EPA/620/R-95/008
August 1995
ENVIRONMENTAL MONITORING AND ASSESSMENT PROGRAM (EMAP)
LABORATORY METHODS MANUAL
ESTUARIES
VOLUME 1 - BIOLOGICAL AND PHYSICAL ANALYSES
edited by
C.J. Strobel3, D.J. Klemm1, L.B. Lobring1, J.W. Eichelberger1, A. Alford-Stevens1, B.B.
Potter1, R.F. Thomas1, J.M. Lazorchak1, G.B. Collins1, and R.L. Graves1
with contributions from
A. Alford-Stevens1, H.E. Allen2, W. Boothman3, D. Cobb3, G.B. Collins1, J.T. Creed1, D. DiToro9,
J.W. Eichelberger1, J. Enriquez5, S. Fink5, J.B. Frithsen6, F. Gongmin2, T. Heitmuller7, M. Jones5,
D.J. Keith3, D.J. Klemm1, J.O. Lamberson3, R.W. Latimer3, J.M. Lazorchak1, L.B. Lobring1, R.
Loebker1, J.D. Mahoney9, N. C. Malof1, T.D. Martin1, W. McDaniel4, D.M. McMullen5, G.E.
Morrison3, J.W. O'Dell1, R. PruelP, M. Smith5, H. Somme5, Q.J. Stober4, R.C. Swartz3, G. Thursby3,
R.M. Valente8, and J. Voit5
Environmental Monitoring Systems Laboratory, Cincinnati, Ohio
2University of Delaware, Newark, Delaware
3Atlantic Eclogy Division, Narragansett, Rhode Island
Environmental Services Division, Region 4, Athens, Georgia
technology Applications, Inc., Cincinnati, Ohio
6Versar, Inc., Columbia, Maryland
7National Biological Survey, Sabine Island, Gulf Breeze, Florida
8Science Applications International Corporation, Narragansett, Rhode Island
9Manhattan College, Bronx, New York
OFFICE OF RESEARCH AND DEVELOPMENT
U.S. ENVIRONMENTAL PROTECTION AGENCY
NARRAGANSETT, Rl 02882
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DISCLAIMER
This document is intended to document analytical methods for use by laboratories
conducting analyses for the Environmental Monitoiring and Assessment Program-Estuaries.
Mention of trade names, products, or services does not convey, and should not be interpreted as
conveying, official EPA approval, endorsement, or recommendation.
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NOTICE
This document replaces the previous EMAP-Estuaries Laboratory Methods Manual
(referenced as U.S. EPA, 1993 or Klemm et a/., 1993) which was never officially released other
than in draft form. That document was split into two volumes to conserve on paper (most
laboratories conducting analyses do not need ALL methods), and reformatted somewhat (for
appearance only). Note that the majority of the text remains unchanged from the 1993 document.
It is distributed unbound but hole-punched so individual sections can be updated and distributed
as necessary.
This document is Volume I of a two-part series. The second volume of the EMAP-Estuaries
Laboratory Methods Manual presents methods for the chemical analyses of sediments and tissue.
This document is AED (Atlantic Ecology Division) contribution Ne 1716.
The appropriate citation for this report is:
U.S. EPA. 1995. Environmental Monitoring and Assessment Program (EMAP): Laboratory
Methods Manual - Estuaries, Volume 1: Biological and Physical Analyses. United States
Environmental Protection Agency, Office of Research and Development, Narragansett, Rl.
EPA/620/R-95/008.
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TABLE OF CONTENTS
NO
1
2
3
4
5
6
iTICE
INTRODUCTION
SEDIMENT TOXICITY TEST METHODS
BENTHIC MACRO IN VERTEBRATE METHODS MACROBENTHIC
COMMUNITY ASSESSMENT
HISTOPATHOLOGY
SEDIMENT SILT-CLAY CONTENT AND GRAIN SIZE DISTRIBUTION
TOTAL SUSPENDED SOLIDS
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EMAP-Estuaries Laboratory Methods Manual Section 1 - Introduction
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SECTION 1
INTRODUCTION
1. BACKGROUND
1.1 The U.S. Environmental Protection Agency is developing the Environmental Monitoring and
Assessment Program (EMAP) to determine the current status, extent, changes and trends in the
condition of our nation's ecological resources on regional and national scales. The nation's
ecological resources are a national heritage, as essential to the country now and in the future as
they have been in the past. Data indicate that regional and international environmental problems
may be endangering these essential resources. The potential threats include acid rain, ozone
depletion, nonpoint-source pollution, and climate change.
1.2 Unfortunately, the status of the national environment is not well documented, rendering it
impossible to assess quantitatively where resources may be degrading and at what rate the
degradation may be occurring. The EPA Science Advisory Board (SAB) recognized this deficiency
and recommended in 1988 that EPA initiate a program that would monitor ecological status and
trends, as well as develop innovative methods for anticipating emerging problems before they reach
crisis proportions. EPA's response to the SAB's recommendations is EMAP.
2. OBJECTIVES
2.1 The primary goal of EMAP is to provide environmental decision makers with statistically valid
interpretive reports describing the health of our nation's ecosystems. Knowledge of the health of
our ecosystems will give decision makers and resource managers the ability to make informed
decisions, set rational priorities, and make known to the public the costs, benefits, and risks of
proceeding or refraining from implementing specific environmental regulatory actions. EMAP's
ecological status and trends data will allow decision makers to assess objectively whether or not
the nation's ecological resources are responding positively, negatively, or not at all, to the
regulatory programs put in place ostensibly to benefit them.
2.2 To accomplish its goals, EMAP is to document the condition of the nation's forest, wetlands,
estuarine and coastal waters, inland surface waters, Great Lakes, agricultural lands, and arid lands
in an integrated manner, on a continuing basis. Although EMAP is designed and funded by EPA's
Office of Research and Development (ORD), other offices and regions within EPA and other federal
agencies have contributed to its development and will participate in the collection and use of EMAP
data. When fully implemented, EMAP will form a complex national monitoring network, with a large
proportion of the data collection and analysis being accomplished by other federal, state, and local
agencies.
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2.3 The following four objectives guide EMAP (from Thornton et a/., 1993):
• Estimate the current status, trends, and changes in selected indicators of the Nation's
ecological resources on a regional basis with known confidence.
• Estimate the geographic coverage and extent of the Nation's ecological resources
with known confidence.
• Seek associations between selected indicators of natural and anthropogenic stresses
and indicators of ecological resources.
• Provide annual statistical summaries and periodic assessments of the Nation's
ecological resources.
3. SCOPE
3.1 EMAP-Estuaries (EMAP-E) is that part of the overall program which is monitoring the health
of estuaries and marine waters along our nation's coastline. Estuaries, which occur in tidally-
influenced portions of rivers and embayments, represent a vital component of near-coastal
ecosystems. Estuaries form a key ecological link between the nation's rivers and the coastal
waters of the continental shelf. Estuarine environments encompass tidal wetlands; submerged
aquatic vegetation communities; and inlets, bays, and lagoons. Estuaries provide critical spawning
and nursery habitat for commercial fish and shellfish, while the land around these ecosystems is
becoming highly populated. These valuable and threatened ecosystems are easily abused.
Estuaries directly receive much of the wastewater, after it has been treated, that is generated by
homes, businesses, and industries in estuarine watersheds. In addition, effluent or runoff that
enters rivers at points far from the coastline eventually can reach estuarine environments.
3.2 EMAP-E has divided all the nation's coastline into discrete regions for study. The first of these
regions to be studied, called the Virginian Province, extends from Cape Cod, Massachusetts to
Cape Henry, Virginia. Study began in Virginian Province bays and estuaries in the summer of
1990. Monitoring in the Gulf of Mexico (Louisianian Province) began in the summer of 1991.
Sampling of other regions will be phased in over the next several years, until all of the coastal
waters in the country are sampled yearly.
4. APPROACH
4.1 Assessing the status and trends for the nation's estuarine and coastal ecological resources
requires data collected in a standardized manner, over large geographic scales, for long periods
of time. Such assessments cannot be accomplished solely by aggregating data from the many
individual, short-term monitoring programs that have been conducted in the past and are being
conducted currently. Differences in the parameters measured, the collection methods used, timing
of sample collection, and program objectives severely limit the usefulness of historical monitoring
data for conducting regional and national status and trends assessments. The EMAP-E program
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will monitor a defined set of parameters (i.e., indicators of estuarine and coastal environmental
quality) on a regional scale, over a period of decades, using standardized field sampling and
laboratory methods with a probability-based sampling design. These characteristics distinguish
EMAP-E from other monitoring programs and will provide the data for preparing the regional and
national scale assessments that are needed to address the environmental issues of the 1990's and
beyond.
4.2 EMAP-E does not have the resources to monitor all ecological parameters of concern to the
public, Congress, scientists, and decision makers. Therefore a defined set of parameters that serve
as indicators of environmental quality are being measured. Categories of indicators identified and
sampled are as follows:
• Biotic condition indicators - Measurements that quantify the integrated response of
ecological resources to individual or multiple stressors. Included are benthic species
composition, abundance and biomass; gross pathology of fish; fish species
composition and abundance; relative abundance of large burrowing bivalves; and
histopathology offish.
• Abiotic condition indicators - Physical, chemical, and biological measurements that
quantify pollutant exposure, habitat degradation, or other causes of degraded
ecological condition. Included are sediment contaminant concentration; sediment
toxicity; contaminants in fish flesh; contaminants in large bivalves; and continuous
and point measurements of dissolved oxygen concentration.
• Habitat indicators - Physical, chemical, and biological measurements that provide
basic information about the natural environmental setting. Included are sediment
characteristics; water salinity, temperature, pH, depth, and clarity; chlorophyll a
fluorescence and the amount of photosynthetically active radiation (PAR) in the water
column.
4.3 Recommended protocols for those indicator parameters that are measured in the laboratory
are presented in the later sections of this document and in Volume II - Chemistry Methods.
Protocols for indicator parameters collected or measured in the field are contained in EMAP-E Field
Operations Manuals (Macauley, 1994; Reifsteckef a/., 1993).
5. REFERENCES
Macauley, J. 1994. Environmental Monitoring and Assessment Program - Estuaries: 1994
Louisianian Province Field Operations and Safety Manual. Environmental Research
Laboratory, Office of Research and Development, U.S. Environmental Protection Agency, Gulf
Breeze, FL.
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Reifsteck, D.R., C.J. Strobel and O.K. Keith. 1993. Environmental Monitoring and Assessment
Program - Estuaries: 1993 Virginian Province Field Operations and Safety Manual. U.S.
Environmental Protection Agency, Office of Research and Development, Environmental
Research Laboratory, Narragansett, Rl. June 1993.
Thornton, K.W., D.E. Hyatt, and C.B. Chapman, eds. 1993. Environmental Monitoring and
Assessment Program Guide. EPA/620/R-93/012, Research Triangle Park, NC: U.S.
Environmental Protection Agency, Office of Research and Development, Environmental
Monitoring and Assessment Program, EMAP Research and Assessment Center.
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SECTION 2
SEDIMENT TOXICITY TEST METHODS
TABLE OF CONTENTS
Subsections
1. Introduction 2
2. Scope and Application 3
3. Summary of Method 4
4. Definitions 4
5. Interferences 5
6. Safety 6
7. Apparatus and Equipment 7
8. Test Organisms 9
9. Dilution Water 11
10. Reagents and Consumable Materials 11
11. Sample Collection, Preservation, and Storage 11
12. Calibration and Standardization 12
13. Quality Assurance/Quality Control 12
14. Test Procedure for Marine Sediments Using Ampelisca abdita 15
15. Test Procedure for Marine Sediments Using Mysids
(Mysidopsis bahia) and Penaeid Shrimp 21
16. Calculations 26
Appendix A 29
Appendix B 32
Appendix C 34
Appendix D 35
17. References 36
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1. INTRODUCTION
1.1 In the EMAP-Estuaries program, the acute toxicity of surface sediments will be assessed as a measure
of the biological effects of sediment contaminants. In general, sediment toxicity tests provide information
that is independent of chemical characterizations and ecological surveys (Chapman, 1988). These tests
have proven to be effective in both marine and freshwater environments (Chapman, 1988; Swartz, 1987)
and have become an integral part of many benthic assessment programs (Swartz, 1989). They have been
used for permitting programs, site ranking for remediation, recovery studies following management actions,
and trends monitoring. A particularly important application is in programs seeking to establish contaminant-
specific effects.
1.2 The marine and estuarine amphipod sediment toxicity tests performed on EM AP-Estuaries will follow
standard ASTM guidelines (ASTM, 1991) and EPA methods (U.S. EPA, 1994). For a typical bioassay, 200
ml_ of surface sediment (top 2 cm) from homogenized grab samples collected at each sampling site will be
placed in one-liter canning jars covered with 600 ml of water. The bioassays will be conducted for 10 days,
under static conditions, at a constant temperature of 20°C and a salinity of 30 ± 2%o. Five replicate test
chambers will be used for the sediment from each station.
1.3 The east coast marine amphipod Ampelisca abdita will be employed as the primary test species in the
sediment toxicity tests. This species has been shown to be both acutely (Breteler et a/., 1989; DiToro et
a/., 1990; Rogerson et a/., 1985) and chronically (Scott and Redmond, 1989) sensitive to contaminated
sediments. In addition, it has been shown to be comparable in sensitivity to more commonly-used test
species like Rhepoxynius abronius and R. hudsoni (DiToro et a/., 1990), and has been successfully tested
using contaminated low salinity sediments. Because Ampelisca is a tube dweller, it is tolerant of a wider
range of sediment types than Rhepoxynius (Long and Buchman, 1989).
1.4 Because the sediment toxicity indicator needed by EMAP-Estuaries must be comparable across all
salinities, all tests will be conducted using Ampelisca abdita at a salinity of 30 ± 2%o.
1.5 In the Louisianian Province (the U.S. Coastal Gulf of Mexico), mysids (Mysidopsis bahia) will also be
used as a primary test species, in addition to A. abdita, for sediment toxicity tests. Mysids are small shrimp-
like crustaceans with demonstrated sensitivity to environmental contaminants. The U. S. EPA
Environmental Research Laboratory at Gulf Breeze, FL (ERL-GB) has developed culturing techniques and
toxicity testing methods (acute and life-cycle) for Mysidopsis bahia (U.S. EPA, 1978; U.S. EPA, 1987). Test
procedures, based on USEPA-approved guidelines for conducting solid-phase bioassays (USEPA/CE,
1977), have been adapted for EMAP-Estuaries and will be 96-hr static tests with aeration.
1.6 Also, during the first year of monitoring in the Louisianian Province, commercially important, penaeid
shrimp (Penaeus duorarum or P. aztecus) will be tested with sediment collected from 16 Indicator Testing
and Evaluation (ITE) sites; cost and logistical constraints prohibit testing penaeid shrimp on a Province-wide
scale. Sediment tests with post-larvae or juvenile penaeid shrimp will be conducted by using
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the same methods as described for mysids. Test results will be compared and evaluated for inferences
linking the observed toxicity to mysids to that for penaeids.
1.7 The sediment toxicity tests also will include clean control sediments with grain size and other natural
features matching those of the test sediment, as well as "water only" 96-hr exposure tests using the
reference toxicant sodium dodecyl sulfate (SDS) at various concentrations to determine LC50 values on
a continual basis (to assess both interlaboratory precision and the relative sensitivity of a given batch of test
organisms).
2. SCOPE AND APPLICATION
2.1 A number of sediment bioassay guides (U.S. EPA/CE, 1977;U.S. EPA, 1994; ASTM, 1991; Swartz et
a/., 1979; Swartz et a/., 1985) describe procedures for obtaining laboratory data concerning the short-term
adverse effects of potentially contaminated sediment, or of a test material experimentally added to
contaminated or uncontaminated sediment, on marine or estuarine infaunal amphipods during static 10-day
exposures. These procedures are useful for testing the effects of various geochemical characteristics of
sediments on marine and estuarine amphipods, and could be used to assess sediment toxicity to other
infaunal taxa, although modifications of the procedures appropriate to the test species might be necessary.
Procedures for 10-day static sediment toxicity tests are described herein for Ampelisca abdita (marine and
estuarine).
2.2 This procedure is applicable to sediments containing most chemicals, either individually or in
formulations, commercial products, and known or unknown mixtures. With appropriate modifications this
procedure can be used to conduct sediment toxicity tests on factors such as temperature, salinity, and
dissolved oxygen, and natural sediment characteristics (e.g., particle size distribution, organic carbon
content, total solids). This method can also be used to conduct bioconcentration tests and in situ tests, and
to assess the toxicity of potentially contaminated field sediments, or of such materials as sewage sludge,
oils, particulate matter, and solutions of toxicants added to sediments. An LC50 or EC50 of toxicants or
of highly contaminated sediment mixed into uncontaminated sediment can be determined. Materials either
adhering to sediment particles or dissolved in interstitial water can be tested.
2.3 This test procedure is not intended to exactly simulate the exposure of benthic amphipods to
contaminants under "natural" conditions, but rather to provide a convenient, rapid, standard toxicity test
procedure yielding a reasonably sensitive indication of the toxicity of materials in marine and estuarine
sediments.
2.4 Amphipods are an abundant component of the soft bottom marine and estuarine benthic community.
They are a principal prey of many fish, birds and larger invertebrate species. Some species are predators
of smaller benthic invertebrates. Others ingest sediment particles and thus are directly exposed to
contaminants. Amphipods are among the first taxa to disappear from benthic communities impacted by
pollution, and have been shown to be more sensitive to contaminated sediments than several other major
taxa (Sax, 1984). The ecological importance of amphipods, their wide geographical distribution, ease of
handling in the laboratory, and their sensitivity to contaminated sediments make them appropriate species
for sediment toxicity testing.
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3. SUM MARY OF METHOD
3.1 The relative toxicity of marine or estuarine sediments can be determined through a 10-day static test
with solid phase sediment and overlying water in aerated 1-L glass test chambers. Mortality and sublethal
effects, such as emergence from sediment, are determined after exposure of a specific number (usually
20) of amphipods to a quantity of test sediment. Response of the amphipods to the test sediment is
compared with response in control sediment. A negative control or reference sediment is used to provide
(a) a measure of the acceptability of the test by providing evidence of the health and relative quality of the
test organisms, and the suitability of the overlying water, test conditions and handling procedures, and (b)
the basis for interpreting data obtained from the test sediments.
3.2 The toxicity of field-collected sediment is indicated by the percent mortality of amphipods exposed to
that sediment compared to those exposed to control sediment. The toxicity of field sediments may also be
assessed by testing dilutions of a highly toxic test sediment with clean sediment to obtain information on
the toxicity of proportions of that sediment.
4. DEFINITIONS
4.1 The term "sediment" is used here to denote a naturally occurring particulate material which has been
transported and deposited at the bottom of a body of water. The procedures described can also be applied
using an experimentally prepared substrate within which the amphipods can burrow.
4.1.1 "Clean" sediment denotes sediment which does not contain concentrations of toxicants which cause
apparent stress to the test organisms or reduce their survival.
4.1.2 "Solid-phase" sediment is distinguished from elutriates and resuspended sediments in that the whole,
intact sediment is used to expose the organisms, not a form or derivative of the sediment.
4.2 "Toxicity" is the property of a material or combination of materials, to adversely affect organisms (see
ASTM 943).
4.3 "Exposure" is contact with a chemical or physical agent (see ASTM 943).
4.4 "Interstitial water" is the water within a wet sediment that surrounds the sediment particles. The amount
of interstitial water in sediment is expressed as the percent ratio of the weight of the water in the sediment
to that of the wet sediment.
4.5 "Overlying water" is the water which is added to the test chamber over the solid phase of the sediment
in a toxicity test.
4.6 "Partial life-cycle test" is a test utilizing a sensitive life-stage(s) of an organism used to assess the
toxicity of contaminants, and is distinguished from a complete life-cycle test which is conducted from egg
to egg or beyond, or for several life-cycles (Rand and Petrocelli, 1985).
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4.7 The LC50 is the statistically or graphically derived best estimate of the concentration of test material
added to or contained in sediment that is expected to be lethal to 50% of the test organisms under specified
conditions within the test period (see ASTM Standard E 943).
4.8 The EC50 is the statistically or graphically estimated concentration of test material in sediment that is
expected to cause a measured sublethal effect (for example the inability of amphipods to rebury in clean
sediment at the end of the test period), in 50% of the test organisms under specified conditions (see ASTM
Standard E 943).
5. INTERFERENCES
5.1 Due to the limited time sediment toxicity tests have been practiced, the methodology continues to
develop and evolve with time and research needs. Because of the developmental nature of sediment
toxicity testing, there are limitations to the method described in this guide.
5.2 Results of acute sediment toxicity tests will depend, in part, on the temperature, water quality, physical
and chemical properties of the test sediment, condition of the test organisms, exposure technique, and
other factors. Factors potentially affecting results from static sediment toxicity tests might include items
5.2.1 to 5.2.11.
5.2.1 Alteration of field sediments in preparation for laboratory testing.
5.2.1.1 Maintaining the integrity of the sediment environment during its removal, transport and testing in
the laboratory is extremely difficult. The sediment environment is composed of a myriad of micro-
environments, redox gradients and other interacting physiochemical and biological processes. Many of
these characteristics influence sediment toxicity and bioavailability to benthic and planktonic organisms,
microbial degradation, and chemical sorption. Any disruption of this environment complicates interpretations
of treatment effects, causative factors, and in situ comparisons.
5.2.1.2 Testing of sediments at temperatures or salinities other than those at which they were collected
might affect contaminant solubility, partitioning coefficients, and other physical and chemical characteristics.
5.2.2 Interactions between the sediment particles, overlying water, interstitial water, and humic substances,
and the sediment to overlying water ratio.
5.2.3 Interactions among chemicals which may be present in test sediment.
5.2.4 Photolysis and other processes degrading test chemicals.
5.2.5 Maintaining acceptable quality of overlying water.
5.2.6 Excess food may change sediment partitioning and water quality parameters.
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5.2.7 Resuspension of sediment during the toxicity test.
5.2.8 Limited opportunity for biological observations during the test because organisms bury in test
sediment.
5.2.9 Natural geochemical properties of test sediment collected from the field which may not be within the
tolerance limits of the test organisms.
5.2.10 Recovery of test organisms from the test system.
5.2.11 Endemic organisms which may be present in field collected sediments including (a) predators, (b)
species which may be the same as or closely related to the test species, and (c) microorganisms (e.g.,
bacteria or molds) or algae colonizing sediment and test chamber surfaces.
5.3 Static tests might not be applicable to materials that are highly volatile or are rapidly biologically or
chemically transformed. Furthermore, the overlying water quality may change considerably from the initial
overlying water. Because the experimental chambers are aerated, the procedures can usually be applied
to materials that have a high oxygen demand. Materials dissolved in interstitial waters might be removed
from solution in substantial quantities by adsorption to sediment particles and to the test chamber during
the test. The dynamics of contaminant partitioning between solid and dissolved phases at the initiation of
the test should therefore be considered, especially in relation to assumptions of chemical equilibrium.
6. SAFETY
6.1 General Precautions
6.1.1 Field sediments to be tested, especially those from effluent areas, might contain organisms that can
be pathogenic to humans. Special precautions when dealing with these sediments might include
immunization prior to sampling and use of bactericidal soaps after working with the sediments.
6.1.2 Sediments collected from the field might be contaminated with unknown concentrations of many
potentially toxic materials. Any potentially contaminated sediments should be handled in a manner to
minimize exposure of researchers to toxic compounds.
6.2 Safety Equipment
6.2.1 Personal Safety Gear
6.2.1.1 Many materials can affect humans adversely if precautions are inadequate. Therefore, skin contact
with all toxicants, overlying water, and sediments should be minimized by such means as wearing
appropriate protective gloves (especially when washing equipment or putting hands into test sediments or
solutions), laboratory coats, aprons, and glasses. Special precautions, such as covering test chambers and
ventilating the area surrounding the chambers, should be taken when conducting tests on volatile
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materials. Information on toxicity to humans (International Technical Information Institute, 1977; Sax, 1984;
Patty, 1963; Hamilton and Hardy, 1974; Goselin et a/., 1976), recommended handling procedures (Green
and Turk, 1978; National Research Council, 1981; Walters, 1980; Fawcett and Wood, 1982), and chemical
and physical properties of the test material should be studied before a test is begun. Special precautions
might be necessary with radiolabeled test materials (National Council on Radiation Protection and
Measurement, 1971; Shapiro, 1981) and with materials that are, or are suspected of being, carcinogenic
(National Institutes of Health, 1981).
6.3 General Laboratory Operations
6.3.1 Mixing of toxic sediments in open containers, and loading of toxic sediments into test chambers
should be done in a well-ventilated area, preferably a chemical fume hood. Face shields or protective
goggles should be worn during any operations that might involve accidental splashing of sediments, such
as sieving, mixing and loading into test chambers.
6.3.2 Health and safety precautions and applicable regulations for disposal of stock solutions, overlying
water from test chambers, test organisms, and sediments should be considered before beginning a test
(see ASTM Standard D 4447). Consideration of cost as well as detailed regulatory requirements might be
necessary.
6.3.3 Cleaning of equipment with a volatile solvent such as acetone, should be performed only in a well-
ventilated area in which no smoking is allowed and no open flame, such as a pilot light, is present.
Cleaning equipment with acids should be done only in a well-ventilated area, and protective gloves and
safety goggles should be worn.
6.3.4 To prepare dilute acid solutions, concentrated acid should be added to water, not vice versa.
Opening a bottle of concentrated acid and adding concentrated acid to water should be performed only in
a well-ventilated area or a chemical fume hood.
6.3.5 Use of ground fault systems and leak detectors is strongly recommended to help prevent electrical
shocks because salt water is a good conductor of electricity.
7. APPARATUS AND EQUIPMENT
7.1 General Requirements
7.1.1 Test chambers containing sediment should be held in a well-lighted (at least 100 lux at the test
sediment surface), constant temperature room, incubator or recirculating water bath to maintain the
experimental temperature. Air used for aeration should be free of fumes, oil and water; filters to remove
oil and water are desirable. The area containing the test chambers must be well ventilated and free of
fumes, both to prevent contamination of test materials and to protect researchers from exposure to toxic
volatile materials which might be released from the test sediments. Enclosures may be needed to ventilate
the area surrounding test chambers.
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7.1.2 The exposure room should be equipped with a timing device for photoperiod control. If a photoperiod
other than continuous light is used, it might be desirable to incorporate a 15 to 30 minute transition period
when lights go on or off to reduce stress to the organisms from sudden large changes in light intensity
(Robinson et a/., 1988; U.S. EPA, 1979a). It is also desirable to have the room temperature and light
controls and the aeration on emergency power to protect the experiment in case of a power failure.
7.2 Construction Materials
7.2.1 Equipment and facilities that contact test solutions or any water or sediment into which test organisms
will be placed should not contain substances that can be leached or dissolved by aqueous solutions in
amounts that adversely affect test organisms. In addition, equipment and facilities that contact stock or test
solutions or sediment should be chosen to minimize sorption of test materials from water.
