DRAFT
EPA-600/4-83-000
REF: 3941C
METHODS FOR USE OF CAGED MOLLUSCS FOR IN SITU BIOMONITORING
OF MARINE SEWAGE DISCHARGES
Edited
by
Cornelius I. Weber, Ph.D.
Biological Methods Branch
ENVIRONMENTAL MONITORING AND SUPPORT LABORATORY
U.S. ENVIRONMENTAL PROTECTION AGENCY
CINCINNATI, OHIO 45268
MAY 1983
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DISCLAIMER
This report has been reviewed by the Environmental Monitoring and
Support Laboratory, U.S. Environmental Protection Agency, and approved
for publication. The mention of trade names or commercial products does
not constitute endorsement or recommendation for use.
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FOREWORD
Environmental measurements are required to determine the quality of
ambient water, the character of effluents, and the effects of pollutants
on aquatic life. The Environmental Monitoring and Support Laboratory-
Cincinnati conducts research to develop, evaluate, standardize and
promulgate methods to:
Measure the presence and concentration of physical, chemical and
radiological pollutants in water, wastewater, bottom sediments,
and solid waste.
Concentrate, recover, and identify enteric viruses, bacteria, and
other microorganisms in water.
Measure the effects of pollution on freshwater, estuarine, and
marine organisms, including the phytopiankton, zooplankton,
periphyton, macrophyton, macroinvertebrates, and fish.
Automate the measurement of the physical, chemical, and
biological quality of water.
Conduct an Agencywide quality assurance program to assure
standardization and quality control of systems for monitoring
water and wastewater.
Section 301(h) of the Clean Water Act of 1977, as amended by the
Municipal Wastewater Treatment Construction Grant Amendments of 1981,
provides publicly owned treatment works (POTW's) currently discharging or
proposing to discharge to coastal and saline estuarine waters an
opportunity to apply for variances from secondary treatment
requirements. This report provides recommended monitoring methods for
use by POTW's seeking to determine compliance with the statuatory
criteria listed under Section 301(h) and applicable Water Quality
Standards.
Robert L. Booth
Acting Director
Environmental Monitoring and Support
Laboratory - Cincinnati
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PREFACE
This manual was prepared with the assistance of the following workgroup
members.
a. U.S. Environmental Protection Agency
Robert Bastian, Office of Marine Discharge Evaluation, Washington, DC
Bruce Boese, Environ. Res. Lab., Corvallis, OR
Donald Phelps, Ph.D., Environ. Res. Lab., Narragansett, RI
Richard Swartz, Ph.D., Environ. Res. Lab., Corvallis, OR
b. Southern California Coastal Water Research Project
David Brown, Ph.D., Long Beach, CA
Henry Schafer, "
c. National Oceanographic and Atmospheric Administration
Paul Dammann, Ocean Acoustics Lab, Miami, FL
Alan Mearns, Ph.D., Office Mar. Poll. Assessment, Seattle, WA
d. California Department of Fish & Game
John Ladd, Sacramento, CA
Michael Martin, Ph.D., Monterey, CA
e. Tetra Tech
Thomas Ginn, Ph.D., Bellevue, WA
f. Dames & Moore, Marine Services
David Young, Ph.D., Los Angeles, CA
Cornelius I. Weber, Ph.D.
Chief, Biological Methods Branch
Environmental Monitoring & Support Laboratory
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ABSTRACT
This manual describes methods for use of caged molluscs in
biomonitoring programs related to Section 301(h), Public Law 95-217.
Molluscs collected at relatively contaminant-free locations are placed in
cages and exposed for one month at a minimum of two stations: (1) in the
plume, within the zone of initial dilution, and (2) at a nearby reference
(control) station, outside of the area of immediate influence of the
discharge. At the end of the exposure period, the organisms are
retrieved, checked for mortality, analyzed for toxic substances, and
examined for indications of sublethal biological effects, including scope
for growth, and the distribution of toxic metals in the detoxification
system.
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CONTENTS
Foreword ill
Preface iv
Abstract v
Figures viii
Acknowledgements ix
1. Introduction 1
2. Exposure Site Selection 3
3. Exposure Depth 4
4. Exposure Systems 5
Anchors 5
Line 5
Buoys 5
Exposure Cages/Bags 6
Gear Configuration 6
5. Test Organisms 8
Recommended Species 8
Size 8
Source and Condition 8
6. Sample Exposure and Retrieval 10
Exposure Procedures 10
Exposure Period 10
Sample Retrieval 10
Field Observations 11
Sample Preservation and Transport 11
7. Chemical Analyses 13
Waste and Vfastewater Analyses 13
Tissue Analyses 13
Sample Preparation 13
Priority Pollutant and Pesticide Analyses ... 13
Metabolite Analysis 14
Analysis for Toxic Substances in the Cytosol 15
8. Biological Analyses 18
Fouling 18
Mortality 18
Incremental Growth 18
Condition Factor 18
Gonadal Index 19
Histopathological Effects 19
Scope for Growth 19
Oxygen:Nitrogen Ratio 25
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9. Quality Assurance 26
10. Data Analysis, Interpretation, and Reporting 27
Chemical Data 27
Water and Wastewater Quality 27
Priority Pollutants in Tissues 27
Metabolites in Tissues 28
Distribution of Metals in the Cytosol 28
Biological Data 28
Fouling 28
Mortality 28
Incremental Growth 29
Condition Factor 29
Gonadal Index 29
Histopathologic Effects 30
Scope for Growth 30
Oxygen: Nitrogen Ratio 30
References 31
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FIGURES
Number Page
1. Examples of exposure systems 7
2. Sephadex G-75 elution profile 17
3. Apparatus for measuring clearance rates and assimilation
efficiency 21
4. Exposure chamber for measuring respiration rates 23
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ACKNOWLEDGEMENTS
Sections of this manual were provided by the following contributors:
Scope for Growth - William Nelson, Environmental Research Laboratory, U.S.
Environmental Protection Agency, Narragansett, R.I.; Metallothionein
Analysis - Dr. David A. Brown, Southern California Coastal Water Research
Project, Los Angles, Calif., and Dr. Kenneth D. Jenkins, California State
University, Long Beach, Calif.; Metabolite Analysis - Dr. David A. Brown
and Richard W. Gossett, Southern California Coastal Water Research
Project, Los Angles, Calif.
In addition to the materials provided by the Workgroup members listed in
the Preface, and the contributions listed above, many helpful review
comments were received from the following: Philip A. Crocker, U. S.
Environmental Protection Agency, Dallas, Texas; Joseph Cummins, U. S.
Environmental Protection Agency, Seattle, Washington; Thomas J. Fikslin,
U. S. Environmental Protection Agency, Edison, New Jersey; Delbert B.
Hicks, U. S. Environmental Protection Agency, Athens, Georgia; William H.
