box (about l"w, 7"1, 4.5"h) is a more
appropriate shape (see Figure IX-IB). The opening of the
exposure chamber should allow the periodic addition of feeding
sediment.
3. Static Exposure System
The simplest static systems are individual aquaria or
beakers filled with water as commonly used in sediment toxicity
tests (e.g., Swartz et al., 1985a). A more common design for
bioaccumulation tests are sets of beakers submerged in aquaria
(e.g., Ferraro et al., 1990). The beakers or aquaria should be
covered to reduce evaporation and gently aerated to maintain
dissolved oxygen levels. Drain tanks about once every two days
by siphoning water from the aquaria, but not the individual
beakers. Retain the waste water in a container for treatment if
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it is suspected of containing unacceptable pollutant levels for
disposal down the sanitary sewer. Gently refill the aquarium
with water of the correct salinity and temperature, and restart
the aeration.
4. Flow-Through Exposure Systems
For flow-through systems, chambers may be sets of beakers
maintained in aquaria (Figure IX-1) or entire aquaria. Flow-
through systems have the advantages of removing waste products
and maintaining oxygen. Though desirable, a flow-through design
is not normally required for a successful bedded sediment test.
To avoid cross contamination, water flowing through one container
must not flow into another container. Water exiting the systems
should be passed through a charcoal filter if substantial
desorption of pollutants from the sediment is anticipated.
Similarly, resuspended sediment should be trapped and retained as
waste. Examples of conducting flow-through tests can be found in
U.S. EPA (1978), Rubinstein et al. (1980), and Rubinstein et al.
(1987).
C. MULTIPLE SPECIES EXPOSURE CHAMBERS
If several species are being tested, it is possible to place
multiple species within each exposure chamber. The advantages of
multiple species per container include reduced space requirements
and a lower cost because of the reduction in the number of
chambers constructed and maintained. The greatest disadvantage
is the potential for negative interactions among the species,
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such as competition or predation, which could have unknown and
variable effects on uptake. For example, uptake could be reduced
in Macoma if siphon clipping by epibenthic shrimp or nereid worms
reduced the clam's feeding rate. Another disadvantage is that
the accidental loss of a chamber reduces the number of replicates
for each of the species.
If multiple species are placed within exposure chambers, the
amount of sediment initially added should at least equal the sum
of the amount required for each individual species. Most of the
potential interactions are density dependent, so increasing the
area of the chambers (while maintaining a sufficient sediment
depth) should reduce the intensity of any negative interactions.
An alternate design is the physical partitioning of the aquarium
with screens to separate the species (e.g., Rubinstein et al.,
1987).
Regardless of the specific design, the same numerical ratio
of one species to another must be maintained in replicate
chambers. It should also be noted that a paired-comparison
approach should be used when statistically comparing the tissue
residues of two species kept in the same chambers.
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CHAPTER X: EXPERIMENTAL INITIATION, MAINTENANCE AND SAMPLING
A. EXPERIMENTAL INITIATION AND MAINTENANCE
1. Pre-Experimental Preparations
Coordinate the collection and acclimation of the bioassay
organisms with the collection of the sediments so the experiment
may begin with a minimum of delay. The glassware, water delivery
system, and any stored water should be ready, as well as sampling
containers, labels and related paraphernalia. Beakers and other
containers should be pre-labeled. A detailed work schedule,
showing daily tasks and persons responsible for accomplishing
them, should be prepared before the sediment arrives. A
prearranged numbering scheme should be agreed upon with the
analytical chemists. It is critical to keep the analytical
chemists well informed of the sampling schedule so they can
prepare for the sample load. Arrange with maintenance personnel
to look for power failures, pump leaks, breakage of aquaria,
inadvertent switching on of lights at night, and other accidents.
Provide telephone numbers for key personnel responsible for
maintenance of the experiment in a prominent location (e.g., on
the door of the laboratory). Any safety warnings should also be
posted at entry points.
2. Experiment Initiation
Weigh all individual organisms or composites of organisms,
while taking care to minimizing exposure of soft-bodied organisms
to the air. To avoid temperature shock, maintain the air
temperature of the room at the experimental water temperature.
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All bivalves should be measured (anterior to posterior valve),
weighed, and individually marked with a random number. When
possible, the number on the clam should be the same as the number
on their exposure chamber. Clams can be marked with a laboratory
marking pen (e.g., SharpieR pen) by first scrubbing the shell
with a KimwipeR or other soft paper towel, blotting the shell,
and then allowing them to dry (about 15 minutes at 12°C). Mark
the same valve (i.e, right or left) in all clams. Discard any
organisms not meeting the criteria for size or condition.
Maintain some extra individuals for potential replacements within
the first 24 hours. Also, randomly choose some specimens for
wet-to-dry weight conversions and for long-term storage for
potential lipid analysis with a different technique (see Chapter
XI) .
Distribute measured aliquots of homogeneously mixed sediment
to each exposure chamber. Weighing the sediment aliquot is
preferable, but sediment volume can be used to estimate mass for
a particular sediment type. During the process of measuring out
aliquots of sediment, periodically re-stir the source to avoid
separation of the fines and interstitial water. If beakers are
used as the exposure chamber, gently tap the beaker to
consolidate the sediment and eliminate air bubbles. To avoid the
loss of surficial fines when filling the beakers, place a plastic
film over the sediment surface, slowly fill the beaker with
water, and then withdraw the film using forceps. Carefully place
the water-filled beakers into filled aquaria and allow any
suspended fines to settle.
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If aquaria or other large containers are used as the
exposure chambers, stir the sediment after adding the appropriate
amount to mix sediment and remove any bubbles. As with the
beakers, a plastic film should be placed over the sediment
surface when filling the aquarium with water. Position any
aerating device so that the induced turbulence does not resuspend
sediment.
Add the organisms after allowing the sediments to
consolidate and any suspended particles to settle, which will
normally take from 15 minutes to a day. Place animals on the
surface of the sediment and allow them to bury. To facilitate
burial, place Macoma nasuta left valve down ("bent-nose" up) on
the sediment. Mobile organisms, such as the polychaetes, should
be observed for a sufficient period to assure that they.bury in
the correct chamber and do not swim into another chamber. For
mobile worms, it may be necessary to place a screen on the tops
of beakers to keep them from swimming out.
2. Experiment Maintenance
Replace any animals whose behavior is abnormal (failure to
bury in the sediment, etc.) within the first 24 hours. Observe
the chambers daily and note any signs of abnormal activity (e.g.,
reduced production of fecal pellets, avoidance of the sediment).
Remove any beakers with dead organisms. It is especially
important to monitor for dead organisms in a static system.
Record temperature, salinity, and other water quality parameters
on a periodic basis (see Chapter IX). Replenish water in static
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experiments according to a preplanned schedule (Chapter IX), and
dispose of drained water in accordance with applicable rules for
hazardous waste.
3. Sediment Renewal
We strongly recommend periodic additions of small amounts of
the appropriate sediment type to each exposure chamber. Because
the bioavailable fraction may only be a small portion of the
total sediment-associated contaminant (see Landrum, 1989), it is
possible for organisms to deplete the available fraction,
especially in organisms which ingest sediment from a restricted
feeding zone (e.g., surface deposit-feeding bivalves). For
example, depletion of the bioavailable fraction may be the reason
that tissue residues of 35 of 37 compounds declined between day
39 and day 79 in Oliver's (1987) study of uptake by oligochaetes.
Also, without organic input from settling phytoplankton and with
low light levels inhibiting benthic microalgae, it is possible
for the nutrient quality of the sediment to decline over the
course of an experiment. Periodic sediment renewal should reduce
these potential laboratory artifacts and help maintain a more
constant pollutant concentration and food supply. The periodic
addition of sediment results in a pulsed-renewal exposure.
Without the addition of new sediment, the exposure is a single-
dose exposure.
The daily amount of sediment added should equal or exceed
the daily sediment processing rate of the organism. Sediment
ingesting clams such as Macoma require about 1 gram wet sediment
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per gram of wet tissue mass per day while arenicolid worms (2-6 g
wet weight) require about 10 g of sediment per day. It is
sufficient to add the sediment two or three times a week (e.g.,
about 3.5 g twice a week for a 1 gram Macoma). Previously, we
had frozen the sediment immediately before its addition to reduce
loss of fines (Specht and Lee, 1989; Boese et al., in press);
however, given the unknown effects of even short-term freezing,
we recommend adding the sediment using a cut-off plastic syringe.
We have successfully used a 3 cc syringe with both a fine sand
and a silt-clay. The volume of sediment in the syringe is a
simple way to estimate mass of sediment, though the volume to
weight ratio has to be determined for each sediment.
For long-term exposures (>28 days), we recommend
periodically replacing all the sediment in the chambers.
Replacement of sediment reduces the possibility of depletion of
the bioavailable fraction of the pollutants and/or food, and
avoids excessive pelletization of the sediment. Additionally,
the periodic addition of surface sediments will overfill most
chambers within a few weeks, requiring a complete replacement of
the sediment. Replacement on a 28-day schedule should suffice,
and coordinates with the long-term sampling schedule (see Chapter
IV). If a field sediment is being tested, all the sediment
should be collected at the same time and the renewal sediment
stored until needed. If a spiked sediment is being tested, it
may be preferable to spike new sediment for replacement.
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Do not feed the test organisms a supplemental source of food
in either 28-day or long-term experiments. By ingesting the
added food, the organisms are presumably ingesting less sediment,
which could result in an underestimation of the bioavailability
of the sediment-associated pollutants. The addition of food is
not required as shown by long-term maintenance (>28 days) of
deposit-feeding bivalves (e.g., Specht and Lee, 1989),
polychaetes (e.g., McElroy and Means, 1988), and crustaceans
(e.g., Landrum, 1989) without supplementing the sediment with an
artificial food source.
B. SCHEDULE FOR ABIOTIC AND BIOTIC POLLUTANT SAMPLES
Samples of sediment, water, and biota should be taken for
pollutant analysis before, during, and after testing. Sampling
techniques and apparatus may vary with the nature of the
sediment, species of test organism, and compound(s) of interest.
As the manner in which samples are taken may affect the analysis,
consistency in sampling for any given parameter is essential.
1. Overlying Water
Although no pollutants are intentionally added to overlying
water in sediment bioaccumulation tests, contaminants may be
introduced from the water supply system, leached from the
sediment, or present on resuspended particulates. The activities
of some species (e.g., Yoldia) can resuspend considerable amounts
of fine-grain material directly into the water column. With a
randomized block design (Figure III-3f), bioturbation may lead to
cross-contamination between treatments. This potential uptake
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from the water needs to be quantified to differentiate it from
uptake from the bedded sediment and to check for possible cross-
contamination among treatments.
At a minimum, overlying water should be sampled for
pollutants from each treatment at the beginning, middle, and end
of the test period (i.e., TQ, T14, and T2Q). A sample from each
aquarium should be analyzed if statistical comparisons are
planned, though in many cases it would be acceptable to composite
water samples from aquaria of the same treatment. If samples are
composited, individual samples from each aquarium should be
archived in case a more detailed analysis is required. Samples
should also be taken during periods of high turbidity or other
unusual water quality.
Overlying water should be sampled at mid-depth from each
exposure unit. When experimental units share the same overlying
water (e.g., test beakers within the same aquarium), overlying
water should be sampled from mid-depth of the entire container.
Care should be taken to avoid disturbing the flocculent material
at the sediment-water interface. Sampling apparatus (pipettes,
sample vials) should be made of materials that do not appreciably
absorb or leach pollutants. To guard against cross-
contamination, rinse off the sampling apparatus after each use.
Sample volumes will depend upon the analytical technique used,
but may range from about 1 to 100 ml.
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2. Sediment and Interstitial Water
Sample all test, control, and reference sediments before the
addition of organisms (tQ sample) and at the end of the exposure
(typically t28). These sediment samples should be analyzed for
pollutant concentrations, TOG, and moisture content. In most
cases, it is adequate to conduct the grain size analysis only on
the initial sample.
One procedure for sampling sediment for organic pollutants
from exposure chambers is as follows:
1. Remove overlying water from the exposure chamber by
siphoning or decanting, taking care not to disturb the surface
floe. Depending on the procedure, interstitial water samples
may be taken at this stage.
2. Remove the test organism(s) from the sediment. Larger
bivalves can be directly removed with forceps. Spread the
sediment out in a tray to remove small bivalves and
polychaetes. Do not use any water to remove the sediment from
the exposure chambers.
3. Homogenize the test sediment from each exposure chamber by
stirring with a TeflonR coated spoon, glass rod, or other
inert utensil. Take a sediment sample from each exposure
chambers, place in a labeled sample vial and freeze. These
individual samples will either be analyzed or archived if
composites are analyzed.
4. If composites are going to be taken, the compositing
strategy will depend upon how the exposure chambers were
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allocated among aquaria. If only one treatment type is placed
in each aquarium, composite all the beakers within an
aquarium. If the exposure chambers are allocated randomly
among aquaria, combine all the sediment from each treatment
(i.e., sediment type) regardless of aquarium. In both cases,
homogenize the sediment, take replicate samples from each
composite, and freeze until analyzed.
Extra sediment samples should be taken from individual
exposure chambers (and from any composites) and frozen in case
there is an analytical failure or greater statistical power is
required.
Because this procedure exposes the sediment to the air,
reduced metal forms will be oxidized. If metal speciation will
be studied, the procedure should be modified, especially steps 2
and 3, to minimize the sediment's exposure to air. One
possibility is to take small sediment cores from the exposure
chambers. Regardless of sampling scheme, interstitial water
should be collected at the same time as the sediment samples.
Interstitial water may be collected by a variety of methods
including centrifugation, sediment squeezing, and dialysis
membranes (see Chapter VI).
C. METHODS OF BIOTIC SAMPLING
Test organisms need to be carefully removed from the
sediment, as described above, and all adhering particles removed.
A gentle rinse with clean seawater will help remove particles
from polychaetes. In general, organisms should be placed in
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control sediment to purge their gut contents for 24 hours before
chemical analysis (see Section D). At the end of the purging
duration, collect the organisms by gently spreading the sediment
out in a tray and removing the organisms using forceps or gently
sieving the sediment.
After collection, rinse the organisms with clean seawater,
blot them dry, and then weigh them. Measure the shell length of
bivalves. Organisms should be analyzed immediately or frozen in
baked-out aluminum foil or glass vials. The entire soft-tissue
of each individual or composite of individuals from an
experimental unit should be prepared for analysis. In many
cases, the tissue from each experimental unit will first be
homogenized and then subsamples taken for organic, metal, and
lipid analyses, and archiving. The type of homogenization
technique will depend upon size and tissue consistency of the
organism, the pollutant of interest, and the analytical
procedures used for pollutant analysis.
D. GUT PURGING
When a whole-body tissue analysis is conducted on a deposit-
feeder, any pollutants associated with the mineral particles and
detritus in the gut are included. Depending on the mass of
sediment and the associated pollutant concentration, the gut
sediment can measurably increase the apparent whole-body tissue
residue. Allowing the organism to purge its gut (i.e., defecate)
in uncontaminated sediment reduces or eliminates this positive bias
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TABLE X-l: Errors Associated With Gut Sediment/Purging
I. Gut Sediment Introduces Greatest Error:
1. In organisms that selectively ingest high organic particles.
2. In organisms with a large gut capacity.
3. During early stages of uptake when tissue residues are low.
4. For compounds not extensively bioaccumulated, especially
high Koc compounds with steric hindrance to uptake.
II. Purging Introduces Greatest Error:
1. For rapidly depurated/metabolized compounds.
2. In organisms which do not clear gut in water.
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However, pollutants will depurate or be metabolized during
purging, resulting in an underestimation of the bioaccumulation.
The type and extent of the error will depend upon many factors,
including the feeding behavior of the organism and the nature of
the pollutant. Factors influencing the errors associated with
gut sediment or purging are summarized in Table X-l.
1. Standard 24-Hour Purge
Organic compounds with high Kow values (e.g., PCBs, DDT,
BaP) are usually the greatest environmental concern in terms of
bioaccumulation. Most of these compounds are slowly depurated,
so a relatively small amount should be lost during purging.
