EPA-600/9-78-010
MARCH 1978 y
PROODUKS
FOR TH£
DKPOSN.
PERMIT
PROGRhM
ENVIRONMENTAL RESEARCH LABORATORY
OFFICE OF RESEARCH AND DEVELOPMENT
U.S. ENVIRONMENTAL PROTECTION AGENCY
GULF BREEZE, FLORIDA 32561
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BIOASSAY PROCEDURES FOR THE OCEAN DISPOSAL
PERMIT PROGRAM
by
Environmental Protection Agency
Ocean Disposal Bioassay
Working Group
U.S. ENVIRONMENTAL PROTECTION AGENCY
ENVIRONMENTAL RESEARCH LABORATORY
GULF BREEZE, FLORIDA 32561
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DISCLAIMER
This report has been reviewed by the Environmental Research
Laboratory, Gulf Breeze, U. S. Environmental Protection Agency, and
approved for publication. Mention of trade names or commercial pro-
ducts does not constitute endorsement or recommendation for use.
ii
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FOREWORD
The Marine Protection, Research, and Sanctuaries Act of 1972, as
amended, (MPRSA) requires that applications for permits for ocean dumping
be evaluated on the basis of their ecological impact on the marine environ-
ment, as well as on other considerations included in the MPRSA. The
International Convention on the Prevention of Marine Pollution from the
Dumping of Wastes and Other Matter, (Convention) for which the MPRSA is the
enabling domestic legislation, requires a similar evaluation and also pro-
hibits the disposal of mercury and its compounds, cadmium and its compounds,
organohalogens, and oils and greases as other than trace contaminants.
The revised EPA ocean dumping regulations and criteria (40 CFR, 220-228),
published January 11, 1977, establish bioassays as the key procedures to be
used to assess the probable ecological impact of particular wastes, and also
establish criteria by which bioassay results are to be used to determine
whether or not a waste is environmentally acceptable for ocean dumping.
Bioassay procedures described in this manual were developed for use by
EPA personnel in carrying out the ocean dumping permit program under the
MPRSA and pursuant to the revised EPA ocean dumping regulations and criteria.
This manual is a revision of EPA-600/9-76-010 published May, 1976, and in-
cludes improvements in bioassay procedures which represent recent advances in
the state-of-the-art in marine bioassay techniques. As such, they should be
considered recommended interim procedures and not as definitive standard
methods.
The procedures presented here cover a wide variety of techniques and
organisms. Selection of appropriate procedures should be made by the
permitting authority on a case-by-case basis, depending on the type and
amount of waste, location of dump site, proposed methods of disposal, and
other appropriate considerations.
Thomas W. Duke
Director
Environmental Research Laboratory
Gulf Breeze, Florida 32561
iii
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ABSTRACT
The bioassay procedures given in this manual were developed to provide
tests tor conducting toxicity evaluations of waste materials considered for
ocean disposal under EPA's Ocean Disposal Permit Program.
i-™ S6 Proc^dures specify the use of various organisms representing several
trophic levels. Both flow-through and static tests are included. Methods
given vary in their utility and complexity of performance. The procedures
in*™ot intended to be considered "standard methods," but, depending on the
th^!
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CONTENTS
Foreword
Abstract ........................... iv
Figures ........................... v
Tables ............................ viii
Contributors .................... ..... ix
I. Introduction .................... 1
II. Bioassay Procedures ................ 2
A. Background Information for the Performance
of Phytoplankton Marine Bioassays ....... 2
B. Static Method for Acute Toxicity Tests
with Phytoplankton ............... 19
C-. Flowing Sea Water Toxicity Test Using
Oysters (Crassostrea virginica) ........ 25
D. Methods for the Culture and Short-Term Bio-
assay of the Calanoid Copepod (Acartia tonsa) . . 28
E. Culturing the Mysid (Mysidopsis bahia) in
Flowing Sea Water or a Static System ...... 59
F. Acute Static Toxicity Tests with Mysid
Shrimp f Mysidopsis bahia) ........... 61
G Entire Life-Cycle Toxicity Test Using Mysids
rMvaidopsis bahia) in Flowing Water ...... 64
H. Culture of the Grass Shrimp (Palaemonetes pugio)
in the Laboratory ............... 69
I. Static Bioassay Procedures Using Grass Shrimp
f Palaemonetes sp.) Larvae ........... 73
j. Entire Life-Cycle Toxicity Test Using Grass
Shrimp f Palaemonetes pugio Holthuis) ...... 83
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K. Static Method for Acute Toxicity Tests
Using Fish and Macroinvertebrates 89
L. Flow-through Methods for Acute Toxicity
Tests Using Fish and Macroinvertebrates 97
M. Laboratory Culture of Sheepshead Minnows
(Cyprinodon variegatus) 107
N. Life-Cycle Toxicity Test Using Sheepshead
Minnows (Cyprinodon variegatus) 109
0. Fish Brain Acetylcholinesterase Inhibition
Assay 118
Appendices 43
«
^ 1-D Synthetic Sea Water Formulation 43
2-D Sea Water and Sterility Enrichment 44
3-D Algal Culture 45
4-D Descriptive Characteristics for Selected
Neritic Copepods 46
vi
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FIGURES
Number Page
1-A Hypothetical relationship between algal
growth and toxicant concentration 10
2-A Relationship between percentage of control
growth rate (0-48 hrs) and copper 13
1-D Mass Copepod Culture Systems (Static) ... 32
2-D Generation Cage 33
3-D Mass Copepod Culture (Flowing) 35
4-D Bioassay protocol 38
1-H A flow-through hatching apparatus for grass
shrimp larvae production 70
1-1 Example of a range-finding bioassay .... 74
2-1 Example of a definitive bioassay 75
vii
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TABLES
Number Page
1-A Sea water and sterility enrichments 4
2-A Synthetic sea water formulation for
algal assays 5
3-A Nutrient enrichments for algal bioassay
medium 7
1-B Composition of mixes to be added to
artificial sea water 21
1-D Composition of algal diet and recommended
concentration for feeding, egg laying, and
naupliar feeding 29
2-D Protocol for mass copepod culture 34
1-K Standard salt water 90
2-K Suggested sea water test temperatures for
vertebrates and invertebrates 93
1-L Maximum sea water test temperatures for
vertebrates and invertebrates 102
viii
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CONTRIBUTORS
Bioassay procedures published in this manual resulted from deliberations
of the Ocean Dumping Bioassay Committee and represent methodology developed
at EPA laboratories. Contributing laboratories and investigators follow:
Static Method for the Performance of_ Phytoplankton
John H. Gentile and Mimi Johnson, Environmental Research Laboratory,
Narragansett, Rhode Island
Flowing Sea Water Toxicity Test Using Oysters (Crassostrea virginica)
Phillip A. Butler, Office of Pesticide Programs at Environmental Research
Laboratory, Gulf Breeze, Florida
Jack I. Lowe, Environmental Research Laboratory, Gulf Breeze, Florida
Methods for the Culture and Short-Term Bioassay of_ the Calanoid Copepod
(Acartia tonsa)
John H. Gentile and Suzanne Sosnowski, Environmental Research Laboratory,
Narragansett, Rhode Island
Culturing the Mysid (Mysidopsis bahia) in_ Flowing Sea Water or_ a_ Static
System
D. R. Nimmo, T. L. Hamaker, and C. A. Sommers, Environmental Research
Laboratory, Gulf Breeze, Florida
Acute Static Toxicity Tests with Mysid Shrimp (Mysidopsis bahia)
Patrick W. Borthwick, Environmental Research Laboratory, Gulf Breeze,
Florida
Entire Life-Cycle Toxicity Test Using Mysids (Mysidopsis bahia) ±n_ Flowing
Water
D. R. Nimmo, T. L. Hamaker, and C. A.Sommers, Environmental Research
Laboratory, Gulf Breeze, Florida
Culture of the Grass Shrimp (Palaemonetes pugio) in the Laboratory
Dana Beth Tyler-Schroeder, Environmental Research Laboratory,
Gulf Breeze, Florida
Static Bioassay Procedures Using Grass Shrimp (Palaemonetes sp.) Larvae
Dana Beth Tyler-Schroeder, Environmental Research Laboratory,
Gulf Breeze, Florida
ix
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Entire Life-Cycle Toxicity Test Using Grass Shrimp (Palaemonetes pugio
Holthuis)
Dana Beth Tyler-Schroeder, Environmental Research Laboratory,
Gulf Breeze, Florida
Static Method for Acute Toxicity Tests Using Fish and Macroinvertebrates
David J. Hansen, Steven C. Schimmel, Del Wayne Nimmo, and Jack I. Lowe,
Environmental Research Laboratory, Gulf Breeze, Florida
Patrick R. Parrish, (formerly Environmental Research Laboratory,
Gulf Breeze; now EG&G, Marine Research Laboratory, Pensacola, Florida)
William H. Peltier, EPA, Region IV, Atlanta, Georgia
Flow-through Methods for Acute Toxicity Tests Using Fish and Macroinverte-
brates
David J. Hansen, Steven C. Schimmel, Del Wayne Nimmo, and Jack I. Lowe,
Environmental Research Laboratory, Gulf Breeze, Florida
Patrick R. Parrish, (formerly Environmental Research Laboratory,
Gulf Breeze; now EG&G, Marine Research Laboratory, Pensacola, Florida)
William H. Peltier, EPA, Region IV, Atlanta, Georgia
Laboratory Culture of_ Sheepshead Minnows (Cyprinodon variegatus)
D. J. Hansen, Environmental Research Laboratory, Gulf Breeze, Florida
Life-Cycle Toxicity Test Using Sheepshead Minnows (Cyprinodon yariegatus)
D. J. Hansen, S. C. Schimmel, and L. R. Goodman, Environmental Research
Laboratory, Gulf Breeze, Florida
Patrick R. Parrish, (formerly Environmental Research Laboratory,
Gulf Breeze; now EG&G, Marine Research Laboratory, Pensacola, Florida)
Fish Brain Acetylcholinesterase Inhibition Assay
David L, Coppage, Environmental Protection Agency, Washington, D. C.
Members of the EPA Ocean Disposal Bioassay Working Group are:
Thomas W. Duke, Chairman, Office of Research and Development (ORD),
Environmental Research Laboratory, Gulf Breeze, Florida
William P. Davis, ORD, Environmental Research Laboratory, Gulf Breeze,
Florida; Bears Bluff Field Station, South Carolina
Jack Gentile, ORD, Environmental Research Laboratory, Narragansett,
Rhode Island
David J. Hansen, and Jack I. Lowe, ORD, Environmental Research Laboratory,
Gulf Breeze, Florida
William E. Miller, ORD, Environmental Research Laboratory, Corvallis,
Oregon
Royal J. Nadeau, Region II, Edison, New Jersey
Carolyn K. Offutt, Office of Water and Hazardous Material (OWHM),
Washington, D. C.
Richard D. Spear, Region II, Edison, New Jersey
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SECTION 1
INTRODUCTION
The bioassays procedures given in this manual were established to
provide procedures for conducting biological evaluation of waste materials
to be disposed of in the ocean. Tests conducted according to these bioassay
procedures will provide information on the toxicity of various materials to
be disposed. However, these bioassay procedures, like all laboratory bio-
assay methods, are attempts at simulation of actual conditions and therefore
suffer all the inaccuracies inherent to simulation systems.
Although these bioassay procedures are not "standard" EPA methods, they
are intended as guides for those involved in evaluating ocean dumping
permits. Accordingly, each method differs in detail and style and does not
conform to a standard format. Permit applicants are expected to modify
bioassay procedures according to both the nature of the waste material and
the type of procedure involved.
The Ocean Dumping Bioassay Committee requires that a minimum of three
species be used in an evaluation of a permit. These species should be
selected from the different taxonomic groups listed in the section on the
flow-through method for acute toxicity tests using fish and macroin-
vertebrates (see page 97)f. We recommend that indigenous organisms be
used whenever possible in addition to those organisms recommended in this
manual.
The EPA bioassay working group intends to revise these bioassay pro-
cedures periodically as new information becomes available. We are coordi-
nating our efforts with the EPA/Corps of Engineers Technical Committee on
Criteria for Dredge and Fill Committee.
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SECTION II
BIOASSAY PROCEDURES FOR ROUTINE APPLICATION
A. BACKGROUND INFORMATION FOR THE PERFORMANCE OF PHYTOPLANKTON MARINE
BIOASSAYS
The primary producer populations of estuaries consist principally of
microscopic phytoplankton. In their role of storing potential energy, via
photosynthesis, these organisms represent the primary energy input into
aquatic ecosystems (Joint Industry/Government Task Force, 1969). Thus, it
is imperative that water quality conditions be favorable to their growth and
reproduction if serious alterations in other components of marine communities
are to be avoided.
Under natural conditions, both the qualitative and quantitative aspects
of phytoplankton population dynamics display a high degree of seasonability,
characterized by well-defined succession patterns. It is essential that not
only the productivity of various systems be maintained, but also the relative
abundance of species according to normal seasonal compositions because
primary herbivore populations exhibit selectivity in their grazing patterns.
Consequently, while a pollutant may seem to have no apparent adverse effect
on the total primary production, it may have drastically altered community
structure and composition. Such alterations often occur when sensitive
indigenous species are replaced by species less desirable ecologically, but
equally active photosynthetically. If the more resistant species is incom-
patible with the feeding and/or nutritional requirements of primary herbivore
populations, then energy transfer to high trophic levels will be affected
and contribute ultimately to significant effects on naturally occurring
desirable populations. Data for the phytoplankton are a necessity to ade-
quately describe and predict the potential effects of a toxicant upon an
estuarine ecosystem response.
1. Species Selection
In the design of a bioassay program, the selection of test species is
pivotal to the acquisition of realistic and meaningful information. Algal
culture techniques historically have focused upon developing suitable culture
media to sustain complete life cycles. Nutritional levels and medium compo-
sition often bore little resemblance to the actual environmental conditions
the organism encountered. Furthermore, research was often limited to a few
species that were readily maintained in the laboratory.
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Within the last decade, culture techniques have greatly broadened the scope
of species available for investigation.
In choosing species for bioassays, the following criteria are useful
guides:
a. Whenever possible, indigenous species representing a diversity of
phylogenetic types from the major seasonal successions should be studied.
b. Since sensitivities vary among species, when possible, more sensi-
tive species should be selected for bioassay.
c. From seasonal and laboratory studies, conditions of greatest vulner-
abilities should also be identified for the species selected.
d. Since a bioassay basically measures the response of an organism to
the product of toxicant concentration and exposure time, the rate of response
of the test species must be considered. Both test species and culture
conditions should permit growth rates of 0.5-1.0 doublings per day under
non-stress conditions.
The above criteria offer maximum flexibility for the experienced re-
searcher. For workers with limited funds and expertise, two species are
recommended if indigenous forms are unavailable: Skeletonema costatum
(species of choice) is an ecologically important phytoplankter that is
common to a wide geographic range of neritic waters. Thalassiosira pseu-
donana, while of lesser ecological significance, is sensitive to heavy
metals and has an 8-hour generation time which offers greater practical
value in the establishment of toxicological responses. It is also recom-
mended that these species be used in conjunction with others to serve as
controls on the systems being tested.