Pierce, U. S. Environmental Protection Agency, San Francisco, California;
H. Ronald Preston, U. S. Environmental Protection Agency, Wheeling, West
Virginia, and Dr. Steven Ferraro, U.S. Environmental Protection Agency,
Newport, OR.
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SECTION 1
INTRODUCTION
Section 301(h) of the Clean Water Act of 1977, as amended by the
Municipal Wastewater Treatment Construction Grant Amendments of 1981,
describes provisions under which publically owned treatment works
(POTW's) may apply for variances from secondary treatment requirements
for discharges to marine waters. A modified NPDES permit may be granted
if the applicant can demonstrate that the less-than-secondary discharge
would not impair the integrity of the marine receiving waters and biota.
Following receipt of a modified permit, the POTW is required to maintain
a monitoring program to demonstrate continuing compliance with applicable
water quality standards and 301(h) requirements.
The major goals of 301(h) monitoring are to identify chemicals in the
discharge that should be controlled and to determine whether discharges
cause adverse biological effects in the receiving v/ater. Monitoring for
priority pollutants discharged by POTWs is an important facet of the
program. Marine organisms do not bioaccumulate all chemicals equally.
Some chemicals may be in low concentration or even below detection limits
in v/astewater, yet accumulate to high and/or toxic levels in marine
organisms. Conversely, some materials in high concentration in the
effluent may not be bioconcentrated. Accordingly, a field monitoring
system is needed that identifies chemicals of biological significance.
Many approaches have been used in monitoring for adverse biological
effects in receiving waters, including studies of natural planktonic and
benthic communities. However, changes in the structure of natural
communities are not as sensitive to pollution as changes in the health of
individual organisms, which can be adversely affected at low but chronic
levels of exposure to toxic chemicals.
Extensive use of filter-feeding bivalve molluscs during the past decade
to determine the distribution and persistence of toxic substances in
marine waters and to detect and measure adverse effects of pollutants on
aquatic life has resulted in the development of methodology which is
applicable to the 301(h) program (Bayne et al., 1978, 1981; Davies and
Pirie, 1980; Goldberg, 1975; Goldberg et al., 1978; Phelps and Galloway,
1980; Phelps et al., 1981; Phillips, 1976, 1977a, 1977b; Stephenson, et
al., 1979, 1980, 1981; Widdows et al., 1981). In recent years, attention
has focused on only a few species, principally in the genus Mytil us.
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This manual describes recommended methods for the exposure of molluscs to
discharges from POTW's to determine the bioaccumulation of toxic
substances, to detect acutely toxic conditions in the plume, and to
measure the degree of stress (sublethal toxicity) to which the test
organisms nay have been subjected. In this protocol, caged molluscs are
collected at a non-polluted site, exposed for one month at a minimum of
two stations—in the zone of initial dilution and at a reference station
—and analyzed for toxic substances in tissues and for sublethal
biological effects.
Caution must be exercised in the use of caged molluscs in biononitoring
programs, such as the 301(h) program. Some organic contaminants which
have a low octanolrwater partition coefficient are not bioconcentrated by
organisms (Gossett et al., 1982). Organisms in the plume may accumulate
organic contaminants reflective of historical rather than current
discharges (Young et al., 1976). A significant portion of contaminants
(e.g. 39 percent for copper, Phillips et al., 1980) found in molluscs may
be associated with sediments in the gut that may not be absorbed.
Therefore, measurements of metals in undepurated organisms may not give a
true measure of actual bioaccumulation of contaminants. Also, often in
these organisms there are large (greater than 3-fold) variations in
concentrations of organic contaminants related to both the stage in the
reproductive cycle, which varies seasonally, and the amount of upwelling
of contaminants from sediments (Brown et al., 1982d), which might make it
difficult to see differences between stations. Seasonal changes also
occur in histology (Reynolds et al., 1980), and in the rates of
metabolism and detoxification of contaminants (Brown et al., 1982d).
These factors must be taken into account when designing the field studies
and interpreting the data.
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SECTION 2
EXPOSURE SITE SELECTION
A minimum of two exposure sites must be used: (1) one in the plume,
within the zone of initial dilution (ZID), and (2) a reference site,
"upstream" from the zone of initial dilution and outside of the area
affected by the discharge.
The plume exposure apparatus should be placed as close as possible to the
outfall diffuser (i.e. at the center of the ZID). Additional exposure
sites may be necessary or desirable to define contaminant gradients in
the vicinity of the outfall, and in the case of receiving waters that are
already stressed, to determine the contribution of other pollutant
sources to bioaccumulation levels.
The control site must have hydrographic and water quality characteristics
similar to those at the outfall. The test organisms are sensitive to
salinity, and the use of more than one control site (i.e. such as
upstream and downstream) may be required in estuarine environments where
salinity gradients are present.
Exposure locations to be avoided include shipping lanes and dredging
sites. Swift currents may preclude the use of some stations, but
exposure gear has been successfully maintained in currents as high as 5
knots.
Station positions may be established by use of surface buoys, visual
sighting (shore transects), use of fathometers to fix depth, acoustic
transducers (pingers), Loran C navigation aid, satellite navigation aids,
and portable navigation aids (e.g. Motorola MinirangerR).
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SECTION 3
EXPOSURE DEPTH
Test organisms are exposed at a minimum of one depth at each station. At
the discharge, exposure cages are placed in the ZID, but at least one
meter above the bottom to avoid the overriding influence of toxic
substances released during the exposure period by the sediments, which
might have been deposited by historical pollution not representative of
the current discharge. At the control station, cages are placed at the
same depth(s) as are used at the discharge. Under some circumstances, it
may be desirable or necessary to expose the organisms at additional
depths to determine concentration gradients or to detect the release of
toxic substances from the sediment. In cases where the plume depth is
expected to vary during the exposure period, it may also be appropriate
to use multiple exposure depths to ensure plume exposures.
To provide meaningful information, it is necessary for exposures to be
conducted at a depth vfhich will ensure maximum potential plume contact.
It is the dischargers responsibility to demonstrate that exposures were
actually conducted in the effluent plume. The spatial distribution of
the plume may be determined by field water quality measurements (e.g.
NH3, turbidity), remote sensing (e.g. acoustic backscatter: Proni et
al.., 1976; Proni and Hansen, 1982), or by mathematical models (See Tetra
Tech, Inc., 1982a and 1982b for examples and application). If models are
used, site-specific v/ater density data (i.e. temperature and salinity)
for the exposure period should be used as input.
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SECTION 4
EXPOSURE SYSTEMS
Gear that has been used successfully in the past and is recommended in
the 301(h) program is described below. For additional information see
Stephenson et al. (1979) and Phelps and Galloway (1980).
Because of the potential for loss of exposure gear due to natural events
(e.g., storms, ice flows) or vandalism, it is recommended that two arrays
be placed at each exposure site. The overall cost increase associated
with an additional array is considerably less than that required for
repeating the entire exposure if the test organisms cannot be recovered.