7.2.2 Glass, Type 316 stainless steel, nylon, high density polyethylene, polycarbonate and fluorocarbon
plastics should be used whenever possible to minimize dissolution, leaching, and sorption, except that
stainless steel should not be used in tests on metals in salt water.
7.2.3 Concrete and rigid plastics may be used for holding tanks and in the water-supply system, but they
should be soaked, preferably in flowing test water (salt or fresh), for a week or more before use (Carmignani
and Bennett, 1976). Brass, copper, lead, cast iron pipe, galvanized metal, and natural rubber should not
contact test seawater, stock solutions, or test sediment before or during the test.
7.2.4 Tubing used in making up test seawater or reconstituted freshwater and in aerating the test chambers
should be nontoxic (Tygon® R-3603 or equivalent). New tubing should be soaked at least one week prior
to use. Separate sieves, dishes, containers, and other equipment should be used to handle test sediment
or other toxic materials and these should be kept and stored separately from those used to handle live
animals prior to testing.
7.3 Test Chambers
7.3.1 Species-specific information on test chambers is given in Appendices C and D. The test chambers
should be placed in a water bath to minimize temperature fluctuations, and should be aerated. Aeration
can be provided as in Subsection 14 and 15.
7.4 Cleaning
7.4.1 Test chambers and other glassware, and equipment used to store and prepare test sea water, stock
solutions, and test sediment should be cleaned before use.
7.4.2 All glassware should be cleaned before each use by washing with laboratory detergent, followed by
three distilled water rinses, 10% nitric v/v (HNO3) or hydrochloric (HCI) acid rinse, and at least two distilled
water rinses.
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7.5 Acceptability of Test Facilities and Equipment
7.5.1 Laboratory and bioassay temperature control equipment must be adequate to maintain recommended
test temperatures. Recommended materials must be used in the fabrication of the test equipment which
comes in contact with the sediment being tested. The acceptability of new holding or testing facilities
should be demonstrated by conducting "non-toxicant" tests in which test chambers contain control sediment
and appropriate overlying water. These tests will demonstrate whether facilities, water, control sediment,
and handling techniques are adequate to result in acceptable control level survival.
7.6 Species Specific Equipment Requirements
7.6.1 See Appendix B for Ampelisca abdita.
8. TEST ORGANISMS
8.1 Species
8.1.1 The organism used in the test described in this manual is the marine amphipod Ampelisca abdita.
All individuals used in the tests should be disease-free and should be positively identified to species.
8.1.2 The species of infaunal amphipod to be used in the sediment toxicity test should be selected based
on availability, sensitivity to test materials, tolerance to ecological conditions (e.g., temperature, salinity, and
grain size), ecological importance, and ease of handling in the laboratory. The source and type of sediment
being tested or the type of test to be implemented might dictate selection of a particular species.
8.1.3 Ideally, species or genera with wide geographical distributions should be selected, so that test results
can be compared among laboratories with similar species. Species used should be identified with an
appropriate taxonomic key, and identifications should be verified by a taxonomic authority.
8.1.4 The appendices to this document give guidance as to requirements and methods of handling for
various species of amphipods. Use of the species listed in the appendices is encouraged to increase
comparability of results.
8.1.5 The environmental requirements and sensitivity of a prospective test species of amphipod to test
materials and to various sediment characteristics should be established before it is widely used in toxicity
tests.
8.1.6 The tolerance of a test species to variations in sediment characteristics such as particle size
distribution, organic enrichment, and interstitial water salinity should be established before responses can
be ascribed to contaminant effects.
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8.1.7 Choice of the scale of the test chamber, density of test organisms, temperature, salinity, and control
sediment may have to be modified to accommodate the requirements of the test species. Required
modifications should be based on conditions at the natural habitat of the species.
8.1.8 If tube-building amphipods are used in sediment toxicity testing, it should be kept in mind that the
amphipods might not be directly in contact with test sediment after their tubes are built, and they might
pump overlying water through their tubes rather than utilizing interstitial water. They might feed on
particulate materials that either are suspended in the water column or have settled on the sediment surface,
while burrowing species might feed on particles or meiofauna found within the sediment. Thus tube builders
and burrowing species might have different routes of exposure to adsorbed or dissolved sediment
contaminants.
8.1.9 Amphipods that emerge from the sediment and either swim in overlying water
or crawl on the sediment surface might not be continually exposed to the test sediment.
8.2 Age
8.2.1 All organisms should be as uniform as possible in age and size. The age or size class for a particular
species should be chosen so that sensitivity to test materials is not affected by such factors as state of
maturity, reproduction, or seasonality. For EMAP-Estuaries sediment toxicity testing, juvenile Ampelisca
abdita in the size range 3 to 5 mm should be used for testing (see appendices).
8.3 Source
8.3.1 All individuals in a test should be from the same source, because different populations of the same
species might have different acute sensitivities to contaminants. Marine amphipods are usually obtained
directly from a wild population in a clean area, although attempts have been made to culture some species.
Collecting permits for field-collected amphipods might be required by some local and state agencies.
8.3.2 If test organisms are cultured or held for an extended period of time in the laboratory, the response
of laboratory-held organisms to test materials should be compared to that of animals recently collected from
the field to assure that laboratory stresses do not affect their sensitivity to test materials (Adams et a/.,
1985). Generally the reference toxicant assay performed with each test satisfies this requirement.
8.4 Quality
8.4.1 All amphipods used in a test must be of acceptable quality. A qualified amphipod taxonomist must
be consulted to ensure that the animals in the test population are all of the same species.
8.4.2 If organisms are collected from the field prior to testing, they should be obtained from an area known
to be free of toxicants and should be held in clean, uncontaminated water and facilities. Organisms held
prior to testing should be checked daily, and individuals which appear unhealthy or dead should be
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discarded. If greater than 5% of the organisms in holding containers are dead or appear unhealthy during
the 48 hours preceding a test, the entire group should be discarded and not used in the test.
8.5 Reference Toxicants
8.5.1 Whenever test organisms are obtained from an outside source (e.g., field collected or obtained from
an outside culture facility), their sensitivity must be evaluated with a reference toxicant in an appropriate
short-term toxicity test performed concurrently with the toxicity tests of actual sediment samples. Short-term
toxicity tests without sediment may be used to generate LC50 values for this purpose. If the laboratory
maintains breeding cultures of test organisms, the sensitivity of the offspring should be determined in a
toxicity test performed with a reference toxicant at least once a month. If preferred, this test also may be
performed concurrently with the toxicity tests of sediment samples.
8.5.2 The reference toxicant sodium dodecyl sulfate (SDS) must be used for EMAP-Estuaries sediment
toxicity testing using Ampelisca abdita.
8.6 Specific Species Requirements
8.6.1 See Appendix B Specific Requirements for Ampelisca abdita.
9. DILUTION WATER
9.1.1 See Appendix A for Ampelisca abdita.
10. REAGENTS AND CONSUMABLE MATERIALS
Reference toxicant (e.g., SDS)
Small-bore pipettes
5% buffered formalin
Rose Bengal stain
11. SAMPLE COLLECTION, PRESERVATION, AND STORAGE
11.1 See appropriate EMAP-Estuaries Field Operations Manual.
11.2 Techniques for sample collection, handling, and storage are described in the EMAP-Estuaries Field
Operations Manuals (Strobel and Schimmel, 1991; Macauley, 1991). Sediment samples for toxicity testing
should be chilled to 4°C when collected, shipped on ice, and stored in the dark in a refrigerator (4°C) for
no longer than 30 days before the initiation of the test.
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11.3 Sample containers should be made of inert materials to prevent contamination, which might result in
artifactual changes in toxicity (Subsection 7, Apparatus and Equipment).
11.4 All sampling devices and any other instruments in contact with the sediments should be chemically
cleaned between stations. Sediment contact with metals (including stainless steel) and plastics (including
polypropylene and low density polyethylene) should be avoided as contaminant interactions (e.g., surface
adsorption) may occur. Only sediments not in contact with the sides of the sampling device should be
subsampled, composited, and subsequently homogenized using instruments composed of non-reactive
(i.e., inert) materials. The adequacy of the field homogenization technique for sediments will be
documented in a special study prior to the start of field work.
12. CALIBRATION AND STANDARDIZATION
12.1 Instruments used for routine measurements must be calibrated and standardized according to
instrument manufacturer's procedures, see EPA methods: 150.1, 360.1, 170.1, and 120.1 (U.S. EPA,
1979a).
12.2 All routine chemical and physical analyses must include established quality assurance practices as
outlined in Agency methods manuals (U.S. EPA, 1979a,b).
13. QUALITY ASSURANCE
13.1 Specific QA/QC guidelines for the Sediment Toxicity Test Methods are found in the most recent
versions of the QA Project Plans developed for each active EMAP-Estuaries Province (Valente and Strobel,
1992; Heitmuller and Valente, 1992).
13.2 Required Controls
13.2.1 Every test requires a control treatment consisting of sediment from the amphipod collection site or
other sediment known to be non-toxic to, and within the geochemical requirements of the test species. The
same water, conditions, procedures, and organisms are used as in the other test treatments, except that
none of the test material is added to the control sediment or water. At least five laboratory replicates of the
control sediment should be included in all tests regardless of whether test sediments are replicated. This
allows comparisons among experiments and among laboratories of the validity of procedures used in
individual tests.
13.2.2 In addition to the standard control, if a field sediment has properties such as grain size or organic
content which might exceed the tolerance range of the test species, it is desirable to include non-toxic
reference sediment controls for these characteristics. The design of field surveys should include an
additional field control involving five replicate samples from an area which is free from sediment
contamination. This provides a site-specific basis for comparison of potentially toxic and non-toxic
conditions, and can account for mortality associated exclusively with subjecting the organisms to non-native
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sediments. The concentrations of chemical contaminants should be measured in these field control
sediments in order to justify the assumption that they are contaminant-free.
13.3 Precision
13.3.1 The ability of the laboratory personnel to obtain consistent, precise results must be demonstrated
with reference toxicants before attempts are made to measure the toxicity of sediment samples.
13.3.2 The single laboratory precision of the Ampelisca test should be determined by performing at least
five or more preliminary tests with a reference toxicant. Short-term (e.g., 96-hr) tests without sediments may
be used for this purpose. Precision can be described by the mean LC50 (determined for each test by an
appropriate method of regression analysis), standard deviation, and percent relative standard deviation
(coefficient of variation, or CV) of the five (or more) replicate reference toxicant tests.
13.3.3 Because single laboratory precision has not been determined previously for the Ampelisca test,
criteria for laboratory acceptance cannot be specified. Continual monitoring of reference toxicant test
precision during the EMAP-Estuaries project, facilitated by the use of control charts (Subsection 13.5,
Control Charts), will provide the basis for establishing such criteria.
13.4 Replication and Test Sensitivity
13.4.1 The sensitivity of the tests will depend in part on the number of replicates, the probability level
selected, and the type of statistical analysis. The recommended minimum number of replicates and the
statistical method(s) useful for addressing the study objectives are detailed in Subsection 14, Test
Procedures for Marine Sediments Using Ampelisca abdita. In general, the number of replicates used in a
test should be adequate for testing hypotheses and detecting departures from the assumptions of the
particular statistical analyses employed.
13.5 Control Charts
13.5.1 A control chart should be prepared for each reference toxicant-organism combination, and
successive toxicity values should be plotted and examined to determine if the results are within prescribed
limits. In this technique, a running plot is maintained for the toxicity values (X,) from successive tests with
a given reference toxicant (Ziegenfuss et al., 1986). For regression analysis results (such as EC50s), the
mean (X) and upper and lower control limits (± 2 standard deviations) are recalculated with each successive
point until the statistics stabilize. Outliers, which are values which fall outside the upper and lower control
limits, and trends of increasing or decreasing sensitivity are readily identified. At the P0 05 probability level,
one in twenty tests would be expected to fall outside of the control limits by chance alone.
13.5.2 If the toxicity value from a given test with the reference toxicant does not fall in the expected range
for the test organisms, the sensitivity of the organisms and the overall credibility of the test are suspect.
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In this case, the test procedure should be examined for defects and should be repeated with a different
batch of test organisms.
13.6 Record Keeping and Reporting
13.6.1 Proper record keeping is required. Bound notebooks should be used to maintain detailed records
of the test organisms such as species, source, age, date of receipt, and other pertinent information relating
to their history and health, and information on the calibration of equipment and instruments, test conditions
employed, and test results. Annotations should be made on a real time basis to prevent loss of information.
Data for all QA/QC variables, such as reference toxicant test results and copies of control charts, should
be submitted by the laboratory as part of the data package.
13.6.2 The record of the results of an acceptable sediment toxicity test should include the following
information either directly or by reference to other available documents.
13.6.2.1 Names of test and investigator(s), name and location of laboratory, and dates of initiation and
termination of the test.
13.6.2.2 Source and method of preparation of water used, its salinity and any other pertinent chemical
characteristics.
13.6.2.3 Source of the control sediments, dates and methods of collection, method of transport and storage
of field sediments, method and dates of treatment of laboratory-prepared sediment, and method of
distribution to test chambers.
13.6.2.4 Source and date of collection of the test organisms, scientific name, method of taxonomic
verification, age, life stage, means and ranges of lengths, observed diseases or unusual appearance,
treatments, holding, and acclimation procedures.
13.6.2.5 Description of the experimental design, test chambers and covers, the depth and volume of
sediment and water in the chambers, the date, time and method of beginning the test, numbers of test
organisms and chambers, temperature, salinity, and lighting regime.
13.6.2.6 The average and range of holding and test temperatures, and the method(s) of measuring or
monitoring or both.
13.6.2.7 Effects used to calculate EC50s (e.g., fecundity, growth) and a summary of general observations
of other effects.
13.6.2.8 A table of the biological data for each test chamber for each treatment (including the control(s))
in sufficient detail to allow independent statistical analyses.
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13.6.2.9 The 10-day LC50s or EC50s and the methods used to calculate them, and their 95% confidence
limits, or the survival or mortality data and their significance relative to the control(s); specify whether results
are based on measured or nominal concentrations of the test material.
13.6.2.10 The mean, standard deviation, and range of the length measurements for the additional group
of animals sorted and preserved at the beginning of each test.
13.6.2.11 Anything unusual about the test, any deviation from these procedures, and any other relevant
information.
13.6.3 Published reports should contain enough information to clearly identify the procedures used and
the quality of the results.
14. TEST PROCEDURE FOR MARINE SEDIMENTS USING Ampelisca abdita
14.1 Dissolved Oxygen (DO)
14.1.1 The concentration of dissolved oxygen (DO) in the water overlying the sediment in the test
chambers should be maintained at or near saturation by gently aerating the water (see appendices). Air
should be bubbled into the test chambers at a rate that maintains a > 90% dissolved oxygen concentration,
but does not cause turbulence or disturb the sediment surface. If air flow to the beakers is interrupted for
more than an hour, DO should be measured in the beakers to determine whether dissolved oxygen
concentrations have dropped to less than 60% of saturation.
14.2 Temperature
14.2.1 The temperature selected should be within the natural range of temperatures in the area from which
the amphipods occur in the field. Within an experiment, individual temperature readings should not vary
by more than 3°C from the selected test temperature, and the time-weighted average measured
temperature at the end of the test should be within 1°C of the selected test temperature. When temperature
is measured concurrently in more than one test chamber, the highest and lowest temperatures should not
differ by more than 2°C.
14.2.2 This species is routinely tested at 20°C, but has been tested from 8 to 25°C. In nature, feeding and
somatic growth occur at temperatures as low as 3 to 5°C (Bousfield, 1973). For comparison with other
Ampelisca abdita test results, 20°C is recommended.
14.3 Salinity
14.3.1 The salinity of the water overlying the test sediment in sediment toxicity tests must be within the
tolerance range of the selected test species.
14.3.2 A. abdita is tolerant of a wide salinity range, but most tests have been conducted at salinities of 28
to 32 %o.
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14.3.3 The salinity of the interstitial water of test sediments from the field should not be adjusted, because
such an operation might change the toxicological properties of the sediment.
14.4 Light/Photoperiod
14.4.1 For sediment toxicity tests involving A. abdita, lights are usually left on continuously. The constant
light increases the tendency of the organisms to remain buried in the sediment, and thus to remain exposed
to the test material.
14.5 Feeding
14.5.1 Infaunal amphipods do not require supplementary feeding during the 10-day toxicity test.
14.6 Beginning the Test
14.6.1 The exposure chamber routinely used to test A. abdita is a 1-L glass canning jar with a narrow
mouth.
14.6.2 Ampelisca abdita has not been tested in the 1-L beaker exposure chamber used in other amphipod
tests, but it is not anticipated that use of beakers would create any problems.
14.6.3 This amphipod inhabits fine-grained sediments, and as with other physical conditions, if it is
suspected that a coarse grain size of a test sediment will stress the animals, a grain size control should be
included.
14.6.4 The toxicity test begins when test organisms are first placed in test chambers containing test
material.
14.6.5 On the day before the test begins, each test sediment sample should be thoroughly homogenized
within its storage container, and an aliquot added to a test chamber to a depth of 3 to 4 cm.
14.6.6 Treatments should be randomly assigned to prenumbered test chambers.
14.6.7 The sediment within the test chamber should be settled by tapping the test chamber against the side
of the hand, or by smoothing the sediment surface with a nylon, fluorocarbon or polyethylene spatula.
14.6.8 A disk cut from 6-mil nylon, Teflon or polyethylene sheeting to fit the inside diameter of the test
chamber, and attached to a length of nylon monofilament for removal, can be placed on the sediment
surface to minimize sediment disruption as prepared toxicity test seawater is added up to the 800 ml mark
on the test chambers.
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14.6.8.1 The disk should be removed and rinsed with seawater between replicates of a treatment, and a
separate disk should be used for each treatment. The test chambers should then be covered, put in
numerical order into a temperature-controlled water bath, and aerated overnight. The system should be
left overnight to allow suspended particles to settle and an equilibrium to be established between sediment
and overlying water before the amphipods are added.
14.6.8.2 With either exposure chamber (1-L jar or beaker), the water column should be gently aerated with
a pipette inserted above the sediment surface.
14.6.9 The toxicity test is initiated (Day 0) when amphipods are distributed to each test chamber. It is
usually not possible to distribute amphipods to all test chambers at the same time, so it is necessary to
select a set of test chambers (usually 10 to 15) to be processed together. If treatments are replicated, each
treatment, including controls, should be represented in each set of test chambers to be processed together.
If treatments are not replicated, selection should be random.
14.6.10 A sufficient number of amphipods should be removed from the holding facility at one time to
provide about one-third more amphipods than are needed for one set of test chambers. This allows
selection of active, apparently healthy individuals.
14.6.10.1 Before amphipods are removed, the temperature and salinity of the water in the holding
containers should be recorded.
14.6.10.2 Amphipods should be sieved from the holding sediment using a 0.5 mm sieve and transferred
to a sorting tray or large Carolina dish containing water of the holding temperature and salinity. Twenty to
thirty amphipods should be tested per replicate.
14.6.10.3 Investigators who purchase amphipods from an outside source and have them shipped to the
testing facility may prefer the following procedures for holding and selecting test amphipods, in lieu of the
holding and sieving methods described above.
14.6.10.4 Amphipods are collected in the field while still remaining in their sediment mats. Disturbance to
the mat is held to a minimum and the mat is packaged in clean, aerated seawater, then shipped to the
testing facility via overnight mail.
14.6.10.5 Upon receipt at the testing facility, the mat containing the amphipods is immediately placed in
chilled (20°C), aerated seawater of appropriate salinity to allow the amphipods to acclimate to test
conditions for at least 48 hours prior to testing.
14.6.10.6 Amphipods are coaxed from the sediment mat by the simultaneous action of gently massaging
a section of the mat while irrigating it with test dilution water to separate the amphipods from their tubes.
The section of loosened mat is then placed onto a 0.5 mm sieve and submerged underwater to rinse the
amphipods clear of the mat. The amphipods tend to become trapped by the surface tension of the water
from which they can easily be collected by scooping with a fine-mesh net or piece of Nitex screen. The
captured amphipods are released into a sorting tray or large Carolina bowl containing sediment-free, test
dilution water.
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14.6.10.7 The isolated animals are then transferred to a lighted table for manual sorting using a small-bore
pipet. Active, apparently healthy amphipods should be impartially selected from the sorting tray and
sequentially distributed among dishes containing approximately 150 ml of prepared toxicity test seawater
until each dish contains the required 20 to 30 immature amphipods. Amphipods should meet the following
criteria to be selected for testing: a) amphipod should have a standard length of 3 to 5 mm when stretched
out or 1 to 2 mm when curled; b) the gut of the animal should be full, as indicated by a dark line located
dorsally on the animal; c) the animal must maintain a healthy appearance. While swimming, it is
outstretched and searching for sediment or cover; at cessation of swimming it immediately curls up. Care
should be taken not to select gravid females or males nearing sexual maturity. The number of amphipods
in each dish should be verified by recounting them into a separate dish containing sediment-free, test
dilution water.
14.6.10.8 At least one additional group of 20 to 30 amphipods should be randomly sorted at the beginning
of each test. This extra group should be preserved in 5 to 10% buffered formalin for later length
measurement as a check to ensure that appropriately-sized amphipods were selected for testing. Length
of each individual in the group should be determined, using a dissecting microscope, by measuring from
the base of the first antennae to the base of the telson. The measurements (mean length, standard
deviation, and range) should be recorded on the data sheets for each test and reported along with the final
results.
14.6.11 Amphipods should be added to test chambers by placing a 6-mil nylon, Teflon or polyethylene disk
on the water surface, and gently pouring the water and amphipods from the sorting dish over the disk into
the test chamber. An alternative method would be to gently pour the organisms onto a Nitex screen and
then gently wash them into the test chamber.
14.6.11.1 For each replicate, the contents of a sorting dish can be rinsed into a plastic cup with a 400- or
500-^m screened base and then into the exposure container. Any animals caught on the water's surface
can be gently pushed under using a glass rod. Any amphipods remaining in the dish should be gently
washed into the test chamber.
14.6.11.2 The water level should be brought up to the final test level in the test chamber (800 ml), the disk
removed, and the chamber replaced in the water bath, covered and aerated.
14.6.12 Amphipods should be given one hour to burrow into the sediment. Any amphipods which do not
burrow within one hour should be removed and replaced.
14.7 Routine Chemical and Physical Analysis
14.7.1 Monitoring the quality of the overlying water (for DO, pH, or for certain chemicals) in the test
chambers can be accomplished without disturbing the sediment, and may be done in the test chambers
containing the test amphipods.
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14.7.2 Temperature from a separate temperature beaker should be recorded throughout the test. If test
chambers are in more than one temperature controlled water bath, a temperature beaker should be set up
in each water bath. Temperature should be monitored at least hourly using a recording thermometer or the
daily maximum and minimum temperatures should be monitored (see ASTM Standard E- 729). Individual
temperature measurements should not vary by more than 3°C and the time-weighted average should not
differ by more than 1 °C from the designated test temperature (see Subsection 13.2, Temperature).
14.8 Duration of Test
14.8.1 The test begins when amphipods are added to test chambers containing test sediment. Amphipods
should be exposed to the test material for ten days. There are no observed substantial effects of starvation
or other laboratory artifacts in this amount of time (Swartz et a/., 1985). An exposure period of less than
ten days is generally not recommended. For some experimental designs, such as comparison of a 96-hr
LC50 between species in the presence or absence of sediment, other exposure periods may be used.
14.9 Observations During the Test
14.9.1 Response criteria indicating toxicity of test sediment include mortality and sublethal effects.
Sublethal effects include an emergence from highly toxic sediment during the course of the test. Response
criteria must be monitored in a "blind" fashion, that is, the observer must have no knowledge of the
treatment of the sediment in the test chambers. This is accomplished through randomization of sample
numbers.
14.9.2 Emergence - Since most infaunal amphipods remain buried during sediment toxicity tests, there is
little opportunity to monitor temporal changes in mortality or sublethal effects. An exception is the temporal
pattern of emergence from highly toxic sediment.
14.9.3 The test should be monitored at least daily (including the day of initiation and the day of termination)
for temperature, aeration, lights, and emergence of the amphipods from the test sediment.
14.9.3.1 Each test chamber should be observed by temporarily turning off the air to the test chambers, and
gently removing the cover from individual chambers with minimal disturbance of the chamber.
14.9.3.2 The number of amphipods observed completely or partially out of the sediment, either on the
sediment surface, swimming in the overlying water, or floating at the water surface, should be recorded.
Amphipods that are caught in the surface film should be gently pushed down into the water.
14.9.3.3 Any pertinent observations on the appearance of the sediment (such as color, presence of non-
test organisms, growth of mold or algae, or depth of oxidized layer) should be recorded.
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14.10 Test Termination
14.10.1 The endpoint for the 10-day test is mortality; dead animals should be counted and removed daily.
An amphipod is considered dead if it does not respond to gentle probing. It is also useful to note any
animals out of their tubes on the sediment or water surface, amphipods which are nearly dead and only
exhibit a muscular pleopod twitch, the presence of molts, and the condition of the tubes built. Emergence
from the sediment and the inability to construct a proper tube are sublethal behavioral responses that would
ultimately result in death.
14.10.2 After checking the assay on the last day, the contents of each exposure container should be rinsed
through a 0.5-mm sieve. (A smaller mesh sieve can be used for the final sieving if there is concern about
losing very small animals, but this will make the sieving process more time-consuming.)
14.10.3 If the experiment is small, the material retained on the sieve can be examined that day. If time
does not permit same-day examination, the retained material from each jar can be preserved in 5% buffered
formalin with Rose Bengal stain for later examination.
14.10.4 Any amphipods which are not accounted for when the sieved material is examined are presumed
to have died during the test. Amphipods which have died in their tubes will generally decompose during
the test or break apart during sieving. Rarely, an individual which has died during the test will be recovered
in the preserved material, and its appearance will be markedly different from those of the amphipods which
were alive when preserved. For instance, there may be little tissue left within its exoskeleton, or it may be
contorted.
14.10.5 If the test species is naturally present in the test sediment, the total number of live and dead
amphipods at Day 10 might exceed the number at Day 0.
14.11 Acceptability of Test
14.11.1 A 10-day sediment toxicity test should usually be considered unacceptable if one or more of the
following occurred:
14.11.1.1 All test chambers were not identical.
14.12.1.2 Treatments were not randomly assigned to test chambers.
14.11.1.3 Test organisms were not randomly or impartially distributed to test chambers.
14.11.1.4 Required negative, reference sediment, positive or solvent controls were not included in the test.
14.11.1.5 All test animals were not from the same population, were not all of the same species, or were
not of acceptable quality.
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14.11.1.6 The average length of the animals used in the test was not within the range 3 to 5 mm, or there
were wide variations in length.