2. Culture Conditions
The culture conditions for all test species generally should reflect
their natural conditions. In order to develop some semblance of uniformity,
two basic regimes are recommended. For temperate species, a temperature of
20° ± 2°C, 2500-5000 lux on a 14-hour light and 10-hour dark cycle (14:10
cycle) is desirable. For cold water forms, a temperature of 8° ± 2°C, 2500-
5000 lux on a 10:14 cycle is recommended. Stock cultures of the test species
are to be maintained in enriched natural (Table 1-A), or synthetic sea
waters (Table 2-A).
The stock cultures should be transferred to the nutritionally dilute
culture medium and allowed to complete two growth cycles prior to use in a
bioassay. This is necessary since nutritional history can have marked
effects upon responses. We have found up to five-fold differences in
responses of bioassay organisms maintained under high and natural nutrient
levels (Gentile et al., 1973). Stock cultures should be maintained with
sterile transfer techniques.
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TABLE 1-A. SEA WATER AND STERILITY ENRICHMENTS
Sea water enrichments for stock algal culture maintenance (After Guillard
and Ryther, 1962):
NaNO,
Vitamins:
Thiamine HCl
Biotin
B12
Trace metals:
ZnS045H20
MnCl2.4H20
Fe-sequestrine
75 mg/liter
5
10 mg/«-
0.10 mg/£
0.50 yg/i
0.50 yg/ft
0.002 mg/fc
0.004 mg/«.
0.002 mg/Jl
0.036 mg/H
0.001 mg/H
1.0 mg (0.13 mg Fe)/£
Buffer:
TRIS-500 mg/£ @ pH 7.8-8.2
Before autoclaving, the following sterility enrichments should be added to
the enriched sea water medium above:
Sodium glutamate
Sodium acetate
Cycline
Nutrient agar
Sucrose
Sodium lactate
L & D alanine
250 mg/£
250 mg/£
250 mg/£
50 mg/£
250 mg/£
250 mg/£
250 mg/£
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TABLE 2-A. SYNTHETIC SEA WATER FORMULATION FOR ALGAL ASSAYS*
Compound Concentration/liter
Nad 24.00 g
Na2SO, 4.00 g
H3B03 0.03 g
CaCl2 . 2H20 1.47 g
MgCl2 . 6H20 10.78 g
Na2Si:03.9H20 t 30.00 mg
KC1 700.00 mg
NaHCO- 200.00 mg
*Adapted from original table, Kester et al., 1967.
tPrepare stock solution in deionized water and adjust to pH 7.8-8.2
3. Sea Water
The choice of sea water is dictated by availability, quality, and cost.
Natural sea water can often be used for bioassays even though inherent
variables in quality may complicate analysis of results. Clean offshore
water is suitable if proper precautions are observed during collection and
processing. In general, synthetic sea water is preferred for the constancy
of its composition and quality even though trace contaminants must be removed
by additional purification. The cost of the required chemicals and purifi-
cation is usually equivalent to the expense of collecting, transporting, and
processing natural sea water.
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a. Natural Sea Water
Sea water is collected from 3-10 meters below the surface (to avoid
surface contamination) with a non-metallic water sampler, and transported in
autoclavable polyethylene carboys. Glass is also suitable if breakage can
be prevented. Within 24 hours, the water is filtered through acid-washed
membrane filters in a non-metallic filtration system. Filtered sea water is
then stored at 4 C in the dark.
b. Synthetic Sea Water
Commercially available synthetic salt water is also acceptable; when
heavy metal toxicity is being tested, an iron-chelate version should be
employed. It is suggested that these sea water mixtures be aged and aerated
for 24 hours before use.
A modified synthetic sea water formulation (Table 2-A) has been
developed from Kester et al. (1967). This sea water is recommended for
fish, invertebrates, and plankton bioassays. This synthetic sea water has
been endorsed by the Environmental Protection Agency, the 14th Edition of
Standard Methods, and the A.S.T.M. Committee on Bioassays.
c. Salinity
A salinity of 30 /oo is recommended for all bioassays. Salinity
adjustments on natural or synthetic sea waters should be made with glass
distilled or deionized water.
d. Sterilization
Sterilization of stock culture maintenance medium can be satis-
factorily achieved by autoclaving since the pH is stabilized by the presence
of TRIS-buffer. Since bioassay medium cannot be autoclaved, two alternative
methods are recommended: 1) positive pressure filtration and/or 2) pasteuri-
zation (60 ± 2 C for 4 hours). These treatments will not appreciably
alter the physico-chemical properties of the sea water but will provide
effective sterilization.
The medium, however, should be filtered (0.45y) through a previously
acid washed (2 N HC1) filter. Removal of residual acid is accomplished by
rinsing the filter with distilled/deionized water and discarding the first
liter of filtered sea water. Medium should be stored in acid-stripped boro-
silicate glass or linear polyethylene carboys, to which a sterile dispensing
tower can be connected to distribute media.
Sterility checks are made weekly on this test medium by inoculating
2 ml aliquot of sea water into 10 ml of sterile water enriched as in Table
3-A. The tubes are incubated at 20 C in the dark for up to one week. Con-
tamination is indicated by turbidity and opalescence of the medium.
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TABLE 3-A. NUTRIENT ENRICHMENTS FOR ALGAL BIOASSAY MEDIUM
Nutrient Amount
Na NO., 4.42 mg/£ (50 yMN)
K2HPO, 0.87 mg/SL (5 yMP)
Thiamine 100.00 g/£
Biotln 0.50 g/fc
B12 0.50 g/X,
Fe* 25.00 g/8,
Mn 10.00 g/X,
Zn 1.00 g/£
Mo 0.50 g/X,
Co 0.10 g/X,
Cu 0.10 g/X,
*Fe as Cl: Dissolve iron sponge or filings in minimum HC1 with warming and
dilute to volume with deionized water.
4. Glassware
All glassware is high grade borosilicate glass (Pyrex/Kimax). The
bioassays, performed in 125-ml Erlenmyer flasks containing 50 ml of medium,
are sealed with foam plugs. Glassware is dry-heat sterilized (170 C for 2
hours) rather than autoclaved, since the steam often carries metal contami-
nants which can interfere with bioassays involving metal toxicity.
Rigorous cleaning is necessary for all glassware to insure against
contamination. Glassware is soaked in detergent, hand or mechanically
brushed, rinsed in deionized water, totally immersed in 10% HNO~ for 2-6
hours, thoroughly rinsed in double glass distilled or deionized water,
and air or oven dried.
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For work involving the toxicity of metals, the glassware should receive
the following post-wash treatment: To eliminate the problems of either
positive or negative contamination, a monolayer of silico-polymer is applied
to all surfaces contacting the sea water. Commercially available SC-87*
prepared as 5% solution in cyclohexane, is poured into the glassware and
drained, leaving a film on the surface. The glassware is then air-dried and
oven-cured at 150-175 C for 4 hours. The result is a completely non-wettable
surface which, after a double glass distilled water rinse, is ready to use.
One coating often lasts two or three assays before recoating is necessary.
Recoating can be done over the old coating or a strong alkali (2N NAOH + 10%
ETOH) can completely strip the old coating prior to recoating. In most
instances, alcoholic-alkali stripping can be avoided by using hot detergent
each time prior to recoating.
5. Bioassay Protocol
The bioassay design consists of three major integrated components:
preparation of log-phase inoculum, nutrient enriched bioassay medium, and
toxicant solutions.
a. Inoculum
Inoculum for the bioassay is prepared by inoculating 0.5 ml (0.1-1.0
ml) of stock culture into triplicate 125-ml flasks containing nutrient
enriched sea water at bioassay level (Table 3-A). At the point of inflect-
ion of the growth curve, inoculate three new flasks from this series and
follow the second growth curve. Cells from this second or later transfers
are suitable for use in the bioassay. These cells now have adapted to the
more natural nutrient levels, and their response will more closely reflect
that expected from a natural population of the test species.
b. Bioassay Medium
Filter sterilized and/or pasteurized, enriched sea water is dispensed
into a presterilized 1-2 liter flask that is compatible with a 50-ml Ace-
dispenser (Cat. no. 8004, Ace Glass Co., Vineland, N.J.). Nutrients (Table
3-A) are aseptically added and inoculum (as described above) is added to
give an initial cell density of 2,500 cells/ml to 10,000 cells/ml. Inocula-
tion of the total medium volume permits the dispensing of a uniform cell
population in all flasks. Initial cell density or biomass is measured.
Fifty milliliters of enriched inoculated medium are dispensed into 125-ml
flasks, using a 50 ml Ace-dispenser in a sterile hood.
*SC-87. Product of General Electric: distributed by Pierce Chemical Co.,
Rockford, Illinois.
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The selection of an initial cell density will be dependent upon the
sensitivity of the biomass parameter measuring system. For example, in ~
clean systems using particle counters, initial cell densities of 2.5 x 10
microscopic counts are employed; initial cell densities of 1 x 1C)4 cells/ml
may be appropriate. For extractive (ATP, Chi "a") or isotope techniques,
the initial cell density can be kept low since the aliquot examined can be
adjusted.
c. Toxicant
Toxicant solutions are prepared in distilled water or suitable
solvent for hydrophobic compounds. Stock solutions or dilutions of a waste
should be prepared to ensure that the same volume is added at all test
levels. This addition should not exceed one milliliter/50 ml of test medium.
When working with waste effluents, a maximum of 5 ml addition is allowed
since this will constitute a 10% maximum alteration in salinity. Toxicant
additions are made to the flasks containing inoculated enriched sea water
and placed in an incubator.
d. Design
The bioassay design is in part determined by the type of toxicant
tested. A general format will include a screening of a broad range of
concentrations from which levels are selected for a definitive evaluation.
Generally, preliminary screenings should cover concentrations at four
orders of magnitude with duplicate cultures at each level. The definitive
assay should include one concentration above and two below the calculated
50% inhibition level, using logarithmic bisection of intervals. Triplicate
cultures should be used for the definitive bioassay.
Parameter measurement should be evaluated at least once every 24
hours for the duration of the experiment. This permits calculation of rates
of response which are important in interpreting the behavior of the toxicant.
The duration of the experiment should be adequate for the control population
to complete its logarithmic growth phase and reach a stationary growth rate.
It is also desirable to determine for the inhibited cultures: the duration
of the lag-phase, maximum rate of growth, and maximum yield (Figure 1-A).
However, not all this information may be readily available from a single
assay and all concentrations.
e. Modifications
The assay system described above uses small volumes (50 ml/125).
This is not meant to frustrate the expansion of assay volumes. The systems
can be easily scaled up to the following dimensions of 125/250; 250/500;
500/1,000. With larger volume systems media, dispensing can be made directly
into the sterile flasks. Nutrients and test species can also be added to
each flask. This increases the potential for variability and contamination
but, with experience, difficulties can be minimized. The larger systems
require more assay medium and space. However, greater volumes will permit
more frequent analysis of a greater number of parameters. This
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10
6 _
10
5 -
CO
z
UJ
o
10'
o
0
HOURS
CONTROL
lOugs Cu/l
- A
2Ougs Cu/l
D - D
4Ougs Cu/l
8Ougs Cu/l
IBOugs Cu/l
O O
144
168
Figure 1-A. Hypothetical relationship between algal growth and toxicant
concentration.
10
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allows a more precise characterization of the anomalies resulting from
specific pollutant exposures.
6. Parameters
There are a variety of parameters available that characterize the
response of the algal cultures. These parameters are measures or indices of
biomass at the time of sampling, which, when plotted against time, produce
a growth response curve. This curve can be used to determine log-phase,
rate of log-growth, and a maximum population density for control and exposed
cultures.
a. Population Density
Microscopic measurements of cell density can be made using a haemo-
cytometer, Palmer-Maloney Chamber, or inverted microscope with settling
chambers. Details of these counting methods are available in the literature
(Schwoerbel, 1970; American Public Health Association, 1975).
The microscopic methods present two problems: they are time-
consuming when done properly and their statistical significance decrease
significantly at cell densities below 1 x 10^. Consequently, when large
numbers of assays and replicates are required, it becomes impractical to
count each assay microspically.
An electronic particle counter offers the most rapid, practical, and
statistically accurate measurement of population density. The initial cost,
while high, is offset by the increased work volume, accuracy, and saving of
time.
b. Population Biomass
Biomass values can be calculated from population density data by
using cell dimensions and assuming the cell is a particular geometrical
shape (i.e., sphere, cylinder, etc.). This method, which depends on cell
counts, is subject to the same limitations mentioned above.
Electronic particle counters can also give volumetric measurements,
but usually such capabilities are obtained at additional cost. It is worth
the expense if large numbers of assays are anticipated.
c. Chlorophyl
Chlorophyl "a" is often used as a measure of algal biomass. Both
spectrophotometric absorbance and fluorescence (in vivo and in vitro)
techniques are available (Strickland and Parsons, 1968). The spectro-
photometric technique lacks the sensitivity particularly at low cell den-
sities. The fluorescent systems, however,.are more sensitive and can be
used at cell densities of less than 1 x 10 cells/ml. The in vivo fluor-
escent technique is particularly useful because it does not require extrac-
tion and is very sensitive.
11
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A potential limitation of this measurement is the general variabil-
ity of cellular chlorophyl "a" as a function of nutrition and environmental
variables (Odum et al., 1959; Yentsch and Ryther, 1957; Yentsch and Menzel,
1963).
d. Carbon-14 Assimilation
Productivity measurements, based upon radioactive carbon assimila-
tion, is a standard technique applicable to both fresh water and marine
algae (Steeman-Neilson, 1952; McAllister, 1961; Jitts, 1963; Jenkins, 1965;
Strickland and Parsons, 1968). This is usually used as a short-term measure
of photosynthetic activity. Culture aliquots may be pulse-labeled for four
hours and C-14 incorporated by cells measured. This relative value may be
used as a biomass index. This latter approach has shown a correlation to
growth rates as measured by changes in cell number or biomass. Transient
changes in C-14 assimilation, not reflecting long-term growth responses,
have also been noted and warrant cautious interpretation of these data.
Adequate C-14 counting procedures may be obtained in Brandsom (1970)
and Chase and Rabinowitz (1967).
e. ATP-Concentration
ATP has been suggested as a sensitive and accurate measure of living
biomass due to a constancy of cellular ATP/carbon ratio (Holm-Hansen and
Booth, 1966; Hamilton and Holm-Hansen, 1967; Holm-Hansen, 1969). Studies
have demonstrated excellent correlation between ATP and direct measures of
biomass (particle counting) and pulse labeling with carbon-14 (Gentile et
al., 1973; Cheer et al., 1974). This technique requires instrumentation
(about $5,000) and costs about $1.00 per analysis. As a measure of living
material, highly contaminated wastes (i.e., sludge) could provide excessive
interference.
The above techniques all offer certain advantages or disadvantages,
depending on the bioassay design, type of effluent tested, facilities, and
personnel.