ANCHORS
Anchors that have been successfully used by various programs include:
(1)
(2)
(3)
(4)
Train wheels (with axles removed), 340 kg.
Degreased automobile engine blocks (use a commercial degreasing
firm). Two blocks are used on each line, chained together.
Cast concrete blocks, 25 - 160 kg.
Fence anchors, auger type, 1 - 2 m length (for use in soft
bottoms).
LINE
Sixteen millimeter (5/8 in) polypropylene line or 8mm (5/16 in)
polypropylene encased steel cable (Rolyan PernaflexR) is reconmended
for surface buoys. Smaller line (6mm; 1/4 in) may be used for subsurface
buoys. The line should be kept bagged and off the deck of the surface
vessel to prevent contamination.
BUOYS
Surface buoys are used primarily as station marker buoys, whereas
subsurface buoys are used to support the mussel cages and/or bags to
reduce losses due to ship damage and vandalism. Surface buoys placed in
navigable waters must be Coast Guard approved (e.g. RolyanR 1352). The
use of spar buoys is recommended.
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Submerged buoys, such as a 30 cm diameter inflatable, phosphorescent
orange, plastic float, plus a 20 cm diameter non-collapsable float, can
be used to support the mollusc cages.
EXPOSURE CAGES/BAGS
Enclosures recommended for use with molluscs include polypropylene or
nylon test tube baskets and bait bags.
(1) Test tube baskets - use non-contaminating material, such as
polypropylene.
(2) Bags -
(a) Nylon mesh bags - 8 cm x 1 m (3 X 36 in) nylon bait bags,
12 mm (1/2 in) mesh, 20 kg test*.
(b) Polypropylene mesh bags - {Vexar*), 15 cm X 225 cm,
12 mm (1/2 in) mesh.
GEAR CONFIGURATION
Examples of gear configuration are shov/n in Fig. 1. A commonly used
configuration is where a USCG approved special purpose buoy (similar to
Rolyan 1352R) is attached by 8 mm (5/16 in) polypropylene encased steel
cable (Rolyan PermaflexR) to a 150 kg concrete anchor. Nylon lines
(6 mm) are run about 6 m to satellite moorings of 25-50 kg each to which
6 mm polypropylene line is attached with 20 cm diameter hard plastic
floats used to suspend mussel baskets 1 meter above the surface of the
sediment. A float placed about 6 m up the nooring cable prevents
entanglement with the subsurface floats, and baskets can be hung on the
cable itself for profile work. Bags containing mussels can also be hung
from a pipe framework as shown in Fig. le.
*Nylon Net Company, P.O. Box 592, Memphis, TN 38101
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. »ES« IACS COITAIIIIK
XUSSELS
SURFACE IUOT
SUtSUIFACE IUOT
THEIHOCLliE
FLASTIC FLMT -
fOLTPROMLEIIE LIKE •
MUSSEL lASttTS
MCHOI 1~—-"j ~
SATELLITE HOOKING
TO SATELLITE «00«1«6
^-SURFACE HARKER RU07
ANCHOHIKfi POOS
Fig. 1. Examples of exposure systems (not to scale): (a)from Stephensen
et al., 1980; (b) From Young et al., 1976; (c) provided by D.
Phelps, USEPA, Narragansett, RI; (d) from Phelps and Galloway,
1980; (e) provided by T. Fikslin, USEPA, Edison, NJ.
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SECTION 5
TEST ORGANISMS
RECOMMENDED SPECIES:
Mussels: Mytilus call form'anus - West coast
Mytil us edulis- West coast, East coast, and Gulf of
Mexico
Oysters: Crassostrea virginica - East coast, and Gulf of Mexico
"gigas - West coast
SIZE:
Mussels: 5-7 cm
Oysters: 7-10 cm
SOURCE AND CONDITION
Test organisms should be collected from an area that is relatively free
of contaminants. If no previous data are available on the level of
contaminants in tissues, and the physiological condition of the organisms
in the proposed collection area(s), representative samples should be
collected and tested. The proposed source of organisms selected for use
in the 301 (h) program should be reviewed by the permitting authority
before organisms are collected for transplanting to the exposure sites.
It should be noted that a state permit may be required for collecting
test organisms.
In the source area, test organisms should be collected from approximately
the same depth, preferrably below mean-tidal level. Test organisms can
be collected by dredge or removed from rocky substrates with stainless
steel pry bars. Organisms should not be collected from steel or man-made
wooden structures. Collectors should wear clean polyethylene gloves at
all times. Care should be taken to avoid contamination of the organisms
during collection and transport. Organisms should be of approximately
the same size, to minimize the natural variation in chemical and
biological parameters.
A random subsample of 20-25 organisms should be removed from the
collection to determine mean length and weight, the condition factor, the
incidence of parasites and disease, the stage in the gametogenic cycle,
and body burden of toxic substances.
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Upon collection, the organisms should be triple bagged in 4-nil cleaned
polyethylene bags and placed in ice chests. The polyethylene bags and
ice chests should be cleaned with detergent (Micro^) and triple rinsed
with distilled water prior to use.
The molluscs should be transplanted to the exposure sites within 48 hours
after collection.
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SECTION 6
SAMPLE EXPOSURE AND RETRIEVAL
EXPOSURE PROCEDURES
At the exposure site, the test organisms are placed In cages or mesh
bags. The bags are constricted every 6-8 Inches with nylon cable ties to
ensure uniform exposure of the organisms to the surrounding water. Four
bags/cages, each containing 25 individuals (total of 100 individuals),
are exposed at each depth (Fig. 1).
The test organisms should be protected from surface contamination by
enclosing them in cleaned 4-mil polyethylene bags until they are hauled
overboard and lowered below the surface. The protective bag is then
removed underwater.
EXPOSURE PERIOD
Organisms are exposed for one month. The choice of dates during which
exposure should take place may vary with location. If pronounced
seasonal changes occur, more than one exposure period is recommended. If
only one exposure period is used, it should be the period of maximum
exposure, i.e., when the sexual organs are v/ell developed, the water
temperature is such that the animals are metabolically active, and there
is the least dilution of the discharge.
The period of maximum stratification and least dilution usually occurs in
late summer. However, contaminant concentrations in tissues during this
period may be the lowest of any time during the year because of
spawning. For this reason, it may also be advisable to expose organisms
during the winter. Exposures during periods of rapidly changing density
gradients should be avoided because of uncertainties in maintaining plume
exposures at a given depth.
Exposure periods greater than one month may be necessary at discharges
where certain toxic substances with relatively slow uptake rates (e.g. Hg
and Ag) are of concern.
SAMPLE RETRIEVAL
The potential for successful retrieval of exposure arrays is enhanced by
the use of electronic navigation aids during deployment and retrieval.