Therefore, we recommend a 24-hour gut purging as the standard
procedure for sediments known or suspected to contain more than
trace amounts of these pollutants. A 24-hour depuration period
is sufficient for organisms to defecate the majority of their gut
contents without introducing substantial errors from pollutant
depuration or metabolism.
Many deposit-feeders require the ingestion of sediment to
completely void their gut contents, so organisms should be placed
in control sediment to assure complete purging. Reference
sediment should not be used as the purging sediment. Maintain
environmental conditions (e.g., temperature, salinity) as during
the exposure phase. The organisms in the control and reference
sediment(s) should undergo the same purging treatment as
individuals exposed to the test sediment. Organisms from
different treatments should be kept in separate containers to
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avoid any possibility of cross-contamination. Observations
should be made on whether feces were produced during the purging
period and on the general health of the organisms.
There are several other techniques or modifications to the
standard 24-hour purge which may be considered in specific cases.
These methods are discussed in Appendix X-l.
2. When Not To Purge
There are certain situations when gut purging may introduce
a greater error than leaving the gut sediment. In the first
situation, the primary focus of the study is comparing laboratory
and field studies. In most cases, it is impractical to purge
field collected organisms. Therefore, to assure that the
laboratory and field results are directly comparable, laboratory
organisms should not be purged. In the second case, the primary
focus is to determine the trophic transport of pollutants. As
deposit-feeders extract sediment-associated pollutants in their
guts (e.g., Lee et al., in press), it is likely that predators
would also extract a certain percentage of the pollutants from
their prey's gut sediment. In the final case, the primary focus
is on lower molecular weight PAHs. These compounds can be
depurated and metabolized rapidly (see Table X-2) so that a 24
hour purge can result in a greater error than leaving the gut
sediment.
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TABLE X-2: Depuration Loss Of Pollutants During 24 and 72 Hour
Gut Purges
COMPOUND
ORGANISM
% LOST (HRS)
REF.
24
72
PCB Crangon septemspinosa
HCB Macoma nasuta
BaP Pontoporeia hoyi
Phe Pontoporeia hovi
3
4
8 McLeese et al., 1980
12 unpublished data
4-28 11-64 Landrum & Poore,
1988
11-54 28-90 Landrum & Poore,
1988
PCB = Aroclor 1254
HCB = Hexachlorobenzene
BaP = Benzo(a)pyrene
Phe = Phenanthrene
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E. ACCEPTABLE LEVELS OF MORTALITY
According to ASTM (1984) guidelines for bioconcentration
tests, a test is unacceptable if "more than 10% of the organisms
in any treatment died or showed signs of disease, stress, or
other adverse effects." This criterion is applicable to studies
of spiked sediments in which it is possible to adjust pollutant
concentrations. Repeat any 28-day spiking experiment at a lower
pollutant concentration if 10% or more of the organisms in any
treatment die or show overt signs of stress. Signs of stress
include avoidance of the sediment, non-burial, casting off of
siphons, abnormal tube construction, and reduced ventilation or
sediment processing rates.
In contrast to most experimental studies of bioavailability,
many of the field sediments or dredge materials of environmental
concern will have moderate to high toxicity. With these
sediments, it may be impossible or difficult to meet the 10%
mortality criterion. However, this may not represent a serious
problem as the purpose of evaluating these sediments is to
determine the extent of bioaccumulation which will result from a
particular sediment. Presumably, the mortality in the laboratory
would mimic the response in the field and so represent the actual
effect of the sediment.
Because of the different purposes of tests conducted on
field versus spiked sediments, we suggest not automatically
rejecting bioassays with greater than 10% mortality in the test
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sediment. The determining factor, in deciding whether to accept a
test treatment with high mortality is whether there are adequate
replicates to obtain sufficient statistical power (see Chapter
III). If the statistical power is insufficient, the experiment
should be repeated. Also, mortality or stress in greater than
10% in the control or reference sediment would indicate initially
stressed organisms, contamination of the system, or unacceptable
control or reference sediment. In such cases, the cause of the
problem should be determined and the experiment repeated.
Consider using a more pollutant-resistant species or diluting the
sediment to reduce toxicity (see Appendix V-l) in any future
tests if the mortality in the test sediment exceeds 25%.
In some regards, high mortality in field sediments is a moot
problem because any sediment sufficiently toxic to kill a
substantial proportion of the recommended test species presumably
would be unacceptable based on toxicity. However, even in cases
where a sediment is rejected on the basis of toxicity, a
bioaccumulation test conducted on the diluted sediment may help
identify the compounds responsible for the toxicity.
F. CHAIN OF CUSTODY
In the event that litigation is expected, it is imperative
to follow proper sample chain-of-custody procedures so that the
results are acceptable for court. We recommend following the
chain-of-custody procedures published by the National Enforcement
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Investigations Center (U.S. EPA, 1986a). Other sources include
"Quality Assurance and Quality Control (QA/QC) for 301(h)
monitoring programs: Guidance on Field and Laboratory Methods"
(Tetra Tech, 1986b) and U.S. EPA Contract Laboratory Program
(U.S. EPA, 1988, 1989).
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CHAPTER XI: POLLUTANT AND LIPID ANALYSIS
A. POLLUTANT ANALYSIS
The specifics of the techniques used to analyze sediment,
water, and tissues for pollutants is a complex subject beyond the
scope of this manual. Discussions of analytical techniques can
be found in Tetra Tech (1985b, 1986f,g,h) and U.S. EPA (1988,
1989). It is possible, however, to offer several guidelines.
First, analytical techniques are media dependent. Thus, time
should be allocated for modifying the procedures for the various
media and any special conditions (e.g., high TOC sediment, low
tissue biomass). Second, a harsh extraction technique should not
be used when analyzing sediments for metals since such a
technique can extract biologically unavailable metals from the
mineral matrix. A discussion of various metal extraction
techniques is found in Luoma and Bryan (1978) and Waldichuk
(1985). Third, to the extent possible, the PCB analysis should
be at the level of identifying and reporting specific congeners
rather than Aroclor equivalents. In particular, the more toxic
planar congeners need to be identified. A thorough review of PCB
congeners, including which to analyze, can be found in McFarland
and Clarke (1989).
The required or desired detection limits will have a major
effect on the choice of analytical techniques and on the ability
to interpret the data. In some cases, the detection limits and
analytical procedures will be specified by the pertinent
regulation while in other cases the decision will be determined
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by the researcher. If no detection limits are specified, we
recommend that, at a minimum, the analytical techniques meet the
requirements of the U.S. Environmental Protection Agency's
Contract Laboratory Program requirements (U.S. EPA, 1988, 1989).
The quantification limits from these documents are summarized in
Table XI-1. These protocols cover both metals and organics in
water and sediment. Although tissues are not covered by these
protocols, it should be possible to obtain the same
quantification limits as with the sediments.
Control samples or samples from relatively clean areas
contain low concentrations of pollutants, and may require lower
detection limits to achieve satisfactory results. The methods
developed for measuring pollutants in samples collected from the
PSDDA control sites in Puget Sound (U.S. ACE, 1988) are
suggested in such cases. The PSDDA values include tissues as
well as water and sediment, and are summarized in Table XI-2.
A complete quality assurance/quality control plan is a
central part of any analytical procedure. Information on
analytical QA/QC procedures are available from several sources
(U.S. EPA, 1988, 1989; U.S. ACE, 1988). An important part of any
QA/QC program is the use of reference samples and standards.
Reference samples and standards are available from the U.S. EPA
in Cincinnati, OH; Las Vegas, NV; and Research Triangle Park, NC,
as well as the National Institute of Standards and Technology
(Office of Standard Reference Materials, Room B311, Chemistry
Building, NIST, Gaithersburg, MD 20899).
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TABLE XI-1: U.S. EPA Contract Laboratory Program Quantitation
Limits for Water and Sediment With Estimates for Tissue Matrices
Organics
Water (a) Sediment (b) Tissue (c)
Volatiles
Semivolatiles
Pesticides/PCB's
5-10
10-50
0.05-1
0.5-10
330-1600
8-160
0.5-10
330-1600
8-160
For individual pollutants - refer to CLP Statement of Work
Metals
Water (a)
Antimony
Arsenic
Cadmium
Copper
Lead
Mercury
Nickel
Silver
Zinc
Metals not listed
20-300
5-100
0.5-10
5-100
5-100
0.2-20
5-100
1-25
0.2-4
refer to CLP Statement of Work
a = ug/L
b = ug/kg wet weight
c = ug/kg wet weight basis. These values were estimated from the
sediment values on the premise that tissue and sediment
pollutant concentrations are of a similar magnitude and are
analyzed by similar techniques.
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TABLE XI-2.
Organics
PSDDA Low Limits of Detection for Water, Sediment
and Tissue Matrices
Sediment (a)
Tissue (b)
Volatiles
Semivolatiles
Pesticides/PCB
Metals
Antimony
Arsenic
Cadmium
Copper
Lead
Mercury
Nickel
Silver
Zinc
a = ug/kg dry
b = ug/kg wet
c = ug/L (ppb)
d = mg/kg dry
e = mg/kg wet
•s
Water (c)
3
1
0.1
1
1
0.2
1
0.2
1
weight (ppb)
weight (ppb)
weight (ppm)
weight (ppm)
10-20
1-50
0.1-15
Sediment (d)
0.1
0.1
0.1
0.1
0.1
0.01
0.1
0.1
0.2
5-10
10-20
0.1-20
Tissue (e)
0.02
0.02
0.01
0.01
0.03
0.01
0.02
0.01
0.20
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B. LIPID ANALYSIS
A number of studies have demonstrated that lipids are the
major storage site for organic pollutants in a variety of
organisms (Roberts et al., 1977; Oliver and Niimi, 1983; de Boer,
1988). Because of the importance of lipids, bioaccumulation
programs have recently attempted to normalize tissue pollutant
concentrations to the tissue lipid concentration. For example,
lipid concentration is one of the factors required in deriving
the Accumulation Factor (AF) (see Appendix 1-1). The approach,
however, has experienced difficulties because of the differences
in the lipid concentrations reported from the wealth of different
lipid methods used (see Kates, 1986 for discussion of lipid
methodology). Work in this laboratory has shown that differences
in lipid technique can result in 3-fold differences in lipid
concentrations. These differences in lipid concentrations
directly translate into a similar variation in the lipid
normalized pollutant concentrations or Accumulation Factors.
To allow lipid normalized tissue residues or AFs to be
compared, it is necessary to either promulgate a standard lipid
technique or to intercalibrate the various techniques.
Standardization on a single method is difficult because the lipid
methodology is often intimately tied in with the extraction
procedure for pollutant analysis. Instead, we recommend that one
lipid technique be chosen as an "intercalibration standard".
Then, regardless of what method is used, the results would be
reported in equivalent units of the standard.
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The difficulty with this approach is deciding upon which
technique to use as the standard. We are presently investigating
the advantages and disadvantages of different techniques, but
have not yet reached a consensus. As in interim solution, we
suggest the Bligh-Dyer lipid method (Bligh and Dyer, 1959) as a
temporary "intercalibration standard". Folch et al. (1957)
developed a total lipid method that extracts the neutral and
polar lipids (i.e. total lipids) from biological samples using
chloroform and methanol as the solvent system. Bligh and Dyer
(1959) improved upon the method by providing a cleanup for the
extracted lipid residues.
The potential advantages of Bligh-Dyer include its ability
to extract neutral lipids not extracted by many other solvent
systems and the use of Bligh-Dyer (or the same solvent system) in
numerous biological and toxicological studies (e.g., Roberts et
al., 1977; Oliver and Nimi, 1983; de Boer, 1988; Landrum, 1989).
Because the technique is independent of any particular analytical
extraction procedure, it will not change when the extraction
technique is modified or changed. Additionally, the method can
be modified for small tissue sample sizes as long as the solvent
ratios are maintained. We have successfully used a modified
technique with tissue samples as small as 1 g wet tissue and
micromethods using chloroform - methanol requiring only milligram
amounts of tissue were developed by Herbes and Allen (1983) and
Gardner et al. (1985).
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One potential disadvantage of the Bligh-Dyer is that by
extracting many of the lipids not extracted by other techniques,
it may be extracting lipids that are not important to the storage
of neutral organic pollutants. Seasonal changes or interspecific
variations in these non-active lipid fraction could obscure the
relationship between lipid content and pollutant accumulation.
However, the standard Bligh-Dyer technique may not solublize
triglycerides (Gagney, P., pers. comm., U.S. EPA, ERL-A, Athens,
GA), thereby underestimating the lipid pool important to
pollutant storage. Other drawbacks are the use of chloroform,
which is a carcinogen, and the need to conduct an additional
analysis instead of measuring the lipids as part of the normal
organic extraction procedure (e.g., Rubinstein et al., 1987; Lake
et al., in review). However, the alternative lipid methods all
have similar limitations, and we believe that Bligh-Dyer is the
best interim calibration method.
If the Bligh-Dyer method is not used as the primary lipid
method, compare the chosen lipid method with Bligh-Dyer for each
tissue type. The chosen lipid method could then be converted to
"Bligh-Dyer" equivalents and the lipid normalized tissue residues
could then be reported in "Bligh-Dyer equivalents". Because of
the interim nature of this suggestion, we also suggest that extra
tissue of each species be frozen for future lipid analysis in the
event that a different technique proves more advantageous.
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C. SAMPLE STORAGE
For organics, the U.S. EPA Contract Laboratory Program (U.S.
EPA, 1988) requires that the samples be protected from light and
refrigerated at 4°C (± 2°C.) from the time of receipt until they
are extracted and analyzed. Water samples shall be extracted
within 5 days of the receipt of the sample. Sediment samples
shall be extracted within 10 days of the receipt of the sample
and if continuous extraction procedures are employed, extraction
of water samples shall be started within 5 days of the receipt of
the sample.
For inorganics, the U.S. EPA Contract Laboratory Program
(U.S. EPA, 1989) requires that soil and sediment samples be
maintained at 4°C. (± 2°C.) until analyzed. Samples for mercury
shall be analyzed within 26 days of the receipt of the sample.
Samples for metals shall be analyzed within 180 days of the
receipt of the sample.
At times, other program priorities (e.g., analysis of
archived samples) do not allow one to abide by the requirements
set by the Contract Laboratory Program. In those cases, it is
suggested that the samples either be frozen (-20°C) in air tight
containers or dried depending on the type of sample and the
analyses required. Purging the container with nitrogen prior to
sealing will delay the degradation of some pollutants as well as
lipids. Sample containers should be as full as practical to
prevent loss of moisture from the sample. Sediment samples so
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preserved are stable for at least 6 months, if not longer (Tetra
Tech, 1986d). Tissue and water samples are expected to be at
least as stable as sediments.
D. REPORTING OF RESULTS
Investigators have reported results on either a dry or wet
basis, usually without a conversion factor between the two and
sometimes without any indication of which was used. This makes
it difficult, or impossible, to compare results from different
studies. In general, a dry-weight basis is preferred for both
sediment and tissue pollutant concentrations. However, certain
analytical techniques use wet tissue or wet sediment,
necessitating the calculation of wet-weight concentrations. To
allow comparisons among studies, the wet-to-dry weight ratios
should be reported for each tissue and sediment type. As
mentioned above, lipid values should be reported in "Bligh-Dyer
equivalents" along with any conversion factor(s) between lipid
methods.
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CHAPTER XII: STATISTICAL ANALYSES
The main objective of statistical testing is to determine
whether the mean tissue residues in animals exposed to the test
sediment are significantly greater from those in the control
and/or reference sediments, or greater than a specified criterion
value such as an FDA Action Limit. Additional statistical tests
comparing the means of other tissue residues (e.g., control vs
reference) or sediment characteristics will also be conducted,
but the same principles and methods apply. A summary of the
standard statistical tests and their interpretation are
summarized in Table XII-1 and Table XII-2.
To perform statistical testing, replicate samples must have
been taken to provide an estimate of variability. Non-replicated
samples (i.e. concentration from a single composite sample)
cannot be compared using these methods. In these tests, the
concentration of each chemical in a tissue or sediment sample is
considered statistically independent and is compared separately.