Automated particle counting, while offering the most rapid, sensitive
and statistically valid method, has limitations. The most restrictive
relates to particle interferences. The test compound or effluent must have
low background in the particle size range of the test species or inevitable
masking and errors will result. This limits the types of effluents to be
evaluated by this technique, unless the particulate fraction can be removed
without jeopardizing the toxic characteristics of the material.
The other methods work well in systems containing particulate
material, but both chlorophyl "a" and carbon-uptake have potentially unde-
sirable response patterns that can make data interpretation difficult.
12
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ATP, on the other hand, appears to be an excellent indicator of living
biomass though it is somewhat expensive to measure routinely and may not
be appropriate for biologically contaminated wastes (i.e., sludge).
All data can be converted to percentage control for any finite
exposure period and the percentage response plotted versus toxicant concen-
tration (Figure 2-A). From this graph, the relationship between toxicant
concentration and degree of inhibition can be determined.
EC-5O=23ugs Cu/l
10
20 40 60 80 100
PERCENT CONTROL RESPONSE
Figure 2-A. Relationship between percentage of control growth rate
(0-48 hrs) and copper.
13
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7. Data Presentation
The design of the bioassay requires a minimum of one observation every
twenty-four hours for the duration of the experiment. Within this schedule,
various options are available to the researcher. The basic data output
represents a growth curve for all concentration examined. This may provide
rate of growth:
k = an —• /AT
o
k: rate of growth
N : population concentration at time zero
N : population concentration at time t
AT: time interval from time zero
and generation time:
AT
k
G: generation time
k: rate of growth
AT: time interval from time zero
and comparisons at maximum population density. Slopes of growth curves
representing the logarithmic growth phase of exposed cultures and population
biomass may be compared with controls by standard statistical analysis.
8. Standard Toxicant
To insure that the technical aspects of the bioassay are properly per-
formed, an internal standard is recommended (LaRoche et al., 1970). We
routinely use sodium dodecyl sulfate (SDS), a surfactant and membrane lytic
agent. This compound produces a very sharp response curve indicating an
almost "total or no" effect at concentrations of 1-2 mg/£. In addition, SDS
is both soluble and stable in aqueous solutions.
While the use of an internal standard can serve as a quality assurance
monitor, it does not, in itself, validate an experiment. There can be
situations where the EC50 concentration for the standard toxicants in two
experiments are essentially identical, but the control growth rates differ
by a factor of two. The deviation of control growth from normal is an
indication of a problem and thus warrants the repetition of the experiment.
In addition, it is recommended that regional offices maintain a Quality
Control Program by requiring contractors to process "blind" reference
samples.
14
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9. Applications
Algal bioassays, with their sensitivity and rapid response, are useful
in many areas of water quality research.
a. The simplest application is for routine screening of potential
toxicants. This represents a well-defined and controlled system where
particle counting is recommended since interferences can usually be mini-
mized. These studies should be designed to produce complete growth curves
with both growth rate and maximum density output.
b. Another application of the algal bioassay is as an evaluation of
water quality. If an impacted area is being investigated, water samples can
be collected along a transect or matrix, depending on hydrographic data. The
water is collected and processed according to techniques described in Sec-
tion 4, and then inoculated with the test species that has been cultured in
enriched water from a control station. Growth rate and population density
can then be compared from station-to-station.
c. The algal assay can also be used to measure the biological impact of
mixed effluents containing suspended solids. In this case, particle count-
ing may not be practicable due to high levels of interference. Consequently,
the growth of the algal culture can be monitored by obtaining daily aliquots
and evaluating the ATP, chlorophyl "a," or measuring C-14 incorporation
after pulse labeling the aliquot (2-4 hours) with NaH1^ C03. The resulting
data, when plotted semi-logarithmically with time, will produce a growth
response curve that may be submitted to the interpretation discussed here.
d. Mention must be made of in situ applications of phytoplankton bio-
assay. Using ATP, C-14 uptake, and chlorophyl "a", both living biomass and
productivity of a water mass may be estimated in situ. These studies can be
made at the site; the samples are preserved and analyzed at a later date.
Such applications, as evaluation of power plant entrainment and point-source
pollution monitoring, commonly use this approach.
10. Remarks
It should be stressed that important advances have been made by the
utilization of phytoplankton bioassays in the establishment of realistic
water quality criteria for marine life.
Fundamental biological anomalies in phytoplankton could impair survival
of high trophic levels and be associated with specific pollutant exposures.
However, it should be noted that problems exist in the application of labor-
atory findings to conditions which may be found in the natural environment.
One scientific discipline greatly neglected in this area is phytoplankton
systematics. As a consequence, in many instances of in situ evaluation of
phytoplankton productivity, identification of species will reveal the
importance of knowing the species present.
15
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REFERENCES
The following literature is recommended to the researcher for detailed
discussions of techniques described in the text. It is not an exhaustive
list but is adequate to acquaint the researcher with the analytical method-
ologies required to perform the assay successfully.
American Public Health Association. 1975. Standard Methods for the
Examination of Water and Wastewater, 14th Ed., 1027 pp.
Bransome, E.D., Jr. CEd.) 1970. The Current Status of Liquid Scintillation
Counting. Grune and Stratton, Inc., New York, 394 pp.
Chase, G.D., and J.L. Rabinowitz. 1967. Principles of Radioisotope
Methodology, 3rd Ed. Burgess Publ. Co., Minneapolis, 633 pp.
Cheer, Sue, J. H. Gentile, and C. S. Hegre. 1974. Improved Methods for ATP
Analysis. Analytical Biochemistry. 60:102-114.
Davey, E.W., J.H. Gentile, S.J. Erickson and P. Betzer. 1970. Removal of
Trace Metals from Marine Culture Medium. Limnol. and Oceanogr. 15:486-
488.
Gentile, J.H., S. Cheer, and P. Rogerson. 1973. The Effects of Heavy Metal
Stress on Various Biological Parameters in Thalassiosira^ pseudonana.
Abstract 34th Annual Meeting, Am. Soc. Limnol. and Oceanogr.
Hamilton, R.D., and 0. Holm-Hansen. 1967. Adenosine Triphosphate Content of
Marine Bacteria. Limnol. Oceanogr. 12:319-324.
Ho-lm-Hansen, 0. and C.R. Booth. 1966. The Measurement of Adenosine
Triphosphate in the Ocean and its Ecological Significance. Limnol.
Oceanogr. 11:510.
Holm-Hansen, 0. 1969. Determination of Microbial Biomass in Ocean
Profiles. Limnol. Oceanogr. 14:740-747.
Holmes, R.W. 1962. The Preparation of Marine Phytoplankton for Microscopic
Examination and Enumeration on Molecular Filters. U.S. Fish Wild.
Serv. Spec. Sci. Rep. No. 433:1-6.
Instruction Manual 760 Luminescence Biometer. 1960. E.I. DuPont De Nemours
and Co., Wilmington, Delaware.
16
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Jackson, H.W. and L.G. Williams. 1962. Calibration and Use of Certain
Plankton Counting Equipment. Trans. Amer. Microscop. Soc. 81:96.
Jenkins, D. 1965. Determination of Primary Productivity of Turbid Waters
With Carbon-14. J. WPCF. 37:1281-1288.
Jitts, H.R. 1963. The Standardization and Comparison of Measurements of
Primary Production by the Carbon-14 Technique. In: Proc. Conf. on
Primary Productivity Measurement, Marine and Fresh Water (M.S. Doty,
ed.) Univ. of Hawaii, Aug.-Sept. 1961. U.S. Atomic Energy Comm. Div.
Tech. Inf. T.I.D. 7633:103-113.
Joint Industry/Government Task Force of Eutrophication. 1969. Provisional
Algal Assay Procedure, pp. 16-29.
Kester, E., I. Dredall, D. Connops, and R. Pytowicz. 1967. Preparation of
Artificial Sea Water. Limnol. & Oceanogr. 12:176-178.
Laroche, G, R. Eisler, and C.M. Tarzwell. 1970. Bioassay Procedures for
Evaluation of Acute Toxicities of Oil and Oil Dispersants to Small
Marine Teleosts and Macroinvertebrates. J. Water Pollut. Control Fed.
42:1982-1989.
Lorenzen, C.J. 1966. A Method for the Continuous Measurement of in vivo
Chlorophyll Concentration. Deep Sea Res. 13:223-227.
Lorenzen, C.J. 1967. Determination of Chlorophyll and Pheopigments:
Spectrophotometric Equations. Limnol. & Oceanogr. 12(2):343-346.
Lund, J.W., C. Kipling, and E.D. Lecren. 1958. The Inverted Microscope
Method of Estimating Algae Numbers and the Statistical Basis of
Estimations by Counting. Hydrobiologia. 11:143-70.
Mackenthun, K.M. 1969. The Practice of Water Pollution Biology. U.S.
Dept. of the Interior, FWPCA. 281 pp.
McAllister, C.D. 1961. Decontamination of Filters in the C-14 Method of
Measuring Marine Photosynthesis. Limnol. & Oceanogr. 6:477-450.
McNabb, C.D. 1960. Enumeration o*f Fresh Water Phytoplankton Concentrated
on the Membrane Filter. Limnol. & Oceanogr. 5:57-61.
Moss, B. 1967. A Spectrophotometric Method for the Estimation of
Percentage Degradation of Chlorophylls to Pheo-pigments in Extracts of
Algae. Limnol. & Oceanogr. 12:355-340.
Mullin, M.M., P.R. Sloan, and R.W. Eppley. 1966. Relationship Between
Carbon Content, Cell Volume, and Area in Phytoplankton. Limnol. &
Oceanogr. 11:307-311.
17
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National Academy of Sciences. - 1969. Recommended Procedures for Measuring
the Productivity of Plankton Standing Stock and Related Oceanographic
Properties. Natl. Acad. Sci., Washington, D.C. 59 pp.
Odum, H.T., W. McConnel, and W. Abbot. 1959. The Chlorophyl "a" of
Communities. Pub. Texas Inst. Mar. Sci. 5:65-95.
Palmer, C.M. and T.E. Maloney. 1954. A New Counting Slide for
Nannoplankton. Am. Soc. Limnol. Oceanogr. Spec. Publ. No. 21, pp. 1-
6.
Schwoerbel, J. 1970. Methods of Hydrobiology (Fresh Water Biology).
Pergamon Press, Hungary, pp. 200.
Steeman-Neilson, E. 1952. The Use of Radioactive Carbon (C-14) for
Measuring Organic Production in the Sea. J. Cons. Cons. Int. Explor.
Mer 18:117-140.
Strehler, B.L. 1968. Bioluminescence Assay: Methods of Biochemical
Analysis. (Glictz, D., Ed.) Interscience, New York. Vol. 14, 99 pp.
Strickland, J.D.H. and T.R. Parsons. 1968. A Practical Handbook of
Sea Water Analysis. J. Fish. Res. Board Can., Bulletin No. 167, 311
pp.
Tailing, J.R. and G.E. Fogg. 1959. Measurements (in situ) on Isolated
Samples on Natural Communities, Possible Limitations and Artificial
Modifications. In: A Manual of Methods for Measuring Primary Pro-
duction in Aquatic Environments, R. A. Vollenweider, ed. IBP Hand-
book, No. 12, F.A. Davis, Philadelphia, pp 73-78.
United Nations Educational, Scientific, and Cultural Organization (UNESCO)
1966. Monographs on Oceanographic Methodology. In: Determination of
Photosynthetic Pigments in Sea Water. UNESCO, Paris. 69 pp.
Utermohl, H. 1958. Zur Vervollkommung der Quantitativen Phytoplankton-
Methodik. Mit. Int. Ver. Theor. Angew. Limnol. 9:1-38.
Weber, C.I. 1968. The Preservation of Phytoplankton Grab Samples. Trans.
Am. Microscop. Soc. 87:70.
Weber, C.I. 1973. Biological Field and Laboratory Methods for Measuring
the Quality of Surface Waters and Effluents. EPA-6704-73-001, U.S.
Environ. Prot. Agency Ecol., Cincinnati, OH.
Yentsch, C.S. and J.H. Ryther. 1957. Short-term Variations in Phyto-
plankton Chlorophyll and Their Significance. Limnol. & Oceanogr.
2:140-142.
Yentsch, C.S. and D.W. Menzel. 1963. A Method for the Determination of
Phytoplankton Chlorophyll and Phaeophytin by Fluorescence. Deep-Sea
Res. 10:221-231.
18
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B. STATIC METHOD FOR ACUTE TOXICITY TESTS WITH PHYTOPLANKTON
J. H. Gentile and Mlmi Johnson
1. Introduction
The method described here is designed for analysis of effects of ocean-
dumped material on growth of marine unicellular algae. It involves addition
of liquid waste or extracts from sludge to algal growth medium, addition of
algae to the medium, and measurement of growth for 96 hours.
Because the capability of calculating EC50 values from bioassay data is
required by law, dilutions of ocean-dumped material are necessary. As it is
impossible to estimate potential algal toxicity or stimulatory action of
each batch of ocean-dumped material, the recommended dilutions may not be
sufficient to yield EC50 values in every case. The logistics of algal
bioassay are complicated and time-consuming. They must be considered care-
fully before requirements are imposed.
2. Maintenance of Test Organisms
The marine unicellular algal species to be used is Skeletonema costatum.
The algae may be obtained from Woods Hole Oceanographic Institution, Woods
Hole, Massachusetts.
The algae are to be maintained in stock culture collections in arti-
ficial sea water medium. The artificial sea water is prepared by dissolving
artificial sea salts (such as Rila Salts, Rila Products, Teaneck, New
Jersey 07666) in glass-distilled water to a salinity of 30 parts per
thousand (30 grams of salt in 1000 ml of artificial sea water). Add 15.0 ml
of metal mix, 1.0 ml of minor salt mix, and 0.5 ml of vitamin mix to each
liter. The compositions of the mixes are given in Table 1-B.
Filter (with suction) the sea water medium through a 0.22y membrane
filter (similar to one manufactured by the Millipore Corporation, Bedford,
Massachusetts 01730, Catalog No. GSWP 047 00). Before filtration, pass 1
liter of 0.1 N HC1 and 5 liters of glass-distilled water through the filter.
Dispense 200 ml of medium into 500-ml Erlenmeyer flasks and use polyurethane
foam plugs to seal the flasks* Autoclave at 121°C and 15 Ib pressure for 15
minutes. The flasks must be washed with detergent, soaked in 10% HC1, and
rinsed 10 times with distilled water.
19
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Equilibrate at room temperature for one day, and check the pH of medium
in a flask especially set up for this purpose as above. The pH should be
between 7.8 and 8.1. If the pH is not within this range, discard all
flasks and make new medium. The pH should fall within this range before a
test is started.
Add 10 ml of stock algal culture to each flask and incubate without
shaking under 45O- to 500-foot candles illumination at 20 ± 2 C with alter-
nating periods of light (16 hours) and darkness (8 hours). Use standard
microbiological techniques for flaming the necks of flasks whenever algae
are transferred.
Stock cultures as described above must be renewed every 10 days. They
need not be shaken during incubation.