The use of such aids is especially important if subsurface buoys are
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used. It is also recommended that an acoustic transducer (pinger) be
attached to each array to aid in location during retrieval. Transducers
are inexpensive (less than $200 each) and operate for 6 months. The use
of acoustic releases is not generally recommended because they are
expensive. Hov/ever, they may be the best alternative for some
applications.
For arrays with subsurface buoys, retrieval can be accomplished by a
combination of electronic positioning and acoustic location, followed by
diver retrieval of the exposure apparatus. The practical limit for diver
retrieval is about 36 m. In situations where the subsurface buoy must he
placed be!ov/ diving depth, or in situations where diver retrieval is not
feasible for other reasons, the exposure array may be retrieved by
snagging a bottom line attached to the anchor (Fig. 1), or use of an
acoustical release device.
FIELD OBSERVATIONS
a. Fouli ng
If fouling is severe, the flow of water to the noil uses may have been
sufficiently reduced to interfere with feeding. The degree of fouling is
observed and reported as the estimated percentage of mesh openings
occluded by fouling organisms (Stephenson et al., 1980).
b. Mortality
Conditions in the ZID may be acutely toxic. Therefore, the percentage of
test organisms surviving to the end of the exposure period should be
determined for each exposure site/depth.
SAMPLE PRESERVATION AND TRANSPORT
Contamination from substances in the surface film can be avoided by
placing the molluscs in polyetheylene bags before surfacing. When
retrieved, the organims may be held briefly in cleaned ice chests until
further processing. Excess v/ater should be drained from the organisms on
ship or after removing to shore.
a. Metal Analyses
Samples collected for trace metal analysis are placed in cleaned
ZiplocR bags, immediately frozen on dry ice and transported to the
laboratory in the frozen state. In the laboratory, samples are stored at
-20C until analyzed.
b. Priority Organic Pollutants and Metabolite Analyses
Samples collected for organic analysis are double-wrapped in precleaned,
hexane-rinsed aluminum foil. The aluminum v/rapped samples are then
placed in ZiplocR polyethylene bags, immediately frozen on
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dry ice and transported to the laboratory in the frozen state. In the
laboratory the samples are maintained at - 20°C until analyzed.
c. Cytosol Analysis
Samples collected for cytosol analysis are placed in cleaned ZiplocR
bags, and immediately frozen on dry ice. They are stored at -80°C upon
return to the laboratory (experiments have shown that metal!othionein is
stable at this temperature, but not at -20°C; Oshi da, 1982).
d. Biological Analyses
Samples collected for biological analyses returned to the laboratory in
cleaned polyethylene ice chests.
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SECTION 7
CHEMICAL ANALYSES
WATER AND WASTEWATER ANALYSES
Methods for v/ater and wastewater analyses are described in USEPA, 1979,
1982.
TISSUE ANALYSES
Whole organism (soft part) composite samples are used for analyses of
toxic substances. Three replicate composite samples (15-20 organisms per
sample) should be analyzed from each exposure site/depth. Tissue samples
are analyzed for (1) the full list of 129 priority pollutants and six
pesticides, (2) for metabolites of toxic organic substances, and (3) for
the distribution of toxic metals and organics in the cytosol
(netallothionein/enzyme/glutathione pool). A subset of the priority
pollutants and pesticides may be analyzed if it can be demonstrated that
only those substances occur in the effluent. Data are reported in ug/g
or ng/g dry weight, with a wet weight conversion factor.
a. Sample Preparation
Immediately prior to analysis, frozen mussels are removed from the bags,
one at a time, scrubbed in deionized water to remove debris (use
polyethylene gloves), and thawed in polyethylene, borosilicate glass, or
stainless steel trays. The adductor muscle is severed with a clean,
stainless steel scalpel, the gonad is excised, and the remainer of the
soft parts are placed in a preweighed acid-cleaned containers. The
quantity of tissue required for analysis is approximately as follows: (1)
Hg - 1 gram; (2) remainder of metals - 5 grams; (3) organic priority
pollutants - 50 grams; (4) metabolites - 5 grams.
b. Priority Pollutant and Pesticide Analyses
Methods for the tissue analysis for priority pollutants and pesticides
are described in USEPA, 1981. The percent lipid (USFDA, 1970) also
should be determined for each sample because it is may help explain the
variability in the concentration of organics. The moisture content of an
aliquot of tissue (dry weight conversion factor) is determined by drying
at 103C for 12 hr (Stephenson et al., 1980).
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c. Metabolite Analysis
Recent studies indicate that metabolites represent the major fora of
xenobiotic orgam'cs in marine organisms (Brown et al., 1982b,c,d). In
addition, it appears that chronic effects of organic compounds are caused
by their metabolic products, while acute effects, which would occur under
only the most extreme circumstances, are caused by parent organic
compounds (Young et al., 1979; McKinney, 1981). However, most studies on
the presence of organic contaminants in the environment do not report
levels of metabolites. These omissions may occur because most
metabolites cannot be extracted by normal procedures since they are bound
to proteins, DNA, glutathione, glucuronic acid, and other substances in
organisms (Reid and Krishna, 1973; Roubal et al., 1977; Varanasi and
Gmur, 1980; Miller and Miller, 1982). Therefore, to determine their
levels, they must first be released from substances to which they are
bound by a heat-catalyzed base hydrolysis (Miller and Miller, 1966;
Miller 1970; Gingell and Wall cave, 1974; Gold et al., 1981; Brown et al.,
1982b). Results obtained by Brown et al. (1982b), indicate the recovery
of metabolites from tissues nay be increased by one to two orders of
magnitude when this procedure is used. Since metabolites appear to be
the predominant form of xenobiotic orgam'cs in organisms, usually
representing over 90% of the total of parent compounds and their
metabolites, it is important that these analyses be included in programs
designed to measure the bioaccumulation of organic compounds. In fact,
it may be that those compounds which are rapidly metabolized after
biological uptake may not be detected by normal procedures.
The methods for extraction of metabolites are similar to EPA standard
procedures (Federal Register, 1979; USEPA, 1981), but with the addition
of a step in which the extract is heated to 90°C for 30 minutes after
extraction of the base/neutral extractable fraction and before extraction
of the acid extractable fraction. The procedure is as follows:
(1) Homogenize 5 g (wet weight) of tissue in 20 ml of deionized (DI)
water in a blender. Rinse the blade twice with DI water.
(2) Dissolve 1.2 g NaOH in the sample homogenate.
(3) Extract the homogenate three times with 50 ml of hexane/
acetone (1:1, V:V). Centrifuge if necessary to obtain complete
separation of the layers.
(4) Take the hexane (top layer) as the base/neutral extractable
fraction and analyze for parent organic compounds.
(5) Heat the remaining aqueous phase to 90C for 30 min to hydrolyze
possible conjugates (Gingall and Wallcave, 1974; Gold et al., 1981).