Comparisons of tissue residues of different chemicals within the
same organisms requires the use of "repeated measures" (Section E)
Standard deviations (SD or s) or standard errors (SE) and
number of the replicates (n) should always be reported in
addition to mean values. When composited values are used, report
the number of organisms per composite (if the composite comprises
the experimental unit) or the number of experimental units per
composite, as well as the number of replicate composites sampled.
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Prior to conducting any statistical analyses, it is
necessary to decide whether the comparisons between means are to
be multiple or pairwise. Pairwise comparisons include
comparisons of a test and control/reference mean for tissue
concentrations, sediment characteristics, etc. Pairwise
comparisons also include the comparison of the control with the
reference mean and comparisons of a mean and a specified
criterion value such as comparison of a test tissue residue with
an FDA action limit. Multiple comparisons involve comparisons of
more than two means simultaneously. Multiple comparisons are
used in cases such as determining whether three or more test
tissue concentration means are equal or whether all the TOC
values for the sediments (test(s), control and reference) are
equal.
After the applicable comparisons are determined, the data
need to be tested for normality to determine whether parametric
statistics are appropriate and whether the variances of the means
to be compared are homogeneous. If normality and homogeneity of
variances are established, t-tests can be performed in the case
of pairwise comparisons or ANOVA in the case of multiple
comparisons.
A. TESTS FOR NORMALITY AND HOMOGENEITY OF VARIANCES
Before conducting parametric statistics, the data need to be
checked for both normality and homogeneity of variances. The
data for each chemical or sediment parameter are tested
separately. Commonly used tests for testing normality are the
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Kolmogorov-Smirnov one-sample test and the chi-square test (Sokal
and Rohlf, 1981). However, these tests are not very powerful,
especially if sample sizes are small (such as 8 replicates).
More powerful, but less common, tests of normality such as
Shapiro-Wilk and K2 tests (D'Agostino and Stephens, 1986) can be
used for small sample sizes.
If the data are not normally distributed, the data can often
can be transformed to achieve normality. The logarithmic and
arcsine are two commonly used transformations for concentrations.
It may be necessary to apply different transformations to
different chemical or sediment parameters. See Sokal and Rohlf
(1981) for a more extensive discussion on transformations. If
normality cannot be established, nonparametric tests for
comparisons of two means, such as the Mann-Whitney test and the
Tukey's Quick test should be used. These non-parametric tests
are usually not as powerful as the more common parametric tests,
such as the t-test or Analysis of Variance (ANOVA). See Daniel
(1978) for a discussion of non-parametric statistics.
The variances of the samples to be compared should be tested
for homogeneity. This is performed using an F-test when
comparing two variances or Bartlett's test when comparing
more than two variances. If the variances are considered
homogeneous, then a t-test or ANOVA is appropriate. If the
variances are heterogeneous, the data can be transformed in an
attempt to achieve homogeneity. Under conditions of variance
heterogeneity, a modified t-test for comparisons of two means or
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approximate tests for multiple comparisons can be performed. See
Sokal and Rohlf (1981) for a more extensive discussion on
appropriate tests when different treatments have unequal
variances.
B. PAIRWISE COMPARISONS
Pairwise comparisons are performed using Student's t-test,
using a pooled variance estimate when variances are homogeneous.
Under conditions of variance heterogeneity, a modified t-test can
be used (see Sokol and Rohlf, 1981). Prior to analysis, it must
be established whether the t-test performed will be a one-tailed
or two-tailed test and whether the Type I error rate should be a
comparison-wise or experiment-wise error rate. These
considerations are discussed below.
1. One-Tailed versus Two-Tailed Tests
In formulating a statistical hypothesis, the alternative
hypothesis can be one-sided (one-tailed test) or two-sided (two-
tailed test). The null hypothesis (Ho) is always whether two
values are equal. A one-sided alternative hypothesis (Ha) is
that there is a specified relationship between the two values
(e.g., one value is greater than the other) versus a two-sided
alternative hypothesis (Ha) which is that the two values are
simply different. 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 test tissue
residue is greater than the control tissue residue. In contrast,
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the two-tailed test is used when the direction of the difference
is not important or cannot be assumed prior to testing. An
example of an alternative two-sided hypothesis is that the
reference sediment TOC is simply different from the control
sediment TOC.
Because control tissue residues and sediment pollutant
concentrations are presumed lower than reference values which are
presumed lower than test values, we recommend conducting one-
tailed tests in most cases. For the same number of replicates,
one-tailed tests are more likely to detect statistically
significant differences between treatments (i.e., have a greater
power). This is a critical consideration when dealing with a
small number of replicates (such as 8 per treatment). The other
alternative to increasing statistical power is to increase the
number of replicates, which increases the cost of the bioassay.
The use of one-tailed tests deviates from the usual
experimental procedure, but is justified where a regulatory
action would be taken only if the tissue residues in organisms
exposed to a test sediment were greater than those in a control
or reference sediment. For example, a dredge material might be
denied disposal in open water if the tissue residues in the test
sediment (i.e., dredge material) were significantly greater than
those in the reference sediment. However, the same regulatory
decision (i.e., allow disposal) would be reached whether the
tissue residues in the test sediment were equal to or less than
those in the reference sediment. The same reasoning would apply
when comparing a tissue residue to a tissue criterion.
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There are cases when a one-tailed test is inappropriate.
When no a priori assumption can be made as to which treatment is
higher than the other, a two-tailed test should be used. For
example, when comparing TOCs of the test and reference sediments,
a two-tailed test should be used. A two-tailed test should also
be used when one regulatory action will be taken when the two
treatments are equal and another when they are not equal,
regardless of which one was larger or smaller. This would be
unusual for tissue residues, but would apply to other benthic
parameters. For example, a two-tailed test should be used when
comparing benthic biomass at a control and test site because both
enhanced and reduced biomass are indicators of organic enrichment
(Pearson and Rosenberg, 1978), so the regulatory question is
whether there is any difference between the two sites. A two-
tailed test should also be used when comparing tissue residues
among different species exposed to the same sediment and when
comparing BAFs or AFs (see Appendix 1-1).
The appropriate one-tailed and two-tailed tests for the
bioaccumulation test are summarized in Table XII-1 and
Table XII-2.
2. Comparison-wise versus Experiment-wise Error Rates
The Type I error rate used in the tests will be chosen
either as a comparison-wise or experiment-wise error rate
depending on whether one decision is made for each pairwise
comparison or from a set of pairwise comparisons. For cases
where test sediments are chosen in a stratified manner or along a
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TABLE XII-1: Summary of Statistical Analyses
Hypothesis
PAIRWISE COMPARISONS
Test (s)*
Comments
Normality
Equality of Variances
Equality of Means
Equality of Means
Equality of a Mean
and a constant
Equality of Means
Chi-square or
Kolmogorov-Smirnov
F-test
t-test
modified t-test
t-test
nonparametric tests
Try transformations
if not normal
Try transformations
if not equal
One-tailed with a
priori knowledge
otherwise two-tailed
If variances are not
equal
One-tailed with a
priori knowledge
otherwise two-tailed
If normality is not
established
Hypothesis
MULTIPLE COMPARISONS
Test(s)*
Comments
Normality
Equality of Variances Bartlett's test
Chi-square or
Kolmogorov-Smirnov
Equality of Means
Equality of Means
ANOVA
nonparametric tests
Try transformations
if not normal
Try transformations
if not equal
If normality is
established
If normality is not
established
Often more than one test can be used for the same hypothesis.
Each test will have different assumptions. Chose the test with the
assumptions most closely matching your specific conditions and
requirements.
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gradient (see examples a and b in Figure XII-1) and any decisions
will be made on a case by case basis, a comparison-wise Type I
error rate of 0.05 should be used for each comparison. For
example, a comparison-wise error would be used in deciding which
specific stations along a gradient were acceptable or not
acceptable.
If the test sediments are selected from a supposedly
homogeneous source {e.g., multiple sediment samples from a dredge
barge, see example c in Figure XII-1) and the decision to accept
or reject the sediment will be made from the results of several
pairwise comparisons, then an experiment-wise error rate of 0.05
should be used. In this example, a regulatory decision will
depend on the results from all the comparisons of the test
treatments to determine if the sediment in the barge is too
contaminated for disposal. Each individual comparison is
performed at a lower error rate such that the probability of
making a Type I error in the entire series of comparisons is not
greater than 0.05. This results in a more conservative test when
comparing any particular sample to the control/reference. Thus,
a single sediment sample from the barge that would have been
rejected at the 0.05 level may not be rejected at the lower
experiment-wise error rate, though the probability of rejecting
Ho for the entire set of samples is still 0.05. Use of
experiment-wise error rates adjusts for the possibility of random
differences when multiple samples are taken from a homogeneous
source (e.g., if 100 samples were taken, a certain percentage
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would be greater than the control/reference because of random
fluctuations). The error rate used in each comparison is a
function of the number of comparisons to be used in the decision
"experiment" and can be computed using the method of Dunn-Sidak
(Sokol and Rohlf, 1981) as:
alpha' = 1-(l-alpha)1/* (1)
where:
alpha = Type I error rate used for each
pairwise comparison
alpha = experiment-wise Type I error rate (0.05)
k = number of comparisons
When an experiment-wise error is used, the power to detect real
differences between any two means decreases as a function of k,
the number of comparisons.
C. MULTIPLE COMPARISONS
For comparisons involving several means, as in the case of
comparing TOG values among all sediment types, an ANOVA is first
performed to establish whether any of the means are different.
The ANOVA also provides a "best" estimate of the variance
(within-treatment error). If there are significant differences,
a series of t-tests can be performed for any planned (a priori)
comparisons (such as between test and control/reference) to
distinguish which means are different. For unplanned (a
posteriori) comparisons, such as between two reference tissue
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FIGURE XII-1
Sampling Schemes
for Comparison—wise (a. and b.)
vs Experiment—wise (c.) Error rates
Stratified selection of test sediments
Any
Harbor,
USA
X ~~ test site
Selection of test sediments
along a gradient
Point source pollutant gradient
X X X X X X
c.
X ~ test site
Selection of test sediments from a
presumably homogenous source
dredge barge
X ~ test site
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residues, tests such as the T-Method or Tukey Kramer procedure
(Dunnett's test) are more appropriate. See Sokal and Rohlf
(1981) for unplanned multiple comparison tests to determine which
is most suited for each case.
It is important to note that an ANOVA is inherently for two-
tailed comparisons. Therefore if the comparisons can be broken
down into a series of one-tailed pairwise comparisons, it is
preferable to perform the analysis in this manner because of the
increase in power. However, if the series of comparisons are
two-tailed, an ANOVA can be performed first to determine whether
any additional comparisons should be made.
D. INTERPRETATION OF COMPARISONS OF TISSUE RESIDUES
If the control mean tissue residues at day 28 are not
significantly greater than the day 0 tissue residues, it can be
concluded that there is no significant contamination from the
exposure system or from the control sediment. If there is
significant uptake, the exposure system and/or control sediment
should be reevaluated as to suitability. Even if there is a
significant uptake in the controls, it is still possible to
compare the controls and treatments as long as the pollutant
concentrations in the test tissue residues are substantially
higher. However, if control values are high, the data should be
discarded and the experiment conducted again after determining
the source of contamination.
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Comparisons between the 28-day control (and/or reference)
tissue residues and 28-day test tissue residues determines
whether there is statistically significant bioaccumulation due to
exposure to test sediment. Comparisons between control and
reference tissue residues at day 28 determines whether there is
statistically significant bioaccumulation due to exposure to the
reference sediment. When test tissue residues are compared with
a one-tailed test with a set criterion value (e.g., FDA Action
Limit), if no significant difference is detected, the residues
must be considered equivalent to the value even though
numerically the mean tissue residue may be lower.
The statistical interpretation of these and other tests are
summarized in Table XII-2.
E. ADDITIONAL ANALYSES
1. Testing BAFs and AFs
Statistical comparisons between ratios such as BAFs or AFs
are difficult due to computation of error terms. Since all
variables used to compute BAFs and AFs have errors associated
with them, it is necessary to estimate the variance as a function
of these errors. This can be accomplished using approximation
techniques such as the propagation of error (Beers, 1957) or a
Taylor series expansion method (Mood et al., 1974). BAFs and AFs
can then be compared using these estimates for the variance. See
Ferraro et al. (1990) for an example of this approach.
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TABLE XII-2: EXAMPLES OF ANALYSES AND INTERPRETATION OF RESULTS
HYPOTHESIS
TEST
INTERPRETATION OF
REJECTION OF NULL HYPOTHESIS
PHYSICAL PARAMETERS
Ho: TOCc=TOCi
Ha:
two-tailed TOC not equal between
t-test control and test sediment i
Ho: TOC =TOCr
Ha: TOCcVTOCr
Ho : TOC =TOCr=TOC1
Ha :
. =TOC
n
two-tailed TOC not equal between
t-test control and reference
ANOVA TOC of one or more
sediment differs
ADEQUACY OF CONTROL
Ho: Ctc=Ctu
Ha: Ctc>Ctu
TREATMENT DIFFERENCES
Ho: Cti=Ctc
Ha: Cti>Ctc
Ho: Cti=Ctr
Ha: Cti>Ctr
Ho: Ctr=Ct
Ha: Ctr>Ctc
Ho : Ct =Ct =Ct., = . . . =Ctn
Ha : Ct
one-tailed
t-test
one-tailed
t-test
one-tailed
t-test
one-tailed
t-test
ANOVA
Exposure system
contaminated
Sig. uptake from test
sediment i above control
Sig. uptake from test
sediment i aboye reference
Sig. uptake from reference
sediment above control
Uptake from one or more
sediment differs
Ho :
Ha:
_
ANOVA Uptake from one or more
test sediment differs
LONG TERM EXPOSURES
Ho: Ct(j)i=Ct(j+l)i=Ct(j+3)i
Ha: Ct()
ANOVA Ct.^ has not reached
steady-state
Ho = null hypothesis
Ha = alternative hypothesis
Ct = concentration of pollutant in tissue at day 28
Subscripts: c = control organisms or sediment
i = l,2,..,n test organisms or sediment
j = last sampling period
n = total number of test treatments
r = reference organisms or sediment
u = unexposed organisms
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2. Comparing Tissue Residues of Different Compounds
In some cases, it is of interest to compare the tissue
residues of different compounds. For example, Rubinstein et al.
(1987) compared the uptake of thirteen different PCB congeners to
test for differences in bioavailability. Because the values for
the different compounds are derived from the same tissue samples,
they are not independent and tend to be correlated, so standard
t-tests and ANOVAs are inappropriate. Rather, a repeated
measures technique (repeated testing of the same individual)
should be used where the individual (experimental unit) is
considered as a random factor and the different compounds as a
second factor. See Rubinstein et al. (1987) and Lake et al. (in
review) for an example of the application of repeated measures to
bioaccumulation data.
3. Analyses for Alternative Test Designs
Long-term exposures require a test to show that steady-state
has been reached. An ANOVA should be performed on the last three
sample sets. ASTM (1984) requires that there be no significant
difference (p>.05) between the means of these sample sets. If
apparent steady-state is reached, the mean of the samples taken
during apparent steady-state should be used for the steady-state
concentration value. For steady-state estimates based on uptake
and depuration tests, see Davies and Dobbs (1984) or Spacie and
Hamelink (1982) for details on the nonlinear parameter estimation
methods required to estimate these rate constants and steady-
state concentrations.
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CHAPTER XIII: REGULATORY STRATEGIES FOR USE
OF BIOACCUMULATION DATA
Bioaccumulation is the link between exposure and effects,
and thus can generate important insights into ecological effects,
human health risks, and the routes and extent of pollutant
exposure. However, its use in a regulatory context is not as
straightforward as acute toxicity data. Death is unequivocal and
undeniably bad for an individual organism. But what of a PCB
tissue residue of 1.5 ppm, or one of 50 ppb? The answer to
questions such as this are often not clear. Yet, many of today's
environmental problems are due to the accumulation and trophic
transport of sublethal concentration of pollutants rather than
major die-offs. Bioaccumulation and consumption of contaminated
seafood is certainly one of the primary environmental concerns of
the public.