3. Preparation of Test Medium
a. Liquid Waste
Liquid waste to be tested must not be modified before use. Liquid
samples taken for analysis, however, must be taken in glass containers with
Teflon-lined lids. The glassware and liners must be washed with detergent,
soaked overnight in 10% HC1, rinsed 10 times with glass-distilled water,
rinsed once with acetone, and again rinsed 10 times with glass-distilled
water.
Prepare dilutions of liquid waste as follows:
(1) Mix 100 ml of liquid waste with 900 ml of artificial sea water
that does not contain trace metal, minor salt, or vitamin mixes. This will
be considered to be undiluted medium.
(2) Add 1 part of (1) to 9 parts of artificial sea water. This is
a 10% solution of undiluted medium.
(3) Add 1 part of (2) to 9 parts of artificial sea water. This is
a 1% solution of undiluted medium.
(4) Add 1 part of (3) to 9 parts of artificial sea water. This is
a 0.1% solution of undiluted medium.
b. Sludge
When sludge is tested, artificial sea water without trace metals,
minor salts, or vitamins are used as extractant. Salinity of the extractant
is 30 parts per thousand and the procedure is:
1. Place a representative portion of the sludge into a 250-ml
capacity graduated cylinder, filling to the 250-ml mark. Let the sludge
settle overnight (approximately 16 hours). Carefully decant and discard the
supernate.
.20
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TABLE 1-B. COMPOSITION OF MIXES TO BE ADDED TO ARTIFICIAL SEA WATER
Mix Amount
Metal mix;
Fe C12 . 6 H2 0* 0.480 g
Mn C12 . 4 H2 0* 0.144 g
Zn S04 . 7 H2 0* 0.045 g
Cu S02 . 5 H2 0* 0.157 nig
Co C12 . 6 H2 0* 0.404 mg
H3B03 0.140 g
Distilled water 1H
Vitamin mix:
Thiamin hydrochloride 50.0 mg
Biotint 0.01 mg
B12t 0.10 mg
Distilled water 100 ml
Minor salt mix;
K3P04 3.0 g
Na N03 50.0 g
Na2 SI03 . 9 H2 0 20.0 g
Distilled water !«,
"Sletal mix should be added after filtration.
*Separate aqueous solutions of these metal salts are maintained at such
concentrations that 1 ml of each is added to 1£ of mix.
tBiotin is maintained as 1 mg/100 ml alcoholic stock solution; B12 in a 10
mg/100 ml aqueous solution.
21
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2. Add 100 ml of the wet settled sludge to a gallon-capacity wide-
mouthed jar and add 900 ml of artificial sea water at room temperature. If
more growth medium will be required, add more settled sludge and artificial
sea water to the jars, but keep the ratio of 100:900 constant. Cap the jars
tightly and shake on an automatic shaker at about 100 excursions per minute
for 30 rain. At the end of the shaking period, remove the jar from the
shaker, stand it in an upright position, and let contents settle for 1 hour.
3. Filter the supernatant fluid through glass wool, a membrane
filter of 5.0y porosity, and then through a membrane filter of 0.22y porosity.
When the filters clog, replace them. The filters must be washed before use
by passing through them one liter of 0.1 N HC1 and 5 liters of glass-dis-
tilled water. All glassware associated with filtration must be washed with
detergent, soaked overnight in 10% HC1, and rinsed with glass-distilled
water before use.
4. The following solutions will be used in the test:
(a) Filtered extract. This will be considered to be undiluted
medium.
(b) Add 1 part of (a) to 9 parts of artificial sea water. This
is a 10% solution of undiluted medium.
(c) Add 1 part of (b) to 9 parts of artificial sea water. This
is a 1% solution of undiluted medium.
(d) Add 1 part of (c) to 9 parts of artificial sea water. This
is a 0.1% solution of undiluted medium.
5. After filtration and dilution of liquid or sludge material, add
30.0 ml of metal mix, 2.0 ml of minor salt mix, and 1.0 ml of vitamin mix to
each liter and record the pH.
6. Add 48.0 ml of each solution to sterile 125-ml volume Erlenmeyer
flasks that were washed with detergent, soaked overnight in 10% HC1, rinsed
10 times with glass-distilled water, rinsed once with acetone, and again
rinsed 10 times with glass-distilled water. Prepare three flasks for each
solution and for each algal species used. Use polyurethane foam plugs to
seal the flasks.
7. Suggested apparatus for extraction, or their equivalent, are:
a. Laboratory shaker, Eberbach 6000 with a 605 Utility Box, or
equivalent, capable of shaking a 1-gallon container at 100 excursions per
minute.
b. Glass jars, wide mouth, 1-gallon capacity with Teflon lined,
screw top lids. Note; If necessary to purchase jars and Teflon sheets
separately, the Teflon lid liners can be prepared by the laboratory personnel.
Jars and lids should be equivalent in quality to those supplied by the
Cincinnati Container Corporation, 2833 Spring Avenue, Cincinnati, Ohio
22
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45225. Jars, Cat. No. 120-400-F-0-0-4 (128 oz.); Lids, Cat. No. 120-400-
White, FTK, PPE. Teflon sheets should be equal in quality to those supplied
by the Cadillac Plastic Co., 3818 Red Bank Road, Cincinnati, Ohio 45227.
4. Bioassay
a. Preparation of algae
Four days before the bioassay test is performed, add 5 ml of algal
stock culture that is at least 5 days old to 45 ml of sterilized artificial
sea water that contains trace metals, minor salts, and vitamins as described
in Section 2, Maintenance of Test Organisms. Place mixture in 125-ml capacity
Erlenmeyer flasks fitted with polyurethane foam plugs. Incubate the new
cultures under 450- to 500-foot candles from cool white fluorescent tubes at
20 ± 2 C. Incubate cultures on rotary shaker platforms (No. G2 shaker
fitted with No. AG2-125 platform from New Brunswick Scientific Co., New
Brunswick, New Jersey 08903, or equivalent) at 140 ± 10 excursions per
minute. The lighting cycle should be 16 hours of light followed by 8 hours
of darkness.
On the first day of testing, add 1.0 ml of algal culture to a
volumetric flask of 25 ml capacity. Bring to approximately half volume with
testing medium, add 2 drops of 10% formalin in growth medium, and bring to
full volume with testing medium. Wait 5 minutes.
Shake each flask to attain a homogeneous suspension of cells. Remove
a sample of the homogeneous suspension quickly with a small pipette and fill
each side of a Spencer Bright-Line haemocytometer. Be sure that the suspen-
sion does not overflow into the troughs of the haemocytometer. At 100X
magnification, count all cells within and impinging upon the 4 1 mm corner
squares and the 1 mm^ central square of each grid. Multiply the count from
the 10 squares by 25,000 to find the number of cells in 1 ml of the original
suspension.
The object of these counts is to determine the dilution required to
attain a final concentration of 100,000 cells per ml in the original cell
culture. For example, if the number of cells in a ml of culture is 200,000,
then the original culture should be diluted 1:1 with test medium to yield
100,000 cells per ml.
b. Growth of algae
Using sterile pipettes, add 2.0 ml of the algal suspension that
contains 100,000 cells per ml to the flasks that were prepared with 48.0 ml
of test medium.
Place the flasks on rotary shaker platforms and set the platform at
140 ± 10 excursions per minute. Illuminate with overhead cool flourescent
lights. Intensity of light should be between 450- and 500-foot candles with
a lighting cycle at 16 hours of light, followed by 8 hours of darkness. The
temperature should be 20 ± 2 C.
23
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Incubate the shaking cultures for 96 hours. At that time, add two
drops of 10% formalin in artificial sea water to each flask, wait five
minutes, swirl the cultures to resuspend the cells to a homogeneous sus-
pension, and count in a haemocytometer as described above.
c. Untreated controls
Control algal cultures must be grown in untreated medium at the time
bioassays on liquid waste or sludge are being done. In this case, untreated
medium, with its full complement of metal, vitamin, and minor salt mixes, is
shaken, filtered, and added to flasks in the same manner as when sludge was
extracted. The cell suspension used to inoculate the untreated growth is
prepared as described above, except untreated growth medium is used for
diluting.
Three flasks are used in growth of controls, and counting is done as
described above.
5. Analysis of Results
Calculate the average values for number of algal cells per mililiter in
control and each dilution of waste-treated flasks.
EC50 value is the dilution at which waste material causes 50% reduction
in growth. In order to estimate this value, inspect the average values to
learn if numbers of algal cells in the waste-treated flasks are (1) less
than half of those in the untreated control flasks, and (2) more than half
of those in the untreated control flasks. To determine an EC50, at least
one point must be greater than and one point less than the EC50. Using a
semilogarithmic coordinate paper, plot the average cell count for a dilution
that yields more than half the average cell count in a dilution that yields
less than half of the average cell count of control flasks. The dilution
should be plotted on the logarithmic axis and the percentage of growth in
relation to the control plotted on the arithmetic axis. Draw a straight
line between the two points. The concentration at which this line crosses
the 50% growth line is the EC50 value.
24
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C. FLOWING SEA WATER TOXICITY TEST USING OYSTERS (CRASSOSTREA VIRGINICA)
P.A. Butler and J. I. Lowe
The following test procedure is included as a "special bioassay" for
evaluating short-term effects of specific wastes on marine mollusks. It
is recommended only for use with the commercial Eastern oyster, Crassostrea
virginica, and requires flowing unfiltered, natural sea water. This test
should be used only with materials which can be dissolved in water or other
solvents. The test has proven valuable at ERL, Gulf Breeze, where it has
been used for several years to evaluate the effect of insecticides, herbi-
cides, and other toxic organics on oysters (Butler, 1965).
This procedure, described below, is reprinted from a report by the
Subcommittee on Mollusks of the Standard Bioassay Committee for the 14th
Edition of Standard Methods for the Analysis of Water and Waste Water. It is
included in this manual by permission of Dr. Philip A. Butler, Chairman of
the subcommittee.
SHELL DEPOSITION TEST
The deposition of new shell in juvenile oysters is directly affected by
changes in ambient water quality. The degree of inhibition in shell deposi-
tion is quantitatively related to the amount of environmental stress.
This 96-hour test demonstrates the comparative toxicity of pollutants to
young oysters. The test is conducted with flowing unfiltered sea water in
the temperature range between 15 and 30 C. Actively feeding oysters extend
their mantle edges to the periphery of the shell or valves. The body can
contract, however, to occupy a much smaller area. If the peripheral valve
edges are ground away mechanically, the oysters respond by depositing new
shell to replace this loss.
The growth of new shell is primarily linear during the first week, and
the rate of deposition is an index of the animal's reaction to ambient water
quality. With acceptable water conditions, 25-mm and larger oysters deposit
as much as 1.0 mm of peripheral new shell per day. Small oysters (less than
50 mm) are more suitable than large ones because typically they form new
25
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shell deposits at temperatures ranging from about 10 to 30 C in contrast to
mature oysters, which tend to become less active at temperature extremes.
Test data are independent of minor fluctuations in temperature and
salinity during the 96-hour exposure, since the simultaneous shell deposi-
tion in control oysters is considered to be the norm or 100 percent.
Procurement and Preparation of Oysters
Oysters, about 25 to 50 mm in height, with reasonably flat,
rounded shape, are culled to singles, cleaned, and maintained in trays in
the natural environment. At the time of the test, oysters are recleaned and
about 3-5 mm of the shell periphery are removed, leaving a smoothly rounded
blunt profile. This is conveniently done by hand-holding the oysters
against an electric disc grinder. Removal of too wide a rim of shell will
make an opening into the shell cavity; damaged oysters should be discarded.
Test aquaria can be fabricated of glass or fiberglassed wood, and
should measure about 64 x 38 x 10 cm deep (25 x 15 x 4 inches) to provide
adequate space for 20 oysters. Such containers permit adequate circulation
of the water, while avoiding physical agitation of the oysters by the water
current.
The unfiltered water supply in a constant head reservoir is
delivered by calibrated siphons to the aquaria via a mixing trough into
which the toxicant is also metered in an appropriate solvent. Stock solu-
tions of the toxicant are prepared so that a delivery of 1 or 2 ml per
minute by means of a calibrated pump will result in the desired concentra-
tion. Baffles in the trough ensure adequate mixing and aeration before the
water enters the test aquaria.
The aquaria contain about 18& at 75 percent capacity and with a
flow rate of lOOfc hour~^ will provide 5& of water hour~l oyster"!. Small
oysters feed and grow readily under these conditions.
Bioassay Procedure
Oysters are prepared and randomly distributed so that each control
and test aquarium contains 20 individuals. Oysters are placed with the
left, cupped-valve down; the anterior hinged ends are oriented in one
direction. One control aquarium is established to receive only the toxicant
solvent; one aquarium is established for each desired concentration of the
toxicant.
At the end of 96 hours, all oysters are removed from the water and
the shell increments are measured. Shell deposition is not uniform on the
periphery, therefore the length of the longest "finger" of new shell on each
oyster, measured to the nearest 0.5 mm, is recorded.
26
-------
Calculation
The ratio of the mean growth of a group of test oysters to the
mean growth of the control oysters provides a percentage index of the toler-
ance of the oysters to a specified toxicant concentration. A 96-hour EC50
(concentration inhibiting shell deposition by 50%) may be calculated from an
appropriate exposure series for the indicated test conditions. These values
are relative and may differ significantly under different salinity or temper-
ature regimes. Appropriate statistical techniques should be used to deter-
mine confidence limits when possible.
A preliminary exposure series is helpful in establishing a suit-
able range of toxicant concentrations. In general, three or four oysters
exposed for 48 hours to appropriate concentrations of the test material will
bracket the range of toxicant concentrations required to determine 96-hour
EC50 data.
REFERENCE
Butler, Philip A. 1965. Reaction of Some Estuarine Mollusks to Environ-
mental Factors. In: Biological Problems In Water Pollution - Third
Seminar - 1962. U.S. Department of Health, Education, and Welfare,
Public Health Service Publication No. 999-WP-25 June, 1965.
27
-------
D. METHODS FOR THE CULTURE AND SHORT TERM BIOASSAY OF THE CALANOID COPEPOD
(ACARTIA TONSA)
John H. Gentile and Suzanne Lussier Sosnowski, Environmental Research
Laboratory, Narragansett, RI
INTRODUCTION
The methodology described in this section is designed to provide bioassay
data on the effects of a toxicant on a marine copepod. The techniques des-
cribed have been used for several years by EPA and represent the synthesis of
many researchers' efforts from both government and universities. Basically,
dose response curves are constructed from mortality data collected from 24-,
48-, 72-, and 96-hour exposure observations. While these observation inter-
vals should be considered a basic requirement, more frequent observations or
longer exposures may be necessary. From the above observations, estimates of
the LC50 and 95 percent confidence limits can be determined (Litchfield and
Wilcbxon, 1949; Finney, 1964, 1971; Standard Methods, 1971, 13th Edition).
COLLECTION AND PREPARATION OF SEA WATER
The sea water for both culture and bioassay, if possible, should be col-
lected from the study area. First, the sea water, when adjusted to 30 /oo.
salinity and 20°C, must support survival of the adult copepod Acartia tonsa
for the 96-hour bioassay period. A second and more demanding requirement is
that, with the proper enrichments, sea water supports growth of the food al-
gae and the complete life cycle of the test species. If no suitable natural
sea water is available, a synthetic sea water formulation may be employed
(Appendix 1-D).