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(6) Allov; the solution to cool, adjust to pH 1 with 6N HC1, and extract
three times with 50 ml of methylene chloride with centrifugation if
necessary.
(7) Take the methylene chloride (bottom layer) to dry ness with a
roto-vaporizer.
(8) Add 10 nl of nethylating agent (5 mg 3-methyl-l-p-tholyl-triazene/l
ml diethyl ether) to the dried sample.
(9) Blow-dry the sample under a stream of nitrogen.
(10) Redissolve the sample in methanol.
(11) Analyze the final methylated extract for the presence of
metabolites using GC/EC, GC/FID or GC/MS (Brown et al., 1982b).
The distribution of metabolites between a site of detoxification, the
glutathione-containing (GSH) pool, and sites of toxic action, including
the metallothionein-containing (MT) pool and the enzyne-containing (ENZ)
pool, can be determined by analyzing the composited cytosolic pools using
the above method, starting at (2) above (Brov/n et al., 1982b).
Both metals and organics share a common site of toxic action, the ENZ
pool, while organic metabolites also appear to act adversly on the MT
pool, reducing metal-binding and detoxification by this pool (Brov/n et
al., 1982b; Jenkins et al., 1982b). When all three cytosolic pools are
analyzed for both metals and organic metabolites, it is possible to
determine which specific contaminants are present at sites of toxic
action and therefore responsible for direct toxic effects. When this
procedure is used in combination with general stress indices, such as
scope for growth, it is possible to ascertain both the sum total of
direct toxic effects and indirect effects related to the metabolic cost
of detoxification.
d. Analysis for Toxic Substances in the Cytosol
The following simple procedures are used to determine the partitioning of
trace metals between a site of detoxification, the metal!othionein-
containing (MT) pool and a site of toxic action, the enzyme-containing
(ENZ) pool (Brown et al. 1982a).
(1) Tissues are thawed and individuals (v/hen practical) or composites
of 15-25 organisms are suspended in three volumes of chilled buffer
(0.05 M Tris-HCl, pH 7.4).
(2) Suspensions are homogenized with an antoxidant (2-mercaptoethanol)
for 15 sec at high speed in a Sorval Omnimix honogenizer at 4°C.
The homogenate is centrifuged for 10 min at lO.OOOxg in a
refrigerated centrifuge, and the resulting supernatant is
recentrifuged for 60 min at 100,000 x g. The final supernatants
15
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(cytosols) from each sanple are combined and rehonogenized for 5
sec to ensure homogeneity. At this point, cytosols can be stored
at -80°C until further processing.
(3) Frozen cytosols are thawed, vortexed and 7 ml applied to a 1.6 x 70
cm column packed with Sephadex G-75 gel. The sample is eluted with
0.05 Tris-HCl (pH 8.2) at a flow rate of 28 mL hr'1, and 3 ml
fractions are collected for metal analysis (Jenkins et al.,
1982c). A standard solution of proteins of known molecular
weights, such as albumin, should be used to characterize the
Sephadex column.
(4) Fractions are analyzed for metals using flame atomic absorption
spectrophotometry when possible (e.g. usually Zn and Cu), or by
graphite furnace atomic absorption spectrophotometry when
necessitated by low metal levels (e.g. usually Cd and Ag).
The first peak to elute, as located by the metal profiles, is the high
molecular weight enzyme-containing (ENZ) pool; the second peak is the
medium molecular weight netallothionein-containing (MT) pool; and the
third peak is the low molecular weight glutathione-containing (GSH) pool
(Fig. 2). To save time for metal analyses, the location of these pools
can be determined by doing a Zn profile, and then combining fractions
constituting each of these pools for the remainder of the metal analysis.
A more rapid procedure has been developed, utilizing HPLC. Whereas each
Sephadex 6-75 column run takes about 8 hours, HPLC runs take only 40
minutes. In the HPLC procedure, 0.1-0.5 mL samples are injected on a
Toya Soda TSK SW 3000 column (5 mm x 600 run) and eluted at one mL/min
with 0.2 M Tris HC1 (pH 7.4). One mL fractions are collected and
analyzed for metals as described above (Jenkins et al., 1982b).
16
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Zn 1.0
0>
V
oT
E
O.08
Cu 0.04
Cd o.oi -
FRACTION #
POOL
0.02 r-
S 10 IS 20 25 30
ENZ MT GSH
Fig. 2. Typical Sephadex G-75 elution profile for a control niissel
(Mytilus caliform'anus) liver shov/ing the concentrations of Zn, Cu and Cd
in individual fractions constituting each of the ENZ: high molecular
weight enzyme-containing pool which contains Zn and Cu as essential
components of netalloenzynes, but is a site of toxic action for excesses
of metals; MT: medium molecular weight metallothionein-containing pool
which serves a storage/detoxification function for essential (e.g. Zn and
Cu) and non-essential (e.g. Cd) metals; and GSH: low molecular weight
glutathione-containing pool which serves as a site of detoxification for
organic metabolites (from Brown et al., 1982d).
17
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SECTION 8
BIOLOGICAL ANALYSES
The recommended biological observations and analyses, arranged in
approximate order of complexity and level of effort required, are as
fol1ows:
Fouli ng
Mortality
Incremental Growth
Condition Factor
Gonadal Index
Histopathological Effects
Scope for Growth
Oxygen:Nitrogen Ratio
FOULING
Fouling is determined in the field at the time of sample retrieval. The
degree of fouling is reported as the estimated percentage of mesh
openings occluded by fouling organisms (Stephenson et al., 1980).
MORTALITY
The percentage of test organisms surviving to the end of the exposure
period is reported for each exposure site/depth.
INCREMENTAL GROWTH
The mean length of the shells (to the nearest 0.1 mm) is determined
(Riisgard and Poulsen, 1981) before and after exposure at the reference
site and in the plume to determine the change in length of the shells
during the exposure period.
CONDITION FACTOR
The condition factor is the v/et weight of the soft body expressed as a
percent of the total organism weight (Bayne and Thompson, 1970; Boalch et
al., 1981).
18
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GONADAL INDEX
The gonads are removed and weighed, and the (gonad weight)/(soft body
weight) ratio is calculated (Ouellette, 1978; Giese and Pearse, 1974).
I-n M. edulis, the gonad develops within the mantle so that physical
separation of the two tissues is difficult. Therefore, the entire
gonadal/mantle complex is taken as gonadal tissue (Lobel and Wright,
1982).
HISTOPATHOLOGICAL EFFECTS
Histopathological analyses will provide useful information regarding the
condition of the organisms and the site of toxic action, which could not
be determined by other means. Methods for tissue preparation are found in
Yevich and Barszcz (1981)
SCOPE FOR GROWTH
"Scope for Growth" (SFG) is a measure of the net amount of energy
available to an organism for growth and reproduction, and is determined
by subtracting the energy used for basic physiological processes from the
food energy assimilated. This index has been found to be statistically
correlated with the concentration of toxic substances in tissues and is
considered a sensitive method to detect sublethal, adverse biological
effects (Bayne et al., 1981; Phelps et al., 1981; Widdows et al., 1981;
Martin et al., 1982a).