In the previous chapter (Chapter XII), the statistical
procedures to test for increases in tissue residues were
discussed. It is important to note that statistical differences
in themselves do not necessarily indicate an environmental or
human health problem. Conversely, the lack of statistically
greater tissue residues in test sediment compared to a proper
control would be strong evidence that the test sediment would not
result in an environmental or human health problem for the
pollutants tested. The lack of a statistical difference between
a test and reference sediment would indicate that the
environmental problems resulting from the test sediment would be
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no worse than those from the existing reference sediment.
Whether this is environmentally acceptable depends on the present
environmental conditions at the reference site.
In this Chapter, we summarize some of the approaches to
interpreting the ecological and human health ramifications of a
significant increase in tissue residues in a test sediment or
dredge material. Some of the approaches are well established,
while other are still in the conceptualization stage. We only
consider bioaccumulation data in this Chapter, whereas an
environmental assessment would normally include sediment toxicity
testing, benthic community analysis, effluent testing, or a
number of other approaches. These various approaches generate
different types of information which complement each other.
A. NO FURTHER DEGRADATION
1. Approach
Compare tissue residues in organisms exposed to a test
sediment to those exposed to an appropriate reference sediment.
If the test tissue residues are not greater than those in the
reference sediment, it is concluded that the test sediment would
not result in a degradation of existing environmental conditions.
2. Advantages and Applicability
The approach is straightforward and does not require any
data other than the bioaccumulation tests. Comparison of
existing or predicted pollution effects to present ecological
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conditions is an established regulatory approach (e.g., 301(h)).
This approach should be applicable in any area in which an
appropriate reference sediment can be found.
3. Disadvantages
One potential problem with any approach that uses a field
comparison is the choice of an appropriate reference site. As
discussed in Chapter II, it is possible to get no statistical
difference between test and reference tissue residues, but still
result in unacceptable degradation. Without national guidelines,
there may be large differences in the "environmental quality" of
reference sites, and hence, in allowable tissue residues.
B: TISSUE RESIDUE EFFECTS
1. Approach
Relate the tissue residues to specific physiological or
biochemical effects on the organism.
2. Advantages and Applicability
The advantage of this approach is that it relates the tissue
residues to an effect on an organism. If a biochemical end-point
or "biomarker" is used, the approach can generate insights into
the mechanisms of stress. If such relationships can be
developed, it would be a relatively simple matter to assess the
environmental quality from tissue residues. The approach is
applicable to all organisms, and depending on the end-points
used, can be adapted for different sensitivities (e.g., early
warning vs. unacceptable impacts).
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3. Disadvantages
Because of the various mechanisms by which different
pollutants can affect organisms, and because the actual
mechanisms of toxicity may vary among taxa, there is often a weak
correlation among tissue residues and effects. The relationship
may also be weak if the stress is due to a small fraction of the
tissue residue (e.g., pollutant affect on nerves). Presently,
most of the techniques are not simple and often require expensive
equipment.
C. WATER QUALITY CRITERION TISSUE LEVEL APPROACH
1. Approach
Tissue residues are compared to those residues that would
occur at a water exposure to the Water Quality Criterion (WQC)
concentration. The tissue concentration which would result from
exposure to pollutants at the WQC are calculated by multiplying
the BCF for a compound by the WQC. This tissue concentration is
the "Water Quality Criterion Tissue Level" (WQCTL). Tissue
residues higher than the WQCTL indicate that the integrated
exposure through all routes was greater than allowed under WQC,
and thus is unacceptable. This approach is similar to the
Equilibrium Partitioning approach to deriving Sediment Quality
Criteria in its use of the WQC as the end-points, except that
tissue residues are used as the measure of exposure rather than
interstitial water concentrations.
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2. Advantages and Applicability
As with the Equilibrium Partitioning Approach, the WQCTL
approach is relatively simple, draws upon extensive previous
toxicological work, and is based on a well established regulatory
approach (WQC). Furthermore, using tissue residues as a measure
of the bioavailable pollutant concentration avoids the numerous
factors which can affect the concentrations and bioavailability
of interstitial water pollutants (e.g., complexation with DOM,
the "solids effects", sampling artifacts, etc.). Also, this
approach does not assume that interstitial water is the only
uptake route or that an organism's tissue residues can not exceed
a thermodynamic maximum. As both of these assumptions appear to
be incorrect for some species (e.g., selective deposit-feeders)
with some compounds (e.g., HCB, BaP), relaxing these assumptions
increases the applicability of using WQC as the basis for
regulating sediments. This approach could be used with any
compound that had a chronic WQC value.
3. Disadvantages
This is a new proposal, and the idea has yet to be
evaluated. As with the Equilibrium Partitioning approach, it
assumes that WQC values are applicable to benthic organisms. The
consistency of BCFs among water column organisms and BAFs or AFs
among benthic species needs to be evaluated. With benthic
species, the actual exposure may depend upon feeding type (Lee et
al., in press) or source of ventilated water (Winsor et al., in
press). Therefore, the exposure calculated using one species may
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not be applicable to a different species. Rapidly metabolized
compounds may give variable results. Finally, to the extent that
the WQC was based on tissue residues, the approach may be
circular.
D. FDA ACTION LIMITS
1. Approach
Compare the tissue residues in the benthic test species with
FDA Action Limits. Tissue residues exceeding an FDA Action Limit
are considered unacceptable.
2. Advantages and Applicability
The FDA Action Limits are a well established regulatory
criteria. The FDA Action Limits are used as end-points in the
evaluation of dredge materials (U.S. EPA/U.S. ACE, 1988).
3. Disadvantages
There are only a few FDA action limits for seafood. There
is concern that the criteria are not sufficiently protective of
human health, especially with high seafood consumption rates.
The Action Limits do not consider ecological impacts. Because of
the human health and ecological limitations, FDA Action Limits
can be considered "one sided" criteria, where exceeding the
limits is unacceptable but failure to exceed the limits is not
strong evidence for acceptability of a sediment.
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E. HUMAN HEALTH RISK ASSOCIATED WITH SHELLFISH
1. Approach
Calculate the excess human cancer risk from the consumption
of contaminated shellfish. The calculation of cancer risk
requires an estimate of the tissue residues, a cancer potency
value, and a lifetime consumption rate of the shellfish. Non-
cancer human health risks could also be calculated, though in
most cases, cancer will be the greater health risk. This
approach differs from using FDA Action Limits in that the risk is
based on an estimate of actual human exposure. Therefore, the
actual allowable concentration would depend upon the rates and
patterns of the consumption of shellfish in an area.
2. Advantages and Applicability
Determining the excess cancer risk associated with'
consumption of seafood is of obvious and direct concern to the
public. Such an approach would be applicable in all areas where
shellfish are harvested recreationally or commercially. The
human cancer model is an established regulatory method. Cancer
potency values are available for many of the environmentally
relevant pollutants. For some compounds (e.g., PCBs, BaP), human
health risk can generate a lower acceptable sediment
concentration than sediment toxicity (Lee and Randall, 1988), so
for these compounds, this approach is protective of the
environment and human health.
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3. Disadvantages.
The local per capita consumption rates of shellfish is
poorly known. Policy decisions must be made about what
constitutes an acceptable human health risk (e.g., 10"5 versus
10"6 excess cancer risks) and whether to regulate on an average
consumption or the maximum consumption rates (e.g., subsistence
fishing by certain ethnic groups). Use of default seafood
consumption rates for a site can generate unrealistically low
safe sediment concentrations in areas with little or no
harvesting of shellfish.
F. TROPHIC TRANSFER OF POLLUTANTS INTO PELAGIC FOOD WEBS
1. Approach
Predict the movement of pollutants from the benthos into
their predators and through the food web, up to and including
human consumers. The acceptability of a benthic tissue
concentration would be determined from the human health effects
associated with consumption of contaminated fishes (or ecological
effects, if end-points were available). The same human cancer
risk models would be used as used in calculating the risk
associated with shellfish. The use of human health risk models
with seafood is discussed in Tetra Tech (1986e). This Trophic
Transfer approach is a generalized form of the Shellfish approach
and the two would normally be combined (see Figure XIII-1).
2. Advantages and Applicability
Trophic transfer of contaminants from the benthos to their
predators is one of the major mechanisms by which certain
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pollutants are introduced into pelagic food webs. This approach
addresses the far-field effects of a contaminant as well as the
near-field effects. The approach would be applicable in any area
where fish are harvested recreationally or commercially. Models
predicting the average increases in various pollutants per
trophic level have been developed (Young et al., 1987; Young,
1988) .
3. Disadvantages
One limitation of this approach is the difficulty in
obtaining the data required to quantify the transfer of
pollutants from the benthos to their predators and through the
food web. The other major limitation is the uncertainty in
assigning human seafood consumption rates.
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FIGURE XIII-1
POSSIBLE REGULATORY STRATEGY FOR
HUMAN HEALTH CRITERIA IN ASSESSING
SEDIMENT CONTAMINATION
measure sediment parameters
predict benthic tissue residues
fail
shellfish harvested?
no
I yes
assess human health
I pass
fisheries feeding site
no
yes
predict extent of trophic transfer
I assess human health
fail I
pass
assess benthic effects
I pass
approve
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APPENDIX 1-1: ADDITIONAL METHODS TO PREDICT BIOACCUMULATION
A. FIELD COLLECTION
The most direct method of assessing tissue residues in
existing sediments is to measure residues in field collected
organisms. The field approach is appealing because it avoids any
laboratory artifacts, as well as the time, expense and facilities
required for laboratory tests. However, use of field collected
organisms has several limitations as a routine method.
The greatest problem is collecting sufficient tissue biomass
^.
of an appropriate species for chemical analysis. This problem is
especially acute at the most contaminated sites because smaller
species tend to dominate stressed communities and during the
early stages of recolonization (Pearson and Rosenberg, 1978;
Rhoads et al., 1978). In addition, benthic densities are reduced
under severe stress (Pearson and Rosenberg, 1978). Even when
sufficient biomass of a particular species can be collected at a
given station, it will often be impossible to collect the same
species from other stations located along a pollution gradient,
seasonally within a single station, or at an estuarine dredge
site and an open ocean disposal site.
One possible approach to collecting sufficient biomass is to
composite the various species collected from each site. Although
mixing species will increase biomass, tissue composites taken
from different stations or seasons are likely to be composed of
substantially different proportions of species and numbers of
individuals. These compounding factors will make it unclear
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whether patterns in tissue residues are due to differences in the
sites or interspecific differences in bioaccumulation. For
example, amphipods have a much greater ability to metabolize PAHs
compared to bivalves (Varanasi et al., 1985). Therefore, a
difference in PAH tissue residues among sites could reflect a
difference in the proportion of amphipods and bivalves rather
than a difference in the bioavailability of PAHs.
Another problem is that the exposure history of field
collected specimens is usually unknown. Many benthic species,
especially amphipods and some polychaetes, are mobile during a
portion of their life-history (e.g., Williams and Porter, 1971;
Williams and Bynum, 1972; DeWitt, 1988) and may have migrated
into a site recently. Although pollutant concentrations in
sediments are usually considered relatively constant,
resuspension events can obscure sediment-bioaccumulation
relationships. For example, deposition of resuspended
contaminated sediments in an uncontaminated site would form a
surface veneer available to surface-deposit feeders or filter-
feeders. In this were the case, a bulk sediment analysis would
underestimate the actual exposure. Also, field organisms are
potentially exposed to contaminated phytoplankton and to
pollutants dissolved in the overlying water. If these water
column routes are important, then relating tissue residues to the
field sediment would generate incorrect conclusions regarding
sediment bioavailability.
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With these limitations, field collections are not as well
suited as laboratory experiments for the routine prediction of
the tissue residues resulting from dredge materials and pollutant
discharges, or for between-site comparisons of sediment
bioavailability. Field collections are, however, a powerful
regulatory tool if used in the context of periodic monitoring of
existing sites. In comparing changes at the same stations over
time, problems with the comparison of different species are
reduced, though there may still be problems with collecting
sufficient biomass. Field collections also complement the
laboratory studies as a quality assurance check and by providing
data on commercially important species difficult to maintain in
the laboratory (e.g., lobster). In some cases, both laboratory
and field assessments of tissue residues are justified by the
size of a discharge or dredging operation or by the high
concentration of pollutants. Guidelines on sampling designs for
field surveys can be found in Green (1979), Elliott (1983), and
NOAA (1988), while Holme and Mclntyre (1984) contains information
on the sampling techniques.
B. BIOACCUMULATION FACTORS AND ACCUMULATION FACTORS
Several approaches have been developed to predict benthic
tissue residues directly from sediment concentrations thereby
obviating the need for field collections or bioassays. The
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simplest of these approaches is the Bioaccumulation Factor (BAF)
which is:
BAF = Ct/Cs (1)
Where:
Ct = tissue concentration (ug/g dry wt)
Cs = sediment concentration (ug/g dry wt)
BAFs are empirically derived either from laboratory
bioassays or field collected organisms. Assuming that BAFs were
constant among species and sediments, multiplying the BAF of a
compound times the sediment concentration would predict the
steady-state tissue residue. BAFs are analogous to the
Bioconcentration Factors (BCF) which are used to predict tissue
residues from water concentrations:
BCF = Ct/Cw (2)
Where:
Cw = concentration in water (ug/g)
Although the formulas are analogous, BCFs are often
calculated using wet tissue concentrations.
Sediment characteristics, such as TOG, have a major
influence on the bioavailability of sediment-associated
pollutants and increase the among-site variation in BAFs. BAF
variability is reduced by normalizing the sediment concentrations
to the TOC content (Rubinstein et al., 1983). Normalizing tissue
residues to tissue lipid concentrations reduces variability in
pollutant concentrations among individuals of the same species
and between species (e.g., Veith, 1975; Clayton, et al., 1977).
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These normalizations are combined in a simple thermodynamic-based
bioaccumulation model for pollutant uptake from sediment (Lake et
al., 1987; Rubinstein et al., 1987). The fundamental assumptions
of this thermodynamic model are that the tissue concentration is
controlled by the pollutant's physical partitioning between
sediment carbon and tissue lipids and that the organism and the
environment are at thermodynamic equilibrium. The method assumes
that lipids in different organisms and TOC in different sediments
partition pollutants in similar manners. The key value in the
model is the Accumulation Factor (AF), which when multiplied by
the TOC normalized sediment pollutant concentration predicts the
lipid normalized tissue residue. (Note: some previous studies
such as Lake et al., (1987) and McElroy and Means (1988) reported
Preference Factors which are the inverse of the Accumulation
Factor).
In its simplest form, the model is:
Ct/L = AF*(Cs/TOC) (3)
or
AF = (Ct/L)/(Cs/TOC) (4)
Where:
L = concentration of lipid in organism (g/g dry wt).
(decimal fraction)
TOC = total organic carbon in sediment (g/g dry wt.)
(decimal fraction)
In theory, AFs should not vary with sediment type or among
species. Based on the relationship between Koc and lipid
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normalized BCFs, the maximum AF for neutral organic compounds has
been calculated at about 1.7 (McFarland and Clarke, 1986).
Measured AFs would be lower than this maximum if metabolism of
the compound by the organism is rapid or the organism fails to
reach steady-state body burdens due to limited exposure
durations. Measured AFs could exceed the calculated
thermodynamic maximum if there is active uptake of the pollutant
in the gut or if there is an increase in the pollutant's gut
fugacity, driving the pollutant from the gut into the body. The
pollutant fugacity in the gut could increase as the volume of
food decreases during digestion or as a result of the reduction
in the lipids (Gobas et al., 1988).
Laboratory and field validation of the thermodynamic
partitioning model suggests that for a large number of organic
pollutants, AF values do not exceed the maximum value (Ferraro et
al., 1990). However, AFs for some highly lipophilic PCB
congeners can exceed the theoretical maximum of 1.7 by as much as
an order-of-magnitude (Rubinstein, et al., 1987). Sediments with
the lowest TOCs tend to have the highest AF values (Rubinstein,
et al., 1987; McElroy and Means, 1988; Ferraro et al., 1990; Lake
et al., in review), which is not explained by the present model.
AFs are also dependent upon the accuracy of the lipid
measurement, and total lipids can vary several fold based on the
extraction technique used. As discussed in Chapter XI, we
recommend the Bligh-Dyer lipid method as interim standard for AF
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determinations. If another lipid extraction technique is used, a
conversion factor should be provided to allow the conversion of
the lipid values to chloroform-methanol extraction values.