Niskin or Van Dorn samplers can be used to collect sea water from three
to ten meters depth to avoid surface contamination. Collected sea water can
be transported to the laboratory in glass or polyethylene carboys that have
been aged with sea water. In the laboratory, the water is filtered through a
1.0 acid washed filter (glass fiber, cellulose, acetate, nylon or poly-
carbonate) to remove particulate matter and stored at 4°C in the above con-
tainers.
Measurements of salinity, dissolved oxygen, and pH should be recorded at
the time of collection.
ALGAL FOOD CULTURES
Although a variety of algal diets have been used for copepod cultures
(Zillioux and Wilson, 1966; Heinle, 1969; Katona, 1970; Nassogne, 1970), the
28
-------
following modification of Wilson and Parish (1971) has been used successfully
in the EPA Environmental Research Laboratory in Narragansett (Table 1-D).
We have added Skeletonema costatum because it is a naturally occurring food
for Acartia tonsa.'
TABLE 1-D. COMPOSITION OF ALGAL DIET AND RECOMMENDED CONCENTRATION
FOR FEEDING, EGG LAYING, AND NAUPLIAR FEEDING
Item
Skeletonema costatum
Thalassiosira psuedonana
Isochrysis galbana
Rhodomonas baltica
Total cells/liter
Adult &
Copepodite
5.0 x 106
7.0 x 106
5.0 x 106
3.0 x 106
2.0 x 107
Naupliar
5.0 x 105
7.0 x 105
5.0 x 105
3.0 x 105
2.0 x 106
Egg Laying
1.5 x 107
2.1 x 107
1.5 x 107
9.0 x 106
6.0 x 107
These algae are grown axenically in filtered natural or synthetic sea
water at 30 loo salinity and 20°C with 2500-5000 lux continuous illumination
or 14L:10D. The nutrient enrichments are modifications of those of Guillard
and Ryther (1962)(Appendix 2-D).
Algal cultures may be grown either in standard test tubes or flash cul-
tures if desired; or in the fill and draw semi-continuous system described
below.
Enriched sea water is dispensed into either screw-capped test tubes
(50 ml) or Erlenmeyer flasks fitted with Teflon lined caps. After autoclav-
ing (15 min @ 15 psi & 250°F), the medium is allowed to cool and equilibrate
with atmospheric gases for 48 hours. Sterility checks are made on each set
of autoclaved medium by randomly selecting a representative number of tubes
or flasks and inoculating one ml of their contents into tubes of sterility
check medium (Appendix 2-D). Caps are tightened and the inoculated tubes
stored in darkness for up to two weeks. The appearance of turbulence or
opalescence in the test medium indicates the presence of contamination.
Tubes or flasks are inoculated with each alga on a regular basis to con-
tinually provide a log-phase, high density food source, the frequency being
determined from interpretation of algal growth curves. The cultures should
be harvested at their maximum log-growth phase cell density. Although this
system works, it is very time consuming since it requires frequent cell
counts and a large turnover of glassware.
29
-------
The recommended algal culture system is of a fill^draw type in which
cultures are easily maintained near their maximum log-phase cell density and
growth rate (Appendix 3-D). It is then a simple matter to draw off a con-
stant volume and replace it with fresh medium so that within 24 hours the
culture will have reached the same cell density. When intervals longer than
24 hours occur between harvests, proportionally greater amounts of cul'ture
are drawn off and replaced. This system can be scaled up or down, depending
on food needs. But most importantly, this system produces algal food that
is physiologically and nutritionally consistent. Thus the nutritional
history of the test species is better controlled. If this system is used, a
series of tube cultures of each of the four algal foods must be maintained
concurrently in case of contamination of the large cultures.
Algal cell densities may be determined in a variety of ways. Direct
microscopic counts can be made with a haemocytomer, Palmer-Maloney chamber,
or Utermohl chamber (inverted scope) (Schwoerbel, 1970) (Standard Methods,
1971) . In addition, an electronic particle counter is an accurate and rapid
method for determining unialgal densities. Finally, manual counts, if
necessary, can be related to chlorophyl absorbance at 440 my or 665 my,
using a spectrometer. A curve that compares cells/ml with absorbancy should
be prepared from serial dilutions of each algal culture. Then a rapid and
simple measure of absorbancy can be used to replace the cell count.
ZOOPLANKTON CULTURE
Collection
Zooplankton (including Acartia tonsa) are collected by slowly (<4 km/hr)
towing a plankton net (aperture 150 to 250 ym at a depth of one to three
meters). Captured animals are carefully transferred to insulated containers
three-fourths filled with ambient sea water. The population density should
not exceed ca. 25/H to assure that the dissolved oxygen concentration
remains adequate if the organisms are not returned to the laboratory within
one to two hours. It is imperative to measure and record the temperature
and salinity at the time of the collection since these parameters must be
maintained during the initial stages of laboratory culture.
Holding
In the laboratory, the samples immediately are transferred to 2.3£ (190
x 100mm) borosilicate crystallizing dishes. Volume is adjusted to 2000 ml
with filtered sea water at ambient temperature and salinity; each dish is
then fed the adult algal diet (Table 1-D). The cultures are incubated at
ambient temperature and 14L:10D cool white illumination of 1000 lux. After
24 hours, acclimation of the cultures to 20°C and 30 loo salinity should
commence. Salinity and temperature increments of 5 /oo and 5°C per day are
satisfactory. Organisms can remain in the original vessel and culture
volumes can be changed by alternately siphoning through 150-ym plankton
netting and adding sea water of a different salinity. Transfers are made by
carefully pipetting or slowly siphoning organisms to new vessels. During
acclimation, a daily feeding schedule is maintained.
30
-------
Holding and acclimation can also be accomplished by adding the tow col-
lections to 4-12£ aspirator bottles equipped with low rpm (<_ 25 rpm) motors.*
Organism density should be adjusted to 1:10 ml of culture volume.
Sorting and Identification
The plankton tow contains a mixture of species from which Acartia tonsa
must be isolated. For basic information on the taxonomy and biology of the
genus Acartia and other coastal calanoids, the following papers are recom-
mended (Conover, 1956; Heinle, 1966, 1969; Wilson, 1932; Rose, 1933; Fraser
and Hansen, Eds., Serie Fiches Identification Zooplancton). (See Appendix
4-D for comparison of calanoid copepods usually occurring with Acartia
tonsa.)
To facilitate capture of organisms, the culture volume is reduced from
2000 ml to 500 ml by slowly siphoning sea water, using 150-ym plankton net-
ting over the siphon intake. Individual adult organisms are attracted to the
edge of the dish with a dim light (440-1400 lux) and carefully drawn up into
a wide-bore (>_ 2mm) transfer pipette. Individual animals are placed in de-
pression slides and identified microscopically. Once the investigator
becomes familiar with Acartia morphology and swimming behavior (short spurts
as opposed to long glides), it will not be necessary to identify animals by
fifth leg. With practice, an investigator can examine several animals simul-
taneously under low magnification. Extraneous species then are removed and
Acartia are transferred to food-enriched filtered sea water at 30 °/oo and
20°C. Contamination of species is prevented by excluding all naupliar and
juvenile forms.
Mass Culture
The objective of this system is to provide large quantities of Acartia
tonsa of standard age for short-term bioassays.
The mass culture unit is derived from culture systems used by Mullin and
Brooks (1967) and Frost (1972). The culture vessel is a pyrex aspirator
bottle whose size can range from 4.0 to 40 liters depending on the number of
copepods needed. The contents are gently mixed by a low rpm motor* (<25 rpm)
mounted above the culture vessel. Thus, algal food is suspended where these
planktonic copepods normally feed. It must be emphasized that water move-
ment is gentle and free of vortices such as produced by magnetic stirrers
(Figure 1-D). Cool white fluorescent lights provide 2000 lux illumination
incident to the culture surface on a 14L:10D cycle.
Acartia tonsa females are capable of producing more than 30 eggs per
female per day when fed the adult algal food ration recommended in Table 1-D
(Wilson and Parrish, 1971). Thus, if 250 or more gravid females are brooded,
theoretically, more than 5,000 eggs will be produced within 24 hours. For
this potential number of adults, a 40-liter culture vessel would be desirable.
Generally, the relationship between culture volume (mis) and organism density
is 10 ml:l.
*W.W. Grainger, Inc. Dayton Shaded Pole Gearmotors, 20 rpm. All angle
Operation #2Z808.
31
-------
Q
NJ
Q\
•LOW RPM MOTORS-
-1/4" DRILL CHUCK-
Q
•PLEXIGLASS RODS-
- ASPIRATOR BOTTLES-
COOL WHITE FLUORESCENT
LAMPS
-SILASTIC TUBING
-HOFFMAN CLAMPS
Figure 1-D. Mass Copepod Culture Systems (Static).
-------
OJ
OJ
-125 X 90 mm PLEXIGLASS CYLINDER
2000ml FILTERED SEAWATER
-PLANKTON NETTING, 250 MICRONS
APERATURE-25 mm FROM BOTTOM
XXXXXXXXXXXXXXXXXXXX X XXXXXXXXX
•2.3 LITER, 190 X 100 mm PYREX CRYSTALIZING DISH
Figure 2-D. Generation Cage (after Heinle) (personal communication).
-------
TABLE 2-D. PROTOCOL FOR MASS COPEPOD CULTURE
Step
1
2
3
4
5
6
7
8
9
10
11
12
Day of Culture
1-3
4
5-6
7
8-9
10
11-12
13
14-15
16
17-18
19
Age- Standardized
Naupliar diet daily
(Table 3)
Replace 50% culture
medium with filtered
s.w. retaining organ-
isms. Feed as in 1
As in 1
Repeat step 2
As in 1
Repeat step 2
Adult diet
Adult diet daily
Repeat step 6
As in 7
Repeat step 6
As in 7
Harvest for
Bioassays
Non- Age-Standardized
Adult diet daily
(Table 3)
As in step 2. Feed
as adults .
As in 1
Repeat step 2
As in 1
Repeat step 2
As in 1
Repeat step 2
As in 1
Harvest 33% of culture
including organisms.
Transfer remaining 67%
to a clean carboy by
siphon* & add filtered
s.w. to volume.
Repeat steps 1-10
___
*Rate of siphoning is controlled by difference in "head pressure". Do not
constrict the siphon tube or animals will be damaged.
34
-------
OJ
_£
-FILTERED SEAWATER
//-NN //^NN
Cx_ —
ALGAL
FOOD
PUMP
j — ^ a
' r — i u
J
CONSTANT HEAD TANK
~i_ .
20 LITER CYLINDRICAL
VESSELS
— •->— - >— — • """IT1
--STANDPIPE
1
1
l—^-K-LOW RPM MOTOR
V— STIRRING ROD
\ rjf-150 MICRON
\ m/ COLLAR
VALVE DRAINS-
STANDPIPE-
DRAIN-
Figure 3-D. Mass Copepod Culture (Flowing).
-------
Fifty to 100 gravid females are placed in each of three to five genera-
tion cages (Figure 2-D), immersed in 2.3/H crystallizing dishes containing
ca. 2000 ml of sea water, and fed at the algal food concentration recommended
for egg-laying (Table 1-D). The generation cage allows the eggs to pass
through the net and hatch, eliminating the possibility of cannibalism by
adults. After 24 hours, the adults are removed by gently lifting each
generation cage out of the dish and quickly immersing it in another dish
with three times the usual food density. The remaining sea water from all
dishes containing eggs and nauplii is carefully siphoned into a glass aspir-
ator bottle containing filtered sea water. The final volume is adjusted and
the naupliar culture is fed as in Table 1-D. If a second mass culture is
desired, the procedure is repeated after 24 hours.
The average length of each developmental stage in the life cycle of
Acartia tonsa at 20°C and 30 °/oo is:
Stage Length in Days
Egg (newly oviposited) 1
Nauplius (6 instars) 7
Copepodite (6 instars) 6
Adult (until gravid) ^3.
Total Life Cycle 17
During the first six days of mass .-culture, only naupliar stages are pre-
sent. Daily feeding should be 2 x 10 cells/*, (Table 1-D) and 50 percent of
the culture medium should be siphoned off and replaced with clean medium on
the third and seventh days. The intake end of the siphon should be covered
with 60 ym netting to prevent loss of nauplii.
After the seventh day, copepodites should be present and, from this
point on, feeding should be 2 x 10? cells/^/day with 50 percent replacement,
of the culture volume with filtered sea water every third day. Within 16 to
17 days, the population will reach maturity and can be bioassayed or used to
start new cultures. Average adult life span at 20°C is _f30 days.
We have also found it useful to maintain a non-age-standardized mass
culture in reserve. Gravid females from the original generation cages are
used to start a 12-liter (3 1/2-gallon) system and fed the adult food ration;
50 percent of their culture water is replaced every third day. In addition,
approximately 1/3 of the culture (including organisms) is harvested periodi-
cally (10-14 days) to keep the population at ca. 50 adults and cope-
podites/£. This precaution is worth the effort since the high density cul-
tures have occasionally "crashed" for no apparent reason. A protocol for
this system is given in Table 2-D.
If a constant source of filtered (l.Oy cartridge filter) sea water is
available, a flowing water mass culture system can be used (Figure 3-D).
This system consists of a constant head tank which feeds two large
36
-------
cylindrical reaction vessels. Dilution water flow is controlled by capillary
restriction or clamps. The four species algal food is proportionally metered
by peristatic pump to provide a constant cell density of 25 x 10? cells/ 'I.
This cell density can sustain culture densities in excess of 100 adults and
copepoditesA , though harvesting is recommended to keep the density at
The reaction vessels are 30 cm high, 30 cm in diameter, and have a 25
cm standpipe. This provides approximately 18& culture volume. The stand
pipe has a collar of 150ym nitex net which effectively retains both eggs and
nauplii even though they are considerably smaller than the pores. Too fine
a net produces excessive clogging. It is likely that bacterial and algal
growths reduce the effective mesh size to occlude particles as small as
50 ym. This net collar requires periodic brushing to maintain effective
drainage. The reaction vessels, illuminated as in the static system, are
equipped with low rpm (^25) motors to maintain the population in suspension.
The dilution rate, approximately 10 ml/min, effectively replaces 50 percent
of the culture volume every 24 hours, although the total volume pumped is 80
percent of the reaction volume. Flow rates > 10 ml/min can be used, with
caution, to avoid washing out eggs, nauplii, or both.
Harvesting
Mass cultures of copepods that have reached the adult stage are harvested
for bioassays as follows: the culture volume is reduced by 75 percent,
using a slow siphon whose intake is covered with 60 ym plankton netting.
The remaining 25 percent of the culture, including organisms, is carefully
transferred .to 2.3X, pyrex crystallizing dishes (ca. 2000 ml/dish). This
transfer is critical and is best performed as follows: because of fragility
of the organism, do not constrict the discharge tube to reduce flow. Dis-
charge flow through the ventral tubulation on the aspirator is controlled by
minimizing the head pressure between the culture vessel and the crystalliz-
ing dish. A slow flow minimizes turbulence and opportunity for organisms to
collide into vessel walls.