14.11.1.7 Amphipods from a wild population were maintained in the laboratory for more than ten days,
unless the effects of prolonged maintenance in the laboratory has been shown to have no significant effect
on sensitivity.
14.11.1.8 The test organisms were not acclimated at the test temperature and salinity at least 48 hours
before they were placed in the test chambers.
14.11.1.9 Temperature, dissolved oxygen and concentration of test material were not measured, or were
not within the range as specified in Section 14.7, Routine Chemical and Physical Analysis.
14.11.1.10 Aeration to the test chambers was off for an extended time such that dissolved oxygen levels
dropped to less than 60% of saturation.
14.11.1.11.2 Response criteria were not monitored in a "blind" fashion, i.e., observers had knowledge of
the treatment of sediments in the test chambers.
14.11.2 Survival of organisms in control treatments should be assessed during each test as an indication
of both the validity of the test and the overall health of the test organism population. Tests are acceptable
if mean control survival is greater than or equal to 90%, and if survival in individual control test chambers
exceeds 80%.
15. TEST PROCEDURE FOR MARINE SEDIMENTS USING MYSIDS (Mysidopsis bahia) AND
PENAEID SHRIMP
15.1 Dissolved Oxygen (DO)
15.1.1 The concentration of dissolved oxygen (DO) in the water overlying the sediment in the test
chambers should be maintained at or near saturation by gently aerating the water (see appendices). Air
should be bubbled into the test chambers at a rate that maintains a > 90% DO concentration, but does not
cause turbulence or disturb the sediment surface. If air flow to the beakers is interrupted for more than an
hour, DO should be measured in the beakers to determine whether dissolved oxygen concentrations have
dropped to less than 60% of saturation.
15.2 Temperature
15.2.1 The sediment tests with mysids and penaeids will be run at a temperature of 20°C. Within an
experiment, individual temperature readings should not vary by more than 3°C from this test temperature,
and the time-weighted average measured temperature at the end of the test should be within 1°C of this
test temperature. When temperature is measured concurrently in more than one test chamber, the highest
and lowest temperatures should not differ by more than 2°C.
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15.3 Salinity
15.3.1 The EMAP-Estuaries tests with mysids and penaeids will be conducted using natural, filtered (< 20
jj,m) seawater with the salinity maintained at 20 + 2 %o. No attempt will be made to adjust the salinity of the
interstitial water of the test sediments.
15.4 Light/Photoperiod
15.4.1 In sediment toxicity tests involving mysids and penaeids, a 14-hour light and 10-hour dark
photoperiod is established with cool-white fluorescent lights adjusted to approximately 1200 microwatts/cm2.
15.5 Test Organisms
15.5.1 Mysids. Newly released juveniles are collected by siphoning water from holding tanks through nylon
screen chosen to allow only the passage of juveniles which are concentrated in a small mesh cage.
Juveniles are transferred to holding tanks where gradual acclimation to test conditions occurs. Within a 24-
hour period, changes in water temperature should not exceed 5°C and salinity changes should not exceed
5%o. During acclimation, mysids should be maintained in facilities with background colors and light
intensities similar to those of the testing areas. Mysidopsis bahia must be 3 to 5 days old and all animals
for any one test must be from the same source; that is, animals cultured in the laboratory and purchased
from a supplier cannot be mixed to form a test population for a sediment or reference toxicant test.
Whenever possible, mysids to be used in toxicity tests should originate from the laboratory to ensure the
individuals are of similar age and known history.
15.5.2 Penaeids (sp). Post-larval or juvenile shrimp should be used. Shrimp may be reared from eggs in
the laboratory or obtained directly from the field as juveniles. As with mysids, all animals for any one test
must be from the same source. Shrimp to be used in a particular test should be of similar age and be of
normal size and appearance.
15.5.3 Upon arrival at the test facility, the juvenile penaeid shrimp should be transferred to water closely
matching the temperature and salinity of the transporting medium. The period of acclimation to ambient
laboratory conditions should be at least 4 to 7 days. During acclimation, shrimp should be maintained in
facilities with background colors and light intensities similar to those of the testing areas. In addition, any
change in the temperature and chemistry of the dilution water used for holding and acclimating the test
organisms to those of the test should be gradual. Within a 24-hour period, changes in water temperature
should not exceed 2°C, and salinity changes should not exceed 5%o.
15.6 Feeding
15.6.1 Mysids and penaeid post larvae will be fed artemia (24 to 48 hours post hatch) once a day
(approximately 100 artemia nauplii per animal per day). Juvenile penaeids will not be fed during the 96-
hour test duration.
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15.7 Test Material
15.7.1 All samples will be chilled during shipment to the laboratory. Immediately after receipt, the sediment
sample(s) must be placed in a cooler and maintained at 4+1 °C. A record of temperature during storage
must be provided as part of the report.
15.8 Glassware
15.8.1 All glassware is detergent-washed, rinsed thoroughly with tapwater, rinsed with acetone, rinsed
thoroughly with deionized water, soaked in 10% HCI for four hours, and rinsed thoroughly with tapwater and
five times with deionized water.
15.9 Beginning the Test
15.9.1 The exposure chambers for the mysid and post larval penaeid test will be 1-L beakers. The
exposure chambers for the juvenile penaeid test (if post larvae are not available) will be 2-L Carolina dishes.
15.9.2 As the sample arrives, each entire test sediment sample should be press-sieved (dry) through a
stainless steel screen (1.0-mm mesh size) to remove predators and larger particles (e.g., rocks, shells).
15.9.3 On the day before the test begins each sediment sample is homogenized using a high speed (>
1600 rpm) stirrer. Sample should freely "roll" while being stirred and all sediment from sides and bottom of
sample container should be mixed into the sample.
15.9.4 The sediment within the test chamber should be settled by tapping the test chamber against the side
of the hand, or by smoothing the sediment surface with a polyethylene spatula.
15.9.5 A petri dish is placed on the sediment surface to minimize sediment disruption as prepared toxicity
test seawater is added up to the 700-ml mark in the test chambers.
15.9.6 The petri dish should be removed and rinsed with seawater between replicates of a treatment, and
a separate dish should be used for each treatment. The test chambers should then be covered, labeled,
transferred into a temperature-controlled water bath or environmental chamber where the tests shall be
conducted and aerated overnight. The system should be left overnight to allow suspended particles to settle
and an equilibrium to be established between sediment and overlying water before the test animals are
added.
15.9.7 The water column should be gently aerated with a glass pipette inserted above the sediment
surface.
15.9.8 The toxicity test is initiated (Day 0) when mysids or penaeids are distributed to each test chamber.
Three replicate test chambers per test organism will be used for each test sediment.
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15.9.9 Before mysids and penaeids are removed from the holding containers, the temperature and salinity
of the water in the holding containers should be recorded. A sufficient number of mysids and penaeids
should be removed from the holding facility at one time to provide about one-third more animals than are
needed for the test chambers.
15.9.10 The mysids or penaeids should be transferred to a sorting tray containing water of the holding
temperature and salinity. Active, apparently healthy mysids or penaeids should be impartially selected from
the sorting tray and sequentially distributed among intermediate holding dishes containing approximately
150 ml of prepared toxicity test water until each dish contains the required number of ten animals.
15.9.11 The animals in the intermediate holding dish should be counted to verify the number of animals
to be loaded into the testing chamber and observed to be sure the animals are active and healthy. The
water in the holding dish is then drawn down to a small volume before the entire lot is transferred to a test
chamber.
15.9.12 The test animals should be impartially distributed among test chambers in such a manner that test
results show no significant bias from the distributions. The test chambers are then positioned randomly.
15.10 Routine Chemical and Physical Analysis
15.10.1 Monitoring the quality of the overlying water in the test chambers can be accomplished without
disturbing the sediment, and may be done in the test chambers containing the test animals.
15.10.2 Temperature, from a separate temperature beaker, should be recorded throughout the test. If test
chambers are in more than one temperature-controlled water bath or environmental chamber, a temperature
beaker should be set up in each. Temperature should be monitored at least hourly using a recording
thermometer or the daily maximum and minimum temperatures should be monitored (see ASTM Standard
E 729). Individual temperature measurements should not vary by more than 3°C and the time-weighted
average should not differ by more than 1 °C from the designated test temperature.
15.11 Duration of Test
15.11.1 The test begins when mysids or penaeids are added to the test chambers containing the test
sediment. Mysids and penaeids should be exposed to the test material for 96 hours.
15.12 Observations During the Test
15.12.1 Mortality and lethargic behavior are the responses that will be monitored during the sediment
toxicity tests with mysids or penaeids.
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15.12.2 The test should be monitored at least daily (including the day of initiation and the day of
termination) for temperature, aeration, lights, and overall behavior of the mysids or penaeids exposed to
the test sediments.
15.12.3 Each test chamber should be observed by temporarily turning off the air to the test chambers, and
gently removing the cover from individual chambers with minimal disturbance of the chamber.
15.12.4 The number of mysids or penaeids observed on the sediment surface, swimming in the overlying
water, or floating at the water surface should be recorded. Mysids or penaeids that are caught in the surface
film should be gently pushed down into the water.
15.12.5 Water quality parameters (i.e., dissolved oxygen and pH) will be measured in at least one replicate
of each test treatment and control on Day 0 and Day 4 of testing.
15.12.6 Any pertinent observations on the appearance of the sediment (such as color, presence of non-test
organisms, growth of mold or algae, or depth of oxidized layer) should be recorded.
15.13 Test Termination
15.13.1 The endpoint for the 96-hour mysid and penaeid test is mortality; dead animals should be counted
and removed daily. A mysid or penaeid is considered dead if it does not respond to gentle probing.
15.13.2 After checking the assay on the last day, the contents of each exposure container should be rinsed
through a 0.5 mm sieve.
15.13.3 If the experiment is small, the material retained on the sieve can be examined that day. If time does
not permit same-day examination, the retained material from each jar can be preserved in 5% buffered
formalin with Rose Bengal stain for later examination.
15.13.4 Any mysids or penaeids which are not accounted for when the sieved test material is examined
are presumed to have died during the test.
15.14 Acceptability of Test
15.14.1 A 96-hour sediment toxicity test should usually be considered unacceptable if one or more of the
following occurred:
15.14.1.1 All test chambers were not identical.
15.14.1.2 Test organisms were not randomly or impartially distributed to test chamber.
15.14.1.3 A required control treatment consisting of clean sediment was not included in the test.
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15.14.1.4 All test animals were not from the same population, were not all of the same species, or were
not of acceptable quality.
15.14.1.5 The test organisms were not acclimated to within ± 5° C of the specified test temperature and
± 5%o of the specified test salinity at least 24 hours before they were placed in the test chambers.
15.14.1.6 Temperature, dissolved oxygen and concentration of test material were not measured, or were
not within acceptable ranges.
15.14.1.7 Aeration to the test chambers was off for an extended time such that dissolved oxygen levels
dropped to less than 60% of saturation.
15.14.1.8 Survival of organisms in control treatments should be assessed during each test as an indication
of both the validity of the test and the overall health of the test organism population. Tests are acceptable
if mean control survival is greater than or equal to 90% and if survival in individual control test chambers
exceeds 80%.
16. CALCULATIONS
16.1 The calculating procedure(s) and interpretation of the results should be appropriate to the
experimental design.
16.2 Procedures used to calculate results of toxicity tests can be divided into two categories: those that
test hypotheses and those that provide point estimates.
16.3 No procedure should be used without careful consideration of a) the advantages and disadvantages
of various alternative procedures and b) appropriate preliminary tests, such as those for outliers and for
heterogeneity. Preprocessing of data may be required to meet the assumptions of the analyses.
16.4 LC50 or EC50 and their 95% confidence limits should be calculated based on (a) the measured initial
concentrations of test material, if available, or the calculated initial concentrations, and (b) percent mortality.
If other LCs or ECs are calculated, their 95% confidence limits should also be calculated (see ASTM
Practice E 729).
16.5 Most acute toxicity tests produce quantal data, i.e., counts of the number of organisms in two mutually
exclusive categories, such as alive or dead.
16.5.1 A variety of methods (Litchfield and Wilcoxon, 1949; Finney, 1964; Finney, 1971; Stephan, 1977;
Hamilton et a/., 1977) can be used to calculate an LC50 or EC50 and its 95% confidence limits from a set
of quantal data that is binomially distributed and contains two or more concentrations at which the percent
dead or affected is between 0 and 100, but the most widely used are the probit, moving average, trimmed
Spearman-Karber and Litchfield-Wilcoxon methods.
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16.5.2 The method used should appropriately take into account the number of test chambers per treatment
and the number of test organisms per chamber. The binomial test can usually be used to obtain statistically
sound information about the LC50 or EC50 even when less than two concentrations kill or affect between
0 and 100 percent.
16.5.3 The binomial test does not provide a point estimate of the LC50 or EC50, but it does provide a
range within which the LC50 or EC50 should lie.
16.6 The results of toxicity tests on field samples without replication may be reported in terms of survival
values. A sample should be considered to be toxic if the single sample value lies outside the 95% tolerance
limits of the survival of the controls.
16.6.1 Alternately, the field result may be compared with the control survival data using outlier detection
methods; the sample may be considered toxic if it would be rejected as an extreme value when considered
as part of the control population.
16.6.2 Another approach is to use the "special case" comparison of a single value against a sample,
described by Sokal and Rohlf. It is strongly recommended that samples be replicated if comparisons
among sites are desired.
16.7 If samples from field stations are replicated, the mean survival at the stations and the mean control
survival should be statistically compared by a one-tailed t-test or analysis of variance (ANOVA) followed by
a multiple comparison test.
16.7.1 Analysis of variance is used to determine whether any of the observed differences among the
concentrations (or samples) are statistically significant. This is a test of the null hypothesis of no difference
among concentrations (or samples).
16.7.2 If the F-test is not statistically significant (P>0.05), it can be concluded that the effects observed in
the toxicant treatments (or field station samples) were not large enough to be detected as statistically
significant by the experimental design and hypothesis test used.
16.7.3 Following a significant F-test result, all exposure concentration effects (or field station samples) can
be compared with the control effects by using mean separation techniques such as those explained by
Chew (1977) orthagonal contrasts, Fisher's methods, Dunnett's procedure and Williams' method.
16.8 The Dunnett's procedure is a multiple comparison test specifically designed to compare several
experimental samples to the concurrent control (Gelber et a/., 1985). A multiple comparison test is a
technique that accounts for the fact that several comparisons are being made simultaneously.
16.9 Daily observations on the number of amphipods which have completely or partially emerged from the
sediment, either lying on the sediment surface, swimming in the water column, or floating at the water
surface, can be used to document an apparent avoidance response to the sediment. Emergence data
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plotted against time can give the observer an impression of the degree of toxicity of the sediment during
the course of the toxicity test, as amphipods often emerge earlier and in greater numbers from more highly
toxic sediment.
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APPENDIX A
DILUTION/CULTURE WATER FOR Ampelisca abdita
A.1 GENERAL REQUIREMENTS
A.1.1 Besides being available in adequate supply, water used in toxicity tests should be acceptable to test
organisms and the purpose of the test. The minimum requirement for acceptable water for use in acute
toxicity tests is that healthy test organisms survive in the water, and in the water with sediment for the
duration of holding and testing without showing signs of disease or apparent stress such as unusual
behavior, changes in appearance, or death. The water in which the test organisms are held prior to the test
should be uniform in quality in that the concentration of contaminants and the range of temperature and
salinity encountered during the holding period do not adversely affect the survival of the test organisms in
the holding tanks or in the control treatments during the test.
A.2 SOURCE
A.2.1 Natural Saltwater- If natural saltwater is used, it should be obtained from an uncontaminated area
known to support a healthy, reproducing population of the test species or a comparably sensitive species.
The water intake should be positioned to minimize fluctuations in quality and the possibility of
contamination, and to maximize the concentration of dissolved oxygen to help ensure low concentrations
of sulfide and iron. A specially designed system might be necessary to obtain salt water from a natural
water source. These precautions are intended to ensure that test organisms are not apparently stressed
by water quality during holding, acclimation and testing and that water quality does not unnecessarily affect
test results.
A.2.2 Reconstituted Salt Water - Reconstituted salt water can be prepared by adding a commercially
available sea salt or specified amounts (Table 1, see ASTM Standard E 729) of reagent-grade chemicals
(American Chemical Society, 1968) to high quality water with (a) conductivity less than 1 uS/cm and (b)
either total organic carbon (TOC) less than 2 mg/L or chemical oxygen demand (COD) less than 5 mg/L.
Acceptable water can usually be prepared using properly operated deionization or distillation units.
Reconstituted salt water should be intensively aerated before use, and aging for one to two weeks might
be desirable. If a residue or precipitate is present, the solution should be filtered before use. The water
should meet the criteria given in Subsection A.1.1.
A.2.3 Chlorinated water must never be used in the preparation of water for toxicity tests because residual
chlorine and chlorine-produced oxidants are highly toxic to many aquatic animals (U.S. EPA 1985).
Dechlorinated water should be used only as a last resort because dechlorination is often incomplete.
Municipal drinking water is not recommended for use because in addition to residual chlorine,it often
contains unacceptably high concentrations of metals, and quality is often highly variable (see ASTM
Standard E 729).
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A.3 PREPARATION
A.3.1 Seawater used in the sediment toxicity test should be passed through a filter effective to 5 um or less
to remove suspended particles and organisms from the water. Water that might be contaminated with
facultative pathogens should be passed through a properly maintained ultraviolet sterilizer (National
Academy of Sciences and National Academy of Engineering, 1973; Federal Register, 1980; Federal
Register, 1985) or a filter effective to 0.45 um or less.
A.3.1.1 If necessary, the salinity should be reduced by diluting the seawater with high quality deionized or
distilled water (see Subsection A.2.2). Salinity can be raised by addition of clean filtered oceanic water or
prepared brine. Common practice is to use a 60 to 90 %o saltwater brine. Such brines have been
successfully prepared using slow, heat-concentration of natural salt water, or by the addition of artificial sea
salts or reagent grade (Americal Chemican Society, 1968) salts to a natural salt water (see Subsection
A.2.2).
A.3.2 Fresh seawater used in the test should be prepared within two days of the test and stored in clean,
covered containers at 4 ± 3°C until sediment and water are added to the test chambers. It might be
necessary to age reconstituted seawater for one to two weeks before use. Sufficient water should be
prepared at one time for all test chambers. Additional water might be required for sieving control sediment
to adjust salinity or for holding the test amphipods prior to the test.
A.3.3 For certain applications the experimental design might require use of seawater from the test sediment
collection site. In other instances, experimental treatments might involve manipulation of the test seawater
conditions.
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TABLE 1. Reconstituted Salt Water (Kesteref a/., 1967; Zaroogian et a/., 1969; Zilliouxef a/., 1973) for
marine and estuarine crustaceans. Add the following reagent-grade (American Chemical Society, 1968)
chemicals in the amounts and order listed to 890 ml of water. Each chemical must be dissolved before the
next is added.A
Chemical Amount
NaF 3 mg
SrCI2- 6H2O 20 mg
H3BO3 30 mg
KBr 100mg
KCI 700 mg
CaCI2-2H2O 1.47g
Na2SO4 4.00 g
MgCI2-6H2O 10.78 g
NaCI 23.50 g
Na2SiCy 9H2O 20 mg
NaHCO, 200 mg
A If the resulting solution is diluted to 1 L, the salinity should be 34 ± 0.5 %o and the pH 8.0 ± 0.2. The
desired test salinity is attained by dilution at time of use. The reconstituted salt water should be stripped
of trace metals (Davey et a/., 1970).
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APPENDIX B
TEST ORGANISM: Ampelisca abdita
B.1 ECOLOGY
B.1.1 Ampelisca abdita is a tube-dwelling amphipod belonging to the family Ampeliscidae, found mainly
in protected areas from the low intertidal zone to depths of 60 m. It ranges from central Maine to south-
central Florida and the eastern Gulf of Mexico (Bousfield, 1973; Mills, 1964), and has also been introduced
into San Francisco Bay (Nichols and Thompson, 1985). It is euryhaline, and has been reported in waters
which range from fully marine to 10%o salinity (Mills, 1967). This species generally inhabits sediments from
fine sand to mud and silt without shell, although it may also be found in relatively coarser sediments with
a sizable fine component (Mills, 1967). A. abdita is often abundant in sediments with a high organic content
(Santos and Simon, 1980).
B.1.2 In the colder waters of its range, A. abdita produces two generations per year, an overwintering
generation which breeds in the spring and a second which reproduces in mid to late summer (Nichols and
Thompson, 1985; Mills, 1967). In New England, breeding of the overwintering generation begins when the
water temperature is about 8°C, but in warmer waters south of Cape Hatteras, breeding may be continuous
throughout the year. Adults mate in the water column, and intense breeding activity is correlated with the
full moon and spring tides. Juveniles are released after approximately two weeks in the brood pouch, at
about 1.5mm in length. It then takes 40 to 80 days for newly released juveniles to become breeding adults
(Mills, 1967). Where A. abdita are present, they are often dominant members of the benthic community with
densities up to 110,000 m~2 (Nichols and Thompson, 1985; Stickney and Stringer, 1957; Santos and Simon
1980). Ampelisca abdita is a particle feeder, feeding both on particles in suspension and on those from the
surface of the sediment surrounding its tube. Gut contents of field-collected specimens have been found
to include algal material, sediment grains, and organic detritus (Bousfield, 1973; Nichols and Thompson,
1985).
B.2 COLLECTION AND HANDLING TECHNIQUES
B.2.1 Ampelisca should be sieved from their native (collection site) sediment as soon as possible after
collection. A 2-mm mesh sieve nesting over a 0.5-mm mesh sieve is useful for this procedure. It is
desirable for the sediment containing the amphipods to be rinsed first through the upper, 2-mm, sieve with
a forceful stream of seawater at the collection temperature and salinity. This will break up the sediment
material and will also force most of the amphipods out of their tubes.
B.2.1.1 The material thus retained on the 0.5-mm sieve should be vigorously shaken and swirled so the
fine sediments pass through and the amphipods are separated from tubes, sediment, and detrital material.
If the sieve is then lifted from the water, allowed to drain, and then slowly lowered into a shallow tray of
seawater, the Ampelisca will be caught on the water's surface tension and can be easily collected with a
fine mesh dip net.
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B.2.1.2 The amphipods can be held temporarily in large culture dishes in a constant temperature bath, and
then separated into two size classes with the use of nested 1.0- and 0.5-mm sieves.
B.2.2 During acclimation, 300-350 Ampelisca can be held in 4-L jars, each containing approximately a 4-cm
deep layer of sieved collection site sediment. If seawater is flowing through the holding containers, a
screened overflow must be used to prevent loss of swimming amphipods.
B.2.3 Amphipods should have food available on a daily basis during acclimation. Research is currently
being conducted to determine optimal food sources for culturing this amphipod. Reasonable growth and
reproduction have been obtained when A. abdita has been fed the diatom Phaeodactylum tricornutum daily
in excess (a suggested amount is 0.5 to 1 L of algae per 4-L jar, or 14 X 106 cells/ml (U.S. EPA, 1994)).
Skeletonema costatum has also been used successfully. Amphipod exposure to the food source will be
increased if, during the feeding period (e.g., overnight), the holding system is static, with aeration to
circulate the algae. Sloping upper sides on the holding containers will aid in movement of algae across the
sediment surface. Photoperiod should be maintained at 16 h light: 8 h dark.
B.2.4 Care should be taken to maintain the temperature with a water bath when seawater is not flowing
through the jars. Approximate density in the holding jars should not exceed 300 amphipods (or one
amphipod per cm2 surface area). Acclimation to the test temperature should not exceed 3°C per day,
amphipods should be used within ten days after collection, and they must be acclimated to laboratory
conditions for at least 48 hours.
B.2.5 Ampelisca abdita may be shipped if this is done within one day of collection. Small plastic
"sandwich" containers (approximately 500 ml) can be used to hold the amphipods. The containers are filled
three-quarters full with a minimum depth of 2 cm of sieved collection site sediment and then to the top with
well-aerated seawater.
B.2.6 No more than 200 amphipods should be added to each container. Amphipods should be allowed
to burrow into the sediment and build tubes before the containers are capped. The capping must be done
underwater to eliminate any air pockets in the containers. Containers should be shipped via overnight
delivery in coolers with a few ice packs to prevent extreme temperature changes during transit.
B.3 OTHER TESTING WITH Ampelisca abdita
B.3.1 Growth of Ampelisca abdita has been measured in 10-day tests; the amphipods must be fed during
the test. Small juveniles in a narrow size range (between 2 and 4 mm in length) should be selected for
testing, and when sorting for the initiation of the test at least one additional group of amphipods should be
sorted. This extra group represents the initial size and should be preserved in 5% buffered formalin for later
measurement at a convenient time. Using a dissecting microscope, length is measured from the base of
the first antennae to the base of the telson.
B.3.2 Chronic tests have also been conducted with this species (Scott and Redmond, 1989) and research
is underway to determine the optimum conditions for those tests.
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APPENDIX C
TEST ORGANISM: Mysidopsis bahia
C.1 NATURAL HISTORY
C.1.1 The estuarine species Mysidopsis bahia, a small (adults, 5 to 8 mm, total length) shrimp-like
crustacean, belongs to the order Mysidacea and is frequently found along the Gulf Coast from southern
Florida (Everglades) to Mexico (San Geronimo). Mysids are found in most bays and estuary systems which
range in salinity from 8-25%o. They may be collected adjacent to tidal marshes or among marsh plants
(Juncus or Spartina) during high tides.
C.1.2 The relative ease with which mysids can be cultured is a contributing factor to their routine use as
an indicator organism. The sexes are separate and externally dimorphic. When sexually mature, females
are easily identified with the use of magnification by their brood pouches. Through a dissecting microscope,
males can be identified by the presence of claspers.
C.2 HANDLING AND CULTURING TECHNIQUES
C.2.1 Holding tanks Temperature-controlled, all-glass aquaria with flow-through or recirculating seawater
maintained at 25 ±2°C.
C.2.2 Aeration Oil-free forced air.
C.2.3 Light A 14-hour light and 10-hour dark photoperiod is established with cool-white fluorescent lights
adjusted to approximately 1200 microwatts/cm2.
C.2.4 Water Natural, filtered (< 20 ^m) seawater or artificial seawater formulated by the addition of sea
salts to deionized water and adjusted to a salinity of 20 to 25%o.
C.2.5 Mysids are fed live Artemia salina nauplii at the rate of approximately 100 Artemia nauplii per mysid
per day; the Artemia nauplii must be < 36 hours old.
C.2.6 To provide an adequate supply of juveniles for a test, mysid cultures should be started at least four
weeks before the test animals are needed. At least 200 mysids should be placed in each culture tank (a
2:1 female to male ratio) to ensure 1000 animals will be available by the time preparations for the test are
initiated.
C.2.7 Newly released juveniles are collected by siphoning water from holding tanks through nylon screen
chosen to allow only the passage of juveniles which are concentrated in a small mesh cage. Juveniles are
transferred to holding tanks where gradual acclimation to test conditions occurs. Within a 24-hour period,
changes in water temperature should not exceed 2°C, while salinity changes shall not exceed 5 %o). During
acclimation mysids should be maintained in facilities with background colors and light intensities similar to
those of the testing areas.