Although laboratory and field evaluations of the AFs have
shown that they are not statistically constant in all cases
(Rubinstein et al., 1987; McElroy and Means, 1988; Ferraro et
al., 1990; Lake et al., in review), AFs are less variable in
predicting sediment uptake than BAFs (Rubinstein et al., 1987;
Ferraro et al., 1990; Lake et al., in review). Because of their
minimal data requirements, AFs have great potential as a cost-
effective, first-order estimate of tissue residues. The
predicted tissue residues can then be used for determining
whether bioaccumulation tests or field surveys are needed.
For these reasons, the data required to calculate AFs should
be collected and reported in all laboratory tests and field
collections. Development of an AF database would be extremely
useful in determining the limits of applicability of this
approach, as well as generating the values for specific
chemicals. After a minimum database has been collected on a
compound, the AFs could be used in deriving a Sediment Quality
Criterion by taking the upper 95% percentile value.
C. TOXICOKINETIC BIOACCUMULATION MODELS:
Toxicokinetic bioaccumulation models are an alternative to
thermodynamic-based partitioning approaches. Toxicokinetic
models assume pollutant uptake is a function of the feeding
behaviors and physiological characteristics of the organism.
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Most of these toxicokinetic models (e.g., Norstrom et al., 1976)
assume that the tissue residue can be predicted as the sum of the
uptake from each individual phase (e.g. interstitial water,
ingested sediment) minus any loss due to depuration or
metabolism.
In its simplest form, uptake from all phases may be
expressed as:
dCt/dt = (Fx*CPx*EPx) - L (5)
Where:
dCt/dt = change in tissue residue with time.
Fx = flux of phase x through organism.
CPx = concentration of pollutant in phase x.
EPx = fraction of pollutant extracted from phase x by the
organism.
L = summation of loss of pollutant through metabolism
and depuration.
x = phase (W = water, F = food, S = sediment)
As an example, the uptake from water would be the product of
the amount of water ventilated across the gills (FW), the
pollutant concentration in the water (CPW), and the efficiency
with which the pollutant is extracted from the water (EPW). As
opposed to the thermodynamic model, the toxicokinetic model
assumes the uptake from each route is independent and additive,
so an organism exposed to two uptake phases (e.g., interstitial
water and sediment) would have a higher steady-state tissue
residue than an organism exposed to one phase. Toxicokinetic
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models usually assume that uptake efficiency values do not change
as body burdens approach steady-state and that loss (L) can be
modeled as a first-order process.
These models have been used to successfully predict PCS,
methylmercury, and kepone levels in marine and freshwater fish
(Norstrom et al., 1976; Jensen et al., 1982; Thomann and
Connolly, 1984). This approach has only recently been applied to
benthic species, and has been used to model the uptake of
hexachlorobenzene by a marine clam (Boese et al., 1988; Boese et
al., in press; Lee et al., in press; Winsor et al., in press). A
slightly different toxicokinetic model has been used to predict
the uptake of various PAHs by freshwater amphipods (Landrum,
1988, 1989; Landrum and Poore, 1988). Landrum used this model to
determine the relative importance of interstitial water versus
ingested particulates as an uptake route for these PAHs.
In contrast to thermodynamic approaches, toxicokinetic
models can predict tissue residues under non-equilibrium
conditions and can account for differences in organism feeding or
ventilatory behaviors due to toxic or natural effects (e.g.,
growth related changes). The models can also predict the time
course of uptake and depuration, which can be important in
certain regulatory contexts. However, the approach requires
relatively sophisticated laboratory experiments to measure the
input parameters. Because of the extensive data needs and the
ongoing process of developing the laboratory methods, this
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approach is not presently suited for the routine prediction of
bioaccumulation. The toxicokinetic models are appropriate when
detailed analysis of sediment or biological effects on
bioaccumulation are required and as a method to test the
assumptions of various Sediment Quality Criteria approaches.
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APPENDIX 1-2. SAFETY AND WASTE DISPOSAL
A. PERSONNEL SAFETY.
Personnel involved in any facet of bioaccumulation testing,
whether sampling in the field or performing tests in the
laboratory, need to be protected from exposure to toxic
chemicals. Exposure to pathogens must also be considered,
especially when working with sediment collected near sewage
discharges. The manner in which personnel will be protected from
these toxics and pathogens must be determined prior to the start
of any work, keeping in mind that exposure can occur from
breathing vapors, from physical contact with the skin, or
ingestion of the polluted materials and/or chemicals. How one is
protected depends on the type of materials and/or toxics involved
and is beyond the scope of the manual. Consult the following
references to determine adequate safety approaches: Sax (1984),
U.S.EPA (1987b, 1988), ACGIH (1987), and U.S. Coast Guard (1986).
IRIS (Integrated Risk Information System) is available to local,
state, and federal public health officials through the Public
Health Network (PHN) of the Public Health Foundation at (202)
898-5600 or through Dialcom, Inc., at (202) 488-0550.
B. HAZARDOUS WASTE DISPOSAL.
Hazardous waste disposal is a serious problem that must be
dealt with properly. Improper shipping or disposal of toxic
materials may result in environmental damage and/or serious legal
consequences. The Federal Government has published regulations
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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, and about
half of them have, 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 the pertinent regulations
applicable in the state in which you are operating and to comply
with them. Refer to The Bureau of National Affairs, Inc., (1986)
for the citations of the Federal requirements.
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APPENDIX III-l: DETERMINING NUMBER OF REPLICATES
Adequate replication is essential for determining
statistically significant differences between treatments with
sufficient power. If there is a question that the eight
replicates recommended (Chapter III) will not provide sufficient
statistical power, then the techniques in this Appendix can be
used to determine the appropriate number. Determining the
appropriate number of replicates requires estimates of the
variability of each treatment and the minimum detectable
difference. The minimum detectable difference is the smallest
difference between two means, or between a mean and a constant
value, that needs to be statistically distinguishable. The
variability is a measure of the within-treatment variation and is
expressed as a standard deviation (SD or s) or coefficient of
variation (CV) and can be obtained from previous experiments or
the literature. This information is needed because treatments
with high variation will require more replication to distinguish
differences between treatments than less variable ones. See
Table III-l for a listing of coefficient of variations for tissue
residues reported for a variety of pollutants.
The number of replicates required is related to the minimum
detectable difference, and detecting a 2-fold increase in tissue
concentrations requires many more replicates than detecting a
100-fold increase. There are no standards for an acceptable
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minimum detectable difference, but we recommend that there should
be sufficient replication to detect 2-fold to 5-fold differences
in tissue concentrations between two treatments (Chapter III).
Additionally, error rates for Type I and Type II errors must
be chosen. A Type I error (alpha) is the probability of
rejecting the null hypothesis when there is no true difference
between treatment means and is usually given a value of 0.05. A
Type II error (beta) is the probability of accepting the null
hypothesis when there is a true difference between treatment
means. As discussed in Chapter III, we recommend a beta of 0.05.
This is equivalent to a power of 0.95, where the power of a test
is the probability of correctly rejecting the null hypothesis.
One equation that can be used to estimate the number of
replicates (n) required to detect a minimum detectable difference
between two means (adapted from Sokal and Rohlf 1981) is:
n > 2*(s/d)2 * v)2 (1)
For the comparison of one mean and a constant (e.g., FDA Action
Limit) the formula becomes:
n > (s/d)2 * (talphafV + t2beta/v)2 (2)
where:
n = sample size for each treatment
s = standard deviation (often a pooled value of the two
sample variances)
d = the minimum detectable difference
v = the number of degrees of freedom [v = 2*(n-l) for the
comparison of two means, v = (n-1) for the comparison
of a mean and a constant]
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alpha = Experiment-wise or comparison-wise Type I error
(see Chapter XII). If a two-tailed test is performed,
each tail will consist of alpha/2. If a one-tailed
test is performed, the single tail is alpha.
beta = Type II error (or 1 - power of test).
^"alpha v = critical value for alpha of Student's t-
distribution with v degrees of freedom. (Use a
two-tailed t-table for a two-tailed test and a
one-tailed table for a one-tailed test.)
fc2beta v = critical value for 2*beta of Student's t-
distribution with v degrees of freedom. (Use a
two-tailed table. If a one-tailed table is used, the
critical value is beta. The critical value is the
same whether the test is one- or two-tailed.)
An iterative approach is used to calculate n since talpha v
and t2beta v are dependent on n through v. The values for
talpha,v t2beta,v alPha» beta, and v are either set by the
investigator or found in tables. Therefore, only the standard
deviation and the minimum detectable difference must be
estimated. Although a minimum detectable difference (d) of 2 is
recommended (see Chapter II), an estimate of the standard
deviation will not be available in many cases. However, the
ratio of the two (s/d) can be described in several ways,
providing different approaches to estimating these parameters.
Three methods of estimating s/d and their advantages and
disadvantages are:
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Method tti.
(s/d) = [s/(Ul-u2)] (3)
Where:
u-^-Uj = difference between mean U]_ and mean u2,
or mean u-^ and a constant.
Advantages: There may be cases when an absolute difference
between two numbers is of interest such as in a comparison of a
measured tissue residue and a regulatory action limit.
Disadvantages: Requires an estimate of the standard
deviation of the sample, a value often difficult to obtain.
Method #2.
(s/d) = [(CY/lOO/n^] (4)
Where:
CV = Coefficient of Variation (expressed as a percent)
m.^ = a multiplicative factor of u-^ that is the
minimum detectable difference between mean u^ and
mean u2 (or criterion value) (e.g., if m-^ = 5,
the minimum detectable difference between u-^
and u2 will be five times the value of u^).
Advantages: The CV is often easier to estimate than the
standard deviation. The CV's in Table III-l can be used as
estimates if no other information is available, though it would
be prudent to consider these values as the minimum estimates of
variation.
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Disadvantages: The value for m-^ will change whether
comparisons are between control and test values or test and a.
criterion value. Control values (tissue residues) will tend to
be low in comparison to the test values (tissue residues) while
test values may be large and close to a criterion value (e.g.,
FDA action limits).
Method #3.
(s/d) = [s/(m2*s)] = [l/m2] (5)
Where:
m2 = a multiplicative factor of s. For example, if
m2= 2, the minimum detectable difference is 2
standard deviations (i.e, u2 will have to be 2
standard deviations from u1 to be able to detect
a difference).
Advantages: No estimates are required of the standard
deviation or CV.
Disadvantages: The value of m2 may have to vary whether
comparisons are between control and test values or test and
action limits.
If a comparison between more than two means is anticipated
(as in the determination of steady-state conditions), see Sokal
and Rohlf (1981) for a modification of this approach or Tetra
Tech (1986a) for tables of estimates.
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APPENDIX IV-1: ADEQUACY OF 10-DAY AND 28-DAY EXPOSURES
Ideally, organisms should be exposed to test sediments for a
period sufficient to attain steady-state tissue residues.
However, cost considerations often prove prohibitive to
conducting tests long enough to document that steady-state has
been attained. As a result, bioaccumulation tests historically
have been conducted for a preset duration. Choosing a single
time period is complicated by the multitude of organic pollutants
and metals found in most field sediments or dredge materials,
each with differing uptake kinetics. To date, a ten-day exposure
to assess "bioaccumulation potential" has been the most commonly
used time period for the testing of marine sediments (primarily
dredge materials) (U.S. EPA/U.S. ACE, 1977). Bioaccumulation
potential is the potential for any uptake of a pollutant by
organisms exposed to a sediment, and the basic premise was that
if there was going to be bioaccumulation it should be possible to
detect it within 10 days. Thus, the original intent of the 10-
day test was as qualitative rather than a quantitative measure.
Since 1977, however, data from 10-day tests have frequently been
extended beyond its original intent and used as a quantitative
result.
Because of the wide-spread use of the 10-day exposures, it
is worth assessing its utility both as qualitative measure of
bioaccumulation potential and as a quantitative method to
generate data for ecological and human health risk assessments.
The percent of steady-state tissue residue obtained after 10 days
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for several organic pollutants was used as a simple measure of
accuracy (Table IV-l). To adequately assess bioaccumulation
potential, the 10-day exposure should result in a sufficient
percentage of the steady-state tissue residues to identify which
sediments could be an environmental problem. Also, the
percentage of the steady-state tissue residue obtained should be
relatively consistent for the same pollutant in different
species. That is, the 10-day exposure should give a strong and
consistent "signal". In the quantitative risk assessments, the
benthic tissue residues will be used to predict the amount of
pollutants transported from the sediment to higher trophic
levels, including man. A large error at the base of the food-web
will result in errors throughout the analysis, especially as some
of the errors may be multiplicative. As a preliminary measure,
we suggest that for data to be acceptable for quantitative risk
assessment, the resulting tissue residues should be within 80% of
the steady-state tissue concentrations. An accuracy of 80% for
each trophic step results in the prediction of tissue residues
being within two-fold of the actual residues for a three step
chain (i.e., sediment to benthos to demersal predator to higher
predator or man; or 0.8*0.8*0.8 = 0.51)
In these studies, only 29% of the organisms approached
within 80% of the steady-state level in ten days (Table IV-1).
Ten-day tissue residues averaged 51% of the estimated steady-
state value, and this average included some rapidly accumulated
PAHs. Tissue residues of PCBs achieved after 10 days averaged
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only about 25% of the steady-state values, and ranged from 100%
to a low of 9%. Other environmentally important compounds with
high Kow compounds, such as DDT, dioxins, and BaP, are expected
to be similar to the PCBs.
Ten days is also likely to generate a relatively low
percentage of the steady-state tissue residues for metals. For
example, mercury levels in fish may not attain steady-state
during the lifetime of the organism (River et al., 1972; Cross et
al., 1973), and the minimum time for lead to attain steady-state
in Mytilus edulis was greater than 230 days (Schulz-Baldes,
1974).
Based on this preliminary review, we reach several
conclusions. First, a 10-day exposure generates a low percentage
of the steady-state tissue residues for PCBs and presumably other
high Kow organics and some heavy metals. These compounds are the
most likely to represent an ecological and human health risk
through bioaccumulation and biomagnification. Second, the
percentage of the steady-state tissue residue obtained varies
several-fold even within a single compound. Third, the amount
accumulated within ten days is such a small percentage of the
steady-state concentration that it may be below detection limits
of standard analytical methods or may not be significantly
different than control values. Thus, the 10-day exposure can
result in false negatives concerning the bioaccumulation
potential of a sediment. Fourth, the percentage of the steady-
state tissue residues accumulated over 10 days is inadequate for
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TABLE IV-1A: Information Gained and Requirements of Different
Approaches to Estimating Benthic Tissue Residues
METHOD
BIOACCUM.
POTENTIAL
FALSE NEG.
BIOACCUM.
POTENTIAL
ESTIMATES
EQUILIBRIUM
RESIDUE
ADDITIONAL
REQUIREMENTS
Accumulation Yes
Factors
10-Day Test Yes
28-Day Test Yes
Kinetic Models Yes
No
Yes
No
No
Yes?
No
Approx.
to Yes
Yes
Long-Term
Exposures
Yes
No
Yes
Sed Cone.,
TOG, Lipids
10 Days Lab
Time, Tissue
Cone.
18 Days Addi-
tional Lab Time
Additional
Tissue Cone.,
Additional Lab
Time?, Develop-
ment of Techni-
ques
23-70 Days
Additional Lab
Time, Additional
Tissue Cone.
Bioaccum. Potential = Qualitative ability to detect uptake.
False Negative Bioaccum. Potential = Amount accumulated is so low that
it is incorrectly concluded that no uptake will occur.
Estimates Equilibrium Residue = Tissue residue data sufficiently
accurate for use in quantitative risk assessments.