Harvested animals can be concentrated in the crystallizing dishes by
further siphoning the culture medium. Capture is facilitated by using posi-
tive phototactive response of the animals.
Short- Term Bioassays
Adult Acartia tonsa (Dana) and culture conditions previously described
are required for the following short-term bioassays. (See Figure 4-D.)
Range-Finding Bioassays
1. Ten adult Acartia are tested per replicate with two replicates required
per test concentration and control. Feeding is omitted for the duration of
the assay. A solvent control must be included when appropriate.
2. Test container must be a suitable flatbottom borosilicate glass dish
containing 100 ml sea water. The depth of medium must be _>2.0 cm.
37
-------
RANGE FINDING BIOASSAY:
Harvested adults (ca. 180)
1
Control
10 10
1
0.1 0.33 1.0 3.3 10
etc. etc. etc. etc. etc.
i
Adults
Evaluate Mortality and Moribundlty at 24-hour
intervals for a 96-hour exposure.
DEFINITIVE BIOASSAYS:
Calculate LC50 for 96-hour data
Harvested Adults (ca. 360)
I 1
Control etc.
1
15
l i
15 15
1
1 1 1
etc. LC50 etc.
etc.
1
1
etc
1
Adults
Evaluate Mortality and Moribundity at 24-hour
intervals for a 96-hour exposure.
Figure 4-D. Bioassay protocol.
38
-------
3. Toxicant concentration selection
Generally, a broad range of concentrations covering at least three
orders of magnitude is chosen initially. This is followed by a progressive
bisection of intervals on a logarithmic scale (see Standard Methods, 1971).
4. Toxicant administration
a. Water miscible toxicants are added immediately prior to the addi-
tion of the test species.
b. Water immiscible toxicants are dissolved in a suitable solvent
prior to addition to the test medium. Solvent evaluation must be performed
to insure solvent concentrations used are not toxic.
5. Ten adult Acartia are captured from stock cultures with a wide-bore
transfer pipette and transferred to a 20-ml beaker containing undosed fil-
tered sea water (ca. 5 ml). Adjust the final volume of this beaker to 15
ml. The animals and the 15 ml of medium are added to 85 ml of toxicant-
dosed medium by immersing the beaker and gently rinsing.
6. Exposure period is 96 hours. The number of dead and moribund copepods
are observed and recorded at 24, 48, 72, and 96 hours of exposure. To
ascertain death, gently touch a motionless animal with a sealed glass capillary
probe. Dead animals are removed at each observation point. Control mortal-
ities in excess of 15 percent invalidate the experiment.
7. At each observation period, dissolved oxygen and pH should be measured,
particularly if wastes contain large amounts of organic matter. Since the
test species is very sensitive to agitation, these measurements at all test
concentrations must be made on a series of concurrently prepared unioculated
samples.
Definitive Short-Term Bioassay
General culture conditions and handling follow previous discussions.
The specifications for this assay are:
(1) Fifteen adults are to be tested in each of three replicates per
toxicant concentration and control.
(2) Test vessels are described above.
(3) Concentration ranges for toxicant must include at least two levels
above and below the 96-hour LC50 determined from range finding bioassays.
(4) Exposure and data collection are described above.
(5) Calculations and data presentation are as described in Standard
Methods (14th Ed.) pp. 565-577. Alternate methods of data presentation are
desirable, particularly the application of confidence limits. (See Litch-
field and Wilcoxon, 1949, and Finney, 1964, 1971.)
39
-------
COMMENTS
The bioassay methodology is, at best, a general framework that is sub-
ject to modifications as determined by the type of toxicant and the experi-
mental design. For example, in assays with toxicants that readily adsorb to
container walls and fail to remain in solution, transfer of organisms to
freshly dosed media is required. The frequency of transfer is determined
after rates of solubility and adsorption are known.-
The mass culture system described can be used as a holding and acclima-
tion system for indigenous populations. For example, in many geographical
areas, A^ tonsa is replaced by A. clausi during the winter months. Using
the above system, we have held _A. clausi at 10°C for several weeks. These
organisms were used in bioassays at 10°C with excellent results. Thus, we
feel that this system, with appropriate modifications, can be used to hold
and culture a variety of zooplankters.
In the event that natural sea water is not suitable for survival,
growth, and reproduction of the test species, the following synthetic formu-
lations are recommended. The formulation in Appendix 1-D has been used for
both whole life history culture and numerous bioassay studies at this labora-
tory. Heinle (1969) found the commercial sea water Instant Ocean, suitable
for the culture of both A^. tonsa and J2. affinis. Data are not available on
the use of Instant Ocean in bioassays or regarding a comparison to natural
sea water. Therefore, Instant Ocean is recommended only for culture, not for
bioassays, until suitable comparative data are available.
40
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REFERENCES
American Public Health Service. 1971. Standard Method for the Examination
of Water and Wastewater. 13th ed. New York. 874 p.
Conover, R.J. 1956. Oceanography of Long Island Sound, 1952-1954. VI. Bi-
ology of Acartia clausi and A_. tonsa. Bull. Bingham Oceanogr. Collect.
Yale Univ. 15:156-233.
Davey, E.W., J.H. Gentile, S.J. Erickson, and P. Betzer. 1970. Removal of
Trace Metals from Marine Culture Medium. Limnol. & Oceanogr. 15:486-488.
Finney, D.J. 1964. Statistical Method in Biological Assay. 2nd ed. Hafner
Publishing Co., New York. 668 p.
. 1971. Probit Analysis. 3rd ed. Cambridge Univ. Press. London.
333 p.
Fraser, J.H., and V. Kr. Hansen, eds. Serie Fiches Identification Zoo-
plancton.
Frost, B.W. 1972. Effects of Size and Concentration of Food Particles on
the Feeding Behavior of the Marine Planktonic Copepod Calanus pacificus.
Limnol. & Oceanogr. 17(6):805-815.
Gentile, J.H., J. Cardin, M. Johnson, S. Sosnowski. 1974. Power Plants,
Chlorine, and Estuaries. Amer. Fish. Soc., 36th Annu. Meeting, Honolulu,
Sept. 9-11.
Guillard, R.R., and J.H. Ryther. 1962. Studies of Marine Planktonic Dia-
toms. I. Cyclotella nana Hustedt, and Detonula confervacia (Cleve)
Grant. Can. J. Microbiol. 8:299-339.
Heinle, D.R. 1966. Production of a Calanoid Copepod, Acartia tonsa, in the
Patuxent River Estuary. Chesapeake Sci. 7:59-74.
. 1969a. Effects of Temperature on the Population Dynamics of
Estuarine Copepods. Ph.D. Thesis, Univ. Maryland, College Park. 132 p.
. 1969b. Culture of Calanoid Copepods in Synthetic Sea Water. J.
Fish. Res. Bd. Can. 26(1):150-153.
Katona, S.K. 1970. Growth Characteristics of the Copepods Eurytemora
affinis and 15. herdmani in Laboratory Cultures. Helgolander wiss.
Meeresunters. 20:373-384.
41
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Kester, E., I. Dredall, D. Connors, and R. Pytowicz. 1967. Preparation of
Artificial Sea Water. Limnol. & Oceanogr. 12(1):176-178.
Litchfield, J.T., and F. Wilcoxon. 1949. A Simplified Method of Evaluation
Dose-Effect Experiments. J. Pharmacol. Exper. Ther. 96(2):99-115.
Mullin, M.M., and E.R. Brooks. 1967. Laboratory Culture, Growth Rate, and
Feeding Behavior of a Planktonic Marine Copepod. Limnol. & Oceanogr.
12:657-666.
Nassogne, A. 1970. Influence of Food Organisms on the Development and
Culture of Pelagic Copepods. Helgolander wiss. Meeresunters. 20:333-
345.
Rose, M. 1933. Faune de France. No. 26. Copepodes Pelagiques. Librairie
de la Facultd des Sciences. Reprinted 1970 by Kraus Reprint, Nendeln
Leichtenstein.
Schwoerbel, J. 1970. Methods of Hydrobiology. Pergamon Press, New York.
Wilson, C.B. 1932. The Copepods of the Woods Hole Region, Massachusetts.
Smithsonian Institute, U.S. National Museum Bulletin 158.
Wilson, D.F., and K.K. Parrish. 1971. Remating in a Planktonic Marine
Calanoid Copepod. Mar. Biol. 9:202-204.
Zillioux, E.J., and D.F. Wilson. 1966. Culture of a Planktonic Calanoid
Copepod through Multiple Generations. Science 151:996-998.
42
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APPENDIX 1-D. SYNTHETIC SEA WATER FORMULATION*
Chemical
Nad 24.00
Na2S04 4.00
CaCl2.2H20 1.47
MgCl2.6H20 10.78
KC1 0.70
H3B03 0.03
NaH003 0.20
*Medium is modified from Kester et al. (1967). Salinity is 34 °/oo and
pH 8.0 and must be adjusted to 30 °/oo with distilled or deionized water.
Trace metal contaminants from major salts must be eliminated by ion exchange
stripping (Davey et al., 1970). Na2EDTA (300 mgs/A) may be used for holding
and culture, but must be omitted in bioassay studies with trace metals.
43
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APPENDIX 2-D. SEA WATER AND STERILITY ENRICHMENT
(A) Sea water enrichments for stock algal culture maintenance (After Guillard
and Ryther, 1962):
Item
Amount
NaNO,
NatLPO, .H.(
242
Vitamins:
Thiamine HC1
Biotin
B12
Trace Metals:
CuSO .5H 0
.
CoCl,.6H,0
Fe-sequestrine
75 mg/liter
5 rag/*
10 mg/fc
0.10 mg/£
0.50 wg/A
0.50 ug/£
0.002 mg/£
0.004 mg/£
0.002 mg/£
0.036 mg/£
0.001 mg/A
1.0 mg
(0.13 mg
Buffer:
TRIS-500 mg/i @ pH 7.8-8.2
(B) Sterility enrichments to be added to enriched sea water medium above
before autoclaving:
Sodium Glutamate
Sodium Acetate
Gycline
Nutrient Agar
Sucrose
Sodium Lactate
L & D Alanine
250 mg/£
250 mg/l
250 mg/£
50 mg/£
250 mg/£
250 mg/A
250 mg/Jl
44
-------
MEDIA
40 L
STOPPER
MEDIA
12 L
T°J/
\°J
40 WATT FLUORESCENT LIGHTS
COOL WHITE
PINCH CLAMP
COTTON PLUG
TO AIR SUPPLY
70% ETOH
>—AIR VENT -
COTTON PLUG
ALUMINUM CLAMP
MEDIA TUBE
TUBING CONNECTOR
PINCH CLAMP
VENT
ALUMINUM CLAMP
ALGAL CULTURE
AIR STONE
SPIN BAR
MAGNETIC MIXER
PINCH CLAMP
STERILE DISPENSING
TUBE
APPENDIX 3-D. Algal culture.
-------
APPENDIX 4-D
DESCRIPTIVE CHARACTERISTICS FOR SELECTED NERITIC COPEPODS
Suzanne Lussier Sosnowski
The purpose of Appendix 4-D is to provide a list of easily recognizable
morphological characteristics which can be used to identify live copepods
from field collections. These characteristics allow identification at less
than lOOx magnification with a dissecting microscope. This list of charac-
teristics is not intended to be a taxonomic key. References are provided if
further identification is required.
Acartia tonsa is placed first in the drawings because it is the recom-
mended species for ocean disposal bioassays at this time. The remaining
species are those normally found with A. tonsa in a plankton collection.
The species are arranged in order of decreasing morphological similarity to
A. tonsa. All illustrations are drawn to scale so that the relative size of
the species may be compared at a glance.
Each illustrated species consists of the following:
A - female dorsal view;
B - male dorsal view;
C - male fifth leg;
D - female fifth leg.
The spatial and temporal distribution of the copepods included in this
appendix also can be used as an aid to identification. Distribution of
these species is governed by both temperature and salinity. There is,
however, considerable seasonal overlap. The following tables can be used to
assist the researcher in anticipating the composition of a plankton tow when
temperature and salinity information is available.
46
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APPENDIX 4-D (Continued)
DESCRIPTIVE CHARACTERISTICS FOR SELECTED NERITIC COPEPODS
Euryhaline
(10-35 °/oo)
Acartia tonsa
Acartia clausi
Acartia longiremis
Eurytemora affinis
Eurytemora americana
Eurytemora herdmani
Pseudodiaptomus coronatus
Temora longicornis
Salinity
Stenohaline
(25-35 /oo)
Centropages typicus
Centropages hamatus
Tortanus discaudatus
Temperature
Not > 20° C
Acartia clausi
Pseudocalanus minutus elongatus
Eurytemora herdmani
Tortanus discaudatus
Not < 10° C
Acartia tonsa
Eurytemora affinis
Pseudodiaptomus coronatus
Centropages typicus
Oithona similis
Species
Descriptive Characteristics
Acartia tonsa
<* 1.00-1.15 mm
? 1.25-1.50 mm
1. Spindle-shaped body
2. Urosome 1/3 length of metasome
3. Caudal rami as long as wide
4. Long hairs on first antennae
5. First antennae nearly straight
on female, but with acute
bend near proximal end on
male
6. No egg sacs present
7. Swims in short spurts
Continued
47
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APPENDIX 4-D (Continued)
DESCRIPTIVE CHARACTERISTICS OF SELECTED NERITIC COPEPODS
Species
Descriptive Characteristics
Acartia clausi
<* 1.00-1.10 mm
9 1.15-1.25 mm
1. Spindle-shaped body
2. Urosome 1/3 length of metasome
3. Caudal rami twice as long as wide
4. Long hairs on first antennae
5. First antennae nearly straight on
female, but with acute bend
near proximal end on the male
6. Three or four pairs of blue dots
on ventral surface of metasome,
visible only on fresh tow
material; preservative causes
pigment to fade
7. No egg sacs present
8. Swims in short spurts
Acartia longiremis
3 0.8-1.0 mm
9 0.9-1.1 mm
1. Spindle-shaped body
2. Urosome 1/3 length of metasome
3. Caudal rami 2-3 times longer than
wide
4. Long hairs on first antennae
5. First antennae of male have small
hinge
6. Fifth segment with delicate spine
on dorsal surface of each
posterior corner
7. No egg sacs
8. Swims in short spurts
Pseudocalanus minutus
elongatus
& 1.00-1.25 mm
9 1.20-1.60 mm
Body shape elliptical
Urosome 1/2 as long as metasome
Caudal rami longer than anal
segment
Short hairs on first antennae
Animals from fresh tow
have reddish color
Female lacks fifth pair of legs
Female often with single egg sac
Continued
48
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APPENDIX 4-D (Continued)
DESCRIPTIVE CHARACTERISTICS OF SELECTED NERITIC COPEPODS
Species
Descriptive Characteristics
Eurytemora affinis
d* 1.4-1.6 mm
9 1.4-1.5 mm
1. Body bullet-shaped
2. Female with large triangular
"fenders" at posterior corners
of fifth segment
3. Very long caudal rami
4. Spines cover the anal segment
and caudal rami
5. Short hairs on first antennae
6. Right first antenna of male is
hinged
7. Female often with single large
egg sac
8. Swims with gliding motion
Eurytemora americana
d 0.75-0.95 mm
9 1.60-1.85 mm
1. Body bullet-shaped
2. Female with large "fenders" with
sharp spines at posterior corners
of fifth segment
3. Very long caudal rami
4. Spines cover the anal segment and
caudal rami
5. Short hairs on first antennae
6. Right first antenna thickened and
hinged
7. Female often with single large egg
sac
8. Swims with gliding motion
Eurytemora herdmani
3 1.2-1.5 mm
9 1.3-1.6 mm
1. Body bullet-shaped
2. Female with large triangular
"fenders" at posterior corners
of fifth segment reaching beyond
genital segment
3. Very long caudal rami
4. Short hairs on first antennae
5. Right first antenna of male hinged
6. Female often with single large
egg sac
7. Swims with gliding motion
Continued
49
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APPENDIX 4-D (Continued)
DESCRIPTIVE CHARACTERISTICS OF SELECTED NERITIC COPEPODS
Species
Descriptive Characteristics
Pseudodiaotomus coronatus
<* 1.00-1.25 mm
9 1.25-1.50 mm
1. Body bullet-shaped
2. Female genital segment swollen with
patches of bristles and spines
protruding ventrally
3. Caudal rami 2-3 times as long as
wide
4. Sparse hairs on first antennae
5. Left first antenna of male
thickened and hinged
6. Female with two egg sacs, the right
sac containing only two eggs
7. Swims with gliding motion
Centropages typicus
°* 1.0-1.60 mm
9 1.25-1.75 mm
6.