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APPENDIX D
TEST ORGANISM: Penaeus SPP.
D.1 NATURAL HISTORY
D.1.1 Penaeid larvae develop at sea and appear to be most active at night, moving vertically through the
water column. Once they have reached the post-larval stage and migrated into the nursery areas, the
animals become totally benthic and remain active both day and night. Penaeids are omnivorous, eating
zooplankton, fragments of higher plants, and ingesting algae as well as sand, mud and organic debris.
D.1.2 Some penaeids can be laboratory cultured, but not with ease. Spawning can be induced in the
laboratory by artificially stimulating egg production through the coupling of regulated lighting and
temperature with eye stalk oblation of adult females.
D.2 HANDLING TECHNIQUES AND HOLDING CONDITIONS FOR POST-LARVAE OR JUVENILES
D.2.1 Holding tanks Temperature-controlled, all-glass aquaria with flow-through or recirculating seawater
maintained at 25 + 2°C.
D.2.2 Aeration Oil-free forced air.
D.2.3 Light A 12-hour light and 12-hour dark photoperiod is established with cool-white fluorescent lights
adjusted to approximately 1200 microwatts/cm2.
D.2.4 Water Natural, filtered (< 20 //m) seawater or artificial seawater formulated by the addition of
seasalts to deionized water and adjusted to a salinity of 20 to 25%o.
D.2.5 Penaeid post-larvae will be fed Artemia. Juveniles will be fed commercial fish flake.
D.2.6 Laboratory-reared, post-larvae will be generated several weeks prior to the initiation of the test and
maintained at the laboratory facility.
D.2.7 Post-larval batches of penaeids will be segregated by age and used for Louisianian Province ITE
samples. These samples will arrive randomly throughout the testing period.
D.2.8 If laboratory-reared post-larvae are not available, laboratory-reared juveniles or wild-caught Penaeus
sp. will be used after an acclimation period of at least four days.
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17. REFERENCES
Adams, W.J., R.A. Kimerle, and R.G. Mosher. 1985. In Cardwell, Purdy, and Bahner, eds., Aquatic
Toxicology and Hazard Evaluation: Seventh Symposium, ASTM STP 854, American Society for Testing
and Materials, Philadelphia pp. 429-453.
American Chemical Society, 1968 Reagent Chemicals: American Chemical Society Specifications.
Washington, DC.
ASTM. 1991. Guide for conducing 10-day static sediment toxicity tests with marine and estuarine
amphipods. ASTM Standard Methods Volume 1104, Method Number E-1367-90.
Bousfield, E.L.1973. Shallow-water Gammaridean Amphipoda of New England, Cornell University Press,
Ithaca, New York.
Breteler, R. J., K. J. Scott, and S. P. Shepurd. 1989. Application of a new sediment toxicity test using the
marine amphipod Ampelisca abdita to San Francisco Bay sediments. In: U. M. Cowgill and L. R.
Williams (eds.), Aquatic Toxicology and Hazard Assessment, Volume 12, American Society for Testing
and Materials, Philadelphia, PA.
Carmignani, G.M. and J.P. Bennett. 1976. Leaching of Plastics Used in Closed Aquaculture Systems,
Aquaculture. 7:89-91.
Chapman, P. M. 1988. Marine sediment toxicity tests. In: J. J. Lichtenberg, J. A. Winter, C. I. Weber and
L. Fredkin (eds.), Chemical and Biological Characterization of Sludges, Sediments, Dredge Spoils, and
Drilling Muds. American Society for Testing and Materials, Philadelphia, PA.
Chew, V. 1977. Comparisons Among Treatment Means in an Analysis of Variance, ARS/H/6, Agricultural
Research Service, U. S. Department of Agriculture.
Davey, E.W., J.H. Gentile, S.J. Erickson, and P. Betzer. 1970. Removal of Trace Metals from Marine
Culture Media. Limnology and Oceanography. 15:486-488.
DiToro, D. M., J. D. Mahony, D. J. Hansen, K. J. Scott, M. B. Hicks, S. M. Mayr, M. S. Redmond. 1990.
Toxicity of cadmium in sediments: the role of acid volatile sulfide. Environmental Toxicology and
Chemistry, 9:1487-1504.
Fawcett, H.H., and W.S. Wood eds. 1982. Safety and Accident Prevention in Chemical Operations, 2nd
Ed., Wiley-lnterscience, New York, NY.
Federal Register, 1980. Vol 45, November 28, 1980, pp. 79318-79379.
Federal Register, 1985. Vol 50, July 29, 1985, pp. 30784-30796.
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Finney, D.J., 1964 Statistical Method in Biological Assay. 2nd ed., Hafner Publishing Co., New York, NY,
668 pp.
Finney, D.J., 1971 Probit Analysis, 3rd ed., Cambridge University Press, London, 333 pp.
Gelber, R.D., P.T. Lavin, C.R. Mehta and D.A. Schoenfeld. 1985. Fundamentals of Aquatic Toxicology:
Methods and Applications. McGraw-Hill.
Goselin, R.E., H.C. Hodge, R.P. Smith, and M.N. Gleason. 1976. Clinical Toxicology of Commercial
Products. 4th Ed., Williams and Wolkins Co., Baltimore, MD.
Green, N.E., and A. Turk. 1978. Safety in Working with Chemicals. MacMillan, New York, NY.
Hamilton, A. and H.L Hardy. 1974. Industrial Toxicology. 3rd Ed., Publishing Sciences Group, Inc., Acton,
MA.
Hamilton, M.A., R.C. Russo, and R.V. Thurston. 1977. Trimmed Spearman-Karber Method for Estimating
Median Lethal Concentrations in Toxicity Bioassays. Environmental Science and Technology. 11:714-
719. Correction Vol. 12, 1978. p. 417.
Heitmuller, T. and R. M. Valente. 1992. EMAP-Estuaries 1992 Louisianian Province Quality Assurance
Project Plan. U.S. Environmental Protection Agency, Office of Research and Development,
Environmental Research Laboratory, Gulf Breeze, FL.
International Technical Information Institute. 1977. Toxic and Hazardous Industrial Chemicals Safety
Manual. Tokyo, Japan.
Kester, D.R., I.W. Duedall, D.N. Connors, and R.M. Pytkowicz. 1967. Preparation of Artificial Seawater.
Limnology and Oceanography. 12:176-179.
Litchfield, J.T., Jr., and F. Wilcoxon. 1949. A Simplified Method of Evaluating Dose-Effect Experiments.
Journal of Pharmacology and Experimental Therapeutics. 96:99-113;
Long, E. R. and M. F. Buchman. 1989. An evaluation of candidate measures of biological effects for the
National Status and Trends Program. NOAA Technical Memorandum.
Macauley, J. 1991. Near Coastal Lousianian Province 1991 Demonstration Project Field Operations
Manual. U.S. Environmental Protection Agency, Office of Research and Development, Environmental
Research Laboratory, Gulf Breeze, FL.
Mills, E.L. 1964. Ampelisca abdita, a New Amphipod Crustacean from Eastern North America. Canadian
Journal of Zoology. 42:559-575.
Mills, E.L. , 1967The Biology of an Amphipod Crustacean Sibling Species Pair. Journal of the Fisheries
Research Board of Canada. 24:305-355.
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National Academy of Sciences and National Academy of Engineering. 1973. Water Quality Criteria. EPA-
R3-73-033. National Technical Information Service, Springfield, VA, pp. 172-193.
National Council on Radiation Protection and Measurement. 1971. Basic Radiation Protection Criteria.
NCRP Report No. 39, Washington, DC.
National Institutes of Health. 1981. NIH Guidelines for the Laboratory Use of Chemical Carcinogens. NIH
Publication No. 81-2385, Bethesda. MD.
National Research Council. 1981. Prudent Practices for Handling Hazardous Chemicals in Laboratories.
National Academy Press, Washington, DC.
Nichols, F.H. and J.K. Thompson. 1985. Persistence of an Introduced Mudflat Community in South San
Francisco Bay, California. Marine Ecology Progress Series. 24:83-97.
Patty, F.A., ed. 1963. Industrial Hygiene and Toxicology. Vol II. 2nd Ed., Interscience, New York, NY.
Rand, G.M. and S.R. Petrocelli. 1985. Fundamentals of Aquatic Toxicology: Methods and Applications.
McGraw-Hill.
Robinson, A.M., J.O. Lamberson, F.A. Cole, and R.C. Swartz. 1988. Effects of Culture Conditions on the
Sensitivity of the Phoxocephalid Amphipod, Rhepoxynius abronius, to Cadmium in Sediment.
Environmental Toxicology and Chemistry. 7:953-959.
Rogerson, P. F., S. C. Schimmel, and G. Hoffman. 1985. Chemical and biological characteristics of Black
Rock Harbor dredged material. U. S. Army COE Tech. Report D-85-9. Vicksburg, MS.
Santos, S.L. and J.L. Simon. 1980. Response of Soft-bottom Benthos to Catastrophic Disturbance in a
South Florida Estuary. Marine Ecology Progress Series. 3:347-355.
Sax, N.I. 1984. Dangerous Properties of Industrial Materials. 6th Ed., Van Nostrand Reinhold Co., New
York, NY.
Scott, K.J. and M.S. Redmond. 1989. The Effects of a Contaminated Dredged Material on Laboratory
Populations of the Tubicolous Amphipod, Ampelisca abdita. In Cowgill, U.M. and L.R. Wlliams, eds.
Aquatic Toxicology and Hazard Assessment: Twelveth Volume, ASTM STP 1027. American Society
for Testing and Materials, Philadelphia, pp. 289-303.
Shapiro, J. 1981. Radiation Protection. 2nd Ed., Harvard University Press, Cambridge, MA.
Stephan, C.E. 1977. Methods for Calculating an LC50. In Mayer, F.L. and J.L. Hamelink, eds. Aquatic
Toxicology and Hazard Evaluation. ASTM STP 634. American Society for Testing and Materials,
Philadelphia, pp. 65-84.
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Stickney, A.P. and L.D. Stringer. 1957. A Study of the Invertebrate Bottom Fauna of Greenwich Bay,
Rhode Island. Ecology. 38:111-122.
Strobel, C. and S.C. Schimmel. 1991. Near Coastal Virginian Province Field Operation and Safety
Manual. Environmental Research Laboratory, Office of Research and Development, U.S.
Environmental Protection Agency, Narragansett, Rl.
Swartz, R. C. 1987. Toxicological methods for determining the effects of contaminated sediment on
marine organisms, pp. 183-198. In: K. L. Dickson, A. W. Maki, and W. A. Brungs (eds.) Fate and
Effects of Sediment Bound Chemicals in Aquatic Systems. Pergamon Press, New York.
Swartz, R. C. 1989. Marine sediment toxicity tests, pp. 115-129, In: Contaminated Sediments-
Assessment and Remediation. National Academy Press, Washington, DC.
Swartz, R.C., W.A. DeBen, and F.A. Cole. 1979. A Bioassay for the Toxicity of Sediment to Marine
Macrobenthos. Journal of the Water Pollution Control Federation. 51:944-950.
Swartz, R.C., W.A. DeBen, J.K.P., Jones, J.O., Lamberson, and F.A. Cole. 1985. Phoxocephalid
Amphipod Bioassay for Marine Sediment Toxicity. In: Cardwell, R.D., R. Purdy, and R.C. Bahner,
Eds., Aquatic Toxicology and Hazard Assessment: Seventh Symposium, ASTM STP 854, American
Society for Testing and Materials, Philadelphia, pp. 284-307.
U.S. EPA/CE. 1977. Ecological Evaluation of Proposed Discharge of Dredged Material into Ocean
Waters. Technical Committee on Criteria for Dredged and Fill Material. Environmental Effects
Laboratory, U.S. Army Engineer Waterways Experiment Station, Vicksburg, MS.
U.S. EPA. 1978. Bioassay procedures for the ocean disposal permit program. Environmental
Research Laboratory, Gulf Breeze, FL. 600/9-78-010.
U.S. EPA. 1979a. Methods for chemical analysis of water and wastes. U.S. Environmental Protection
Agency, Environmental Monitoring and Support Laboratory, Cincinnati, Ohio, EPA-600/4-79/020,
revised March 1983.
U.S. EPA. 1979b. Handbook for analytical quality control in water and wastewater laboratories. U.S.
Environmental Protection Agency, Environmental Monitoring and Support Laboratory, Cincinnati,
Ohio, EPA/600/4-79/019.
U.S. EPA. 1985. Ambient Aquatic Life Water Quality Criteria for Chlorine-1984. EPA 440/5-84-030,
National Technical Information Service, Springfield, VA.
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U.S. EPA. 1987. Short-term methods for estimating the chronic toxicity of effluents and receiving
waters to marine and estuarine organisms. Environmental Monitoring and Support Laboratory,
Cincinnati, OH. EPA/600/4-87/028.
U.S. EPA. 1994. Methods for Assessing the Toxicity of Sediment-associated Contaminants with
Estuarine and Marine Amphipods. EPA/600/R-94/025. United States Environmental Protection
Agency. Office of Research and Development. Washington, DC 20460.
Valente, R. M. and C. J. Strobel. 1992. EMAP-Estuaries 1992 Virginian Province Quality Assurance
Project Plan. U.S. Environmental Protection Agency, Office of Research and Development,
Environmental Research Laboratory, Narragansett, Rl.
Walters, D.B., ed. 1980. Safe Handling of Chemical Carcinogens, Mutagens, Teratogens and Highly
Toxic Substances. Ann Arbor Science, Ann Arbor, Ml.
Zaroogian, G.E., G. Pesch, and G. Morrison. 1969. Formulation of an Artificial Sea Water Media
Suitable for Oyster Larvae Development. American Zoologist. 9:1144.
Ziegenfuss, P.S., W.J. Renaudette, and W.J. Adams. 1986. In: Poston and Purdy, eds., Aquatic
Toxicology and Environmental Fate: Ninth Volume, ASTM STP 921, American Society for Testing
and Materials, Philadelphia, pp. 479-493.
Zillioux, E.J., H.R. Foulk, J.C. Prager, and J. Cardin. 1973. Using Artemia to Assay Oil Dispersant
Toxicities. Journal of the Water Pollution Control Federation. 45:2389-2396.
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SECTION 3
BENTHIC MACROINVERTEBRATE METHODS
MACROBENTHIC COMMUNITY ASSESSMENT
TABLE OF CONTENTS
Subsections
Introduction 2
Laboratory Safety 2
Training 3
Sample Storage and Treatment 4
Sample Sieving, Sorting, Species Identification, and Enumeration 5
Macrofaunal Biomass Determination 11
Quality Assurance and Quality Control 14
Data Management 19
Data Forms 19
References 20
Appendix A 23
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1. INTRODUCTION
1.1 This chapter describes the laboratory procedures used by the EMAP Estuaries Resource Group to
measure the species composition, abundance, and biomass of macroinvertebrate fauna found in estuarine
sediments.
1.2 Estuarine bottom-dwelling (benthic) assemblages have many attributes that potentially make them
reliable and sensitive indicators of ecological condition and pollution stress (Boesch and Rosenberg 1981;
Bilyard 1987). First, benthic organisms have limited mobility and cannot avoid exposure to pollutants and
adverse conditions. Second, benthos live in bottom sediments where chemical contaminants accumulate
and stress from low dissolved oxygen is most severe (e.g., Mirza and Gray 1981; Chapman et al. 1987).
Third, benthic assemblages are taxonomically diverse and include multiple feeding modes and trophic
levels. As a result, they display a broad range of physiological tolerances and responses to multiple types
of stress (Pearson and Rosenberg 1978; Rhoads et al. 1978; Sanders et al. 1980; Boesch and Rosenberg
1981; Swartz et al. 1986). Finally, benthic organisms are important links between primary producers and
higher trophic levels including economically important species (Virnstein 1977; Holland et al. 1980), and
their feeding and burrowing activities affect oxygen, carbon, nutrient, and sediment cycles (Rhoads and
Young 1970; Rhoads 1974; Cloern 1982).
1.3 The procedures outlined in this chapter are designed to produce data of consistent quality meeting the
measurement quality objectives (MQO) of 10% total error for the extraction of organisms from sediments
and 10% total error for the identification and enumeration of extracted fauna. To ensure that these
objectives are met, specific quality assurance steps are included as part of these procedures.
1.4 Laboratory procedures are based upon currently accepted practices in benthic ecology (Holmes and
Mclntyre 1984). Changes are not permitted without prior approval from the EMAP-Estuaries Technical
Director. Such changes will be incorporated into subsequent versions of this manual in a timely manner.
1.5 The fixative for EMAP benthic samples will be formalin in all cases. Samples will be preserved in a
10% formalin solution for at least one month prior to processing. Subsequent preservation in a 70% ethanol
solution is recommended.
1.6 These procedures assume that benthic samples arrive at the laboratory in pre-labeled, plastic screw-top
jars, preserved in a rose bengal stained buffered, 10% formalin solution. Procedures used to prepare
benthic samples in the field are given in the appropriate EMAP Estuaries Field Operations Manual and will
not be repeated here.
2. LABORATORY SAFETY
2.1 Safe laboratory practices must be followed at all times, and should be outlined in a laboratory safety
manual that is kept in the laboratory at all times.
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2.2 Eye protection (i.e., laboratory safety glasses or approved prescription glasses) must be worn when
pouring solvents or preservatives (e.g., formalin) handling chemicals, or when performing any procedures
that require use of a fume hood. Safety glasses are available in the laboratory.
2.3 Repeated exposure to formaldehyde can increase sensitivity to this potentially harmful chemical.
Exposure should be minimized by handling formaldehyde and preserved samples with gloves, conducting
all work involving formaldehyde under a fume hood, using portable ventilation devices over microscopes
when sorting preserved samples, and storing samples in a well-ventilated room away from areas in which
personnel are working.
2.4 Waste Disposal
2.4.1 No chemicals are to be disposed through laboratory drains leading to a municipal or private sewer
or septic system. Chemicals must be disposed in appropriate containers available in the laboratory.
2.4.2 Diluted formaldehyde may, in certain cases, be disposed through laboratory sewer lines. Permitted
discharge volumes and concentrations will depend upon state and local regulations.
2.5 Work areas are to be kept clean and neat at all times.
2.6 Any conditions that hinder the accomplishment of work or represent a possible safety hazard should
be brought to the attention of the laboratory supervisor immediately.
3. TRAINING
3.1 A program the size of EMAP-Estuaries involves many field and laboratory technicians. Standard
procedures must be followed to ensure that data produced by multiple laboratories and different personnel
are comparable.
3.2 All laboratory personnel will have a minimum basic level of training. Instruction will include evaluation
and proficiency testing to insure the mastery of skills.
3.3 Training will be provided by experienced personnel in established laboratories. New employees will
learn laboratory techniques using practice or "dummy" samples and in no case will handle, without
supervision, real EMAP samples without prior demonstration of acceptable proficiency.
3.4 The overall proficiency of laboratory personnel will be continuously evaluated using the various quality
assurance and quality control procedures outlined in the methodologies given below. The status of
personnel demonstrating substandard performance will be reevaluated, and, if necessary, personnel will
be retrained or removed from the laboratory to maintain the consistent, high quality production of data.
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4. SAMPLE STORAGE AND TREATMENT
4.1 Sample processing starts with the reception of samples from field crews. Samples are bar coded with
a unique sample number. Upon receipt, a bar code reader will be used to log samples into computer files;
sample identification numbers will be checked against a master list provided by the EMAP-Estuaries
Province Field Operations Center. In addition, the data files containing the records of all samples received
by the laboratory will be transferred to the Province Field Operations Center at regular intervals for the
purpose of sample tracking. Discrepancies, missing, or damaged samples will be reported to the Province
Manager and/or Field Coordinator immediately.
4.2 Within the laboratory, all samples will be carefully tracked by sample number using a sample log. It
will be the responsibility of each laboratory to keep accurate and timely records of the location and status
of all samples in their custody.
4.3 Stored samples must be easily retrieved and protected from environmental extremes. Samples cannot
be allowed to freeze and should be stored above 5°C to prevent the formation of paraformaldehyde (Jones
1976). Temperatures greater than about 30°C should be avoided to retard evaporative losses. Stored and
archived samples should be checked once every three months to avoid excessive evaporative losses due
to loosely fitting or cracked container lids or inadequately sealed jars. Exposure to direct sunlight should
be minimized since long-term exposure can degrade the vital rose bengal stain.
4.4 Previously fixed samples that are being processed in the laboratory should be represerved in a 70
percent (volume/volume) ethanol solution. Samples should not remain in water without preservative for
more than 24 hours since microbial degradation may occur. Degradation may impact subsequent biomass
measurements.
4.5 Differential Treatment of Samples Due to Salinity:
4.5.1 Sample treatment is dependent upon the salinity of the habitat from which samples were taken.
Benthic samples from tidal fresh and oligohaline salinity zones (0 to 5 %o) will be treated differently than
samples from mesohaline and polyhaline salinity zones (> 5 %o).
4.5.2 Oligochaetes and chironomids from tidal fresh and oligohaline salinity zones will be identified to
species, where possible, whereas individuals of these groups from higher salinities will not be further
differentiated.
4.5.3 The rationale for this inconsistent treatment of samples is that the additional taxonomic effort
necessary to identify oligochaetes and chironomids to the species level produces important and useful
information for the interpretation of benthic communities in tidal fresh and oligohaline regions, but produces
information of marginal value in mesohaline and polyhaline environments. This is largely due to the greater
diversity of these two groups in lower salinity environments.
4.5.4 Salinity categories (tidal fresh and oligohaline vs. mesohaline and polyhaline) will be determined
using bottom salinity values collected at the time benthic grabs were taken. Laboratory processing of
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benthic samples will not proceed without salinity data for the sampling station or area from which benthic
samples were collected.
4.5.5 In cases where the salinity for a sample is unknown and the species assemblage is insufficient to
place the sample in a specific salinity class, the sample will be treated as a tidal fresh/oligohaline sample.
This will insure that no potentially useful information will be lost due to the treatment of samples.
4.5.6 Sample replicates from the same station will be treated similarly. All replicates will be processed as
tidal fresh/oligohaline or as mesohaline/polyhaline/samples.
5. SAMPLE SIEVING, SORTING, SPECIES IDENTIFICATION, AND ENUMERATION
5.1 The objective of sorting benthic samples is to completely remove from sample debris all fauna of
interest that were alive at the time of collection. Sample debris includes primarily sediment, but also detritus
and the remnants (death assemblage) of the hard parts of various benthic organisms (for example, the
shells of bivalve mollusks or the exoskeletons of crustaceans).
5.2 Benthic Fauna of Interest
5.2.1 The fauna of interest to EMAP-Estuaries are benthic macrofauna operationally defined here as those
metazoan organisms retained by a 0.5 mm (500 urn) mesh sieve. All fauna retained on the 0.5 mm sieve
will be identified, enumerated, and included as macrofauna, except those meiofaunal and pelagic groups
specified in paragraphs below. No upper size limit for macrofauna will be used by the program.
5.2.2 Faunal groups, typically termed meiofauna, for which the majority of individuals are too small to be
retained on a 500 /^m sieve will not be identified, enumerated, or processed for biomass measurements.
These groups include the following:
Turbellarian flatworms
Kinorhynchs
Nematodes
Harpacticoid copepods
Cyclopoid copepods
Ostracods
Halacarids.
5.2.3 Faunal groups that are clearly pelagic and not benthic will not be identified, enumerated, or measured
for biomass. Examples of pelagic groups include the following:
Cladocerans
Calanoid copepods.
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5.2.4 Eggs or egg cases will not be identified, enumerated, or measured for biomass. Only individuals that
have fully separated from their eggs will be counted.
5.2.5 Flying insects that may have been collected in samples, will not be identified, enumerated, or
measured for biomass.
5.3 Sieving:
5.3.1 Benthic samples were previously sieved through 0.5 mm mesh sieves in the field to remove most
sediment particles less than 0.5 mm (see Field Operations and Methods Manual). All samples will be
resieved in the laboratory using 0.5 mm mesh sieves to ensure each sample is consistently and completed
processed. Samples will be sieved in the following step-wise manner:
5.3.2 Throughout laboratory processing, all samples will be tracked by the sample number given prior to
field sampling.
5.3.3 Sieves will be cleaned and backwashed thoroughly before processing each sample.
5.3.4 Under a fume hood, the sample will be poured through the sieve. The filtrate has a 3.7%
concentration of formaldehyde and will be saved in a properly labeled container for later disposal or reuse.
5.3.5 Using tap water, any portion of the sample remaining in the jar will be rinsed into the sieve.
5.3.6 The sieve containing the sample will be placed in a wash basin. The basin will be filled with water
and agitated to wash fine material through the sieve. This procedure minimizes mechanical damage to
fragile fauna. A gentle spray of water may also be used to wash material through the sieve, but direct,
heavy jets of water should not be used.
5.3.7 Using a gentle spray of water, material will be transferred from the sieve into a labeled container in
preparation for sorting.
5.3.8 Sieves will be examined after rinsing to ensure that all organisms have been removed and to
minimize cross contamination with the next sample.
5.4 Sorting:
5.4.1 All macrofauna that were alive at the time of collection must be separated and removed from organic
debris and sediment particles remaining after sieving. All organisms, including significant body fragments,
must be saved to obtain correct estimates of density and to make accurate biomass determinations.
Unidentifiable material must be saved until positively identified.
5.4.2 Sorting commences by pouring sieved material into white enamel or plastic trays for initial removal
of larger organisms. Finer material will be transferred to a grided petri dish for sorting using a
stereomicroscope. Samples must be evenly distributed over the tray or petri dish and the water level must
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be low enough to prevent sloshing back and forth as the tray or dish is moved.
5.4.3 Organisms will be removed as the tray or petri dish is systematically searched. Organisms will be
divided into major taxa, typically polychaetes, oligochaetes, bivalves, gastropods, crustaceans, etc.
5.4.4 Laboratory personnel completing the initial sorting normally do not complete identifications to the
species level. Species identifications are completed by taxonomists who have more training and experience
than sorters.
5.4.5 Organisms may be identified by taxonomists to species level immediately or may be saved for later
identification. Fauna will be sorted into major taxonomic groups, placed into small screw-top vials, and
preserved in ethanol. Vials will contain labels indicating the sample number and taxon. Vials from the
sample will be bound together with rubber bands.
5.4.6 The number of vials for each sample will be recorded in the sample log (sample tracking sheet) along
with the initials of the sorter completing this part of the laboratory processing.
5.4.7 Sample debris that remains after sorting will be transferred from the petri dish to a jar, preserved in
a 70% (volume/volume) ethanol solution, labeled, and for each technician, saved in batches of 10. A log
will be kept of all archived samples. Ten percent of each batch will be resorted as a quality control check
on each sorter's efficiency.