The size of the test organisms and the stage in the gametogenic cycle are
important sources of natural variation in SFG measurements. At least 10
organisms of similar size should be used to minimize the experimental
error.
SFG is determined by measuring four parameters - clearance (food uptake)
rate, food energy absorption efficiency, respiration rate and ammonia
excretion. The values for these parameters are converted into energy
units (Joules), and substituted into the following equation:
SFG = (C x A) - (R + E)
where: SFG = Net energy available for growth and reproduction
C = Food energy consumed
A = Food energy absorption efficiency (%)
R = Energy lost through respiration
E = Energy lost through excretion
The methods used for measuring these parameters for Mytil us edulis are
described below. For the sake of consistency, physiological measurements
are completed in the following sequence for each group of organisms
tested:
19
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Day 1: AM - Collection
PM - Clearance rate
Day 2: AM - Absorption efficiency
- Respiration rate
PM - Ammonia excretion rate
a. Clearance Rate
Clearance rate is the volume of v/ater cleared of particles (diameter
greater than Sum) per unit tine. Seawater, filtered to 1 urn, flows to a
mixing chamber where algae (such as Tetralsemis suecica, Martin et al.t
1982a) is added continuously to obtain a concentration of approximately
8-15x1O3 cells nL-1. This water then flows at a rate of 50-75 ml
min'T through separate supply lines to ten 1-liter chambers, each
containing one mussel (Fig. 3) . Each chamber is gently aerated to
ensure uniform mixing and prevent settling of the algae. The algal
uptake measurements are not initiated until after the animals have opened
their values and ventilated for approximately 60 minutes. The inflowing
and outflowing algal concentrations are determined at one hour intervals
for three hours. To complete the algal counts in the timely manner, the
use of an electronic counting instrument, such as a Coulter Counter
(Model TA11, 100 urn aperture tube, using channels 4-13), is recommended.
The clearance rate (CR) is calculated for each mussel at each hourly
interval using the following equation:
f - r
-1 1 9
CR (L h ') = ] r * X F
L2
where: C] and £-2 - incoming and outflowing algal concentrations,
respectively.
F = flow rate through each chamber in Lh~l.
The mean of the three hourly rates is then calculated and used as the
representative clearance rate for each mussel.
b. Food Energy Absorption Efficiency
Mussels are maintained in clearance chambers overnight to allow for a
sufficient amount of feces to be deposited by each individual. The
following morning a sample of feces is collected from the bottom of the
chamber with a pipette, deposited on a washed, ashed and weighed glass
fiber filter using a Mi Hi pore system, and viashed with 2.4% ammonium
formate to remove any salts from the fecal pellets and the filter. The
filter is dried overnight at 70°C and weighed to provide the dry weight
of the material. The filter is then ashed in a muffle furnace at 500°C
for at least four hours and weighed to provide an ash weight. The food
material is also collected on a washed, ashed and weighed glass filter,
20
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B
Fig. 3. Apparatus for measuring clearance rates and assimilation
efficiency. A. Clearance rate apparatus. B. Enlarged side view of
feeding chamber: (a) inflov/ing algae; (b) inflowing filtered seawater;
(c) mixing chamber; (d) running water bath; (e) feeding chamber; (f)
algae; (g) air; (h) overflow; (i) mussel; (j) pedestal; (k) stirring bar;
(1) magnetic stirrer (from Martin et al., 1982a).
21
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and treated in the same manner as the feces. The absorption efficiency
(AE) is calculated as follows (Conover, 1966):
AE = : ~: ___ X TOO
ML (1-EHF) * IUU
where: F = ratio, (ash-free dry wgt)/(dry wgt), for food.
E = ratio, (ash-free dry wgt)/(dry wgt), for feces.
c. Respiration Rate
Respiration rates are measured by isolating individual mussels in closed
respirometer vessels equipped with a dissolved oxygen electrode (Fig. 4).
After it is placed in the chamber, the organism is allowed to acclimate
for a period of approximately 30 minutes, during which time seawater
containing algae is pumped through the apparatus as described above.
This ensures the measurement of routine (feeding) metabolism. At the end
of the acclimation period, the chamber valves are closed, isolating the
mussel. The decline in P02 is recorded using a Radiometer blood gas
analyzer strip chart recorder. Water in the container is stirred
continuously to give an accurate reading at the oxygen probe. The oxygen
tension should not be allowed to decline below about 115 mm Hg, at which
concentration mussels become oxygen conformers.
The respiration rate (RR), expressed in ml 02 per animal per hour, is
determined as follows (use period of steady decline only):
Convert nm Hg into nL D£ L~l :
RR - (T - T ) X W (1) - VA (D x DOS
KK UQ I.|J A t A UUi
Where: DO TQ = initial dissolved oxygen concentration, ml L~l
DO T-] = final dissolved oxygen concentration, ml L'1
RR = respiration rate in nL Og animal "^hour~l
VV = volume of respiration vessel, mi's
VA = volume of animal, mi's
DOS = Saturation value for dissolved oxygen at the
specified temperature and salinity, obtained from
a table.
22
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Fig. 4. Exposure chamber for measuring respiration rates: (a) inflow of
water; (b) outflow of water; (c) 62 probe; (d) v/ater bath; (e) experi-
mental chamber; (f) mussel; (g) pedestal; (h) stirring bar; (i) magnetic
stirrer; (j) radiometer (from Martin et al., 1982a).
23
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d. Ammonia (Nitrogen) Excretion Rate
Mussels are placed in individual beakers containing 300 ml of 0.45um
filtered seav/ater. The beakers are placed in a water bath at ambient
temperature, and the mussels are left undisturbed for a period of three
hours. One control beaker, lacking a mussel, is used to determine
background ammonia levels.
At the end of the three-hour period, the seawater in each beaker is
gently mixed, sampled and analyzed in duplicate for ammonia, using the
salicylate-hypochlorite method of Bower and Holm-Hanson (1980).
The procedure is as follows:
(1) A 50 nL syringe is rinsed by filling it with water from an exposure
beaker, and discarding the contents.
(2) The syringe is filled with a second 50 mL sample from the beaker, and
a filter holder containing a glass fiber filter is attached to the
end of the syringe
(3) A 10 ml sanple is flushed through the filter and discarded.
(4) The remaining 40 ml are filtered into a pre-washed, HCl-rinsed,
plastic container, covered and frozen at -20°C until the sample is
processed. It is more convenient to delay the NH4 analyses until a
large number of samples are available.