Exper. Techniques = Resources devoted to determining the correct uptake
and depuration periods for specific compounds and organisms
Lab Time = Laboratory time required for biological exposure
Lipids = Tissue samples analyzed for lipid content
Sed. Cone. = Sediment samples analyzed for pollutants
Tissue Cone. = Tissue samples analyzed for pollutants
TOC = Sediment samples analyzed for TOC
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a quantitative risk assessment. Lastly, the 10-day exposure does
not generate any additional insights into bioaccumulation
potential that are not generated by use of the Accumulation
Factors (see summary in Table IV-1A)
For these reasons, we conclude that an exposure duration
longer than 10 days is required. Based on the use of 28 day
exposures in the bioconcentration tests (ASTM, 1984), we
recommend a 28-day exposure as a practical compromise between
cost, data accuracy, and data utility. When 28-day organic
pollutant levels were compared to observed or estimated steady-
t
state levels (Table IV-l), steady-state tissue residues were
approached (i.e.^80% of steady- state) in 76% of the tests, and
the mean steady-state pollutant tissue level increased to 86% of
the steady-state maximum. An average of 82% of the PCB steady-
state tissue residues was obtained after 28 days, though in one
instance the value was only 25% of the steady-state residue.
This level of accuracy should be sufficient in nearly all cases
to test for bioaccumulation potential with a reasonable level of
statistical certainty. In most cases, the data should be
sufficiently accurate for quantitative risk analysis. In cases
when more accurate estimates are required, either a long-term
exposure or a kinetic approach can be used (Chapter IV).
Besides underestimating tissue residues because of
insufficient duration, single point tests can underestimate
maximum tissue residues when a compound reaches a maximum value
before the sampling period and then declines. For example,
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phenanthrene approaches its maximum tissue residue in freshwater
amphipods after about 10 days and then declines (Landrum, 1989).
In this case, a 28-day test would generate a lower value than a
10-day test. Presumably, the decline is the result of an
increase in the metabolic degradation rate of the pollutant, and
should be most common with the lower molecular weight PAHs.
Because the ability to degrade PAHs varies among taxa (Varanasi
et al., 1985), a decline in tissue residues should be most
pronounced with amphipods and less so with bivalves. If low
molecular weight PAHs or other rapidly metabolized compounds are
of interest, time series samples should be taken before day 28
(see Chapter IV).
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APPENDIX IV-2: ALTERNATIVE TEST DESIGNS
A. SHORT-TERM TEST
Some compounds (e.g., volatiles) may attain steady-state in
less than 28 days (see Table IV-1), so that a 28-day exposure may
not be necessary. Generally, 10-day tests should be acceptable
with organic compounds which have log Row's <3 that have been
spiked into sediments. Even with these compounds, a 10-day test
should only be used after it has been documented to approach
steady-state in phylogenetically similar species in less than ten
days, or that the depuration rate (k2) in phylogenetically
similar species is >0.5/day. When determining the
bioaccumulation of pollutants from field sediments, however, a
28-day test should be used because nearly all field sediments
contain some pollutants with slow uptake kinetics. Biotic and
abiotic samples should be taken at day 0 and day 10 following the
same protocol as used for the 28-day tests. If time-series
biotic samples are desired, sample on days 0, l, 3, 5, 7, 10.
B. ESTIMATING STEADY-STATE FROM UPTAKE RATES
In theory, it is possible to estimate both kl and k2 from
the uptake phase alone if the experiment continues past the point
when the tissue residues begin to "bend over", indicating that
depuration is sufficient to slow net uptake (Figure IV-1). This
approach obviates the need to run a separate depuration
experiment, as is required in kinetic approach discussed in
Section C of Chapter IV. However, since both kl and k2 are
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estimated from the fitting of mathematical models, this method is
less reliable than the kinetic approach which uses independent
measures of kl and k2. Nonetheless, this approach has utility
when time or analytical support is limited, or if a long-term,
time-series uptake test is terminated before steady-state is
attained. In this design, the sampling schedule should follow
closely that of the uptake phase of the kinetic approach using
both uptake and depuration rates (Chapter V). Refer to Branson
et al. (1975) and Foster et al. (1987) for the specifics of
estimating kl and k2.
C. GROWTH DILUTION
If test organisms grow during an experiment, growth
dilution, the dilution of pollutant concentrations in the tissues
by the increase in tissue mass, will occur. Taking an extreme
example, if an organism doubled its weight during a depuration
study, it would appear that half of pollutants had been depurated
even if none of the pollutants were excreted from the organism.
Without correction for growth, the depuration rate (k2)
calculated from this experiment would be incorrect for an
organism growing at a different rate. Many experiments have not
taken growth dilution into account, which may contribute to the
variation among measured depuration rates (see Niimi et al.,
1981).
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In 28-day experiments, growth dilution is not a problem if
growth is relatively slow and kinetic rate constants are not
derived from the data. However, for the kinetic approach, growth
dilution can cause errors in estimating uptake and depuration
parameters, resulting in errors in predicting steady-state
concentrations and time to steady-state.
If substantial growth occurs during experiments to determine
the rate constants, uptake rate constants will be underestimated
and depuration rate constants will be overestimated. If these
erroneous constants are used in the kinetic model (Equation l of
Chapter IV) under conditions of no growth, both steady-state
tissue concentrations and time to steady-state will be
underestimated. Conversely, an error occurs when correct (i.e.,
derived under no growth) uptake and depuration rate constants are
used in this kinetic model when the organisms are growing. In
this case, both the steady-state concentrations and time to
steady-state will be overestimated because the model does not
compensate for growth dilution.
If possible, experiments should be conducted with organisms
that grow very slowly or under environmental conditions that keep
growth at a minimum (such as low temperatures). If growth can
not be avoided, then growth dilution must be taken into
consideration if a kinetic approach is used. Assuming that
growth dilution is a first-order process and that growth occurs
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at a constant rate, the kinetic model (Equation 1 of Chapter IV)
becomes:
Ct(t) = kl*Cs/(k2+k3)*[l-e"(k2+k3)*t] (1)
where:
Ct = concentration in the organism at time t
Cs = concentration in the sediment
kl = the uptake rate constant [days"1]
k2 = the depuration rate constant [days' ]
k3 = the growth rate constant [days'1]
t = time [days]
The growth rate constant (k3) can be measured from the
change in weight during the exposure experiment or during a
separate growth experiment under similar environmental
conditions. Equation l assumes that the kl and k2 values are
true uptake and depuration constants measured under conditions of
no growth or, if growth occurs, then growth dilution was taken
into account. If the depuration rate is measured while organisms
are growing, the rate measured will actually be a function of
growth and depuration and can be modeled as k2+k3.
Under conditions of growth, and using an estimated growth
constant (k3), the maximum tissue residues becomes
ctmax = kl*Cs/(k2+k3) (2)
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APPENDIX IV-3: CALCULATION OF TIME TO STEADY-STATE
Having an estimate of the time to reach steady-state tissue
residues is very helpful in designing long-term studies and
assessing the adequacy of a 28-day test. If no estimate for a
pollutant in phylogenetically similar organisms is available, the
time required to approach steady-state can be estimated from a
linear uptake, first-order depuration model (see Chapter IV,
Section C). This model is an approximation for benthic
invertebrates as it was developed for fish exposed to dissolved
organic contaminants.
Uptake of organic pollutants from water (dissolved phase)
has been modeled in fish species using a linear uptake, first-
order depuration model (Spacie and Hamelink, 1982):
Ct(t) = kl*Cw/k2*(l-e"k2*t) (1)
Where:
Ct = pollutant concentration in tissue at time t
Cw = dissolved pollutant concentration in water.
kl = uptake rate constant. [days'1]
k2 = depuration rate constant. [days'1]
t = time [days].
This model predicts that equilibrium would be reached only
as time becomes infinite. Therefore, for practical reasons,
apparent steady-state is defined here as 95% of the equilibrium
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tissue residue. The time to reach steady-state can be estimated
by:
S = In[l/(1.00-0.95)]/k2 = 3.0/k2 (2)
Where:
S = time to apparent steady-state (days)
Thus, the key information is the depuration rate of the
compound of interest in the test species or phylogenetically
related species. Unfortunately, little of this data has been
generated for benthic invertebrates. When no depuration rates
are available, the depuration rate constant for organic compounds
can then be estimated from the relationship between Row and k2
for fish species (Spacie and Hamelink, 1982):
k2 = antilog[1.47-0.414*log(Row)]. (3)
The relationship between S and k2 (using Equation 2) and
between k2 and Row (using Equation 3) is summarized in Table IV-
3A. This table may be used to make a rough estimate of the
exposure time to reach steady-state tissue residues if a
depuration rate constant for the compound of interest from a
phylogenetically similar species is available. If no depuration
rate is available, then the table may be used for estimating the
S of organic compounds from the Row value. However, as this data
was developed from fish bioconcentration data, its applicability
to the kinetics of uptake from sediment-associated pollutants is
unknown and the estimated S values should be considered as
minimum time periods. Also, Equation 2 does not account for
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Table IV-3A: Estimated Time to Obtain 95% of
Steady-State Tissue Residue
Estimated time (days) to reach 95% of pollutant steady-state
tissue residue (S) and depuration rate constants (k2) calculated
from octanol-water partition coefficients using a linear uptake,
first-order depuration model (Spacie and Hamelink, 1982). k2
values are the amount depurated (decimal fraction of tissue
residue lost per day). Note that the calculated k2 values for
log Row values <3 are expressed in hours and are based on
extrapolation from the relationship between k2 and Kow, as
described by Spacie and Hamelink (1982). As a result, caution
should be excercised in using these particular k2 values.
Log Kow k2 S (days)
1 °-48! °-3
2 0.17 0.7
3 0.07 1.8
4 0.65 4.6
5 0.25 12
6 0.097 31
7 0.037 80
8 0.014 208
9 0.006 540
* k2 values expressed in hours, all other k2 values
expressed in days.
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growth dilution (Appendix IV-2). To correct for growth dilution,
Equation 3 becomes
S = In[l/(1.00-0.95)]/(k2+k3) = 3.0/(k2+k3) (4)
Where:
k3 = growth rate constant [days"1]
Using a linear uptake, first-order depuration model to
estimate exposure time to reach steady-state body burden for
metals is problematical for a number of reasons. The kinetics of
uptake may be dependent upon a small fraction of the total
sediment metal load that is bioavailable (Luoma and Bryan, 1982).
Depuration rates may be more difficult to determine, as metals
bound to proteins may have very low exchange rates (Bryan, 1976).
High exposure concentrations of some metals can lead to the
induction of metal binding proteins, like metallothionein, which
detoxify metals. These metal-protein complexes within the
organism have extremely low exchange rates with the environment
(Bryan, 1976). Thus the induction of metal binding proteins may
result in decreased depuration rate constants in organisms
exposed to the most polluted sediments. Additionally, structure-
activity relationships that exist for organic pollutants (e.g.,
relationship between Row and BCFs) are not well developed for
metals.
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APPENDIX V-l: TECHNIQUES FOR SEDIMENT MANIPULATION
In this appendix we summarize techniques to experimentally
manipulate sediment for bioaccumulation tests. This is an area
of much interest, and these guidelines may need revision in the
future based on the ongoing research.
A. SEDIMENT SPIKING
1. Methods Used To Add Pollutants To Sediment
A variety of methods have been used to dose or "spike"
sediments with pollutants. Toxicants can be added to overlying
water and allowed to partition with the sediment (Breteler and
Saksa, 1985; Pritchard et al., 1986), added to dry sediment and
mixed by stirring or agitation (Adams et al., 1985; Foster et
al., 1987), added to wet sediment and mixed by stirring or
agitation (Stein et al., 1987), added with a solvent carrier, or
evaporated on the sides of jars and the sediment mixed by rolling
in the jars (Swartz et al., 1986; McElroy and Means 1988; Boese
et al., in press). Sediments are also spiked by suspending them
in aqueous solutions of the pollutants that contain carrier
solvents (McLeese et al., 1980). Alternatively, carrier-free
aqueous solutions of pollutants can be prepared using generator
columns (Veith et al., 1975), which then can be used to spike the
suspended sediment.
Solvents are often used as carriers to add hydrophobic
pollutants to water and sediment. However, carriers can alter
pollutant bioavailability (Dalela et al., 1979; Hughes et al.,
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1983) as well as pollutant partitioning (Nkedl-Kizza et al.,
1985) . Although the exact effects of solvents are currently
being debated, there is sufficient evidence to recommend that
carrier solvents not be added directly to the sediment whenever
possible. If a carrier solvent must be used, the amount should
be minimized.
The effects of these various techniques on bioaccumulation
have not been tested, and recent work indicates that DOC may
increase proportionally with the magnitude of disturbance
(mixing), continue to increase after the disturbance, and fail to
return to previous DOC concentrations after 10 weeks (DeWitt, T.,
pers. comm., OSU, Mar. Sci. Ctr., Newport, OR). However, tests
have also shown that bioaccumulation of a PCB was not affected by
a long rolling time (McElroy and Means, 1988). Until standard
methods are developed, it is recommended that appropriate caution
be exercised in comparing spiked and field results and when
comparing results from sediments spiked by different techniques.
2. Spiking Methodology
The following is a summary of the method we have used to
spike two sediment types, a fine-grained sand and a silt, with
hydrophobic pollutants. These techniques should work well with
similar sediments, but those with drastically different
properties, such as cohesive muds, may require modification. The
advantages of this method are that it avoids the addition of
solvents to the sediments and assures good physical mixing.
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Dissolve the pollutant with an appropriate solvent and place
an aliquot of the mixture into glass jars. Roll jars in a fume
hood while evaporating the solvent with a gentle stream of
nitrogen or purified air. After the solvent has evaporated, add
wet sediment with a sufficient water content to allow adequate
movement of the sediments. We have spiked 2 kg of wet sandy
sediment in 1 gallon jars, and up to 13 kg of wet sandy sediment
in 3 gallon jars. Leave adequate space in the jar for the
sediment to roll. Cap jars and roll the sediment slurry
continuously at approximately 12 rpm. The rolling apparatus can
either be home-built or a commercial ball mill. Place a catch
basin under the rolling apparatus to contain any contaminated
sediment in case of leakage or breakage. The sediment can be
rolled at room temperature or 4°C, but the cooler temperature
will slow microbial degradation of carbon and organic pollutants.
At the end of the rolling period, place the jar in a fume hood
and stir the sediment with a large TeflonR-coated stainless steel
or polypropylene spoon.
Appropriate solids to water ratios are essential when
spiking sediment in order to assure pollutant homogeneity. Sandy
sediments with water content of about 30% and fine grained
sediments with water contents of 60% have been mixed
successfully. However, sediments higher in organic carbon or
clays may require higher water contents for successful mixing.
Low water content is indicated by "balling up" of the sediment
and/or inadequate movement of the sediment in the rolling jar.
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The sediment needs to slide on the sides of the jar, if only
slightly, while it is being rolled. If additional water is
added, the excess can be removed by pipetting or siphoning after
the sediment has settled. Overnight settling is usually
sufficient, but longer periods may be required for particularly
fine sediments. Sediment can also be centrifuged to remove the
overlying water, though this may result in a sediment with too
low a water content. If that occurs, water may be added back to
the solids by stirring the sediment or sieving it through a 1 mm
(or appropriate size) screen with a portion of the decanted
water.
The mixing times required to assure a homogeneous dispersion
of the spiked pollutant may vary, but sediments spiked with
hydrophobic pollutants (e.g., HCB, PCBs) are physically
homogeneous within one to three days. Although the pollutant
concentration is homogeneous in a few days, we normally roll the
sediments for ten days to allow the time for the pollutant to
partition among the various sediment phases. At the end of ten
days, at least 3-8 replicate samples of the spiked sediment
should be analyzed to confirm pollutant homogeneity. If the
sample is not homogeneous, the mixing should continue until
homogeneity is achieved.
Sediment can be spiked with a water soluble (hydrophilic)
compound by first dissolving the pollutant in water of
appropriate salinity and then following the previously described
steps of adding and rolling the sediment slurry. This procedure
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differs only in that the water (solvent) is not evaporated prior
to adding sediment. Hydrophilic pollutants may not require a 10
day rolling period to achieve partitioning among the water and
the solid phase.
B. INCREASING SEDIMENT TOC.
The organic content of a sediment can be increased to
determine effects of organic enrichment on the benthos or effects
of organic matter on pollutant bioavailability. The organic
content can be increased by adding sewage sludge, manure, humics,
natural detritus or other organic-rich materials. Such
experiments can generate important insights, but the
investigator must be aware of several potential confounding
factors. First, though sediments may be equal in TOC
concentrations, not all organic matter reacts in the same manner.