5.4.8 Sample debris in each sample batch may be discarded after quality control procedures have been
completed and approved.
5.4.9 Sorting efficiencies meeting or exceeding the measurement quality objective of 90% will be assured
by several measures: only qualified technicians trained to process EMAP-Estuaries samples will sort
samples; the sample sorting protocol will be documented and uniformly applied to all samples; and all
sorting will be closely and continuously monitored by supervisory staff. In combination, the application of
training, supervision, and controlled laboratory procedures will ensure that all samples will be processed
correctly, and that resulting data will not be invalidated by contamination, loss of vials, or incomplete
removal of organisms from the sample.
5.5 Species Identification and Enumeration:
5.5.1 The identification of biological specimens to the species level requires specialized taxonomic training,
experience, and a familiarity with current taxonomic literature. The validity of taxonomic identifications
affects the quality of subsequent population and community analyses, as well as the comparability of the
research to other studies; therefore, only qualified and experienced technicians will perform species
identifications. Typically, no one person can completely master the taxonomic complexities of all benthic
macrofaunal groups. For example, in any laboratory, one technician might develop an expertise in
identifying polychaetes; another may be better at identifying gastropods. All technicians will be trained to
recognize commonly occurring species. Rarer species will require the taxonomic expertise of specialists
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within the laboratory, and may, at times, require the assistance of expert taxonomists in other laboratories.
A list of taxonomic experts will be maintained by EMAP-Estuaries.
5.5.2 The objective of species identification and enumeration is to accurately identify all organisms found
in a sample to the lowest possible taxonomic category consistent with study objectives and to accurately
count the number of organisms in each taxon. For EMAP-Estuaries, specimens will be identified to the
species level whenever possible. The usefulness of such efforts in environmental monitoring programs is
well established (Grassle and Grassle, 1984).
5.5.3 Due to various taxonomic difficulties, certain groups expected to be encountered by EMAP-Estuaries
will not be identified to species. These groups and the associated level of expected taxonomic classification
include the following:
Degree of
Taxonomic Group Expected Classification
Phylum Nemertinea (Proboscis worms) Phylum
Phylum Sipuncula Phylum
Class Oligochaeta Class*
Class Hirudinea Class
Class Anthozoa Class
Family Chironomidae Family*
*ln marine, polyhaline, and mesohaline regions only. Identifications will be made to species level, if
possible, in oligohaline and tidal fresh regions.
5.5.4 The number of individuals counted for each taxon must reflect the number of organisms alive at the
time of sampling; therefore, when organism fragments are recovered, counts will be based upon only the
number of heads found. Posterior body fragments will not be counted, but must be retained with the
appropriate taxonomic group for biomass determinations. If only posterior fragments are present (no
heads), they will be counted as one individual unless a greater number of individuals can be positively
identified. In that case, the greater number of individuals will be recorded.
5.5.5 All identifications will be performed under a high quality dissecting microscope with sufficient
magnification for clear resolution of morphological details; a microscope with 5 to 50x power is usually
sufficient. On occasion, a compound microscope capable of higher magnification may be required and
should be available for use.
5.5.6 Wherever possible, different taxonomists will identify specimens in replicate samples taken at the
same station and time. Alternatively, the same taxonomist will process replicates on different days.
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5.5.7 Sample processing for identification and enumeration commences by retrieving sample vials for a
particular sample from the previous sorting procedure. At that time, a species identification data sheet is
started. The sample number on the vials will be checked with that recorded in the sample log; the number
of vials should match the number noted in the log.
5.5.8 Specimens from each vial will be rinsed into separate petri dishes. Care must be taken to remove
all specimens from the vial, both the vial and cap will be checked for remaining specimens.
5.5.9 Specimens will be identified, counted, and removed from the petri dish one at a time.
5.5.10 Identified organisms will be grouped by the categories to be used in biomass determinations
(Subsection 6.6), placed in vials, and preserved in 70% ethanol. Vials will be labeled with sample number
and biomass group. The appropriate biomass category will be marked with a check on the biomass data
sheet which accompanies each abundance data sheet.
5.5.11 Petri dishes will be thoroughly inspected for missed specimens and then will be rinsed to minimize
cross contamination between vials and samples.
5.5.12 Identified species will be checked for suspicious mismatches as part of the identification process.
An example of a suspicious mismatch is the identification of a freshwater benthic species in a sample from
a polyhaline salinity region.
5.5.13 Specimens that are difficult to identify will be set aside in vials and preserved in ethanol for future
study. Some specimens will require the expertise of more experienced technicians in the same laboratory.
Other specimens may require further laboratory processing (for example, oligochaetes and chironomids will
need to be mounted on microscope slides) before species determination can be made. Still other
specimens may need to be sent to other laboratories to complete species identifications. The location of
all specimens for a particular sample will be tracked using the species identification data sheet and the
laboratory sample log.
5.5.14 Treatment of Oligochaetes and Chironomids
5.5.14.1 In general, all specimens will be identified and enumerated from visual inspection using a
stereomicroscope. However, certain taxonomic groups will require special handling to optimize both species
identification and biomass determination. Those two groups are the Class Oligochaeta and the Family
Chironomidae. Specimens of both groups in samples from tidal fresh and oligohaline regions will need to
be inspected under high magnification using a compound microscope to properly complete species
identifications. Sample processing of oligochaetes and chironomids will proceed in the following manner
for each group.
5.5.14.2 For each sample and for each group (oligochaetes and chironomids), if less than 20 individuals
are found in a sample, all individuals will be permanently mounted on a microscope slide and identified to
the species level when possible. No biomass determination will be made for these samples. Since most
oligochaetes and chironomides are relatively small, this will not cause underestimation of a significant
amount of biomass.
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5.5.14.3 If the number of specimens is between 20 and 400, then approximately 50% of the specimens
will be mounted and the remaining portion will be measured for biomass. Samples will be split in the
following manner:
5.5.14.3.1 Specimens will be distributed in a grided tray as evenly as possible. Grids will be selected at
random until exactly half of the total number of grids are selected. The specimens in these grids will be
mounted and identified. Due to the random distribution of fauna, half of the sample grids will not
necessarily contain 50% of the specimens.
5.5.14.3.2 Specimens in the remaining grids will be enumerated, combined, and processed for biomass.
5.5.14.4 If the number of specimens is greater than 400, samples will be treated in the following manner:
5.5.14.4.1 Samples will be split as in 5.5.14.3.1. Instead of using half of the total number of grids to select
specimens to be identified, grids will be randomly selected until at least 200 specimens are mounted. Any
remaining specimens in the last selected grid will also be identified. The total number of identified
specimens in these samples will usually be greater than 200.
5.5.14.4.2 Specimens in the remaining grids will be enumerated, combined, and processed for biomass.
5.5.15 Count of the number of species in each sample.
5.5.15.1 Each species will be included in the count of the total number of species in each sample.
5.5.15.2 Specimens that can be identified only to genus, family, or order will also be included in the total
number of species in each sample (e.g., specimens identified to be within family Spionidae will be counted
as one species). If a specimen identified to genus, family, or order can be identified as one of several
species already identified in the sample, that specimen will not be counted as an additional species. This
procedure eliminates double counting of certain species. For example, a specimen is identified as being
Tellinidae, but the taxonomist believes it to be either Macoma balthica or Macoma mitchelli, both of which
are present in the sample. The specimen would be recorded as Tellinidae and would not be included in
the species count for that sample.
5.5.15.3 Identified specimens that will not be included in species counts will be so marked by recording a
'1' in the appropriate column on the abundance data sheets.
5.5.15.4 Fragments of organisms generally will not contribute to the count of number of species per
sample; however, if a fragment contributes to abundance by meeting the criteria outlined in Subsection
5.5.4, then fragments will also contribute to the count of the number of species per sample.
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5.5.16 Additional quality control procedures for species identification and enumeration, including
maintenance of a laboratory taxonomic reference collection, are given in Subsection 7.
6. MACROFAUNAL BIOMASS DETERMINATION
6.1 Biomass is an additional measure of the overall health, status, and history of benthic communities.
This section describes the methods to be used to measure benthic biomass. Biomass will be reported as
formaldehyde dry weight since samples were fixed and initially preserved in formaldehyde. To ensure that
measurements are standardized and that comparisons may be made between samples and between
taxonomic groups, two factors must be considered. First, biomass measurements must be made only after
samples have been preserved for a certain minimum time, and, second, soft bodied organisms and those
having significant inorganic body parts must be treated separately.
6.2 Biological samples lose weight when preserved in formalin due to the precipitation of proteins (Jones
1976). Weight loss is variable and is related to organism composition and duration of preservation. In
general, weightless decreases exponentially and is negligible after two months (Howmiller 1972; Schram
etal. 1981; Mills et a/. 1982). In order to standardize measurements, all samples will be preserved in a 10%
formalin solution for at least one month before biomass measurements are made. Subsequent transfer to
and represervation in ethanol is not thought to impact biomass measurements.
6.3 Biomass measurements cannot be made until all specimens in a sample have been properly identified
and all quality control procedures have been implemented. Measuring biomass is a destructive process
after which no further taxonomic identifications or checks are possible.
6.4 Soft-bodied organisms (e.g., amphipods and polychaetes) may be dried and weighed immediately after
identification. Hard-bodied organisms (e.g., bivalves, gastropods, and echinoderms) will need to be
acidified prior to drying and weighing. Acidification removes the calcium carbonate present leaving behind
organic carbon. Acidification procedures are given in Subsection 6.10.
6.5 An analytical balance with an accuracy of 0.1 mg will be used to measure biomass.
6.6 Biomass will be individually determined for the most dominant macrofaunal species or group of species
in each Province. Dominant species or groups of species are selected prior to laboratory processing based
upon a review of published studies. Remaining species are grouped into categories having ecological or
taxonomic relevance. Biomass categories selected for the Virginian Province are as follows:
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Amphipod Biomass Groups:
Ampelisca spp.
Corophium spp.
Gammaridae
Haustoriidae
Leptocheirus spp.
Monoculodes spp.
Unciola spp.
Other and unidentifiable amphipods
Arthropod Biomass Groups:
Cyathura spp.
Other and unidentifiable isopods
Decapods
Chironomids
Polychaete Biomass Groups:
Glycera spp.
Heteromastus filiformis
Leitoscoloplos spp.
Maldanidae
Marenzelleria viridis
Mediomastus ambiseta
Neanthes succinea
Nephtys spp.
Paraprionospio pinnata
Polydora spp.
Streblospio benedicti
Other polychaetes - sub-surface deposit
feeders
Other polychaetes - surface deposit/suspension
feeders
Other polychaetes - carnivores/omnivores
Other and unidentifiable polychaetes and
polychaete fragments
Oligochaete Biomass Group
All oligochaetes and oligochaete fragments
Bivalve Biomass Groups
Corbicula fluminea
Ensis directus
Gemma Gemma
Mercenaria mercenaria
Mulinia lateralis
Nucula spp.
Rangia cuneata
Tellinidae
Yoldia limatula
Other bivalves - deposit feeders
Other bivalves - suspension feeders
Other and unidentifiable bivalves and bivalve
fragments
Gastropods Biomass Groups:
Acteocina canaliculata
Hydrobia spp.
Other and unidentifiable gastropods
Miscellaneous Species and Groups:
Echinodermata
Hemichordata
Miscellaneous Species and Groups:
Phoronis spp.
Nemertinea
Other species and unidentifiable macrofauna
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6.7 The measurement of biomass for each of the groups above will commence with the collection of the
species identification data sheet and taxon storage vials for an individual sample. Biomass data sheets will
accompany abundance sheets and will note how many and which biomass vials should be present. Any
discrepancies between data sheets and vials will be corrected at this time.
6.8 In the steps that follow, the treatment for soft-bodied and hard-bodied macrofaunal organisms differs.
The treatment for soft-bodied organisms will be given first and will be followed by the treatment for hard-
bodied organisms.
6.9. Soft-Bodied Macrofauna:
6.9.1 Soft bodied organisms will be placed in preweighed, numbered weighing pans. An appropriately
sized pan will be selected for each taxonomic category according to the amount of material to be
processed. The pan number will be recorded on the biomass data sheet along with the taxonomic group
to be measured.
6.9.2 Care will be taken to check that all organisms are rinsed from the vials into the weighing pan.
6.10 Hard-Bodied Macrofauna:
6.10.1 Hard bodied organisms will be placed in preweighed, numbered, porcelain crucibles. A crucible of
appropriate size will be selected for each taxonomic category according to the amount of material to be
processed. The crucible number will be recorded on the biomass data sheet along with the taxonomic
group to be measured.
6.10.2 Care will be taken to check that all organisms are rinsed from the vials into the crucible.
6.10.3 Large bivalves (length > 2 cm) will be shucked instead of being acidified; acidification of large shells
is time consuming and uses an excessive amount of acid.
6.10.4 Crucibles will be acidified in a fume hood using 10% HCI. Acidification will continue until all visible
traces of shell material are removed. Additional acid may be added as needed to bring about the complete
dissolution of shell material.
6.10.5 When no traces of shell material remain, the acid will be removed with a pipette and the remaining
material will be rinsed with distilled water.
6.11 In subsequent steps, soft-bodied and hard-bodied macrofauna will be similarly treated.
6.12 Weighing pans and crucibles will be placed in carrying trays and dried in an oven at 60°C.
6.13 Pans and crucibles to be weighed will be grouped in batches. The batch number will be recorded on
biomass data sheets. Each batch will contain weighing blanks. Blanks are pans and crucibles which have
been treated as biomass samples but to which no fauna have been added. Blanks will be used to
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determine the overall accuracy of the weighing procedure and will help detect errors due to the
contamination of biomass samples. Approximately 5 to 10% of the number of pans and crucibles in a batch
will be blanks.
6.14 Prior to weighing pans and crucibles, the balance will be zeroed and a standard weight will be used
to test its calibration. Each standard is individually numbered and this number will be noted in the biomass
log book. Subsequent weighings will use the same standard weight.
6.15 Dried samples awaiting measurement will be stored in a desiccator to avoid absorbing moisture from
the atmosphere. Desiccator storage also allows samples to cool to room temperature prior to weighing.
6.16 Typically, 24 to 48 hours at 60 °C is sufficient for the dry weight of benthic samples to stabilize,
however, some samples may take longer to dry. As a check, all samples will be weighed after 24 hours,
the weight will be recorded on the biomass data sheets, and the samples will be returned to the drying
oven. Dry weight will be measured again after 24 hours. If the second sample weight differs from the first
by more than 5%, then the sample will be returned to the drying oven for an additional 24 hours. This cycle
will be repeated until a stable dry weight measurement is obtained. All weights will be recorded on the
biomass data sheet.
6.17 Approximately 10% of all pans and crucibles will be reweighed by a second technician as a quality
control check of biomass measurements.
6.18 Biomass data sheets will contain a record of all weights; however, only the tare weight of weighing
pans and crucibles and the final weight of the sample (sample weight + crucible or pan weight) will be
entered into the data base. Net (sample) weights will be calculated within the data base.
6.19 Weighing blanks should vary by no more than 0.3 mg. If greater variations are found, the balance
and the weighing procedures used by the technician should be checked, and, as necessary, the balance
will be repaired or the technician will be retrained.
7. QUALITY ASSURANCE AND QUALITY CONTROL
7.1 Various quality control (QC) procedures will be implemented to ensure consistent production of high
quality data. In addition to the QC procedures included in this chapter, the following procedures will be
periodically conducted as part of data quality control.
7.2 Sorting:
7.2.1 A minimum of 10% of all samples sorted by each technician will be resorted to monitor technician
performance and provide feedback necessary to maintain acceptable standards. Resorts will be conducted
on a regular basis on batches of 10 samples, and all results will be documented and recorded in the QA/QC
logbook for the laboratory.
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7.2.2 The QC resort procedure is designed to provide effective and continuous monitoring of sorting
efficiency. For EMAP-Estuaries, the minimum acceptable sorting efficiency is 90%. Based upon the
experience of other programs using similar methods (Holland et a/. 1988), however, sorting efficiencies are
expected to be greater than 95%.
7.2.3 Samples sorted by a particular technician will be randomly selected for resorting from a sample batch.
7.2.4 The archived sample residues will be retrieved and the sample number will be recorded in the QC
log book.
7.2.5 The residue will be resorted using the sorting procedures given in Subsection 5.4.
7.2.6 Sorting efficiency (%) will be calculated using the following formula:
# organisms originally sorted 100
# organisms originally sorted + additional # found in resort
7.2.7 The results of sample resorts may require that certain actions be taken for specific technicians. If
sorting efficiency is greater than 95%, no action will be required. If sorting efficiency is 90 to 95%, the
technician will be retrained and problem areas identified. Laboratory personnel and supervisors must be
particularly sensitive to systematic errors (i.e., consistent failure to represent specific taxonomic groups) that
may suggest the need for further training. Resort efficiencies below 90% will require resorting all samples
in that batch and continuous monitoring of that technician to improve efficiency.
7.2.8 If sorting efficiency is less than 90%, organisms found in the resort will be added to the original data
sheet and placed in the appropriate biomass group vial. If sorting efficiency is 90% or greater, the results
will be recorded in the QC log book; however, the animals should be kept separate from the original sample
and should not be used for biomass determinations.
7.2.9 If a sample batch fails to meet the 90% efficiency sorting criteria, all samples within the batch will be
resorted. An additional sample from the batch will be randomly selected and used to check the sorting
efficiency of the resorted batch.
7.2.10 After resorting, and if quality control criteria are met, sample residues may be discarded.
7.2.11 Resort results will be summarized for each technician on a QC resort summary sheet.
7.3 Species Identification and Enumeration:
7.3.1 Only senior taxonomists are qualified to complete identification quality control checks. A minimum
of 10% of all samples processed by each taxonomic technician will be checked to verify the accuracy of
species identifications and enumerations. This control check establishes the level of accuracy with which
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identification and counts are performed and offers feedback to taxonomists in the laboratory to maintain
a high standard of accuracy. Samples will never be rechecked by the technician who originally processed
the sample.
7.3.2 Approximately 10% of each sample batch will be checked. A sample batch consists of 10 samples
and ideally is made of samples from a similar habitat type (all oligohaline stations, for example). Rechecks
will be performed in a timely manner so that subsequent processing steps (e.g., biomass determinations)
and data entry may proceed.
7.3.3 The vials containing specimens from the randomly selected sample will be retrieved along with the
original species identification sheet and information will be recorded in the QC log book.
7.3.4 The specimens in each vial will be reidentified and enumerated using the procedures given in
Subsection 5.5 of this chapter.
7.3.5 As each taxon is identified and counted, results will be compared to the original data sheet.
Discrepancies will be double-checked to verify that the final results are correct.
7.3.6 Following reidentification, specimens will be returned to the original vials and set aside for biomass
determination.
7.3.7 When the entire sample has been reidentified, the total number of errors will be computed. The total
number of errors will be based upon the number of misidentifications and miscounts. Numerically, accuracy
will be represented in the following manner:
Total # of organisms in QC recount - total no. of errors *
Total # of organisms in QC recount
* Three types of errors are included in the total number of errors:
1) Counting errors (for example, counting 11 Gemma gemma as 10)
2) Identification errors (for example, identifying a Nucula annulata specimen as Nucula proxima,
where both are present)
3) Unrecorded taxa errors (for example, not identifying Phoronis spp. when it is present)
7.3.8 For EMAP-Estuaries benthic samples, the minimum acceptable taxonomic efficiency will be 90%.
If taxonomic efficiency is greater than 95%, no action will be required. If taxonomic efficiency is 90 to 95%,
the taxonomist will be consulted and problem areas will be identified. Taxonomists and laboratory
supervisors must be particularly sensitive to systematic errors (i.e., repeated errors for specific taxonomic
groups) that may suggest the need for further training. Taxonomic efficiencies below 90% will require
reidentifying and enumerating all samples in that sample batch and additional monitoring of the taxonomist
to improve efficiency.
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7.3.9 Any species identification changes resulting from quality assurance procedures will be recorded on
the original data sheet; however, the numerical count for each taxonomic group will not be corrected unless
the overall accuracy for the sample is below 90%.
7.3.10 Treatment of the results of quality control audits are illustrated in the following examples.
7.3.10.1 Example 1: Ten Mulinia lateralis individuals were recounted as eleven. The sample had a
greater than 90% overall efficiency, therefore, the original count of ten Mulinia would be recorded.
7.3.10.2 Example 2: One individual of the species Prionospio steenstrupi was misidentified as Streblospio
benedicti. On the final data sheet, one Prionospio steenstrupi and no Streblospio benedicti would be
recorded.
7.3.10.3 Example 3: Ten Nucula annulata and no Nucula proxima were originally recorded. During the
QA/QC check, one N. annulata was found to be N. proxima. Providing the overall efficiency was greater
than 90%, nine N. annulata and one N. proxima would be recorded on the final data sheet.
7.3.10.4 Example 4: Five Nucula annulata and ten Mulinia lateralis were originally recorded. During the
QA/QC check, one M. lateralis was found to be a N. annulata. Providing the overall efficiency was greater
than 90%, six N. annulata and nine M. lateralis would be recorded on the final data sheet.
7.3.10.5 Example 5: One Onuphidae spp. (juvenile) was recorded on original data sheet. During the
QA/QC check, this individual was not found. On the final data sheet, one Onuphidae spp. (juvenile) would
be recorded.
7.3.10.6 Example 6: Terebellidae spp. (juvenile) was found in the annelid fragment category during the
QA/QC check. No Terebellidae were previously recorded on the data sheet. On the final data sheet, one
Terebellidae spp. would be recorded.
7.3.11 The results from all QC rechecks of species identification and enumeration will be recorded in the
QC log book that will become a part of the documentation for EMAP-Estuaries.
7.3.12 All corrections to data sheets will be initialed and dated by the person making the changes.
7.4 Taxonomic Reference Collection:
7.4.1 Taxonomic identifications should be consistent within a given laboratory and with the identifications
of other regional laboratories. Consistent identifications are achieved by implementing the procedures
described above and by maintaining informal interaction among the taxonomists working on each major
group.
7.4.2 A voucher specimen collection should be established by each laboratory processing EMAP samples.
This collection should consist of representative specimens of each species identified in samples from an
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individual province. For some species, it may be appropriate to include in the voucher specimen collection
individuals sampled from different geographic locations within the Province.
7.4.3 New species added to a laboratory's voucher specimen collection should be sent to recognized
experts for verification of the laboratory's taxonomic identifications. The verified specimens should then
be placed in a permanent taxonomic reference collection. The reference collection should be used to train
new taxonomists. Participation of the laboratory staff in a regional taxonomic standardization program (if
available) is recommended, to ensure regional consistency and accuracy of identification.
7.4.4 All specimens in the reference collection should be preserved in 70% ethanol in labeled vials that are
segregated by species and sample. More than one specimen may be in each vial. The labels placed in
these vials should be made of waterproof, 100-percent rag paper and filled out using a pencil. Paper with
less than a 100-percent rag content or that is not waterproofed will disintegrate in the 70-percent alcohol
mixture. It is important to complete these labels because future workers may not be familiar with details of
the work in progress.
7.4.5 To reduce evaporation of alcohol, the lids of vials and jars can be sealed with plastic tape wrapped
in a clockwise direction. The species (and other taxonomic designation) should be written clearly on the
outside and on an internal label. Reference specimens should be archived alphabetically within major
taxonomic groups.
7.4.6 Reference collections are invaluable and should be retained at the location where the identifications
were performed. In no instance should this collection be destroyed. A single person should be identified
as curator of the reference collection and should be responsible for its integrity. Its upkeep will require
periodic checking to ensure that alcohol levels are adequate. When refilling the jars, it is advisable to use
full-strength alcohol (i.e., 95 percent), because alcohol tends to evaporate more rapidly than water in a 70-
percent solution.
7.4.7 The laboratory will maintain a log pertaining to the taxonomic reference collection. This log will
contain the species name, the name and affiliation of the person who originated the reference sample, the
location of the reference sample, the status of the sample if it has been loaned to outside experts, and
information about the confirmation of identification by outside experts. The log may also contain references
to pertinent literature describing the species in the reference sample.
7.5 Biomass Measurements:
7.5.1 A minimum of 10% of all pans and crucibles in each batch will be reweighed to monitor technician
performance and to provide the feedback necessary to maintain acceptable standards. Reweighings will
be conducted on a regular basis on sample batches, and all results will be documented and recorded in the
laboratory QA/QC log book for the laboratory.
7.5.2 Samples to be reweighed will be randomly selected from a sample batch.
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7.5.3 Selected samples will be reweighed and the results compared against the final weight recorded on
the biomass data sheet.
7.5.4 Weighing efficiency will be calculated using the following formula:
Original Final Weight - Reweighted Final Weight\
Reweighed Final Weight
7.5.5 If weighing efficiency is 95% or greater, the sample has met the acceptable quality control criteria and
no further action is necessary. If the weighing efficiency is between 90% and 95%, the sample has met
acceptable criteria, but the technician who completed the original weighing will be consulted and proper
measurement practices reviewed. If the weighing efficiency is less than 90%, the sample has failed the
quality control criteria and all samples within the sample batch must be reweighed. Additionally, the
performance of the original technician will be reviewed and the technician will be retrained.
7.5.6 Correction to the original data sheet will be made only in those cases where weighing efficiency is
less than 90%.
7.5.7 The results of all QC reweighings will be recorded in the QC log book that will become a part of the
documentation for EMAP-Estuaries.
8. DATA MANAGEMENT
8.1 All data generated in the laboratory will be recorded directly onto standardized data forms. Data forms
will be designed so that all necessary information will be recorded clearly and unambiguously. All data will
be recorded in ink. Data forms will be linked to specific samples using the bar coded sample numbers that
were assigned by the Province Field Operations Center prior to field sampling. Completed data forms will
be kept in bound notebooks arranged by type. Completed data forms will be made available to the Province
Field Operations Center upon demand. Laboratories will contact the EMAP-Estuaries Technical Director
prior to the disposal of any data sheet, QA/QC forms, or laboratory notebooks pertaining to the generation
of EMAP data. The length of time these materials must be archived by contract laboratories may be
determined by contract.
9. DATA FORMS
9.1 This section lists data forms that will be used for the identification, enumeration, and determination of
biomass for benthic macrofaunal samples collected by EMAP-Estuaries in the Virginian Biogeographic
Province (Cape Cod, MA to Cape Henry, VA). Sample data forms are included in Appendix A. Data forms
are presented in the following order:
• Abundance data sheet for tidal fresh and oligohaline regions (1 page)
-------
EMAP-Estuaries Laboratory Methods Manual Section 3 - Benthic Macroinvertebrates
Volume 1 August 1995
Page 20 of 35
• Abundance data sheet for mesohaline regions (1 page)
• Abundance data sheets for southern polyhaline regions (2 pages)
• Abundance data sheets for northern polyhaline region (2 pages)
• Biomass data sheets for tidal fresh and oligohaline salinity regions (2 pages)
• Biomass data sheets for mesohaline and polyhaline salinity regions (2 pages)
• Laboratory Sample Tracking form (Benthos log sheet 1 page)
• QC Sample Batch Listing form (1 page)
• QC Sample Resort sheet (1 page)
• QC Sample Reidentification sheet (1 page)
• Biomass log sheet (1 page)
• QC Biomass Reweighing and Blank sheet (1 page)
10. REFERENCES
Bilyard, G.R. 1987. The value of benthic infauna in marine pollution monitoring studies. Marine Pollution
Bulletin 18: 581-585.