7.
Body rectangular with well defined
head region
Female with large unequal "fenders"
(right side larger) on posterior
corners of fifth segment
Male with smaller unequal spines
(left side larger) on posterior
corners of fifth segment
Short hairs on first antennae
First antennae reach beyond tips of
caudal rami
Tooth-like spines on -the first,
second, and fifth segments of male
and female first antennae
Right first antenna of male
thickened and hinged
No egg sacs present
Female genital segment with several
stiff spines
Continued
50
-------
APPENDIX 4-D (Continued)
DESCRIPTIVE CHARACTERISTICS OF SELECTED NERITIC COPEPODS
Species
Descriptive Characteristics
Centropages hamatus
o" 0.9-1.2 mm
9 1.0-1.4 mm
1. Body rectangular with well defined
head region
2. Female with unequal spines on pos-
terior corners of fifth segment,
the right turned outward
3. Male with symmetrical spines on
posterior corners of fith seg-
ment
4. Short hairs on first antennae
First antennae reach beyond tips of
caudal rami
No tooth-like processes present on
first antennae
5. Right first antenna of male
thickened and hinged but not as
pronounced as in C_. typicus
6. No egg sacs present
Tortanus discaudatus
rf 1.75-2.00 mm
9 2.00-2.25 mm
1. Very large spindle-shaped body
2. Female has symmetrically curved
spines on posterior corners of
fifth segment
3. Urosome very asymmetrical in male
and female
4. Female right caudal ramus twice as
wide as left
5. Male caudal rami unequal
6. Male urosome curved to right
7. First antennae reach caudal rami
8. Right first antenna thickened and
hinged in male
9. No egg sacs present
Continued
51
-------
APPENDIX 4-D (Continued)
DESCRIPTIVE CHARACTRISTICS OF SELECTED NERITIC COPEPODS
Species
Descriptive Characteristics
Temora longicornis
<* 1.00-1.35 mm
9 1.00-1.50 mm
1. Body shaped like bear's paw; wide
at head, tapering rapidly to
fifth segment
2. Caudal rami very long
3. Male urosome longer and narrower
than female urosome
4. First antennae have very short
hairs
5. Right antenna on male thickened
and hinged
6. No egg sacs present
Oithona similis
tf 0.6-0.70 mm
9 0.7-0.95 mm
1.
2.
3.
4.
5.
Body spindle-shaped
Urosome 3/4 length of metasome
First antennae have very long hairs
First antennae of male hinged twice
Female has two ovisacs appressed to
sides of urosome
Oithona nana
tf 0.48-0.57 mm
9 0.50-0.65 mm
1. Spindle-shaped body
2. Urosome 3/4 length of metasome
3. First antennae have very long hairs
4. First antennae hinged twice on male
52
-------
ACARTIA TONSA
ACARTIA CLAUSII
ACARTIA LONGIREMIS
PSEUDOCALANUS MINUTUS ELONGATUS
2mm
APPENDIX 4-D-l
-------
EURYTEMORA AFFINIS
EURYTEMORA AMERICANA
Ul
EURYTEMORA HERDMAN
0
i i
PSEUDODIAPTOMUS CORONATUS
I 2mm
i i I i i i i I i i i i I
APPENDIX 4-D-2
-------
CENTROPAGES TYPICUS
CENTROPAGES HAMATUS
Ui
TORTANUS DISCAUDATUS
TEMORA LONGICORNIS
2mm
APPENDIX 4-D-3
-------
OITHONA SIMILIS
OITHONA NANA
B
B
0 I 2mm
i i i i I i i i i I i i i i I i i i i I
APPENDIX 4-D-4
-------
REFERENCES
Katona, S.K. 1971. The Developmental Stages of Eurytemora affinis (Poppe,
1880)(Copepoda, Calanoida) Raised in Laboratory Cultures, including a
Comparison with the Larvae of Eurytemora americana Williams. 1906, and
Eurytemora herdmani Thompson and Scott, 1897. Crustaceana 21(1):5-20.
Eurytemora affinis; A, B, p.10, Fig. 47, 54, 49, 53 and p.14,
Fig. 93, C, D, p. 13, Fig. 88, 85.
Mori, Takamochi. 1964. The Pelagic Copepoda from the Neighboring Waters
of Japan. The Soyo Company, Inc* Tokyo, Japan. 150 pp. 80 pi.
Acartia clausi; B, PI. 51, Fig. 9.
Acartia longiremis; B, PI. 51, Fig. 6
C, PI. 51, Fig. 9 '
Tortanus discaudatus: A, B, PI. 52, Figs. 4, I.
Oithona similis; A, B, PI. 62, Figs. 4, 8.
Oithona nana; A, B, PI. 63, Figs. 1, 2
Sars, G.O. 1903. An Account of the Crustacea of Norway. Vol. IV. Copepoda.
Bergen Museum. Alb. Cammermeyer's Forlag, Christiana. 171 pp. 108 pi.
Acartia longiremis; A, PI. XCIX
Acartia clausi; C, PI. CI
Pseudocalanus minutus elongatus: A, B, C
PI. X, XI
Centropages typicus; A, B, PI. XLIX, LI
Centropages hamatus; A, B, PI. LII
Temora longicornis; A, B, PI. LXV
Thompson and Scott. 1897. Proceedings, Liverpool Biological Society.
Vol. XII, PI. V p. 78.
Eurytemora herdmani; A, B, C, D, p. 78, Figs. 1, 2, 9, 11, 10, 8.
Wilson, Charles B. 1932. The Copepods of the Woods Hole Region Massachu-
setts. Smithsonian Institution. United States Government Printing
Office. Bulletin 158. 635 pp.
Acartia tonsa; A, B, C, D, p. 161, Fig. 109 a, b, c, d.
Acartia clausi; A, D, p. 164, Fig. 112 a, b.
Acartia longiremis; D, p. 165, Fig. 113 c.
Eurytemora americana; A, B, C, D, p. 109, Fig. 72 a, b, c.
Pseudodiaptomus coronatus; A, B, C, D, p. 102, Fig. 68 a, c, b.
57
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Centropages typicus; C, D, p. 88, Fig. 60 d, e.
Centropages hamatus; C, D, p. 89, Fig. 61 e, f.
Tortanus discaudatus: C, D, p. 167, Fig. 114 f, g.
Temora longicornis; B, C, D, p. 105, Fig. 70 b, d, e.
Acknowledgment: Special appreciation is extended to Ms. Lianne Armstrong
for the illustrations in this appendix.
58
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E. CULTURING THE MYSID (MYSIDOPSIS BAHIA) IN FLOWING SEA WATER OR A STATIC
SYSTEM
D.R. Nimmo, T.L. Hamaker, and C.A. Sommers
INTRODUCTION
Many freshwater but few estuarine or marine animals have been found
practical for life-cycle toxicity tests. Life cycles of certain marine
species are complex: many require an estuarine existence as larvae or juve-
niles, followed by adult migration to deeper waters offshore to reproduce.
Culture and maintenance of estuarine and marine species entail elaborate and
expensive equipment with temperature or salinity controls, anticorrosion
surfaces, and if necessary, special filtration systems. We have cultured
the bay mysid, Mysidopsis bahia, for life-cycle toxicity tests at ERL, Gulf
Breeze, in (1) flowing sea water and (2) a re-circulating aquarium. Both
methods are described below; however, the re-circulating method is appropri-
ate for laboratories not equipped with flowing seawater.
FLOWING SEA WATER METHOD
Mysids, collected from Santa Rosa Sound near Pensacola, Florida, are
cultured in the laboratory in 38-liter glass aquaria supplied with filtered
(20y) flowing water (10 to 27 parts per thousand salinity) at 18 to 28°C.
Mysids are fed 48-hour-old Artemia salina larvae daily. Overflow from each
aquarium exits through a standpipe, where an attached ring of screen Nitex^
prevents escape of mysids and Artemia. Thus, this species can be cultured
continuously for 4-5 months without fluctuations in population density.
STATIC, RECIRCULATING METHOD
Although culture of mysids is more efficient in flowing water, less
maintenance is required for cultures in the re-circulating aquaria. We
maintained two cultures for 13 months in aquaria without changing the water.
This method, now being refined, does not depend on large quantities of
natural or artificial salt water. At least four aquaria are recommended to
ensure sufficient production for continuing experiments. The most critical
step in establishing a viable culture is the conditioning or aging of cul-
ture water.
TJitex is a registered trademark of Tobler, Ernst, and Trabor, Inc., Murray
St., New York, NY. Reference to commercial products does not constitute
endorsement by the Environmental Protection Agency.
59
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Physical System—
A 10-gallon (38-liter) glass aquarium, equipped with a MetaframeR under-
gravel filter and MetaframeR filter light, provides the basic system. Air-
lift tubes are attached to each undergravel filter base to circulate water
in the aquarium. To each air-lift tube, set in its filter base, is attached
to a small glass chamber (vented into the atmosphere) mounted directly above
the aquarium. This device collects air and water exiting from the airlift
tube. Thus, air is vented and water is recycled immediately through a small
glass tube directed toward the aquarium's center. Another option utilizes a
u-tube attached to the air-lift tube to direct water and air downward toward
the aquarium's center. The former design offers the advantage of an uninter-
rupted flow of water recirculated to the culture with minimal splash.
Currents created by resultant water flow are necessary to orient adults
especially during feeding.
Substratum—
Substratum can be either: Coquina sp. shell (mined in Florida for
limited distribution) or PilotR brand crushed oyster shell (pullet size).
For either substratum, 4.5 kilograms (10 Ibs) are required per aquarium.
Culture Water—
If available, natural seawater is recommended; salinity must be adjusted
to 22-26 °/oo with deionized water. The water, after filtered through a 20-
micron filter, is added to the aquarium containing the shell. We substitute
for natural sea water RilaR marine mix salts which are autoclaved as a
precaution against the presence of pathogenic microorganisms. The water in
the aquarium should be aged for at least 2-3 weeks before mysids are intro-
duced.
Water-Conditioning—
Water-conditioning is required, although no explanation can be offered
concerning the changes that occur in quality of water. After adding water
to each aquarium: (1) circulate the water, (2) illuminate the fluorescent
light, and (3) introduce living biological material to facilitate the con-
ditioning process. We shortened this process with these aids: algal mats
from previous cultures; living, unfed Artemia, and mysids, if available.
After mysids release their young and survive for 48 hours, the culture
should be viable.
Food—
Mysids are fed ad libitum 48-hour-old Artemia nauplii. M. bahia is
carnivorous, and, if food is not available, will cannibalize their young.
^etaframe Corporation, Elmwood Park, NJ 07407 and Comptom. CA 90220.
rilot brand is a registered trademark of Oyster Shell Products, subsidiary
of Southern Industries Corp., Mobile, AL; Houston, TX; Baltimore, MD.
is a registered trademark of Rila Products, P.O. Box 114, Teaneck, NJ
60
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F. METHODS FOR ACUTE STATIC TOXICITY TESTS WITH MYSID SHRIMP
(MYSIDOPSIS BAHIA)
Patrick W. Borthwick
INTRODUCTION
f Mysidopsls bahia is a shrimp-like estuarine crustacean that has been
shown to be very sensitive to toxic substances and used successfully in acute
static toxicity tests with complex wastes. M. bahia is recommended as a test
species due to its sensitivity, short life-cycle, small size, and practical
culture methods (Nimmo et al., 1977 and 1978). Results from toxicity tests
with mysids can be used to estimate the impact of ocean-dumped materials on
other salt water crustaceans.
SELECTION OF TEST CONTAINERS
Based on comparable toxicity tests with mysids exposed in different
containers, control survival is best when glass 2-liter CarolinaR dishes are
used instead of 4-liter, wide-mouthed jars. These cylindrical, stackable
culture dishes provide a large surface-to-volume ratio and ample horizontal
space to minimize cannibalism. When filled with test medium to 1 liter,
culture dishes allow easy visual examination of the mysids. Observation is
hampered if the test medium is turbid or dark. Stocking density should not
exceed 10 mysids/liter to insure minimal loading and ease of counting. Cul-
ture dishes may be stacked and placed in a temperature-controlled incubator.
Minimal disturbance and continual lighting help prevent mysids from "jumping
out" of the test medium and "sticking" to the sides of the test container. If
evaporation is evident (especially at high temperatures), distilled or
deionized water should be added to the test medium daily to prevent hyper-
saline conditions.
SALT WATER AND TEST MEDIA PREPARATION
Materials considered for ocean disposal vary in solubility and complex-
ity. Thus, several approaches are necessary for testing various types of
wastes.
Effluents from ocean outfalls may contain a mixture of wastes that are
substantially diluted with fresh water. To achieve a desired salinity in the
test medium without further diluting the effluent, it is necessary to add
dry, autoclaved artificial sea salts to the effluent. This is best accom-
plished by stirring the effluent with a magnetic stirrer while the dry salt
mix dissolves.
R0btained from Carolina Biological Supply Co., Burlington, NC 27215
61
-------
Materials tested for ocean disposal (e.g. liquid, solid, concentrate, or
sludge) should be diluted or suspended with filtered natural sea water that
can be adjusted to a desired salinity by adding artificial salts or deionized
or distilled water.
If soluble, single compounds, or substances of relatively simple or
known composition, should be added directly to the test solution. Insoluble
materials should be tested as a suspension, rather than with a carrier, to
solubilize test material. Filtered natural sea water, if available, should
be used in lieu of artificial sea water, particularly if the mysids were
cultured in natural sea water.
Test media are prepared, stirred to uniformity, and allowed to equili-*
brate to the test temperature for at least 30 minutes before test animals
are introduced.