A highly labile organic source (e.g., dried baby food) is more
likely to drive a sediment to anoxia than a refractory carbon
(e.g., natural detritus). Second, addition of organic matter
will probably change physicochemical properties such as grain
size, Eh, sorption capacity, and texture of the sediment.
Lastly, the effect of the organic matter addition on
pollutant bioavailability can vary with the feeding behavior of
the test species (Lee et al., in press). Figure V-1A illustrates
the effect selective feeding can have on ingested pollutant dose
in natural sediment and in organically enriched sediment. In
this case, the organism selects particles similar in size and
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Legend to Figure V-1A: Effect of Sediment Selection on Ingested
Pollutant Dose in Natural and Organically Enriched Sediments:
Figure V-1A illustrates a surface deposit-feeding clam, Macoma
nasuta. in a clambox (see Appendix IX-1). The clambox separates
the inhalant (left side) and exhalant (right side) siphons,
allowing the collection of fecal pellets for a determination of
the size and pollutant concentration of ingested particles. In
the upper illustration, the clam is exposed to a natural sandy
sediment, but selectively ingests the finer, high TOC particles.
In the lower illustration, the clam is exposed to a sediment
enriched with finer, high TOC particles. The clam ingests fine
particles with a similar size and TOC content as in the sandy
sediment. Because the clam ingests particles with a similar TOC
content, and hence concentration of organic pollutants, the
clam's ingested pollutant dose is similar in the two sediments
even though the bulk sediment pollutant concentrations are
dissimilar.
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FIGURE V-1A
Effect of Sediment Selection on Ingested Dose
in Natural and Organically Enriched Sediments
Inhalant
siphon
« «
Exhalant
r siphon
t CLAMBOX
Natural sediment
Fecal
" pellets
Dental dam
Organically Enriched Sediment
SJLND GRAINS, LOW TOG, LOW POLLUTANT CONC.
ORGANIC PARTICLE, HIGH TOG, HIGH POLLUTANT CONC.
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organic content in the two sediment, resulting a smaller
difference in the ingested pollutant dose than would be predicted
from bulk sediment analysis.
Probably the most "natural" way of augmenting sediment TOC
is either by enriching a sediment with the fine fraction
collected from the same sediment type (Swartz et al., 1985a) or
by using the fine fraction directly (DeWitt et al., 1988; Boese
et al., in press). These methods use naturally occurring organic
matter, though grain size distributions of the test sediment is
substantially altered. The fine fraction can be obtained by wet
sieving the sediment through a series of sieves. After sieving,
the fine fraction can be dewatered by allowing the solids to
settle for 16-24 hours before siphoning off excess water. Excess
water can also be removed by centrifugation and decanting.
Similarly, other high organic materials such as sewage sludge
solids can be dewatered by centrifugation and then added to the
test sediment (Swartz et al., 1984).
C. DECREASING SEDIMENT POLLUTANT AND/OR TOC CONCENTRATIONS
Bulk sediment pollutant concentration and organic content
can be decreased by adding control sediment, clean sand, ashed
portions of the same sediment, and other inert inorganic
materials, or by chemical extraction techniques. Such a
manipulation can be used to reduce the toxicity of a field
sediment (Swartz et al., 1989) so that its pollutant
bioavailability or LC50 can be determined. These techniques,
however, can unexpectedly change the pollutant sorption capacity
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and the bioavailability of both metals and organics. Also,
depending on the feeding behavior of the test species, addition
of particles which are not ingested may have little affect on the
ingested pollutant dose (see Fig. V-lA).
The least disruptive method of diluting sediments is not
clear. Using a control sediment with a grain size similar to
that of the test sediment will help maintain the physical
characteristics of the sediment. However, by adding many
partitioning sites, the addition of a control sediment may
totally disrupt the distribution of the contaminant. Instead,
the addition of clean sand, with few binding sites, may be a
better alternative (Landrum, P., pers. comm., NOAA, Great Lakes
Enviorn. Res. Lab., Ann Arbor, MI). The advantages and
disadvantages of the various approaches requires further
investigation.
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APPENDIX VII-1: SELECTION CRITERIA FOR TEST SPECIES
A. INDIGENOUS VERSUS SURROGATE SPECIES
The first decision in choosing test species is whether to
use a representative indigenous species or a surrogate species.
The supposed advantage of indigenous organisms is that they are
the same species which will be impacted by the dredge material or
discharge. However, benthic communities can undergo drastic
fluctuations in species composition in response to natural (e.g.,
Frankenberg and Leiper, 1977) and pollution events (Pearson and
Rosenberg, 1978) and during recolonization (Rhoads et al., 1978).
Because of this variation, the indigenous species chosen for
laboratory testing may not be closely related phylogentically or
ecologically to the species at the impacted site.
Many of the common indigenous species do not meet the
criteria for use as a bioaccumulation test species, negating any
advantage of using a native species. Even when an indigenous
species is acceptable, established surrogate test species offer
several advantages. There is considerable information on the
maintenance and biology of the recommended test species.
Additionally, as more tests are conducted on these species, a
database will develop allowing the comparison of bioaccumulation
under different environmental conditions. As ASTM (1984) has
pointed out, it is more advantageous to gather detailed
information on a few species rather than a smattering of
information on a large number of species.
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Our recommendation is to use surrogate species for routine
monitoring of sediments and discharges. If there are local
species which appear to meet the various criteria discussed
below, they can be tested along with the recommended
bioaccumulation species. If the local species prove acceptable
and the results intercalibrate with the results from the standard
species, the local species could be substituted for the standard
species in future tests. Local species that do not meet the
criteria but are of special concern (e.g., lobster) can be tested
in addition to the surrogate species but should not be
substituted for them.
B. REQUIRED CRITERIA:
As ingested sediment can be a major uptake route for higher
Kow compounds (Landrum, 1989; Boese et al., in press), test
species must ingest sediment. Using a filter-feeder, in which
the only route for bedded-sediment exposure is from interstitial
water, may underestimate the total uptake from sediments. Many
benthic invertebrates are flexible in their feeding mode, and
this requirement does not preclude the use of facultative filter-
feeders (e.g., Macoma spp) as long as the only route of uptake
during the exposure is from bedded sediment (i.e, no resuspended
particles) and as long as the bedded sediment supplies adequate
nutrition.
The requirement for a sediment ingesting species excludes
obligate filter-feeders, such as Mercenaria. Mya. oysters (e.g.,
Crassostrea) and mussels (Mytilus spp), as well as obligate
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predators such as Glycera spp. If there is concern about the
human health consequences of the tissue residues in edible
filter-feeding bivalves, they should be tested in addition to but
not substituted for the standard test species. To accurately
predict the tissue residues in filter-feeders, a resuspension
exposure system is required (see Appendix IX-1).
Test species must be sufficiently pollutant resistant to
survive the duration of the exposure with a minimum level of
mortality. For bioconcentration tests, ASTM (1984) states that a
test is unacceptable if more than 10% of the organisms "died or
showed signs of disease, stress, or other adverse effects." This
requirement is based on deriving BCFs for single compounds in
which it is possible to control the pollutant concentration.
This requirement appears too strict for testing of environmental
sediments in which it is difficult to meaningfully manipulate
toxicity. Nonetheless, if a pollution-sensitive species is
sufficiently stressed to inhibit normal feeding, the resulting
tissue residues may underestimate the amount bioaccumulated by a
more hardy species. Additionally, excessive mortality can create
problems in the statistical analysis of the data.
Environmentally collected sediments display a wide range of
toxicities. If sediment is extremely toxic, then it may be
necessary to use a highly pollution-tolerant species (e.g.
Capitella spp.). As a rule, however, these pollution-tolerant
species are small. Most field collected sediments will be
moderately toxicity, allowing the use of the moderately
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pollution-tolerant species though not the sensitive species
commonly used in sediment toxicity tests (e.g., phoxocephalid
amphipods). A general indication of the relative hardiness of
various species can be obtained from the lists of opportunistic
species listed in Pearson and Rosenberg (1978), the species used
to calculate the "Infaunal Trophic Index" (Word, 1978), as well
as from multispecies bioassays (Swartz et al., 1979)
C. DESIRABLE CHARACTERISTICS:
Besides the required criteria, there are a number of
desirable characteristics which would make conducting the tests
easier, interpreting the results more straightforward, or allow
the results to be applied to a wider range of habitats.
On important characteristic, especially if repeated tests
are planned, is the ease of obtaining the test species in
sufficient numbers at the correct season. The ease of collecting
specimens is related to a species' abundance, habitat (intertidal
vs subtidal vs offshore), robustness to collection techniques,
depth in the sediment, and seasonality. It is important not to
underestimate the time required to collect sufficient numbers of
healthy individuals. In general, it is prudent to collect twice
the number required, especially polychaetes which are prone to
breakage. Information on collecting and transporting specimens
is given in Chapter VIII. As an alternative, test organisms may
be purchased from biological supply houses or local collectors
(See Appendix VIII). Local bait suppliers may sell species such
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as Nereis and Callianassa.
If a large number of bioaccumulation tests will be conducted
over an extended time period, culturing of test organisms may be
cost effective. Culturing will provide a ready supply of
organisms of known history, but maintaining a culture to supply
sufficient biomass for bioaccumulation tests will be time
consuming. A few sediment ingesting polychaetes (e.g., Capitella
capitata and Neanthes arenaceodentata) can be cultured with
relatively simple equipment (Reish and Richards, 1966; Reish,
1974, 1985; Dean and Mazurkiewicz, 1975), as can Palaemonetes
(Tyler-Schroeder, 1976a,b). Although these organisms are
generally suitable test species, most of the species are small,
making it difficult to obtain sufficient biomass. Culture of
bivalves, larger polychaetes, and most crustaceans is impractical
except for experimental studies.
Regardless of how test species are obtained, they should be
amenable to laboratory conditions and not require elaborate
holding facilities. Fortunately, most pollutant-resistant
species are relatively hardy and adaptable to laboratory
conditions. Most of the bioaccumulation test species listed in
Table VII-1 are reasonably easy to maintain and do not require
flowing seawater.
Whether field-collected or laboratory-cultured specimens are
used, gravid individuals or individuals which are likely to
become gravid during a test should be avoided if possible. The
reduction in tissue lipids that often occurs with spawning
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(Gabbot, 1976: Davis and Wilson, 1983) can result in a
corresponding reduction in the associated pollutants. Spawning
may also result in unacceptable mortalities. Certain species,
such as Macoma nasuta in Oregon, have a reasonably well defined
spawning cycle and size at reproductive maturity, making it
possible to minimize the collection of reproductive individuals.
Other species, such an Neanthes virens. change appearance when
reproductively mature. In extended tests, it may be impossible
to completely avoid gravid individuals, though the occurrence of
the reproductive state should be noted.
A very important characteristic is organism size. Test
species need to be small enough to be easily maintained, yet
large enough to supply sufficient biomass for chemical analysis.
The amount of biomass required depends upon the analytical
procedures used as well as the types of analyses required (e.g.,
metals, organics, lipids). At least 1 gram of wet tissue is
required in nearly all cases, and commonly up to 5 grams tissue
will be required. Ideally, the species should be large enough to
allow chemical analysis on individuals. Chemical analyses on
individuals are possible using the more "sophisticated"
analytical procedures with the larger benthic organisms, such as
Macoma nasuta and lug worms. Depending on the techniques, it may
be impossible to conduct both metals and organic analyses on an
individual, even when using large species, thereby necessitating
twice as many exposure chambers if both types of pollutants are
required.
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An alternative approach to obtaining sufficient biomass is
to composite individuals (see Chapter III). Even when
compositing individuals, the size of the individual is an
important consideration. It is simpler to handle and count a few
larger individuals (e.g., Nereis) than dozens or even hundreds of
smaller specimens (e.g., Capitella).
Species suitable for measures of sublethal stress or
experimental manipulations offer the advantage that toxicokinetic
or toxicological data can be collected concurrently with the
bioaccumulation data. Growth is the simplest measure of
sublethal stress. Measuring changes in wet weight is possible
with both polychaetes and bivalves, but wet weight measurements
are prone to error. Johns et al. (1989) gives techniques for
conducting a growth bioassay with Neanthes arenaceodentata. some
of which could be adapted to measurement of growth during
bioaccumulation tests. With bivalves, growth can also be
measured as changes in shell length. Although shell length has
the limitation of only showing positive growth, the comparison of
shell growth in treatments to the controls is a simple sublethal
measure. Scope-for-growth, a measure of the amount of energy
available for growth, has been used frequently as a measure of
stress in mussels (Bayne et al., 1981). Scope-for-growth
measurements have been conducted on infaunal polychaetes (Johns,
et al., 1985) and presumably could be adapted to other bivalves
and larger crustaceans, though the techniques are not simple.
The arenicolid worms and the bivalves, particularly Macoma
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nasuta. lend themselves to exposure chambers in which it is
possible to collect ventilated water and processed sediment
(i.e., feces) for bioenergetic measurements and for
determinations of the pollutant uptake rates (e.g., Pelletier et
al., 1988a,b; Specht and Lee, 1989). Construction of these
exposure chambers is discussed in Appendix IX-1.
The more tolerant a species is to sediment, temperature, and
salinity variations, the more types of sediments in which it can
be used. Using a few widely adaptable species allows a direct
comparison of sediment bioavailability from a variety of
environments or biogeographic regions. Also, collecting and
maintaining a few widely adaptable species is simpler than
developing techniques for a larger number of less adaptable
species. The approximate salinity and temperate ranges of
potential bioaccumulation species are given in Table VII-1.
These ranges are estimates of the ranges in which the organisms
could be used in bioaccumulation test and are not the
physiological limits. For many of the species, the ranges are
based on the general literature and discussions with other
researchers rather than extensive experimentation. A preliminary
survival test would be advisable before initiating a large
bioaccumulation test using species near the limits of the ranges
given in Table VII-1.
Because the goal of bioaccumulation tests is to estimate the
maximum likely tissue residues, it is important to chose species
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which have a high bioaccumulation potential. Unfortunately,
insufficient numbers of multi-species tests have been conducted
to adequately compare the bioaccumulation potential of a range of
species over a range of compounds. In general, tissue residues
will be higher in species with a higher lipid contents, which can
vary as much as 10-fold among species (e.g., Rubinstein et al.,
1987). If PAHs are of concern, at least one test species should
be a bivalve as they have a lower ability to metabolize PAHs than
either polychaetes or crustaceans (Varanasi et al., 1985).
Infaunal species are preferable over epibenthic deposit-
feeders because the latter are only intermittently exposed to
interstitial water. Interstitial water is thought to be the
major uptake route for compounds with a Kow less than about 5
(Adams, 1987) and possibly for metals as well (Assanullah et al.,
1984), so the potential uptake of these compounds could be
underestimated. This criterion limits the use of Palaemonetes.
the only well established crustacean on the list of
bioaccumulation species (Table VII-1). Infaunal crustaceans
(e.g., Callianassa) have not been used extensively as
bioaccumulation species, and appear to be more difficult to
maintain in the laboratory than Palaemonetes.
Compatibility with other species or with the same species is
important if multiple species or multiple individuals of the same
species are exposed in the same chamber. Several of the nereid
worms are aggressive to members of the same sex (Reish and Alosi,
1968; Johns et al., 1989). Some nereids also prey on smaller
species and Palaemonetes may crop the siphons of bivalves.
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D. RECOMMENDED AND SECONDARY SPECIES:
Based on these various criteria, we have identified five
recommended bioaccumulation test species and another eight
"secondary" test species (Table VII-l). The "recommended" species
meet all or nearly all of the desired criteria and are well
established bioaccumulation test species in both regulatory and
experimental studies. The recommended species are the
polychaetes Nereis diversicolor and Neanthes virens. and the
bivalves Macoma nasuta. Macoma balthica. and Yoldia limatula.
Within their tolerance levels, these species should serve as
suitable test species, and we recommend using at least one of
these species in all tests, at least until the suitability of
other species has been demonstrated locally.