Boesch, D.F. and R. Rosenberg. 1981. Response to stress in marine benthic communities. In: Stress
effects on natural ecosystems, pp. 179-200. G.W. Barret and R. Rosenberg (eds). NY: John Wiley.
Chapman, P.M., R.N. Dexter and E.R. Long. 1987. Synoptic measures of sediment contamination, toxicity
and infaunal community composition (the sediment quality triad) in San Francisco Bay. Mar. Ecol. Prog.
Ser 37:75-96.
Cloern, J.E. 1982. Does the benthos control phytoplankton biomass in south San Francisco Bay? Mar.
Ecol. Prog. Ser. 9:191-202.
Grassle, J.P. and J.F. Grassle. 1984. The utility of studying the effects of pollutants on single species
populations in benthos of mesocosms and coastal ecosystems. In: Concepts in Marine Pollution
Measurements, pp. 621-642. H.H. White, ed. Maryland Sea Grant College, College Park, MD.
Holland, A.F., AT. Shaughnessy, L.C. Scott, V.A. Dickens, J.A. Ranasinghe.and J.K. Summers. 1988.
Progress report: Long-term benthic monitoring and assessment program for the Maryland portion of
Chesapeake Bay (July 1986 ~ October 1987). Prepared for the Maryland Department of Natural
Resources, Power Plant Research Program and Maryland Department of the Environment, Office of
Environmental Programs. PPRP-LTB/EST-88-1.
-------
EMAP-Estuaries Laboratory Methods Manual Section 3 - Benthic Macroinvertebrates
Volume 1 August 1995
Page 21 of 35
Holland, A.F., N.K. Mountford, M.H. Hiegel, K.R. Kaumeyerand J.A. Mihursky. 1980. Influence of predation
on infaunal abundance in upper Chesapeake Bay, USA. Marine Biology 57:221-235.
Holmes, N.A. and A.D. Mclntyre. 1984. Methods for Study of Marine Benthos. Blackwell Scientific
Publications, London.
Howmiller, R.P. 1972. Effect of preservatives on weight of some common macrobenthic invertebrates.
Trans. Am. Fish. Soc. 101:743-746.
Jones, D. 1976. Chemistry of fixation and preservation with aldehydes. In: Zooplankton fixation and
preservation, pp. 155-171. H.F. Steedman (ed.). Paris: UNESCO Press.
Mills, E.L., K. Pittman and B. Munroe. 1982. Effect of preservation on the weight of marine benthic
invertebrates. Can. J. Fish. Aquat. Sci. 39:221-224.
Mirza, F.B. and J.S. Gray. 1981. The fauna of benthic sediments from the organically enriched Oslofjord,
Norway. J. Exp. Mar. Biol. Ecol. 54:181-207.
Pearson, T.H. and R. Rosenberg. 1978. Macrobenthic succession in relation to organic enrichment and
pollution of the marine environment. Oceanogr. Mar. Biol. Ann. Rev. 16: 229-311.
Rhoads, D.C. 1974. Organism-sediment relations on the muddy sea floor. Mar. Biol. Ann. Rev.
12:263-300.
Rhoads, D.C., and O.K. Young. 1970. The influence of deposit feeding organisms on sediment stability
and community trophic structure. J. Mar. Res. 28:150-177.
Rhoads, D.C., J.Y. Yingst and W.J. Ullman. 1978. Seafloor stability in Central Long Island Sound: Part I.
Temporal changes in erodibility of fine-grained sediment. In: Estuarine Interactions, pp. 221-244. M.L.
Wiley (ed.). NY: Academic Press.
Sanders, H.L., J.F. Grassle, G.R. Hampson, L.S. Morse, S. Garner-Price and C.C. Jones. 1980. Anatomy
of an oil spill: Long-term effects from the grounding of the barge Florida off West Falmouth,
Massachusetts. Jour. Mar. Res. 38:265-380.
Schram, M.D., G.R. Ploskey and E.H. Schmitz. 1981. Dry weight loss in Ceriodaphnia lacustris (Crustacea,
Cladocera) following formalin preservation. Trans. Am. Micros. Soc. 100:326-329.
Swartz, R.C., F.A. Cole, D.W. Schults and W.A. DeBen. 1986. Ecological changes in the Southern
California Bight near a large sewage outfall: Benthic conditions in 1980 and 1983. Mar. Ecol. Prog.
Ser. 31: 1-13.
Virnstein, R.W. 1977. The importance of predation by crabs and fishes on benthic infauna in Chesapeake
Bay. Ecology 58: 1199-1217.
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Volume 1 August 1995
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THIS PAGE INTENTIONALLY LEFT BLANK
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EMAP-Estuaries Laboratory Methods Manual Section 3 - Benthic Macroinvertebrates
Volume 1 August 1995
Page 23 of 35
APPENDIX A
DATA FORMS
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EMAP-Estuaries Laboratory Methods Manual
Volume 1
Section 3 - Benthic Macroinvertebrates
August 1995
Page 24 of 35
Page
of
NEAR-COASTAL EMAP 1993
Species Abundance Data Sheet
Tidal Freshwater / Oligohaline
EXAMPLE
SAMPLE: EXAMPLE Priority 1
LAB: Versar ESN Operations
SORT DATE: SORTED BY:
m d v
MOUNT DATE: M. ID BY:
BH
GP
46
29
28
38
38
38
34
27
27
27
27
27
27
27
27
27
27
27
27
25
18
2
3
3
3
Species
Turbellaria
Corbicula fluminea
Macoma mitcheLli
Musculium spp.
Musculium transversum
Pisidium spp.
Rang i a cuneata
Aulodrilus limnobius
Aulodrilus pigueti
Dero digitata
Dero spp.
I Lyodri lus templetoni
Limnodrilus cervix
Limnodrilus hoffmeisteri
Limnodrilus udekemianus
Quistadrilus multisetosus
Tubificidae w/o cap
Tubificidae wth cap
Tubificoides heterochaetus
Hobsonia f Lori da
Marenzelleria viridis
Coroptiium lacustre
Gamma r us daiberi
Gamma r us fasciatus
Gamma rus spp.
Code
TURBELLA
CORBFLUM
MACOMITC
MUSCULIU
HUSCTRAN
PISIDIUM
RANGCUNE
AULOLIMN
AULOPIGU
DEROOIGI
DERO
ILYOTEMP
LIMNCERV
LIHNHOFF
LIMNUDEK
QUISMULT
TUBIFIHI
TUBIFIUO
TUBIHETE
HOBSFLOR
MAREVIRI
COROLACU
GAMMDAIB
GAMMFASC
GAHMARUS
Nui
*
OLIGO. SPLIT: / CHIRON. SPLIT: /
ID DATE: ID BY:
m it y
SYSTEM: Chesapeake Bay Maryland
BM
Gp
5
6
12
12
12
12
12
12
12
12
12
12
46
10
9
Species
Leptocheirus plumulosus
Monoculodes sp. 1 Wat I ing
Chironomidae
Chironomus spp.
Coelotanypus spp.
Cryptochironomus fulvus
Crypt ochironomus spp.
PolypediLum convictum
Polypedi tun spp.
Procladius spp.
Procladius sublettei
Tanytarsus spp.
Chaoborus punctipennis
Chiridotea almyra
Cyathura polita
Code
LEPTPLUM
HONOSPE1
CHRNMDAE
CHIRONOM
COELOTAN
CRYPFULV
CRYPTOCH
POLYCONV
POLYPEDI
PROCLADI
PROCSUBL
TANYTARS
CHAOPUNC
CHIRALMY
CYATPOLI
Nui
Dir Mnt
*
Temporary Record - Do not enter into data base
Oligochaetes - Total Split
Chironomids - Total Split
xxxxxxxx
xxxxxxxx
*: Write '!' in this column if taxon is NOT to be considered
Num=# of Individuals
when counting species in sample
of Bionass Vials
-------
EMAP-Estuaries Laboratory Methods Manual
Volume 1
Section 3 - Benthic Macroinvertebrates
August 1995
Page 25 of 35
Page
of
NEAR-COASTAL EMAP 1993
Species Abundance Data Sheet
Mesohaline
EXAMPLE
SAMPLE: EXAMPLE
LAB: Cove Corporation
ID INFORMATION:
Priority 1
| LAB SAMPLE *:
| SYSTEM: Chesapeake Bay Maryland
BM
Gp
45
33
38
28
28
35
38
38
34
38
28
39
41
41
27
19
26
14
26
23
23
18
13
21
17
Species
Nemer tinea
Gemma germa
Lyons! a hyalina
Macoma balthica
Hacoma mitchelli
Mulinia lateral is
Mya arenaria
Parvilucina multilineata
Rang i a cuneata
Tagelus plebeius
Tellina agi lis
Acteocina canaliculate
Acteon punctostriatus
Haminoea solitaria
Oligochaeta
Glycera dibranchiata
Glycinde solitaria
Heteromastus filiformis
Hypereteone heteropoda
Leitoscoloplos robustus
Lei toscoloplos spp.
Marenzelleria viridis
Mediomastus ambiseta
Neanthes succinea
Paraprionospio pinnata
*: Write '1' in this column if taxon
Num=# of Individuals
Code
NEMERTIN
GEMMGENM
LYONHYAL
MACOBALT
MACOMITC
MUL1LATE
MYAAREN
PARVMULT
RANGCUNE
TAGEPLEB
TELLAG1L
ACTECANA
ACTEPUNC
HAH I SOL I
OLIGOCHA
GLYCDIBR
GLYCSOLI
HETEFILI
ETEOHETE
LEITROBU
LEI TOSCO
MAREVIRI
MED IAMB I
NEANSUCC
PARAPINN
NUB
*
BM
GP
24
26
47
16
26
25
25
15
2
5
6
9
10
46
44
42
Species
Pectinaria gouldii
Podarkeopsis levifuscina
Polychaeta
Polydora cornuta
Pseudeurythoe pauci branch iata
Scolelepis texana
Spiophanes bombyx
Streblospio benedict i
Corophium lacustre
Leptocheirus plumulosus
Monoculodes sp. 1 Wat I ing
Cyathura polita
Edotea triloba
Neomysis americana
Phoronis spp.
Leptosynapta tenuis
is NOT to be considered when counting species in sample
Code
PECTGOUL
PODALEVI
POLYCHAE
POLYCORN
PSEUPAUC
SCOLTEXA
SPIOBOMB
STREBENE
COROLACU
LEPTPLUM
MONOSPE1
CYATPOL1
EDOTTRIL
NEOMAMER
PHORONIS
LEPTTENU
Nun
*
Number of Bionass Vials
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EMAP-Estuaries Laboratory Methods Manual
Volume 1
Section 3 - Benthic Macroinvertebrates
August 1995
Page 26 of 35
Page
of
NEAR-COASTAL EMAF 1993
Species Abundance Data Sheet
Southern Folyhaline
EXAMPLE
SAMPLE: EXAMPLE
LAB: Cove Corporation
ID INFORMATION:
Priority 1 Abundance:
| LAB SAMPLE #:
| SYSTEM: Chesapeake Bay Maryland
Page 1
BM
Gp
46
45
38
48
36
33
28
35
31
38
38
28
28
32
39
41
41
41
41
27
26
25
25
26
20
Species
Turbellaria
Nemertinea
Anadara transverse
Bivalvia
Ens is directus
Gemma gemma
Macoma tenta
Mulim'a lateral is
Nucula annul at a
Parvilucina multi lineata
Pi tar morrhuanus
Tellina agi I is
Tellinidae
Yoldia limatula
Acteocina canaliculata
Anachis lafresnayi
Astyris lunata
Odostomia engonia
Turbonilla interrupts
Oligochaeta
Ancistrosyl lis hartmanae
Aricidea catherinae
Asabe Hides oculata
Bhawania heteroseta
Clymenella torquata
Code
TURBELLA
NEMERTIN
AN AD IRAN
BIVALVIA
ENSIDIRE
GEHMGEHM
MACOTENT
MULILATE
NUCUANNU
PARVMULT
PITAMORR
TELLAGIL
TELLINID
YOLDLIMA
ACTECANA
ANACLAFR
ASTYLUNA
ODOSENGO
TURBINTE
OLIGOCHA
ANCIHART
ARICCATH
ASABOCUL
BHAUHETE
CLYMTORQ
NUB
*
BM
Gp
26
19
19
19
26
14
26
23
23
26
25
25
20
13
21
22
17
24
26
26
47
16
25
26
26
Species
Orilonereis longa
Glycera americana
Glycera dibranchiata
Glycera spp.
Glycinde solitaria
Heteromastus filiformis
Hypereteone heteropoda
Lei toscoloplos robustus
Leitoscoloplos spp.
Lepidametria commensal is
Loimia medusa
Magelona spp.
Maldanidae
Hediomastus ambiseta
Neanthes succinea
Nephtys pi eta
Paraprionospio pinnata
Pectinaria gouldii
Phyllodoce arenae
Podarkeopsis levifuscina
Polychaeta
Polydora cornuta
Prionospio perkinsi
Pseudeurythoe pauci branchiate
Sigambra tentaculata
Code
DRILLONG
GLYCAMER
GLYCDIBR
GLYCERA
GLYCSOLI
HETEFILI
ETEOHETE
LEITROBU
LEITOSCO
LEPICOMM
LOIMMEDU
MAGELONA
HALDANID
MED IAMB I
NEANSUCC
NEPHPICT
PARAPINN
PECTGOUL
PHYLAREN
POOALEVI
POLYCHAE
POLY CORN
PRIOPERK
PSEUPAUC
SIGATENT
Nun
*
*: Write '1' in this column if taxon is NOT to be considered when counting species in sample
Num=# of Individuals
-------
Page
of
NEAR-COASTAL EMAP 1993
Species Abundance Data Sheet
Southern Polyhaline
EXAMPLE
SAMPLE: EXAMPLE
Priority
Abundance: Page 2
SYSTEM: Chesapeake Bay Karyland
BM
GP
25
25
15
25
1
1
8
2
8
8
7
46
46
46
11
11
10
44
Species
Spiochaetopterus costarum
Spiophanes bombyx
Streblospio benedict i
Tharyx sp. A Morris
Ampelisca abdita-vadorum cpx
Ampelisca verrilli
Batea catharinensis
Corophium tuberculatum
Listriella barnardi
Listriella clymenellae
Unciola serrata
Balanus improvisus
Leucon americanus
Oxyurostylis smith i
Ogyrides alphaerostris
Pinnixa spp.
Edotea triloba
Phoronis spp.
Code
SPIOCOST
SPIOBOMB
STREBENE
THARSPA
AMPEABVA
AMPEVERR
BATECATH
COROTUBE
LISTBARN
LISTCLYM
UNCISERR
BALAIMPR
LEUCAMER
OXYUSMIT
OGYRALPH
PINNIXA
EDOTTRIL
PHORONIS
Nun
* 1 BM
I <*
Species
Code
Nun
*
*: Write '1' in this column if taxon is NOT to be considered when counting species in sample
Num=# of Individuals Nunber of Bionass Vials
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EMAP-Estuaries Laboratory Methods Manual
Volume 1
Section 3 - Benthic Macroinvertebrates
August 1995
Page 28 of 35
Page
of
NEAR-COASTAL EMAP 1993
Species Abundance Data Sheet
Northern Polyhaline
EXAMPLE
SAMPLE: EXAMPLE Priority 1 Abundance: Page 1
LAB: Cove Corporation
ID INFOMUTION:
LAB SAMPLE #:
SYSTEM: Chesapeake Bay Maryland
BM
GP
46
45
36
33
38
28
35
38
31
38
28
32
39
41
41
41
41
27
25
25
25
25
20
24
19
Species
Turbellaria
Nemertinea
Ensis directus
Gemma gemna
Lyons i a hyalina
Macoma tenta
Mulinia lateral is
My til us edulis
Nucula annulate
Pi tar morrhuanus
Tel Una agilis
Yoldia limatula
Acteocina canaliculate
Anachis lafresnayi
Astyris lunata
Nassarius trivittatus
Turbonilla interrupta
Oligochaeta
Ampharete arctica
Ampharetidae
Aricidea catherinae
Asabe Hides oculata
Clymenella torquata
Cossura longocirrata
Glycera americana
Code
TURBELLA
NEMERTIN
ENSIDIRE
GEMHGEMM
LYONHYAL
MACOTENT
MULILATE
MYTIEOUL
NUCUANNU
PITAMORR
TELLAGIL
YOLDLIMA
ACTECANA
ANACLAFR
ASTYLUNA
NAS5TRIV
TURBINTE
OL1GOCHA
AHPHARCT
AMPHARTD
ARICCATH
ASABOCUL
CLYMTORQ
COSSSOYE
GLYCAMER
Nun
* 1 BM
I Gp
19
19
26
26
14
23
23
26
25
25
20
13
21
26
22
22
17
24
26
26
47
25
16
25
Species
Glycera dibranchiata
Glycera spp.
Glycinde solitaria
Goniadidae
Heteromastus filiformis
Leitoscoloplos robustus
Leitoscoloplos spp.
Lepidametria commensal is
Levinsenia gracih's
Loimia medusa
Maldanidae
Mcdiomastus ambiseta
Neanthes succinea
Nephtyidae
Nephtys incisa
Nephtys picta
Paraprionospio pinnata
Pectinaria gouldii
Phyl lodoce arenae
Podarkeopsis levtfuscina
Polychaeta
Polycirrus spp.
Polydora cornuta
Prionospio steenstrupi
Sigambra tentaculata
Code
GLYCD1BR
GLYCERA
GLYCSOL1
GON1A01D
HETEFIL1
LEITROBU
LEI TOSCO
LEPICOMM
LEV1GRAC
LOIMHEDU
MALDAN1D
MED IAMB I
NEANSUCC
NEPHTYID
NEPHINCI
NEPHPICT
PARAPINN
PECTGOUL
PHYLAREN
POOALEVI
POLYCHAE
POLYCIRR
POLY CORN
PRIOSTEE
S1GATENT
Nun
*
*: Write '11 in this column if taxon is NOT to be considered when counting species in sample
Num=# of Individuals
-------
Page
of
NEAR-COASTAL EHAP 1993
Species Abundance Data Sheet
Northern Polyhaline
EXAMPLE
SAMPLE: EXAMPLE
priority 1 Abundance: Page 2
| SYSTEM: Chesapeake Bay Maryland
BH
Gp
25
25
15
25
1
1
2
5
7
7
46
1,6
11
11
11
10
44
Species
Spiochaetopterus costarum
Spiophanes bonfcyx
Streblospio benedicti
Tharyx sp. A Morris
Ampelisca abdita-vadorum cpx
Ampelisca verrilli
Corophium spp.
Leptocheirus pinguis
UncioLa irrorata
Unciola spp.
Leucon americanus
Oxyurostylis smith!
Cancer spp.
Crangon septemspinosa
Pirmixa spp.
Edotea tritoba
Phoronis spp.
Code
SPIOCOST
SPIOBOMB
STREBENE
THARSPA
AMPEABVA
AMPEVERR
COROPHIU
LEPTPING
UNCIIRRO
UNCIOLA
LEUCAMER
OXYUSMIT
CANCER
CRANSEPT
PINNIXA
EDOTTRIL
PHORONIS
Nu>
*
BH
GP
Species
Code
Nm
*
*: Write '!' in this column if taxon is NOT to be considered when counting species in sample
Num=# of Individuals Number of Bio
ss Vials
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EMAP-Estuaries Laboratory Methods Manual
Volume 1
Section 3 - Benthic Macroinvertebrates
August 1995
Page 28 of 35
Page
of
NEAR-COASTAL EMAP 1993
Biomass Data Sheet
EXAMPLE
SAMPLE: EXAMPLE Priority 1 Biomass: Page 1
LAB: Versar ESM Operations OLIGO. SPLIT: CHIRON. SPLIT:
TARE 1 DATE: BY: TARE 2 DATE: BY:
TARE
Vial
V
3 DATE: BY: LAB SAMPLE *:
BH
GP
1
2
3
4
5
6
7
8
9
10
11
12
49
13
14
15
16
17
18
19
20
21
22
23
24
25
26
Species
Ampelisca spp.
Corophium spp.
G airman dae
Haustoriidae
Leptocheirus spp.
Monoculodes spp.
Unciola spp.
Amphipoda - Other
Cyathura spp.
Isopoda - Other
Decapoda
Chironomidae - Fragments
Chironomidae - Heads
Mediomastus ambiseta
Heteromastus fHiformis
Streblospio benedicti
Polydora spp.
Paraprionospio pinnata
Marenzelteria viridis
Glycera spp.
Haldanidae
Neanthes succinea
Nephtys spp.
Leitoscoloplos spp.
Polychaetes - Other:
Sub-surface Deposit Feeders
Surf .Depst/Suspensn Feeders
Carnivores/Omni vores
Code
AMPELISC
COROPHIU
GAMMARID
HAUSTIDA
LEPTOCHE
MONOCULO
UNCIOLA
AMPHIPOD
CYATHURA
ISOPOOA
DECAPOOA
CHRNHDAE
HEDIAMBI
HETEFILI
STREBENE
POLYDORA
PARAPINN
MAREVIRI
GLYCERA
MALDANID
NEANSUCC
NEPHTYS
LEITOSCO
POLYCSUB
POLYCSUR
POLYCCAR
Pan*
Pan Ut
Cg)
Dry Ut 1
(9)
Dry Ut 2
(9>
Dry Ut U 1
(9)
-------
Page
of
NEAR-COASTAL EMAP 1993
Biomass Data Sheet
B EXAMPLE
SAMPLE: EXAMPLE priority 1 Biomass: Page 2
LAB: Versar ESM Operations LAB SAMPLE #:
Vial
BM
47
27
50
28
29
30
31
32
33
34
35
36
37
38
48
39
40
41
42
43
44
45
46
Species
Polychates - Unident + frags.
Oligochaeta - Fragments
Oligochaeta - Heads
Tellinidae
Corbicula fluminea
Mercenaria mercenaria
Nucula spp.
Yoldia limatula
Gemma gemma
Rangia cuneata
Mulinia lateralis
Ens is directus
Bivalvia: Other
- Deposit Feeders
- Suspension Feeders
- Unidentified
Acteocina canaliculate
Hydrobia spp.
Gastropoda - Other
Echinodermata
Hemichordata
Phorom's spp.
Nemert i nea
Miscellanea
Code
POLYCHAE
OLIGOCHA
TELLINID
CORBFLUM
HERCMERC
NUCULA
YOLDLIMA
GEMMGEMM
RANGCUNE
MULILATE
ENSIDIRE
BIVALDEP
BIVALSUS
BIVALVIA
ACTECANA
HYDROBIA
GASTROPO
ECHINODE
HEMICHOR
PHORONIS
NEMERTIN
MISCELLA
Pan *
Pan Ut
(9)
Dry Ut 1
(9)
Dry Ut 2
Cg)
Dry Ut U
(9)
Number of Biomass Viais
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EMAP-Estuaries Laboratory Methods Manual
Volume 1
Section 3 - Benthic Macroinvertebrates
August 1995
Page 29 of 35
Page
of
NEAR-COASTAL EMAF 1993
Biomass Data Sheet
EXAMPLE
SAMPLE: EXAMPLE Priority 1
LAB: Versa r ESM Operations
TARE 1 DATE: BY:
TARE 3 DATE: BY:
Biomass: Page 1
LAB SAMPLE *:
TARE 2 DATE: BY:
Vial
/
BH
GP
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
47
Species
Ampelisca spp.
Corophium spp.
Ganmaridae
Haustoriidae
Leptocheirus spp.
Honoculodes spp.
Unciola spp.
Amphipoda - Other
Cyathura spp.
Isopoda - Other
Decapods
Chironomidae
Mediomastus ambiseta
Heteromastus filiformis
Streblospio benedict i
Polydora spp.
Paraprionospio pinnata
Marenzel leria viridis
Glycera spp.
Maldam'dae
Neanthes succinca
Nephtys spp.
Lei toscoloplos spp.
Polychaetes - Other:
Sub-surface Deposit Feeders
Surf .Depst/Suspensn Feeders
Carnivores/Omnivores
Unidentified (inc fragnmts)
Code
AMPELISC
COROPHIU
GAMHARID
HAUSTIDA
LEPTOCHE
MONOCULO
UNCIOLA
AMPHIPOD
CYATHURA
ISOPODA
DECAPODA
CHRNHDAE
MED IAMB I
HETEFILI
STREBENE
POLYDORA
PARAPINN
MAREVIRI
GLYCERA
MALDANID
NEAN5UCC
NEPHTYS
LEI TOSCO
POLYCSUB
POLYCSUR
POLYCCAR
POLYCHAE
Pan *
Pan Ut
(9)
Dry Ut 1
(9)
Dry Ut 2
(9)
Dry Ut U
(9)
-------
Page
of
NEAR-COASTAL EMAP 1993
Biomass Data Sheet
B EXAMPLE
SAMPLE: EXAMPLE Priority 1
LAB: Versar ESM Operations
Biomass: Page 2
LAB SAMPLE *:
Vial
1
BH
Gp
27
28
29
30
31
32
33
34
35
36
37
38
48
39
40
41
42
43
44
45
46
Species
Oligochaeta
Tellinidae
Corbicula fluminea
Mercenary a mercenarja
Nucula spp.
Yoldia limatula
Gemna gemma
Rang i a cuneata
MuLinia lateral is
Ens is directus
Bivalvia: Other
- Deposit Feeders
- Suspension Feeders
- Unidentified
Acteocina canal iculata
Hydrobia spp.
Gastropoda - Other
Echinodermata
Hemichordata
Phoronis spp.
Nemertinea
Miscellanea
Code
OLIGOCHA
TELLINID
CORBFLUM
HERCMERC
NUCULA
YOLDLIMA
GEMHGEMM
RANGCUNE
MULILATE
ENSIDIRE
BIVALDEP
BIVALSU5
BIVALVIA
ACTECANA
HYDROBIA
GASTROPO
ECHINOOE
HEMICHOR
PHORONIS
NEMERTIN
MISCELLA
Pan*
Pan Ut
(g)
/^
Dry Ut 1
(9)
\
Dry Ut 2
Cg)
Dry Ut U
(g)
Mutter of Biomass Vials
-------
Data Form - Laboratory Sample Tracking Form
Laboratory Sample Tracking Form
Project
Station Replicate
Serial No.