The NH4-N excretion rates (NER) are calculated as follows:
NER (ug NH4-N h"1) = (TC - CC)(VSW) X 14 X h"1
Where: 1 uM NH4 - N = 14 ug M
TC = Concentration of NH4 (uM/L) in test
beakers containing mussels
CC = Concentration of NH4 (uM/L) in control
beaker
VSW = Volume (L) of seav/ater in exposure vessels
If 300 ml are used in each beaker, and the exposure period is three
hours, (TC - CC) x 1.4 = ug NH4-N h'l
e. Algal Cultures
Algae used in SFG experiments are from unialgal cultures (e.g. Tetralsemis
suecica, Martin et al., 1982a) maintained in a healthy, log-growth phase
by adding small pure cultures (400 ml) and commercially available
nutrients to five gallon carboys of autoclaved seawater (collected on
incoming tide). The energy content of the algae is determined using the
wet oxidation method of Maciolek (1962).
24
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f. Calculation of Scope for Growth Index (SFG)
Scope for growth is calculated as follows:
P = (C x A) - (R + E),
where: P = Energy (Joules h~l ) available for growth and reproduction
C = Food energy (Joules h~^) consumed
A = Food energy absorption efficiency (%)
R = Energy (Joules h~l ) lost through respiration
E = Energy (Joules H~l ) lost through excretion
(1) C = Joules of food energy consumed per hour
Where: (a) Clearance rate (In'1) x food concentration
(cells L'1) = total No. cells consumed per hour.
(No. cells h'Tj/tNo. cells ng'1 ash-free wgt) =
mg algae h'^
mg algae h~l x Joules mg~l algae = Joules h~l
The energy content (Joules ng~M of the algae is
determined according to the method of Maciolek, 1962.
(2) A = Food energy absorption efficiency (%)
(3) R = ml 02 consumed h'1 x 20.08 Joules per ml 02
(4) E = mg NH4-N h'1 x 24.81 Joules per mg NH4-N
Additional information required to complete the calculations includes
determining mussel volume and dry weight, to standardize calculations for a
1 gram animal. Calculation of standardized rates is only recommended when
using animals that vary widely in length. If the lengths of the animals
fall within a narrow range, weight-specific rates should be calculated for
clearance, respiration, and ammonia excretion before calculating the SFG.
OXYGEN: NITROGEN (0:N) RATIO
The 0:N ratio, which is the ratio of oxygen consumed to nitrogen excreted,
is another useful physiological index of stress. This value can be
calculated from the above data as follows (Bayne, 1975; VMddows, 1978b):
a. Convert ml Oh"1 to mg Oh"1 by multiplying by 1.428.
b. o:N
mg 09h-1 mg NH.-N h"1
25
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SECTION 9
QUALITY ASSURANCE
A quality assurance plan must be prepared as an integral part of the
301(h) monitoring plan. Factors in the field that will affect the
quality and utility of the data include the condition, uniformity in
size, and stage in the gametogenic cycle of the organisms, the care taken
in avoiding contamination and injury of the organisms during collection
and transport of the test organisms, the attention given to the depth and
positioning of the exposure gear, and the v/ater quality conditions, such
as salinity, at the exposure sites.
Laboratory quality assurance practices include the regular calibration of
instrumentation, the use of duplicate analyses (i.e. every tenth
analysis) and reference materials, and participation in interlaboratory
studies such as round robins and performance evaluations. Detailed
laboratory quality assurance guidelines are described in USEPA (1979,
1982). Reference materials for water, v/astewater and tissue analyses are
available fron the Quality Assurance Branch, Environmental Monitoring and
Support Laboratory, U.S. Environmental Protection Agency, Cincinnati,
Ohio.
26
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SECTION 10
DATA ANALYSIS, INTERPRETATION AND REPORTING
The overall objective of the caged mollusc biononitoring progran is to
provide the permittee and the regulatory agency with data useful in
identifying discharges that cause adverse effects on marine life or
otherwise pose a threat to the marine environment. Differences observed
in the chemical and biological parameters at reference and plume stations
must be analyzed for statistical significance to account for natural
variations in chemical and biological data, which are often large. The
use of composite samples is generally not recommended because it obscures
the variation in the individual organims, and prevents an adequate
determination of the precision of the analyses. However, compositing is
sometimes necessary to obtain sufficient material for analysis, or to
reduce an otherwise overwhelming analytical burden. The selection of
test organisms of similar size and stage of gametogenesis will tend to
reduce the variation in the biological data (Bayne et al., 1981), and
enable the investigators to detect smaller differences in population
responses between stations than otherwise possible.
Upon the completion of the statistical analyses (t-test, ANOVA, etc.),
parameters which fail to show a significant difference between the
reference and exposed stations are reported, together with an appropriate
discusssion if the results were unexpected or otherwise unusual.
Parameters which are significantly different are further evaluated to
determine the magnitude of the difference, whether any FDA, EPA, or state
criteria for standards have been exceeded, and what reduction in the
concentrations must be achieved to reach acceptable biological conditions.
CHEMICAL DATA
a. Water and Wastewater Quality
Water quality data should be reported to document conditions at the
reference and plume stations, and should be evaluated in terms of the
environmental requirements of the test organisms and confounding effects,
if any, on the interpretation of the biological data.
b. Priority Pollutants in Tissues
Data on priority pollutants in tissues of organisms exposed at the
reference and plume stations should be compared and evaluated in terms of
27
-------
differences in biological responses at the stations. FDA action levels
should also be considered if test species are being harvested for human
consumption from the polluted zone.
c. Metabolites in Tissues
As mentioned above, it appears that chronic effects of organic compounds
are caused by their metabolic products, while acute effects, which would
occur under only the most extreme circumstances, are caused by parent
organic compounds (Young et al., 1979). Body burdens of metabolites
should be checked against data on toxicity of metabolities and parent
compounds.
d. Distribution of Metals in the Cytosol
The metal levels in each pool of cellular proteins are added and
expressed as an amount of metal per unit weight of tissue. The metal
concentration in the MT pool can be compared to the loading capacity of
this pool as determined by laboratory exposures. In this v/ay, the degree
of utilization (saturation) of the detoxification capacity of the
organism can be determined (Brov/n et al., 1982a). Using this
information, predictions can be made as to how much additional metal
could be loaded into the biota before spillover of trace metals from the
MT pool to the ENZ pool would occur, with resultant toxic effects. These
toxic effects occur because excesses of essential metals or non-essential
metals in the ENZ pool result in disruption of normal enzyme function.