The secondary bioaccumulation species meet the required
characteristics but either are deficient in one or more of the
important desired characteristics and/or there is insufficient
information to make a final evaluation. Some of the secondary
species offer potential advantages such as including additional
phylogenetic groups (i.e., crustaceans), adaptability to
culturing (e.g. Neanthes arenaceodentata). and high pollution
tolerance (Capitella spp). The importance of these various
characteristics will depend upon the site specific situation
(e.g., level of toxicity of sediment). We recognize that the
list of secondary bioaccumulation species is not exhaustive, and
there may be other suitable test species.
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This list of recommended and secondary test species includes
mostly estuarine species, reflecting both the hardiness and
relative ease of collecting estuarine species. As many of these
species extend into shallow marine habitats, they should serve as
suitable surrogates species for near-coastal environments. There
are no open ocean or brackish water (<10 ppt salinity) species
included, and the species listed generally do not extend into
these habitats. However, there does not seem to be a priori
reason why estuarine species should underestimate the tissue
residues of off-shore or brackish water species, though this
assumption should be tested. Discussion of fishes or
megainvertebrates is beyond the scope of this work, but a list of
recommended species for field monitoring of sewage discharges can
be found in Tetra Tech (1985a).
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APPENDIX VIII-1: SOURCES FOR TEST ORGANISMS
BIOLOGICAL SUPPLY HOUSES
COMPANY
PERTINENT SPECIES
Gulf Specimen Company
PO Box 237
Panacea, Florida 32346
904-984-5297
Pacific Biological Supply
P.O. 536
Venice, CA 90291
213-822-5757
Sea Life Supply
740 Tioga Ave.
Sand City, CA 93955
408-394-0828
Wood Hole Marine Biological Laboratory
Marine Resources Dept.
Woods Hole, MA 02543
508-548-3705
Nereis. Palaemonetes
Uca. Callinectes
Palaemonetes. Capitella.
Uca. Callianassa
OTHER SOURCES
Don Reish
Dept of Biology
California State U.
Long Beach, CA 90840
213-985-4845
Maine Bait Co.
Newcastle, Maine 04553
207-563-3000
Local Bait suppliers
University Biology or
Marine Sciences Depts.
Neanthes arenaceodentata
Capitella sp.
Nereis virens
nereid worms, Callianassa.
clams,
wide variety
Note: Inquire as to the availability of specific species well
ahead of the time the organisms are required.
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APPENDIX IX-1: SPECIAL PURPOSE EXPOSURE CHAMBERS
A. CLAMBOX
This exposure chamber is designed to separate the inhalant
and exhalant siphons of sediment-ingesting clams having
independent siphons (see Figure IX-1A). The technique is
applicable for Macoma spp. and other tellinids, though in most
bivalves the two siphons are fused together to form the "neck".
The apparatus allows the isolation and collection of the feces
from the parent sediment and ventilated (pumped) water from the
input supply. This allows a direct measure of short- and long-
term ventilation and sediment processing rates (the Fx terms of
Equation 5, Appendix 1-1) (Specht and Lee, 1989). By analyzing
the pollutant content in the feces or the ventilated water, the
amount of pollutant extracted by the clam (the EPx term' of
Equation 5, Appendix 1-1) can be estimated. The chamber has been
used to determine the efficiency of uptake of dissolved
hexachlorobenzene (HCB) by the gills (Boese et al., 1988), the
efficiency of HCB uptake through the gut from ingested sediment
(Lee et al., in press), uptake from ventilated interstitial water
(Winsor, et al., in press), and the passive sorption of HCB to
the soft-tissues (Lee et al., 1988).
B. WORMTUBES
These exposure chambers are tubes open on each end, which
simulate the burrow of sediment-ingesting polychaete worms such
as Abarenicola pacifica and Arenicola marina. The worms pump
199
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FIGURE DC-1A
Clambox Exposure Chamber
Dental dam membrane
o
o
Inhalant siphon
Exhalant siphon
Contaminated sedlmenft
Fecal pellets
100 ml Beaker
Overflow
standpipe
-------
water and sediment in one direction through the tubes (Figure IX-
1B). As with the clamboxes, the feces can be collected and
separated from the parent sediment, allowing the measurement the
sediment processing rate and the collection of the feces for
chemical analysis. These systems have been used to study the
effects of crude oil on sediment processing rates (Augenfeld,
1980) and on the uptake rate of cadmium as a function of the
addition of sewage carbon to sediment (Pelletier et al., 1988b).
Some versions also allow the simultaneous measurement of
ventilation rate and oxygen consumption (Kristensen, E., 1981;
Pelletier et al., 1988a).
C. SEDIMENT RESUSPENSION SYSTEMS.
This flow-through device automatically maintains a constant
suspended sediment load in the water column, using an electro-
optical feedback mechanism (U.S. EPA, 1978) which employs an
airlift dosing system, a transmissometer to measure particle
concentration, and a microcomputer which calculates the dose
required to achieve a programmed turbidity (Sinnett and Davis,
1983; Lake et al., 1985; Pruell et al., 1986). This system has
been used in several studies on the uptake and effects of
pollutants from resuspended sediments using the mussel, Mytilus
edulis. and the infaunal polychaete Nephtys incisa (Lake et al.,
1985; Nelson et al., 1987; Yevich et al., 1987). Other systems
201
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for maintaining suspended sediments are given in Rubinstein et
al. (1980) and Peddicord (1980). These chambers should be used
when there is concern about bioaccumulation in obligate benthic
filter-feeders (e.g., Mercenaria. Mya. Mytilus). or facultative
filter-feeders (e.g., Macoma) via resuspended sediments. This
mode of exposure is important in areas where current or wave
action periodically resuspend sediments and in areas with a
flocculent surface layer.
202
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FIGURE IX-IB
Wormtube Exposure Systems
w
a
b
incurrent
a.
b.
c.
Worm in sediment, 1 L glass box
Worm in glass tube, 30 L aquarium
Expanded view of ventilated water collection
and monitoring device
203
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APPENDIX X-l: ADDITIONAL TECHNIQUES FOR CORRECTING
FOR GUT SEDIMENT
A. MODIFICATIONS TO 24-HOUR PURGE AND DISSECTION
There are a number of other techniques or modifications to
the standard 24-hour purge in control sediment (Chapter X) which
should be considered in specific cases. When it is unclear
whether a species is voiding all its gut contents within 24
hours, a marker "sediment" can be added to the control sediment
during the purging. Marker sediments are inert particles of a
contrasting color or phosphorescence under UV radiation added to
the control sediment. Observation of feces composed of these
marker sediments is indicator that the gut has been voided.
Techniques for marking sediments for use as tracers are given in
Ingle (1966). In cases when it is critical not to have any
sediment in the gut, such as in certain studies of metals, it may
be necessary to purge the organisms in clean water without
sediment. Before using this approach, it is necessary to
determine whether the test species will satisfactorily void its
gut in the absence of sediment.
Another approach is to remove the gut sediment by
dissection. Dissection avoids the problems with the loss of
tissue pollutants during the purging, but is limited to the
larger test species (e.g., Abarenicola). Care has to be taken to
minimize loss of body fluids and to avoid contamination,
especially with the metals. General instructions for minimizing
contamination are available in Lauenstein and Young (1986).
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B. CALCULATING POLLUTANT MASS OF GUT SEDIMENT
It is possible to calculate the mass of pollutant associated
with the gut sediment if both the mass and the pollutant
concentration of gut sediment can be estimated. For selective
deposit-feeders, the pollutant concentration of the ingested
sediment may be several fold greater than the concentration of
the bulk sediment (see Lee et al., in press), so the bulk
sediment concentration should not be used as an estimate of the
gut sediment. Instead, the gut concentrations can be estimated
either from the pollutant concentrations of the ingested sediment
or the feces. Using the fecal pellet concentrations as the input
parameter, the whole body tissue residue (Ctw, including both the
tissue and gut sediment pollutants) can be expressed as:
Ctw =
(Mg*CPSf) + (Mt*Ct)
Mg + Mt
Expressed on a tissue residue only basis (i.e., no gut
sediment), the formula becomes:
Ctw*(Ms + Mt) - (CPSf*M )
ct = (2)
Mt
Where:
Ctw = whole body tissue concentration (tissue and
gut sediment) (ug/g)
Mg = mass of gut sediment (g)
CPSf = pollutant cone, in feces (ug/g)
Mt = mass of tissue (g)
Ct = tissue concentration without gut sediment (ug/g)
205
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If the ingested pollutant concentration (CPSi) is used, the
formula is the same except that CPSi is substituted for CPSf.
Use of fecal pellet pollutant concentration underestimates the
average gut pollutant content because some of the pollutants are
extracted from the sediment before defecation. Conversely,
ingested sediment overestimates the average gut pollutant content
because some of the pollutants have been extracted. These errors
are not expected to be large, but both methods could be
calculated and the results averaged for the most accurate
estimate. Fecal pellets can be collected for chemical analysis
by using special exposure chambers such as the clambox with
Macoma or wormtubes with polychaetes (see Appendix IX-1).
A method to estimate ingested dose is given in Lee et al. (in
press).
C. USE OF CONSERVATIVE TRACE ELEMENTS
Another approach to correcting for gut sediment is to use
the concentration of a conservative, non-biologically active
element as a means to determine sediment mass in the gut
(Kennedy, 1986). Knowing the sediment pollutant concentration,
it is then theoretically possible to calculate the amount of
pollutant associated with gut sediment. Some of the conservative
elements common in minerals but not typically found in more than
trace amounts in tissues include silicon, aluminum, and iron
(Kennedy, 1986). The difficulty with this approach is that the
elemental content of gut sediment in selective deposit-feeders
206
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may differ from that of the bulk sediment, especially if the
organism selectively ingests organic rather than mineral
particles. Additionally, this method will underestimate the gut
pollutant mass unless the pollutant concentration of the ingested
sediment (CPSi) is used rather than the bulk sediment
concentration.
207
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GLOSSARY
Accumulation Factor fAF) - Ratio of lipid normalized tissue
residue to carbon normalized sediment pollutant
concentration. (Appendix I-I).
ACE - Army Corps of Engineers.
Alpha - see Type I error.
Apparent Steady-State - See Steady-State.
ASTM - American Society for Testing and Materials.
BAF - See Bioaccumulation Factor.
BaP - Benzo(a)pyrene
BCF - See Bioconcentration Factor.
Bedded Sediment - Consolidated sediment (i.e., not suspended).
Beta - see Type II Error.
Bioaccumulation - Uptake from all phases, including water, food
and sediment.
Bioaccumulation Factor - Ratio of tissue residue to sediment
pollutant concentration. (Appendix 1-1)
Bioaccumulation Potential - Qualitative assessment of whether a
pollutant in a particular sediment is bioavailable.
(Appendix IV-1)
Bioconcentration - Uptake from water.
Bioconcentration Factor (BCF) - Ratio of tissue residue to
to water pollutant concentration. (Appendix 1-1)
Block - Group of homogeneous experimental units. (Chapter III)
Coefficient of Variation - A standardized variance term; the
standard deviation divided by the mean and expressed as a
percent. (Chapter III)
Comparison-wise Error - Type I error applied to a single
comparison of two means. Contrast with Experiment-wise
error. (Chapter XII)
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Compositing - The combining of separate tissue or sediment
samples into a single sample. (Chapter III)
Control Sediment - Sediment with very low pollutant concentrations
which is compared with reference and/or test sediments.
(Chapter II)
Control Treatment - Treatment (i.e., sediment type) that is
chosen to give a baseline value for comparison with results
from test treatments. May consist of either control or
reference sediments. (Chapter III)
Degradation - As used in the manual, it refers to the metabolic
breakdown of parent pollutant by a test species. Along
with depuration, it is one of the processes by which
pollutants are removed from an organism.
Depuration - Loss of the parent pollutant from an organism. See
Degradation.
DDT - Common environmental pollutant. Metabolites include DDD
and DDE.
DOC - Dissolved organic carbon. (Chapter VI)
DOM - Dissolved organic matter. (Chapter VI)
Eh - Redox potential, which is a measure the oxidation state of
a sediment. (Chapter VI)
EPA - Environmental Protection Agency
Experiment-wise Error - Type I error (alpha) chosen such that
the probability of making any Type I error in a series of
tests is alpha. Contrast with Comparison-wise error.
(Chapter XII)
Experimental error - Variation among experimental units given
the same treatment. (Chapter III and XII)
Experimental unit - Organism or organisms to which one trial of a
single treatment is applied. (Chapter III)
FDA - Food and Drug Administration.
Fines - Silt-clay fraction of a sediment. (Chapter VI)
Gut Purging - Voiding of sediment contained in the gut. (Chapter X
and Appendix X-l)
209
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Ha - The alternate hypothesis. (Chapter III)
Hg. - The null hypothesis. (Chapter III)
Hydrophobic pollutants - Low water solubility pollutants with a
high Row, and usually a strong tendency to bioaccumulate.
Interstitial water - Water between the particles (i.e., interstices)
in sediment. (Chapter VI)
kl - Uptake rate constant. (Chapter IV)
k2 - Depuration rate constant. (Chapter IV)
Kinetic Bioaccumulation Model - Any model that uses uptake
and/or depuration rates to predict tissue residues. In
this manual, it refers to the linear uptake, first-order
depuration model. (Chapter IV)
Koc - Organic-carbon partitioning coefficient.
Row - Octanol-water partitioning coefficient.
Long-Term Uptake Tests - Bioaccumulation tests with an exposure
period greater than 28 days. (Chapter IV)
LOI - Loss on ignition. (Chapter VI)
Multiple Comparisons - Statistical comparison of several
treatments simultaneously such as with ANOVA. (Chapter XIII)
Metabolism - see Degradation.
Minimum Detectable Difference - The smallest (absolute)
difference between two means that is statistically
distinguishable. (Chapter III)
NOAA - National Oceanic and Atmospheric Administration.
No Further Degradation - Approach by which a tissue residue is
deemed acceptable if it is not greater than those at a
reference site. (Chapter XIII)
PAH - Polyaromatic hydrocarbons.
Pairwise Comparisons - Statistical comparison of two
treatments. Contrast with multiple comparisons
(Chapter XII)
PCS - Polychlorinated biphenyls. Consists of over 200 congeners.
210
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Power - Probability of detecting a difference between the
treatment and control means when a true difference
exists. (Chapter III)
Pseudoreplication - Incorrect assignment of replicates, often due
to biased assignment of replicates. (Chapter III)
Reference sediment - Sediment used as an indicator of background
pollutant levels and resulting tissue residues. May have
moderate levels of pollutants. (Chapter II)
Replication - Assignment of a treatment to more than one
experimental unit. (Chapter III)
Sampling unit - The fraction of the experimental unit that is
to be used to measure the treatment effect. (Chapter III)
Spiking - Experimental addition of pollutants to a sediment.
(Chapter V and Appendix V-l)
Standard Reference Sediment - Standardized sediment and pollutant
used to determine the variability due to variation in the
test organisms. (Chapter II)
Steady State - A "constant" tissue residue as determined
by no statistical difference in three sampling periods
(Chapter IV)
TC - Total carbon, including organic and inorganic carbon
(Chapter VI)
Test Sediment - The sediment or dredge material of concern. This
is the sediment on which the regulatory decision will be
made. Contrast with Test Treatment. (Chapter II)
Test Treatment - Treatment that is compared to the control
treatment. It may consist of a test sediment (compared to a
reference or control sediment) or a reference sediment
(compared to the control sediment). (Chapter III)
Thermodynamic Partitioning Bioaccumulation Model - Bioaccumulation
model based on pollutant equilibrium partitioning among
lipids and sediment carbon. (Appendix 1-1)
Tissue residues - Pollutant concentration in the tissues.
TOC - Total organic carbon. (Chapter VI)
211
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Toxicokinetic Bioaccumulation Model - Bioaccumulation model based
on the feeding and ventilatory fluxes of the organism.
(Appendix 1-1)
Treatment - The procedure (type of sediment) whose effect is to
be measured. (Chapter III)
TVS - Total volatile solids. (Chapter VI)
Type I Error - Rate at which Ho is rejected falsely. (Chapter III)
Type II Error - Rate at which Ho is accepted falsely. (Chapter III)
212
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