Sorting*
Personnel
Vials
Generated
General ID **
Collection Date
Oligochaete
Preparation **
Chironomid
Preparation**
Chironjnid ID
QC**
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EMAP-Estuaries Laboratory Methods Manual
Volume 1
Section 3 - Benthic Macroinvertebrates
August 1995
Page 31 of 35
Data Form - QC SAMPLE BATCH LISTING FORM
Necessary Remedial Action:
Comments:
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EMAP-Estuaries Laboratory Methods Manual
Volume 1
Section 3 - Benthic Macroinvertebrates
August 1995
Page 32 of 35
QC SAMPLE RESORT SHEET
Taxa
Additional Org
Number
anisms Found
Taxa
Number
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EMAP-Estuaries Laboratory Methods Manual
Volume 1
Section 3 - Benthic Macroinvertebrates
August 1995
Page 33 of 35
QC SAMPLE REIDENTIFICATION SHEET
0/ E _ Total No. Original Inds. - Total No. Errors
Total No. Inds.
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EMAP-Estuaries Laboratory Methods Manual
Volume 1
Section 3 - Benthic Macroinvertebrates
August 1995
Page 34 of 35
Data Form - BIOMASS LOG SHEET
BIOMASS LOG
Batch Number
Sample ID
Date Pan
Weight
Date First
Biomass
Weight
Date Second
Biomass
Weight
Initials
Complete
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EMAP-Estuaries Laboratory Methods Manual
Volume 1
Section 3 - Benthic Macroinvertebrates
August 1995
Page 35 of 35
QC Biomass Reweighing and Blanks
Batch Number
Blanks
Crucible
Number
Crucible weight
Dr2 Weight #1
D2 Weight #2
Quality Control
Crucible
Number
Original
Weight
Reweigh
Weight
Initials
-------
EMAP-Estuaries Laboratory Methods Manual Section 4 - Histopathology
Volume 1 August 1995
Page 1 of 9
SECTION 4
HISTOPATHOLOGY
1. INTRODUCTION
1.1 Fish quality in EMAP-Estuaries will be measured as a composite index of the incidence of diseases and
contaminant body burdens in selected resident species. Both of these factors are key elements of the
public perception of the environmental quality of estuarine habitats. The assessment and improvement of
fish quality relates directly to the Clean Water Act goal of protection and maintenance of fishable waters.
While gross fish pathology is a potential response indicator of environmental status that is easy and
economical to measure, it may not provide insight into the potential cause of the pathological abnormalities
or may not be related to environmental quality. To address this concern, EMAP-Estuaries will perform
detailed histopathological examinations of randomly selected individuals of target and non-target fish
species at the indicator testing and evaluation sites. All individuals of each target species that "fail" the field
gross pathology examination and up to 25 randomly selected individuals of each target species that "pass"
the field examination at the indicator testing and evaluation sites will undergo a detailed gross and
histopathological examination. In addition up to ten randomly selected individuals from non-target species
collected at these sites will be examined similarly. Detailed histopathology exams will also be conducted
on collected fish that have gross pathological disorders. The results of this microscopic examination will
be used to assess the relationship between the incidence of external abnormalities and internal
histopathological abnormalities, to characterize the types of external/internal pathologies and to create a
baseline of histopathological information for the Virginian and Louisianian Provinces.
2. LABORATORY EXAMINATION OF FISH
2.1 The laboratory examination of finfish for pathological abnormalities will be conducted using two
somewhat different procedures corresponding to the two types of field sampling (i.e., base sampling and
indicator validation sampling) conducted during the Near Coastal EMAP Virginian Pilot Demonstration. The
corresponding pathology methods are described below.
2.2 Base Samples
2.2.1 Fixed specimens will be unpacked, logged in, and placed in 70% ethyl alcohol (EtOH) for at least 48
hours prior to examination. The specimens from the Base Sampling sites will be subjected to a critical gross
examination as described below and findings will be compared to the findings from the field examination.
2.2.1.1 A careful visual inspection will be made of the fins and body surfaces. Any discolorations of body
surfaces, hemorrhaging, raised scales, white spots, parasites visible to the naked eye, lumps, bumps, or
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EMAP-Estuaries Laboratory Methods Manual Section 4 - Histopathology
Volume 1 August 1995
Page 2 of 9
other growths, ulcerations, fin erosion, deformities of the vertebral column and/or mandibles, swelling of the
anus, or any other abnormal conditions will be noted.
2.2.1.2 Eyes will be examined noting any hemorrhage, exophthalmia (i.e., pop eye), microphthalmia (i.e.,
depression into the orbits), or cataracts.
2.2.1.3 A thorough examination of the branchial chamber and buccal cavity will be done identifying any
pathological abnormalities such as opercular perforations or deformities, lumps, bumps, ulcerations, gill
erosion, clubbing or other deformities, and/or parasitic infestations.
2.2.1.5 A thorough gross examination of the visceral organs will be performed and any abnormalities will
be noted.
2.2.2 Representative tissue samples from any of the gross pathologies will be taken and placed in properly
labeled tissue-processing cassettes for possible future evaluation.
2.3 Indicator Validation Samples
2.3.1 Specimens from the Indicator Validation sites will be examined grossly as described for the Base
Samples.
2.3.2 Representative tissue samples from any noted gross pathologies will be taken and placed in properly
labeled tissue-processing cassettes. All tissue specimens of gross pathologies will be processed for routine
paraffin histological evaluation.
2.3.3 Tissue samples of liver and spleen will be taken from each specimen from Indicator Validation sites
and placed in properly labeled tissue-processing cassettes. All tissue specimens will be processed for
routine paraffin histological evaluation.
3. HISTOPATHOLOGICAL PROCESSING
3.1 All tissue specimens selected for histological evaluation will be placed in properly labeled tissue-
processing cassettes.
3.2 Tissue specimens in labeled cassettes will be dehydrated, cleared, and infiltrated with Paraplast®X-
TRA using an automated tissue processor (Shandon Hypercenter® 2 Tissue Processing System). The
processing schedule is as follows:
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EMAP-Estuaries Laboratory Methods Manual
Volume 1
Section 4 - Histopathology
August 1995
Page 3 of 9
Step Reagent Temp
Vacuum Immersion Time
Drain Time*
1
2
3
4
5
6
7
8
9
10
11
12
70% EtOH
80% EtOH
95% EtOH
95% EtOH
100% EtOH
100% EtOH
100% EtOH
xylenes
xylenes
xylenes
Paraplast
Paraplast
*
A
Y
N
A
A
A
A
A
A
A
A
A
A
60
60
Seconds
Ambient
Yes
No
N
N
N
N
N
N
N
N
N
Y
Y
Y
01:00:00
01:00:00
01:00:00
01:00:00
01:00:00
00:50:00
00:50:00
01:00:00
01:00:00
01:00:00
01:00:00
01:00:00
120
120
120
120
120
120
120
120
120
120
120
120
3.3 Tissue specimens will be embedded in Paraplast®X-TRA. Labeled tissue-processing cassettes are
used in the embedding process so that the tissue sample and cassette are molded together with the
hardened Paraplast®X-TRA thus retaining the processing code with the tissue sample.
3.4 Representative sections will be cut from each tissue specimen using a rotary microtome. Sections will
be cut at 6 ^m and then floated on a waterbath containing 6 ml of Surgipath® Stayon. Sections will be
collected on slides labeled with the appropriate processing code and then placed in a drying oven (48°C)
overnight. Two slides containing two to five sections each will be prepared from each specimen.
3.5 Slides will be stained with Harris' hematoxylin and eosin for routine histological examination.
staining schedule utilized is as follows:
The
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EMAP-Estuaries Laboratory Methods Manual
Volume 1
Section 4 - Histopathology
August 1995
Page 4 of 9
Station
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
Reagent
Xylenes
Xylenes
Xylenes
100% EtOH
100% EtOH
95% EtOH
70% EtOH
Running Water
Hematoxylin
Running Water
Acid Alcohol
Running Water
Ammonia Water
Running Water
70% EtOH
Eosin Y
95% EtOH
95% EtOH
100% EtOH
100% EtOH
100% EtOH
Xylenes
Xylenes
Xylenes
Time
3 min.
3 min.
3 min.
1 min.
1 min.
1 min.
1 min.
1 min.
9 min.
1 min.
1 sec.
1 min.
1 min.
1 min.
1 min.
5 sec.
5 sec.
1 min.
1 min.
1 min.
1 min.
1 min.
1 min.
Following staining, slides will be cover-slipped using 24 x 50 mm or 24 x 55 mm, No. 1 1/
coverslips and Permount®, and dried overnight in a drying oven.
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EMAP-Estuaries Laboratory Methods Manual Section 4 - Histopathology
Volume 1 August 1995
Page 5 of 9
4. HISTOLOGICAL EXAMINATION
4.1 Slides will be examined using a compound research microscope. Notes on the condition of the tissue
specimens will be recorded on data sheets. Diagnoses of pathologic conditions will be made and the coded
data will be entered into a database management program for subsequent transfer to the EMAP Near
Coastal database. Diagnostic codes will consist of three parts, a condition identification (Table 1), a tissue
site identification (Table 2), and , where appropriate, an intensity identifier (i.e., 1 - 4 with 4 being the most
severe condition).
Table 1. Diagnostic Codes Used for Histopathologic Examination of EMAP Specimens*
DIAGNOSIS CODE
No pathologic changes NPC
GENERAL PATHOLOGIC ABNORMALITIES
Traumatic injury Tl
Granulomatous inflammation Gl
Inflammatory focus (foci) IF
Fatty degeneration FD
Autolysis AL
Congestion C
Dysplasia DY
Enteritis E
Necrosis N
Hyperplasia H
Megalocytosis M
Fibrosis FB
INFECTIOUS DISEASES
Lymphocystis L
Bacterial infections Bl
Fungal infections FU
(continued)
* This list of coded diagnoses is by no means a complete listing of possible conditions and
may be amended as necessary.
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EMAP-Estuaries Laboratory Methods Manual
Volume 1
Section 4 - Histopathology
August 1995
Page 6 of 9
Table 1. continued.
DIAGNOSIS
CODE
PARASITIC DISEASES
Protozoans
Monogeneans
Digeneans
Cestodes
Nematodes
Aeanthocephalans
Crustaceans
MISCELLANEOUS RESPONSES
Ductal proliferation
Rodlet cell response
Spongiosis hepatis
Spongiosis, kidney
PRENEOPLASTIC LIVER LESIONS
Vacuolated cell focus (foci)
Clear cell focus (foci)
Eosinophilic focus (foci)
Basophilic focus (foci)
NEOPLASTIC LESIONS
LIVER
Adenoma
Hepatocellular carcinoma
Cholangioma
Cholangiocellular carcinoma
Pericytoma
PZ
MG
DG
CT
NT
AT
CR
DP
RCR
SH
KS
VCF
CCF
EF
BF
A
HC
CL
CC
PC
(continued)
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EMAP-Estuaries Laboratory Methods Manual
Volume 1
Section 4 - Histopathology
August 1995
Page 7 of 9
Table 1. continued.
DIAGNOSIS
CODE
PANCREAS
Atypical acinar cell focus (foci)
Acinar cell adenoma
Acinar cell carcinoma
Adenocarcinoma
CARDIOVASCULAR
Hemangioma
Hemangioendothelioma
Hemangioendotheliosarcoma
Hemangiopericytoma
BONE AND CARTILAGE
Hyperostosis
Osteoma
Osteochondroma
Osteosarcoma
Chondroma
Chondrosarcoma
SOFT TISSUES
Lipoma
Liposarcoma
Fibroma
Fibrosarcoma
Leiomyoma
Leiomyosarcoma
Rhabdomyoma
Rhabdomyosarcoma
Mesothelioma
AAFC
ACA
ACC
ADC
HM
HE
HES
HP
HO
O
OC
OS
CD
CS
LP
LPS
F
FS
LM
LMS
RM
RMS
MT
(continued)
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EMAP-Estuaries Laboratory Methods Manual
Volume 1
Section 4 - Histopathology
August 1995
Page 8 of 9
Table 1. continued.
DIAGNOSIS
CODE
KIDNEY AND URINARY TRACT
Polycystic kidney
Nephroblastoma
Renal adenocarcinoma
HEMATOPOIETIC TISSUES
Lymphoma
Malignant fibrous histiocytoma
GASTROINTESTINAL TRACT
Adenoma
Adenocarcinoma
NEURAL
Neurofibroma
Schwannoma
INTEGUMENT
Papilloma
Melanoma
Squamous cell carcinoma
PK
NB
RA
LY
MFH
AD
IA
NF
SW
PL
ME
SCC
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EMAP-Estuaries Laboratory Methods Manual
Volume 1
Section 4 - Histopathology
August 1995
Page 9 of 9
Table 2. Tissue Codes
TISSUE
Barbel, mandibulary
Barbel, maxillary
Barbel, mental
Barbel, nasal
Bile duct
Blood
Blood vessel
Body (trunk), xsec
Bone
Brain
Buccal cavity
Cartilage
Caudal peduncle
Dermis (epidermis)
Dermis (site 1)
Dermis (site 2)
Dermis (site 3)
Eye
Fin, anal
Fin, caudal
Fin, dorsal
Fin, pectoral
Gall bladder
Gills
Gill arch
Gill arch, right
Gill arch, left
Used for
CODE
BD
BM
BB
BN
Bl
BL
BV
BX
BO
BR
BU
CA
CP
DM
D1
D2
D3
EY
AF
CF
DF
PF
GB
Gl
GL
G1
G2
Histopathologic Examination of EMAP
TISSUE
Gonad
Gut
Heart
Kidney
Kidney, head
Kidney, trunk
Kidney interstitium
Liver
Liver surface
Meninges
Mesentery
Mouth
Muscle
Nares
Ovary
Pancreas
Pseudobranch
Pyloric caeca
Renal tubules
Skin
Spleen
Swimbladder
Testis
Thymus
Thyroid
Visceral mass
Specimens.
CODE
GO
GT
HT
KD
KH
KT
Kl
LV
LS
MN
ME
MO
MS
NA
OV
PA
PS
PC
RT
SK
SP
SW
TE
TH
TY
VM
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EMAP-Estuaries Laboratory Methods Manual Section 5 - Sediment Grain Size
Volume 1 August 1995
Page 1 of 28
SECTION 5
SEDIMENT SILT-CLAY CONTENT
SEDIMENT GRAIN SIZE DISTRIBUTION
TOTAL ORGANIC CARBON CONCENTRATION
LABORATORY PROCEDURES
TABLE OF CONTENTS
Subsections
Introduction 2
Laboratory Safety 2
Training 3
Sample Storage and Processing 3
Procedures for Silt-Clay Content Determination 3
Procedures for Percent Water Content 6
Procedures for Sediment Grain Size Distribution Determination 9
Procedures for Sediment Total Organic Carbon Determination 16
Quality Assurance and Quality Control Procedures for Sediment Analyses 17
Data Forms 18
References 18
Appendix A 21
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EMAP-Estuaries Laboratory Methods Manual Section 5 - Sediment Grain Size
Volume 1 August 1995
Page 2 of 28
1. INTRODUCTION
1.1 Habitat indicators provide important information about the environmental setting of a sample site.
Salinity and temperature are among the most important factors controlling the distribution of biota and
ecological processes in estuaries. Organic content and grain size distribution are major sediment
characteristics that influence sediment quality and processes, as well as benthic invertebrate distributions.
Cumulatively, these parameters define the major habitats sampled by EMAP-Estuaries and information on
these habitat indicators will be essential for normalizing responses of exposure and response indicators to
natural environmental gradients. They will also be used to define subpopulations for analysis and
integration activities.
1.2 This chapter describes the laboratory procedures used to determine the silt-clay content, water content,
grain size distribution, and total organic carbon concentration of sediment collected for EMAP-Estuaries.
The procedures are designed to yield reliable, reproducible results and incorporate specific quality
assurance/quality control procedures.
1.3 The laboratory procedures are based upon currently accepted practices in benthic ecology and
sedimentology (Buchanan 1984; Plumb 1981). Although these practices are fairly standard throughout the
research community, certain procedures may require modification during a long-term (decades) program
such as EMAP. Such modifications will be kept to a minimum to ensure the long-term comparability of data
and will not occur without prior consultation with the Near Coastal Technical Director. Modifications will be
incorporated into this manual in a timely manner.
2. LABORATORY SAFETY
2.1 Safe laboratory procedures must be followed at all times and are outlined in a laboratory safety manual
which is posted in the laboratory.
2.2 Eye protection (i.e., laboratory safety glasses or approved prescription glasses) must be worn when
handling chemicals or when performing procedures that require use of a fume hood. Safety glasses are
available in the laboratory.
2.3 Respirators capable of filtering small particles must be used when grinding sediments that may contain
high concentrations of chemical contaminants.
2.4 No chemicals are to be poured down laboratory drains connecting to public or private sewer systems.
Exceptions to this rule depend upon local and state regulations concerning the disposal of hazardous
wastes.
2.5 All work areas are to be kept clean and neat at all times.
2.6 Any conditions hindering accomplishment of work or presenting a safety hazard should be brought to
the attention of the laboratory supervisor immediately.
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EMAP-Estuaries Laboratory Methods Manual Section 5 - Sediment Grain Size
Volume 1 August 1995
Page 3 of 28
3. TRAINING
3.1 A program the size of EMAP involves many field and laboratory technicians. To ensure that data
produced by all workers are comparable, standard procedures must be followed.
3.2 All personnel will have a minimum level of training. Instruction will include evaluation and proficiency
testing to insure the mastery of basic skills.
3.3 Training will be provided by experienced personnel in established laboratories. New employees will
learn laboratory techniques using practice samples and, will not handle real EMAP samples without
supervision and demonstration of acceptable proficiency.
3.4 The overall proficiency of laboratory personnel will be evaluated using the various QA/QC procedures
outlined in the methods described below. The status of personnel demonstrating substandard performance
will be reevaluated, and personnel will be retrained or removed from the laboratory, if necessary to maintain
consistent, high quality production of data.
4. SAMPLE STORAGE AND PROCESSING
4.1 Sediment samples may be chilled at 4 to 5 °C prior to processing, but samples should not be allowed
to dry before grain size analyses are conducted (Plumb, 1981).
4.2 Sieves used in the determination of sediment grain size will not be used for other purposes (e.g.,
benthic sorting). All wet sieving procedures are to be carried out using stainless steel screens. Fine
screens (63 ^m mesh) will be cleaned with copious amounts of water to prevent clogging of mesh openings.
Screens will not be cleaned with brushes, which may distort openings. Sediments will not be forced through
screens.
4.3 An analytical balance accurate to 0.1 mg will be used for all weighing. Prior to each use, the balance
will be zeroed, and its calibration will be checked using a standard weight. The same standard weight (each
standard is numbered) will be used for all weight measurements for a particular batch of samples.
5. PROCEDURES FOR SILT-CLAY CONTENT DETERMINATION
5.1 The following procedures from Folk, 1968; Lewis, 1984; and Lewis and McConchie, 1994 are used to
determine the percent by weight of silts and clays in sediment samples. Silts and clays are those particles
that pass through a 63 /^m mesh sieve. Materials retained on the sieve used in this procedure are generally
sands (>63 ^m but < 2 mm) but may include gravel sized particles (> 2mm but < 64mm -- size classification
according to the Wentworth-Lane scale, Pettijohn 1975).
5.2 Sediment samples will be retrieved from cold storage and brought to room temperature. Sample
numbers will be recorded on a silt-clay analysis data sheet upon retrieval from storage.
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EMAP-Estuaries Laboratory Methods Manual Section 5 - Sediment Grain Size
Volume 1 August 1995
Page 4 of 28
5.3 Sediments will be removed from storage bags, placed in a clean 250 ml glass beaker and
homogenized. Homogenization will be accomplished by stirring the sediment with a small spatula with a
small amount of deionized water added for lubrication (if necessary) for at least three minutes. After stirring,
rinse sediment from the spatula back into beaker using deionized water.
5.4 How Much Sediment to Use For Analysis?
5.4.1 The amount of sediment to be processed depends upon sediment type. The technician will visually
classify samples as primarily sand or primarily mud based upon the texture of sediments.
5.4.2 The best amount of sample for processing is approximately 15-20 grams of mud (i.e., sample in the
< 63 jj,m fraction). With more sample, the grains interfere with each other too much during settling and may
flocculate; with too little sample, the experimental error in weighing becomes large with respect to the
sample size.
5.4.3 For sandy sediments, approximately 45-50 g wet weight will be removed from the 250 ml glass
beaker and placed into a clean 100 ml glass beaker for wet sieving. Note the importance of Section 5.5
because of the coarseness of the sample.
5.4.4 For muddy sediments, approximately 20-25 g wet weight will be removed from the 250 ml glass
beaker to a 100 ml glass beaker for wet sieving.
5.4.5 The remaining sediment will be returned to the original storage bag and held in cold storage until all
QA/QC checks for this sample have been passed.
5.5 Dispersion of Clay Fraction of Sediments:
5.5.1 Make-up a 5g/L stock solution of dispersant. Add 5 grams of sodium hexametaphosphate "Calgon"
to 1 liter of deionized water.
5.5.2 Add 20 ml of the dispersant solution (100 mg of hexametaphosphate) and 30 ml of distilled water to
the sample. Stir, using a magnetic stirrer for one to five minutes to break-up sediment aggregrates.
5.6 Wet Sieving the Sample:
5.6.1 After stirring, the sample will be wet sieved through a 63 /^m mesh sieve into a large evaporation dish
using as little distilled water as possible.
5.6.2 Place the sieve over the large evaporation dish and wash all fines into the sieve using as little distilled
water as possible.
5.6.3 The volume of sediment + water in the evaporation dish must be < 900 ml to allow for rinsing the
sample into a 1000 ml graduated cylinder.
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EMAP-Estuaries Laboratory Methods Manual Section 5 - Sediment Grain Size
Volume 1 August 1995
Page 5 of 28
5.7 Analysis of the Silt and Clay Fraction (Particles < 63
5.7.1 Carefully transfer the mud in the evaporation dish to a 1000 ml graduated cylinder. Carefully rinse
the mud (generally medium-coarse silt-size (16-63 urn particles) found at the bottom of the dish into the
graduated cylinder using deionized water, being careful not to exceed the 1000 ml mark. See Section 5.8
for handling the >63 urn fraction that remains on the sieve.
5.7.2 Fill, with deionized water, up to the 1000 ml mark. Using a metal stirring rod, vigorously stir the water
column from bottom to top, using short strokes, starting at the base of the column and working upwards.
Keep stirring until the material is distributed uniformly throughout the column. End up stirring with
long,smooth strokes the full length of the column. Be careful not to break the water surface as material
could be lost. Place a beaker with tap water next to the cylinder and insert a thermometer to record water
temperature.
5.7.3 Immediately (20 sec) after stirring, withdraw 40 ml of sample using a 40 ml volumetric pipette. Expel
sample into a tared 50 ml glass beaker. Rinse pipette with a small volume of deionized water, and add the
rinse to the 50 ml beaker. If the sample is taken in two parts (i.e., two 20-ml samples), the cylinder will be
stirred between extractions and samples withdrawn after each stirring and added to the beaker.
5.7.4 The 50 ml glass beaker will be placed in an oven at 100°C until dry. Typically, 24 hours is sufficient
for sediment samples to come to a stable dry weight. A randomly selected subsample of each batch will
be reweighed after an additional 24 hour drying period, as a check for the stability of the dry weight
measurement.
5.8 Treatment of >63 urn Fraction Retained on Sieve:
5.8.1 If necessary, remove shell and shells fragments, pieces of wood and algae. Place in plastic weigh
pans to air dry. Record weights. Note that these weights are not part of the sand or silt/clay fraction but are
part of the sediment record.
5.8.2 The fraction retained on the sieve (>63 ^m) will be transferred to a tared 50 ml glass beaker and
placed in an oven at 100 °C until dry. Typically, 24 hours is sufficient for the dry weight of sediment
samples to stabilize. All samples will be weighed after 24 hours; the weight will be recorded on the biomass
data sheets, and the samples will be returned to the drying oven. A randomly selected subsample of each
batch will be reweighed after an additional 24-hour drying period, as a check for the stability of the dry
weight measurement.
5.9 Storage of Unused Samples:
5.9.1 Unused sediment from each sample will be stored at 4 to 5 °C for QA/QC analyses and other
sediment analyses.
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EMAP-Estuaries Laboratory Methods Manual Section 5 - Sediment Grain Size
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5.10 Weighing Samples:
5.10.1 After drying, remove beaker from the oven and let equilibrate with the atmosphere for at least 1.5
hr before weighing. Weigh to nearest 0.001 grams and record weight.
5.11 Calculations for Sand and Silt-Clay (Mud) Determinations:
5.11.1 Sand weight calculation:
Sane/ wt. = Gross wt. (sample + beaker) - tare wt. (beaker)
5.11.2 Silt-clay (mud) weight calculation:
Silt-clay wt. =
(Gross wt. - beaker wt.) x «°tal volume in
(sample volume from cylinder)
dispersant
weight
Note: The total volume in cylinder is 1000 ml. The sample volume from cylinder is 40 ml. Using the
prescribed methods, this results in a dispersant weight of 4 mg.
5.11.3 Percent silt-clay (mud) calculation:
% silt-clay - - wt x 100
sane/ wt. + silt-clay wt.)
5.11.4 Percent sand calculation:
% sand = 100 - %mud
Note: (100 - % mud) is not, in all cases, equal to the percent sand, since gravel sized particles (>
2mm but < 64 mm) may be present in some samples.
6. PROCEDURES FOR PERCENT WATER CONTENT
6.1 The following procedures are used to determine the percent by weight of water in sediment samples.
The percent water content of sediment samples is needed to correct sediment dry weights for salt content,
since salts are left behind in the drying process.
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6.2 Sample Retrieval:
6.2.1 Sediment samples will be retrieved from cold storage and brought to room temperature. Sample
numbers will be recorded on a data sheet upon retrieval from storage. Samples that have dried cannot be
processed, since dried sediments yield erroneous results.
6.3 Sample Preparation:
6.3.1 Sediments will be removed from storage bags, placed in a clean 250 ml glass beaker and
homogenized. Homogenization will be accomplished by stirring sediment with a small spatula for at least
three minutes. Do not rinse sediment from the spatula into the beaker and do not add water to the
beaker during the homogenization process!
6.3.2 Approximately 5-10 grams wet weight of sediment will be placed in a clean, tared 50 ml glass beaker
(or 3.3 ml assuming a wet weight density of 1.5).
6.4 Recording Sediment Weight:
6.4.1 Weigh sample immediately. The sample must not be allowed to stand for more than a few minutes,
since evaporation at room temperature will affect the percent water content measurement.
6.5 Drying the Sample:
6.5.1 The sample will be placed in a drying oven at 100°C until dry. Typically, 24 hours is sufficient for the
dry weight of sediment samples to stabilize. Dry samples will be stored in a desiccator containing a hydrous
silica gel until cooled to touch (approximately 1 hr). (Note: Dry sediment samples may absorb moisture
from wet sediment samples, thus, dry samples should be removed before placing moist samples in the
oven). All weights will be recorded on data sheets to the nearest 0.001 grams. Ten percent (10%) of
randomly selected subsamples of each batch will be reweighed after an additional 24-hour drying period,
as a check for stability for the dry weight measurement (a change of less than 0.1% is expected).