It should be noted that a certain anount of Cu and Zn will always occur
on the ENZ pool because these metals are essential components of
metal loenzymes (Brov/n and Chatel, 1978; Jenkins et al., 1982c). However
essential metals present in excess of that required in the metalloenzymes
must be partitioned onto MT or they will have a toxic effect. Further
discussions of the analytical methods and significance of the
metallothionein data can be found in the following references: Brown et
al., 1982c; Jenkins et al., 1982b,c,d,e; Kohler and Riisgard, 1982;
Noel-Lambot et al., 1980; Piscator, 1964; Shiakh and Lucis, 1971; Simkiss
and Taylor, 1981; Simkiss et al., 1982; Squibb et al., 1974; and
Viarengo et al., 1980, 1981;
BIOLOGICAL DATA
a. Fouling
If severe fouling is observed at the end of the exposure period, the flow
of water to the molluscs may have been sufficiently reduced to interfere
with feeding. The degree of fouling, therefore, should be taken into
consideration in evaluating data on growth and condition.
b. Mortality
The percentage of test organisms surviving to the end of the exposure
period is reported for each exposure site/depth. Conditions in the ZID
28
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may be acutely toxic. If less than 90 percent of the organisns exposed
at the reference site survive, the test would be considered invalid and
should be repeated. If survival at the reference site exceeds 90
percent, but survival at the plume site(s) is less than 90 percent, acute
toxicity nay be present. Under these circumstances, the bi©accumulation
data would be invalid. To obtain adequate bioaccumulation results, it
v/ould be necessary to repeat the test using additional exposure sites.
c. Incremental Growth
This index is simply the increase in the mean shell length during the
exposure period. Shell growth is dependent upon water temperature,
available food, and other environmental factors, in addition to the
presence of pollutants. Under normal conditions, a growth of several mm
v/ould be expected in 30 days. Riisgard and Poulson (1981), starting with
organisms 2.26 mm in length, reported a increase in length of as much as
6.6 mm in tl. edulis in 18 days. Additional observations on the growth of
M. eduli s were reported by Kautsky (1981). If the mean increase in shell
length of organisms exposed in the plume is significantly (Po.05) less
than at the reference station(s), the likelihood of adverse environmental
conditions is indicated.
d. Condition Factor
The condition factor is the wet weight of the soft body expressed as a
percent of the total organism weight (Bayne and Thompson, 1970). Boalch
et al., (1981) observed a mean condition factor of 6% in a composite of
20 n. edulis with a mean length of four cm. In their study, the condition
factor was significantly correlated (Po.05) with the logio of the
metal concentration for all metals except copper. They observed that the
use of a composite sample reduced the variation from three orders of
magnitude, for individual organisms, to approximately 50%. Stephenson et
al.. (1980), using a slightly different form of the condition factor,
(soft body weight)X(length), observed that mussels with the highest
condition factor were collected away from heavily industrialized areas.
A statistically significant (P().05) decline in the condition factor
ratio during the exposure period, or a significantly lower CF at the
plume site compared to the reference site(s), v/ould indicate the
likelihood of adverse environmental conditions at the exposure site.
e. Gonadal Index
The gonadal index varies with the stage in the gametogenic cycle of the
organism. Within the gametogenic cycle, the proliferation of gonadal
tissue and maturation of the gametes will be affected by the
physiological condition of the organism, which in turn will be determined
by the availability of food and other environmental conditions, including
the concentration of toxic substances. Stephenson et al. (1980) reported
gonadal indices ranging from 0.21 to 0.39. The low indices observed at
some stations in their study v/ere assumed to be related to high metal
concentrations in tissues.
29
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f. Histopathological Effects
Histopathological analyses provide information on the general condition
of the organisms and the site of toxic action which could not be
determined by other means (Barry and Yevich, 1975; Lov/e and Moore, 1978;
Lowe et a!., 1981; Mix and Schafer, 1979; Mix et al., 1977; Mix et al.,
1979a; Mix et al., 1979b; Reynolds et al., 1980: Thompson et al., 1978;
Yevich and Barszcz, 1977). This information includes:
Identity of Specific tissues affected by the toxic substances (site
of toxic action)
Whether the effects are reversible or irrersible
The sex, stage in the reproductive cycle, and condition of the gonads
Whether the poor condition of organisms that appear to be stressed
was caused by parasites, pollutants, or nutrition
g. Scope for Growth
"Scope for Growth" (SFG), which is a measure of the net amount of energy
available to an organism for growth and reproduction, has been found to
be inversely related to the concentration of toxic substances in mollusc
tissues and is considered a sensitive method to detect sublethal, adverse
biological effects of toxic substances on molluscs (Phelps et al., 1981;
Widdows et al., 1981; Martin et al., 1982a). A decline in SFG results in
less rapid growth and a reduction in fecundity (Bayne et al., 1975, 1978;
Bayne and Widdows, 1978; Bayne and Worrell, 1980). Gillfillan et al.
(1976) observed an inverse correlation between SFG and the concentration
of aromatic hydrocarbons in tissues, and a similar relationship was
reported between SFG and the concentration of the water-accommodated
fraction of North Sea crude oil by Widdows et al. (1982), and between SFG
and tissue burdens of metals and organics by Phelps and Galloway (1980).
An excellent discussion of factors related to the collection and analysis
of SFG data can be found in Bayne et al. (1981).
g. Oxygen;Nitrogen Ratio
The OxygenrNitrogen (0:N) ratio provides information on the relative
utilization of protein in energy metabolism compared to other carbon
sources. A high rate of protein utilization, compared to carbohydrates
and lipids, results in a low 0:N ratios, which are generally indicative
of a stressed condition (Widdows, 1978). According to Bayne (1973a,b),
low 0:N ratios (i.e. 20 or less) may result from low food concentrations
(starvation), whereas at high food concentrations (1.5 mg/L or greater),
0:N ratios will fall in the range of 40 to 50 in the absence of other
adverse environmental conditions. They indicated that food levels
available during most of the year support 0:N ratios in the range of
25-30. Widdows et al. (1981), however, stated that 0:N ratios of less
than 30 were indicative of organisms that were very stressed. They
observed 0:N ratios of 50-75 in organisms that were well nourished and
living under generally favorable environmental conditions. Although some
variability in the 0:N data is indicated, they may be useful in detecting
stress.
30
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by temperature and nutritive stress. J. Mar. Biol. Assoc. U.K.
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Bayne, B.L. 1973b. Aspects of the metabolism of Mytil us edulis during
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Bayne, B.L. 1975. Aspects of physiological condition of Mytil us
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ed., Aberdeen Univ. Press, p. 213-238.
Bayne, B.L. 1978. The potential of bivalve molluscs for monitoring the
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Bayne, B.L., P.A. Gabbott, and J. Widdov/s. 1975. Some effects of
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Bayne, B.L., and R.J. Thompson. 1970. Some physiological consequences
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Bayne, B.L., and J. Widdov/s. 1978. The physiological ecology of two
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Bayne, B.L., and C.M. Worrall. 1980. Growth and production of mussels
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Brov/n, D.A., J.F. Alfafara, S.M. Bay, G.P. Hershelman, K.D. Jenkins,
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