6.6 Disposal of Unused Sediment
6.6.1 Unused sediment from each sample will be stored at 4 to 5°C for QA/QC analyses and other
sediment analyses.
6.7 Calculation of Sediment Water Content:
6.7.1 The following procedures will be used to calculate the water content of sediment. Because dry salts
are included in the dry sediment weight, a correction must be applied to account for this (section 6.8.3).
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EMAP-Estuaries Laboratory Methods Manual Section 5 - Sediment Grain Size
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6.7.2 Sediment water content calculation:
% water = ( (Gross wet wt ~ fare) " (Gross dry wt ~ fare)] x 100
{ Gross wet wt. - tare }
6.7.3 Correction of dry weight for salt content.
6.7.3.1 Equations:
Corrected dry weight = (Gross Dry wt. - tare) - (Salt weight))
Salt weight (g) = Water loss (ml) x Salinity (mg/ml [ = %o = ppt ])
Water loss (mI) = Gross wet weight (wet sample + pan)-Gross dry weight (dry sample + pan)
assuming a water density of 1 g/ml for the water that evaporates (fresh water)
6.7.3.2 Example calculation of corrected dry weight:
Sample salinity = 12.4 %o (as a first approximation, the salinity of interstitial water is assumed to
be equal to the salinity of bottom water as measured by
EMAP CTD casts).
Beaker Weight = 31.144 g
Wet Sediment + Beaker Weight (Gross Wet Wt) = 39.219 g
Dry Sediment + Beaker Weight = 37.135 g
Known: Water Density = 1g/ml
Calculations:
Water Loss (ml) = (Wet Sdmt Wt + Bkr Wt) - (Dry Sdmt Wt + Bkr Wt)
= 39.219 g - 37.135 g
= 2.084 g
= 2.084ml
Salt Weight (g) = Water Loss (ml) x Salinity (mg/ml)
= 2.084ml x 12.4 mg/ml
= 25.84mg
= .0258g
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Corrected Dry Weight (g) = ((Dry Sdmt + Bkr Wt) - Bkr Wt) - Salt Weight
= (37.135g-31.144g) - 0.0258g
= 5.991 g - 0.0258g
= 5.965g
Corrected Percent Water = ((Wet Sdmt + Bkr Wt) - Bkr Wt) - Corrected Dry Wt.
xlOO
((Wet Sdmt + Bkr Wt) - Bkr Wt)
= (39.219-31.144)-5.965
x 100 = 26
39.219-31.144
7. PROCEDURES FOR SEDIMENT GRAIN SIZE DISTRIBUTION DETERMINATION
7.1 The following procedures from Folk, 1968; Lewis, 1984; and Lewis and McConchie, 1994 are used to
determine the percent by weight of silts and clays in sediment samples. Silts and clays are those particles
that pass through a 63 ^m mesh sieve. Materials retained on the sieve used in this procedure are generally
sands (>63 /^m but < 2 mm) but may include gravel sized particles (> 2mm but < 64mm -- size classification
according to the Wentworth-Lane scale, Pettijohn 1975). The procedures allow determination of weight
percent quantiles for sediments, as well as the quantile deviation of skewness.
7.2. Sediment samples will be retrieved from cold storage and brought to room temperature. Sample
numbers will be recorded on a sediment grain size data sheet upon retrieval from storage.
7.3 How Much Sediment to Use For Analysis?
7.3.1 The best amount of sample for processing is approximately 15-20 grams of mud (i.e., sample in the
< 63 ^m fraction). With more sample, the grains interfere with each other too much during settling and may
flocculate; with too little sample, the experimental error in weighing becomes large with respect to the
sample size.
7.3.2 For sandy sediments, approximately 45-50 g wet weight will be removed from the 250 ml glass
beaker and placed into a clean 100 ml glass beaker for wet sieving. Note the importance of Section 5.5
because of the coarseness of the sample.
7.3.3 For muddy sediments, approximately 20-25 g wet weight will be removed from the 250 ml glass
beaker to a 100 ml glass beaker for wet sieving.
7.3.4 The remaining sediment will be returned to the original storage bag and held in cold storage until all
QA/QC checks for this sample have been passed.
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EMAP-Estuaries Laboratory Methods Manual Section 5 - Sediment Grain Size
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7.4 Sample Handling:
7.4.1 Sediments will be removed from storage bags, placed in a clean 250 ml glass beaker, and
homogenized. Homogenization will be accomplished by stirring sediment with a spatula and a small amount
of deionized water for at least three minutes.
7.5 Removal of Organic Matter for the Determination of "True" Particle Size Distribution (Lewis, 1986):
7.5.1 If the sample is equal to or less than 20% mud (silt-clay), proceed to Section 7.6
7.5.2 If the sample is greater than 20% mud, the organics in the sample must be removed.
7.5.3 Initially, add enough deionized water to cover the sample. Add small quantities of 30% H2O2 to the
sample, stirring until any effervescence ceases. Cover beaker with large watch glass cover if frothing is
excessive. If the solution heats excessively, cool the beaker in a water bath. Continue adding H2O2 until
frothing ceases, then slowly heat to 60-70°C (H2O2 decomposes above 70°C). Observe for 10 mins. to
ensure that the possibility of a strong reaction has passed. Add H2O2 until no further reaction occurs.
7.6 Dispersion of the Clays
7.6.1 Make-up a stock solution of dispersant. Add 5 grams of sodium hexametaphosphate "Calgon" to 1
liter of deionized water.
7.6.2 Add 20 ml of the dispersant solution (100 mg of hexametaphosphate) and 30 ml of distilled water to
the sample. Stir, using a magnetic stirrer for one to five minutes to break-up sediment aggregrates.
7.7 Wet Sieving the Sample:
7.7.1 After stirring, the sample will be wet sieved through a 63 /^m mesh sieve into a large evaporation dish
using as little distilled water as possible.
7.7.2 Place the sieve over the large evaporation dish and wash all fines into the sieve using as little distilled
water as possible.
7.7.3 The volume of sediment + water in the evaporation dish must be < 900 ml to allow for rinsing the
sample into a 1000 ml graduated cylinder.
7.7.4 Carefully transfer the mud in the evaporation dish to a 1000 ml graduated cylinder. Carefully rinse
the mud (generally medium-coarse silt-size (16-63) urn particles) found at the bottom of the dish into the
graduated cylinder using deionized water, being careful not to exceed the 1000 ml mark. See Section 7.9
for handling the sand-size and greater fraction that remains on the sieve (>63 urn fraction).
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EMAP-Estuaries Laboratory Methods Manual Section 5 - Sediment Grain Size
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7.7.5 Fill, with deionized water, up to the 1000 ml mark. Using a metal stirring rod, vigorously stir the water
column from bottom to top, using short strokes, starting at the base of the column and working upwards.
Keep stirring until the material is distributed uniformly throughout the column. End up stirring with
long,smooth strokes the full length of the column. Be careful not to break the water surface as material
could be lost. Place a beaker with tap water next to the cylinder and insert a thermometer to record water
temperature.
7.8 Analysis of the Silt and Clay ( < 63 urn) Fraction :
7.8.1 Stir the cylinder to suspend the sample in accordance with procedures in Section 7.7.5. As soon as
the stir rod emerges for the last time, start the timer. At the end of 20 seconds, insert the pipette to a depth
of 20 cm and withdraw exactly 20 ml. This is the most important single step in this exercise as
subsequent analyses are based on the calculation of the total mud weight. Continue to withdraw 20-ml
samples with the 20 ml volumetric pipette at the depths and times indicated on Table 1 for the recorded
water temperature (from Lewis and McConchie,1994).
7.8.2 Transfer the pipette sample fractions to separate tared 50 ml glass beakers. Each pipette withdrawal
should be rinsed with a small volume of deionized water which is then added to the 50 ml sample beaker.
7.8.3 Each beaker should be placed in an oven at 100° C for 24 hrs. All weights are to be recorded on the
data sheet.
7.8.4 Unused sediment from each sample will be stored at 4 to 5 °C for QA/QC analyses and other
sediment analyses.
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EMAP-Estuaries Laboratory Methods Manual
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Section 5 - Sediment Grain Size
August 1995
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EMAP-Estuaries Laboratory Methods Manual
Volume 1
Section 5 - Sediment Grain Size
August 1995
Page 13 of 28
Table 1. continued. Calculations for determining sample withdrawal times for pipette analyses.
Template for Sediment Grain Size Analysis:
Sample withdrawal times for pipette analysis are based upon Stoke's law which can be written as:
T = Depth/[1500*A*(d2)] (Folk, 1968)
where T is time in minutes,
Depth is in centimeters,
A is a contant, and
d is the particle diameter in millimeters.
The A value is a function of temperature, gravity, and density of particles. We will assume a density of 2.65
(associated with quartz or clay minerals). The following table relates various temperatures to the A
constant.
Temperature
(degrees °C) A
20
21
22
23
24
25
26
3.57
3.66
3.75
3.84
3.93
4.02
4.12
Temp ? 25 =>Enter temperature.
A value = 4.02 =>The A value will automatically be determined.
Time? 0:00:00 =>Enter the start time for the analysis.
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EMAP-Estuaries Laboratory Methods Manual Section 5 - Sediment Grain Size
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7.9 Treatment of the Sand Fraction (> 63 |jm) Retained on the Sieve
7.9.1 Transfer the >63 /^m fraction to a 250 ml glass beaker and place in a drying oven at 100°C until dry
for 24 hrs.
7.9.2 Transfer the dried sediment into the top of a stack of clean, stainless steel sieves composed of 500
^m(1.00), 355 ^m(»1.50), 250//m (2.00), 180 //m (»2.5 0), and 125 ^m (3.00), 90 ^m (-3.5 0) and
63 //m (4.0 0) sieve with a closed pan on the bottom. Shake on a rotary tapper (Ro-tap) for 15 minutes.
7.9.3 Weigh each sieved fraction as follows: Tare a 100 ml beaker to zero; add the 500 /^m (1.0 0)
sediment fraction and weigh to 0.001 grams. Next add the 355 ^m (~ 1.5 0) fraction to the beaker. Proceed
to add subsequent fractions until all weighed. Record the individual and cumulative weights of the > 63 /^m
fraction to 0.001 grams.
7.10 Removal of Carbonates (if necessary):
7.10.1 If a sediment contains >50%, by weight, of calcareous material, the sample is described as a
carbonate sediment (Blatt et a/., 1975). The following steps will be followed to process carbonate
sediments. Record the weight of the sand fraction to 0.001 grams. A 10% (by volume) HCI solution will be
added to the dried and weighed sediment. Cover the sediment completely with HCI and let sit for four
hours. Additional acid will be added and if foaming is apparent, the sample will be left to stand for several
more hours. This process will be repeated until no further reaction occurs with subsequent additions of HCI.
7.10.2 The sample will be transferred to a 63 /^m sieve and washed using copious amounts of deionized
water. This will remove any salts formed during the acidification step.
7.10.3 The sample will be transferred to a 100 ml glass beaker, dried, and weighed to 0.001 grams.
Proceed with steps in Section 7.9.
7.10.4 Calculate the sediment carbonate content:
{wt. of beaker and sed. (before acid) - wt. of beaker and sed. (after acid)}
% Carbonate =
{wt. of beaker and sed. (before acid) - beaker wt.}
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EMAP-Estuaries Laboratory Methods Manual Section 5 - Sediment Grain Size
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7.11 Calculations for Sediment Grain Size Distributions
7.11.1 Calculate the total weight of mud (silt-clay) in the sample (obtained @ 20 sec withdrawal).
Total mud weight (g) = [((beaker+ sed wt.) - beaker wt.) x 50] - dispersant weight
Dispersal weight =0.1 gram, if procedures in Section 7.6.1 were followed.
(note: the amount of mud in each 20 ml withdrawal is equal to 1/50 of the total amount of mud
remaining in the 1000 ml cylinder at the withdrawal time and at the withdrawal depth)
7.11.2 Calculate the total sample weight:
Total sample weight (g) = total mud weight + total sand weight
7.11.3 Determination of cumulative percentages for each mud (<63 urn )size fraction:
Each pipette sample represents material in the column finer than a certain grain size. To begin, multiply
each size fraction by 50, subtract the weight of dispersant:
50 fraction wt. (g) = (50 sample weight X 50) - dispersant weight (g)
60 fraction wt. (g) = (60 sample weight-X 50) - dispersant weight (g)
70 fraction wt. (g) = (70 sample weight X 50) - dispersant weight (g)
80 fraction wt. (g) = (80 sample weight X 50) - dispersant weight (g)
90 fraction wt. (g) = (90 sample weight X 50) - dispersant weight (g)
Then divide each fraction by the total sample weight, subtract from 1, and multiply the product by 100:
Cumulative % (at 5 0)= [1 - (5 0 fraction wt. (g)/ Total sample weight (g))] x 100
Cumulative % (at 6 0)= [1 - (6 0 fraction wt. (g)/ Total sample weight (g))] x 100
Cumulative % (at 7 0)= [1 - (7 0 fraction wt. (g)/ Total sample weight (g))] x 100
Cumulative % (at 8 0)= [1 - (8 0 fraction wt. (g)/ Total sample weight (g))] x 100
Cumulative % (at 9 0)= [1 - (9 0 fraction wt. (g)/ Total sample weight (g))] x 100
7.11.4 Determination of cumulative percentages for each sand (<63 urn) fraction:
The weight of each sand fraction is:
% Wt. of each sand fraction = (wt. of sand for each 0 size / total sample weight) x 100
Add these percentages incrementally to obtain cumulative weight percentages.
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EMAP-Estuaries Laboratory Methods Manual Section 5 - Sediment Grain Size
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7.12 Determination of Statistical Parameters of Grain Size:
7.12.1 Plot the cumulative curve of the sample and read the 0 values which correspond to the 24th (025),
50th (i.e., median (Md0)) and 75th (075) percentiles by linear interpolation.
7.12.2 Calculate the Phi Quartile Deviation (QD0):
Phi Quartile Deviation = (075 - 025) / 2
7.12.3 Calculate the Phi Quartile Skewness (Skq0):
Phi Quartile Skewness = ((025 + 075) - (2 x Md0)) / 2
7.12.4 Record the Md0, QD0, and Skq0 for each sample.
8. PROCEDURES FOR SEDIMENT TOTAL ORGANIC CARBON DETERMINATION
8.1 The following procedures are used to determine the total organic carbon concentration in sediment.
Total organic carbon will be determined by combusting pre-acidified samples at high temperature and
measuring the volume of carbon dioxide gas produced (Salonen 1979b). Analytical procedures are
generalized. Although the basic method used by various laboratories may be the same, instrumentation
differences prohibit giving specific, step-by-step procedures.
8.2 Sample Preparation:
8.2.1 Sediment samples will be retrieved from cold storage and brought to room temperature. Sample
numbers will be recorded on a data sheet upon retrieval from storage. Unlike other procedures, samples
that have dried can be analyzed to determine total organic carbon concentration.
8.2.2 Sediments will be removed from storage bags, placed in a clean beaker, and homogenized.
Homogenization will be accomplished by stirring sediment with a small metal spatula for at least three
minutes.
8.2.3 A sample of at least 5 g wet weight (3.3 ml) will be placed in a tared evaporating dish and covered
to protect the sample against contamination with carbon from other sources.
8.2.4 The sample will be placed in a drying oven at 60 °C until dry. Typically, 48 hours is sufficient for the
dry weight of sediment samples to stabilize. A randomly selected subsample of each batch will be
reweighed after an additional 24 hour drying period as a check for the stability of the dry weight
measurement. All weights will be recorded on the data sheet.
8.2.5 The dried sediment will be transferred to a porcelain mortar and thoroughly ground with a porcelain
pestle. Obvious large shell fragments should be removed with metal forceps, being careful not to
contaminate the sample with foreign material.
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8.2.6 The ground sediment will be placed in cleaned glass scintillation vials with foil-lined caps. All labels
should remain on the exterior of the vials to minimize carbon contamination.
8.2.7 Samples prepared as above may be stored at room temperature indefinitely.
8.3 Sample Analysis:
8.3.1 Approximately 20 to 30 mg of the dried and ground sediment samples will be placed into small
beakers. The samples will be acidified to remove sources of inorganic carbon (e.g., shell fragments). This
is accomplished by suspending the beakers over a concentrated pool of HCI in a sealed desiccator
(Lambert and Oviatt, 1986). Samples are typically fumed for at least 15 hours.
8.3.2 After fuming, the samples are transferred to an oven and dried for an additional two hours (100°C).
For muddy sediment, 5 to 10 mg of sample is placed inside an aluminum vial and crimpled into a small ball.
For sandy sediment, as much as 50 to 100 mg of sample may be required due to the low organic content
generally associated with sands.
8.3.3 For analyses, samples are typically placed in a tube furnace at approximately 950°C and exposed
to a precombusted stream of oxygen. The CO2 evolved is measured by an infrared gas analyzer, and the
resulting gas peak is integrated. Integrator units are compared to a standard curve to convert to organic
carbon.
8.3.4 Standard curves for analyses are typically made with a high purity organic compound such as D-
glucose dissolved in low organic distilled water. Curves are checked with sediment standards (MESS and
BCSS) from The Marine Analytical Chemistry Standards Program of the National Research Council of
Canada.
9. QUALITY ASSURANCE AND QUALITY CONTROL PROCEDURES FOR
SEDIMENT ANALYSES
9.1 Quality control for the sediment analysis procedures will be accomplished by reanalyzing samples that
fail either a range check or recovery check. Quality assessment will include reanalysis of 10% of the
samples. Quality assessment samples will be selected randomly. Reanalysis will consist of repeating the
sediment analysis procedure on the archived sample collected from the same grab as the sample failing
QC.
9.2 For the range check, any sample results that fall outside expected ranges will be reanalyzed. For
example, any percentage that totals greater than 100% will be reanalyzed. For the recovery check, if the
total weight of the recovered sands is 10% (by weight) less or greater than the starting weight of sands, the
sample will be reanalyzed.
9.3 For Quality Assurance, samples will be selected randomly from each batch and reanalyzed. A batch
of samples is a set of samples of a single textural classification (i.e., mud), processed by a single
technician, using a single procedure (i.e., complete sediment analysis).
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9.4 Sediment sample reanalysis will be done in the following manner:
9.4.1 Approximately 10% of each batch completed by the same technician will be reanalyzed.
9.4.2 A random selection of the samples will be processed in the same manner as the original sample
batch.
9.4.3 If the absolute difference between the original number (silt-clay percentage, for example) and the
second number is greater than 10% then a third analysis will be completed by a different technician.
9.4.4 The values closest to the third value will be entered into the data base.
9.4.5 If more than 10% of the data from a batch are in error, then the whole batch will be reprocessed using
the archived sediment. A third check of the reanalyzed samples will be completed by a different technician
to assure that the reanalyzed values are correct.
9.4.6 Reanalysis and QA checks must be accomplished within 30 days from the date the original sediment
analysis was conducted.
9.4.7 Reanalysis and QA checks are dependent upon having enough sediment to complete the various
reanalyses.
10. DATA FORMS
10.1 Blank copies of data forms used for the analysis of sediment samples are presented in Appendix A
in the following order:
• Sediment data sheet for silt-clay analysis
• Sediment data sheet for grain-size distribution analysis
• Example probability paper for plotting grain-size distributions
11. REFERENCES
Blatt, H., G. Middleton, and R. Murray. 1972. The Origin of Sedimentary Rocks. Prentice-Hall, Inc.,
Englewood Cliffs, New Jersey, 634 p.
Folk, R.L. 1968. Petrology of Sedimentary Rocks. Hemphill Publishing Company, Austin, Texas, pp 33-46.
Lambert, C.E. and C.A. Oviatt. 1986. Manual of biological and geochemical techniques in coastal areas.
MERL Series Report No. 1, Second Edition. University of Rhode Island, Kingston, Rl.
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EMAP-Estuaries Laboratory Methods Manual Section 5 - Sediment Grain Size
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Lewis, D.W. 1984. Practical Sedimentology. Hutchinson Publishing Company, Stroudsburg, Pennsylvania,
pp 80-100.
Lewis, D.W. and D. McConchie 1994. Analytical Sedimentology. Chapman and Hall. New York , p. 92-109.
Pettijohn, F.J. 1975. Sedimentary Rocks. Harper and Row, New York.
Plumb, R.H. 1981. Procedure for handling and chemical analysis of sediment and water samples.
Prepared for the U.S. Environmental Protection Agency/Corps of Engineers Technical Committee on
Criteria for Dredge and Fill Material. Published by Environmental Laboratory, U.S. Army Waterways
Experiment Station, Vicksburg, MS. Technical Report EPA/CE-81-1.
Salonen, K. 1979a. A versatile method for the rapid and accurate determination of carbon by high
temperature combustion,. Limnol Oceanogr. 24:177-183
Salonen, K. 1979b. The selection of temperatures for high temperature combustion of carbon. Acta
Hydrochim. Hydrobiol. 1-10.
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APPENDIX A
DATA FORMS
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(See SED-DATA worksheet in Sec5.xls)
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(See GRAIN SIZE worksheet in Sec5.xls)
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EMAP-Estuaries Laboratory Methods Manual Section 6 - Suspended Solids
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SECTION 6
RESIDUE, NON-FILTERABLE
(SUSPENDED SOLIDS)
(Gravimetric, Dried at 103-105°C)
1. SCOPE AND APPLICATION
1.1 This method is applicable to drinking, surface, and saline waters, domestic and industrial wastes.
1.2 The practical range of the determination is 4 mg/L to 20,000 mg/L.
2. SUM MARY OF METHOD
2.1 A well-mixed sample is filtered through a glass fiber filter, and the residue retained on the filter is dried
to constant weight at 103-105°C.
2.2 The filtrate from this method may be used for Residue, Filterable.
3. DEFINITIONS
3.1 Residue, non-filterable is defined as those solids which are retained by a glass fiber filter and dried to
constant weight at 103-105°C.
4. SAMPLE HANDLING AND PRESERVATION
4.1 Non-representative particulates such as leaves, sticks, fish, and lumps of fecal matter should be
excluded from the sample if it is determined that their inclusion is not desired in the final result.
4.2 Preservation of the sample is not practical; analysis should begin as soon as possible. Refrigeration
or icing to 4°C, to minimize microbiological decomposition of solids, is recommended.
5. INTERFERENCES
5.1 Filtration apparatus, filter material, pre-washing, post-washing and drying temperature are specified
because these variables have been shown to affect the results.
5.2 Samples high in Filterable Residue (dissolved solids), such as saline waters, brines and some wastes,
may be subject to a positive interference. Care must be taken in selecting the filtering apparatus so that
washing of the filter and any dissolved solids in the filter (7.5) minimizes this potential interference.
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6. APPARATUS
6.1 Glass fiber filter discs, without organic binder, such as Millipore AP-40, Reeves Angel 934-AH, Gelman
type A/E, or equivalent.
NOTE: Because of the physical nature of glass fiber filters, the absolute pore size cannot be
controlled or measured. Terms such as "pore size" collection efficiencies and effective retention are
used to define this property in glass fiber filters. Values for these parameters vary for the filters listed
above.
6.2 Filter support: Filtering apparatus with reservoir and a coarse (40-60 m) fritted disc as a filter support.
NOTE: Many funnel designs are available in glass or porcelain. Some of the most common are
Hirsch or Buchner funnels, membrane filter holders and Gooch crucibles. All are available with
coarse fritted disc.
6.3 Suction flask.
6.4 Drying oven, 103-105°C.
6.5 Desiccator.
6.6 Analytical balance, capable of weighing to 0.1 mg.
7. PROCEDURE
7.1 Preparation of glass fiber filter disc: Place the glass fiber filter on the membrane filter apparatus or
insert into bottom of suitable Gooch crucible with wrinkled surface up. While vacuum is applied, wash the
disc with three successive 20 ml volumes of distilled water. Remove all traces of water by continuing to
apply vacuum after water has passed through. Remove filter from membrane filter apparatus or both
crucible and filter if Gooch crucible is used, and dry in an oven at 103-105°C for one hour. Remove to
desiccator and store until needed. Repeat the drying cycle until a constant weight is obtained (weight loss
is less than 0.5 mg). Weigh immediately before use. After weighing, hand the filter or crucible/filter with
forceps or tongs only.
7.2 Selection of Sample Volume
For a 4.7 cm diameter filter, filter 100 ml_ of sample. If weight of captured residue is less than 1.0 mg,
the sample volume must be increased to provide at least 1.0 mg of residue. If other filter diameters
are used, start with a sample volume equal to 7 ml/cm2 of filter area and collect at least a weight of
residue proportional to the 1.0 mg stated above.
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NOTE: If during filtration of this initial volume the filtration rate drops rapidly, or if filtration time
exceeds five to ten min, the following scheme is recommended: Use an unweighed glass fiber filter
of choice affixed in the filter assembly. Add a known volume of sample to the filter funnel and record
the time elapsed after selected volumes have passed through the filter. Twenty-five ml increments
for timing are suggested. Continue to record the time and volume in increments until filtration rate
drops rapidly. Add additional sample if the filter funnel volume is inadequate to reach a reduced rate.
Plot the observed time versus volume filtered. Select the proper filtration volume as that just short
of the time a significant change in filtration rate occurred.
7.3 Assemble the filtering apparatus and begin suction. Wet the filter with a small volume of distilled water
to seat it against the fritted support.
7.4 Shake the sample vigorously and quantitatively transfer the predetermined sample volume selected
in Sect. 7.2 to the filter using a graduated cylinder. Remove all traces of water by continuing to apply
vacuum after sample has passed through.
7.5 With suction on, wash the graduated cylinder, filter, non-filterable residue and filter funnel wall with
three portions of distilled water allowing complete drainage between washing. Remove all traces of water
by continuing to apply vacuum after water has passed through.
NOTE: Total volume of wash water used should equal approximately 2 ml/cm2. For a 4.7 cm filter
the total volume is 30 ml_.
7.6 Carefully remove the filter from the filter support. Alternatively, remove crucible and filter from crucible
adapter. Dry at least one hour at 103-105°C. Cool in a desiccator and weigh. Repeat the drying cycle until
a constant weight is obtained (weight loss is less than 0.5 mg).
8. CALCULATIONS
8.1 Calculate non-filterable residue as follows:
Non-filterable residue (mg/L) = (^ 5) x 1,000
where:
A = weight of filter (or filter and crucible) + residue in mg
B = weight of filter (or filter and crucible) in mg
C = ml of sample filtered
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9. PRECISION AND ACCURACY
9.1 Precision data not available at this time.
9.2 Accuracy data on actual sample cannot be obtained.
10. REFERENCE
NCASI Technical Bulletin No. 291, March 1977. National Council of the Paper Industry for Air and Stream
Improvement, Inc., 260 Madison Ave., New York.
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