CARE AND HANDLING
Mysids are easily mishandled; special care in transferring animals from
cultures to culture dishes is mandatory.
Mysids are removed from cultures with a glass tube (300 x 9.0 mm i.d.),
fire-polished at both ends. By placing the index finger over the end of the
empty tube and submersing the tip, a single mysid can be captured gently by
breaking the finger seal. Then the mysid can enter the tube in surrounding
water. Ten newly hatched individuals are assigned randomly to a series of
30 mi glass beakers containing sea water. Sea water volume in each beaker
is reduced to 5 m£, and a beaker containing ten mysids is gently submersed
into each culture dish until the test animals swim into the test medium. It
is often difficult to remove a beaker that has no mysids on the glass sur-
face. Therefore, beakers must be carefully inspected and the number of my-
sids confirmed in each test container. Mysids are handled, observed, and
counted over a waterproof lighted table; sudden movements or disturbances
must be avoided.
SELECTION OF TEST ANIMALS
Newly hatched juvenile mysids (_4-hour-old) are used because of their
uniform size and proven success in toxicity tests. Test results are con-
founded if brooding mysid females release young into the test medium—thus
affecting loading, uptake of toxicant, and competition for food and space.
To obtain juveniles, isolate several brooding females in a large beaker the
day before the test, and harvest the young on the day of the test. Care in
handling is essential to a successful toxicity test.
FEEDING
Mysids have a short lifecycle, and their metabolic demands are high.
They seem to thrive best when fed living 48-hour-old Artemia nauplii. For
96-hour acute static toxicity tests, I recommend that mysids be fed 10 to 20
nauplii per mysid per day to minimize cannibalism. Although it is generally
undesirable to feed most fish and macroinvertebrates during static toxicity
62
-------
tests, an exception is necessary for M. bahia.
EXPERIMENTAL DESIGN
The recommended test procedure for 96-hour acute static toxicity tests
with mysids should include a sea water control, carrier control (if appli-
cable), and at least five concentrations of test media. When materials of
unknown toxicity are tested, a range-finding test may be necessary to approx-
imate the range of concentrations for the definitive test. Two replicate
tests of 10 mysids each are desirable for each concentration in definitive
tests. Animals should be randomly assigned to the culture dish test con-
tainers 30 minutes after 1 liter of medium is added. For additional details,
consult the section, "Static Method for Acute Toxicity Tests Using Fish and
Macroinvertebrates."
OBSERVATIONS
Mysids are observed at 24, 48, 72, and 96 hours to determine the number
of dead or affected individuals. Dead animals should be removed when ob-
served. Observations should note erratic swimming, loss of reflex, molting,
cannibalism, unusual behavior, discoloration, and ability of individuals to
capture live Artemia during feeding.
CALCULATIONS AND REPORTING
For definitive tests, the 96-hour LC50 and 95-percent confidence limits
must be calculated, using Probit Analysis (Finney, 1971). For range-finding
data, the LC50 can be estimated by linear interpolation.
Reports should follow the outline in the section titled, "Static Method
for Acute Toxicity Tests Using Fish and Macroinvertebrates."
REFERENCES
Finney, D.J. 1971. Probit Analysis, 3rd. ed., Cambridge Univ. Press,
London and New York.
Nimmo, D.R., L.H. Bahner, R.A. Rigby, J.M. Sheppard, and A.J. Wilson, Jr.
"Mysidopsis bahia; an Estuarine Species Suitable for Life-Cycle Toxicity
Test to Determine the Effects of a Pollutant," Aquatic Toxicology and
Hazard Evaluation, ASTM STP 634; F.L. Mayer and J.L. Hamelink, Eds.,
American Society for Testing Materials, 1977, pp. 109-116.
Nimmo, D.R., R.A. Rigby, L.H. Bahner, and Jim Sheppard. 1978. Acute and
Chronic Effects of Cadmium on the Estuarine Mysid, Mysidopsis bahia,
Bull. Env. Contain. Toxicol. 19(1) (in press).
63
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G. ENTIRE LIFE CYCLE TOXICITY TEST USING MYSIDS (MYSIDOPSIS BAHIA) IN
FLOWING WATER
D.R. Nimmo, T.L. Hamaker, and C.A. Sommers
INTRODUCTION
The purpose of this method is to determine effects of continuous exposure
of a pollutant on the survival, reproduction, growth, and behavior of this
crustacean through a life cycle. Among the advantages of using this species
in toxicity tests are: (1) ease of culture and maintenance; (2) short gener-
ation time (14-17 days depending on the temperature); and reproduction data
based on actual count (of juveniles) rather than estimates. Further, this
species is representative of many intermediates in estuarine food webs.
Data on toxicity, reproduction, and growth, using a modification of the
procedure described here, have been published (Nimmo et al., 1977).
Mysidopsis bahia is an estuarine species and three reports in the liter-
ature (Molenock, 1969; Odum, 1972; Nimmo et al., 1977) suggest that its
range is from Calveston, Texas to Miami, Florida. We have successfully
captured mysids from small shallow ponds fed by salt water from Santa Rosa
Sound near Pensacola, Florida. A small fish net, used by tropical fish
retailers, or a 3-4 foot push net with very small mesh, is sufficient to
capture the adult mysid shrimp.
PHYSICAL SYSTEMS
Test Water—
1. The source of test water should be a natural water with a salin-
ity jy.5 /oo, although mysids can live at much lower salinities. We have
observed that mysids have survived for 72 hours at 2-3 /oo but reproduction
is affected at 6-8 /oo if maintained for prolonged periods.
2. Sea water must be filtered to remove particles 15u or larger to
remove planktonic larvae that grow, then prey on mysids or their food during
the test.
3. The water source must be analyzed for pollutants such as pesti-
cides, PCB's, and metals. Special determinations should be made for those
chemicals being investigated in the toxicity tests.
Dosing Apparatus—
We suggest that all tests be conducted in intermittent flows from, a
diluter or in continuous flow with the toxicant added by an infusion pump.
Further, we recommend the procedures of Mount and Brungs (1967) or Hansen
64
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et al. (1974) if the toxicant can be added without a solvent; the device
described by Hansen et al. (1974) if a solvent is necessary; or procedure of
Bahner et al. (1975) if pumps are required for continuous flow.
Aquaria—
Glass aquaria 34 x 72 x 18 cm with a water depth of 6 cm are pre-
ferred, but we have used smaller aquaria (12 x 24 x 12 cm) with good results.
When each aquarium receives the maximum volume of salt water from a diluter
or continuous flow apparatus, a self-starting siphon reduces the volume to
about one liter. Therefore, water levels, fluctuating at intervals of about
30 min, ensure an exchange of salt water within each aquarium and the small
chambers devised to retain the mysids.
Retention Chambers—
The chambers consist of a standard, 10-cm glass petri dish (or
cover) to which a 15-cm-high cylinder of NitexR screen (mesh number 210) is
attached by silicone cement.
Test Procedures
Flow Rate of Test Water—
Flow rates to each aquarium should (1) provide 90 percent replacement
in 8-12 hours (Sprague, 1969); (2) maintain dissolved oxygen 60 /o satura-
tion; and (3) maintain the toxicant concentration. Our flow rate is
25&/hour/test aquarium for the continuous-flow system; for the diluter,
6fc/hour/test aquarium. We suggest that dissolved oxygen determinations be
made twice weekly.
Lighting--
Lighting is continuous, using flourescent bulbs.
Temperature—
Test temperatures should be maintained above 20°C, although we have
conducted a test successfully with temperatures above 15°C. We maintain
temperature > 20° and < 30°C by heating or cooling procedures. '
Cleaning and Aeration—
We do not clean any test chambers used to retain the animals during
the test. Instead, we transfer mysids to a pre-cleaned chamber. For aera-
tion, a small stream of compressed air is delivered into each chamber to
safeguard against possible anoxic conditions and to create a current that
apparently aids animal orientation.
%itex is a registered trademark of Tobler, Ernst and Trabor, Inc., Murray
St., New York, NY Reference to commercial products does not constitute
endorsement by the Environmental Protection Agency.
65
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e. Concentrations of Toxicants
(1) When we use a diluter to deliver the toxicants, we use a minimum
of four concentrations of the toxicant and a control with carrier; and a
carrier control, a control without carrier, and four concentrations of toxi-
cant for the continuous flow system.
(2) In many instances, a carrier is necessary to disperse and dis-
solve the toxicant in the test water. Therefore, we employ either acetone
or triethylene glycol (Banner et al., 1975).
(3) Concentrations selected must adversely affect at least one, but
not all life stages of the mysid and this is usually noted as death. Concen-
trations for chronic toxicity tests should be based on results of acute
flow-through toxicity tests. Selection of test concentrations is difficult
because chronic effects on survival, growth, or reproduction of mysids can
occur at concentrations that range from 0.5 to 0.0001 of the 96-hour LC50.
The accuracy of the selection process can be improved by some preliminary
tests such as (a) acute, 96-hour, flow-through tests using different life
stages (e.g. adult, juvenile) and (b) acute tests to determine incipient
LC50 (Sprague, 1969). The highest concentration in life-cycle tests gener-
ally should be the lowest concentration affecting survival or growth in
preliminary tests.
(4) The material and water should be analyzed in this test. Water
from each aquarium should be analyzed at least twice during the 96-hour
test; water samples should also be analyzed from one duplicate weekly during
the life-cycle test. Cost and complexity of analyses, as well as conclusions
and decisions based on test results, should dictate frequency and number of
samples.
Test Procedure—
At the outset, we isolate 250 to 350 gravid female mysids in a 5-
liter glass battery jar and allow an extremely slow flow of salt water
(about 4 drops/second) to drip into the jar. The outflow exit is an auto-
matic siphon, whose inlet is covered by Nitex screen (see retaining chambers).
We maintain a constant supply of Artemia nauplii to mysids for 24 hours and
remove juvenile mysids about every 3-4 hours until their number is sufficient
to begin a test. Thus, we have a synchronous population (within 24 hours of
age) of mysids for our tests. We begin the test with eight retention cham-
bers, five juveniles each or 40 animals per concentration.
Food—
All mysids in the retaining chambers are fed 48-hour-old Artemia
nauplii ad_ libitum daily.
Test Progression and the F^ Generation—
(1) In monitoring daily changes in survival or populations, the
retaining chamber is lifted gently from the aquaria, water is drained from the
66
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Nitex cylinder to the level of the Petri dish, and the chamber is placed on
a lighted counter top. We record live animals by sex, number, females with
or without brood pouches, their young, and any other germane criteria.
Dead mysids are removed.
(2) On days 8-12, mysids are categorized by sex and number per
retention chamber. Juveniles which are usually released beginning on day 14
are counted and removed.
(3) Our life cycle test can be completed in 12 days by maintaining
the temperature at 29°C, but we recommend a test temperature between 22° and
25°C, if possible; otherwise, >_20° and £30°C. We also recommend that testing
continues for 26-28 days. Recommended time and temperature allow the females
tested to complete multiple broods (about three broods with number of juve-
niles varying per brood); thus, number of young per female or the reproduc-
tive success is more easily observed. On the basis of seven toxicants
tested to date, reproductive success (number of juveniles per female) is the
most consistent criterion of sublethal effects; the pesticide Kepone, how-
ever, affected growth of females at a lower concentration.
(4) To test whether the F^ generation is susceptible to the toxi-
cant, we isolate 20 juveniles in separate chambers (5 per chamber) and
follow development of these animals until the onset of reproduction (F2).
Statistical Analysis
The LC501s and the 95% fiducial limits are calculated by linear regres-
sion analysis after probit transformation. We employ Dunnett's "t" test,
comparing mean brood size (number of young per female) in multiple treat-
ments to control. Data such as growth, determined by measurement of length,
may be amenable to analysis of variance, or chi-square tests. We use
« <0.05 as significant difference.
67
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REFERENCES
Bahner, L.H., C.D. Craft, and D.R. Nimmo. 1975. A Saltwater Flow-through
Bioassay Method with Controlled Temperature and Salinity. Progressive
Fish-Culturist 37:126-129.
Hansen, D.J., S.C. Schinmel, and J. Forester. 1974. Aroclor 1254 in Eggs
of Sheepshead Minnows (Cyprinodon variegatus). Effect of Fertilization
Success and Survival of Embryos and Fry. Proc. 27th Ann. Conf. Southeast,
Assoc. Game Fish Comm. Oct. 1973. Hot Springs, Arkansas: 420-426.
Hansen, D.J., P.R. Parrish, J.I. Lowe, A.J. Wilson, Jr., and P.D. Wilson.
1971. Chronic Toxicity, Uptake, and Rentention of AroclorR 1254 in Two
Estuarine Fishes. Bull. Environ. Contarn. Toxicol. 6:113-119.
Molenock, J. 1969. Mysidopsis bahia, New Species of Mysid (Crustacea:
Mysidacea) from Galveston Bay, Texas. Tulane Studies in Zoology and
Botany, 15(3):113-116.
Mount, Donald I., and William Brungs. 1967. A Simplified Dosing Apparatus
for Fish Toxicology Studies. Water Research 2:21-29.
Nimmo, D.R., L.H. Bahner, R.A. Rigby, J.M. Sheppard, and A.J. Wilson, Jr.
1977. Mysidopsis bahia; an Estuarine Species Suitable for Life-cycle
Toxicity Tests to Determine the Effects of a Pollutant. F.L. Mayer and
J.L. Hamelink, Eds., American Society for Testing and Materials STP
634:109-116.
Odum, W.E., and E.J. Heald. 1972. Trophic Analysis of an Estuarine Mangrove
Community. Bull. Mar. Sci. Gulf and Caribbean 22(3):671-738.
Sprague, J.B. 1969. Review Paper: Measurement of Toxicity to Fish. 1.
Bioassay Methods for Acute Toxicity. Water Research 3(11):793-821.
68
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H. CULTURE OF THE GRASS SHRIMP (PALAEMONETES PUGIO) IN THE LABORATORY
Dana Beth Tyler-Schroeder
The grass shrimp, Palaemonetes pugio, a useful organism in assessing tox-
icity of various materials, is (1) easily cultured in the laboratory, (2)
sensitive to toxicants, and (3) can be exposed in flow-through systems
throughout a life cycle. Culture and holding procedures for the grass shrimp
are described below.
INDUCTION OF SPAWNING
Laboratory spawning of P_. pugio was first described by Little (1968).
Deposition of eggs began five to eight weeks after initiation of a photo-
period and temperature regime. Egg production is directly proportional to
rostrum-telson length of females greater than 18 to 20 mm (Jensen, 1958;
Wood, 1967). Shrimp are sexed by examination of the second pleopod (Meehean,
1936), but field data show a 50/50 ratio of sexes (Wood, 1967).
I have found that spawning can be induced at a constant temperature of
25°C, or above, with appropriate increase in photoperiod. A minimum photo-
period of 10 hours light:14 hours darkness per day at 25°C is necessary to
activate ovarian development and spawning. Egg deposition usually follows
within two to four weeks after this regime is established. Continued spawn-
ing of laboratory populations has been observed when the light portion of the
photoperiod is increased by a 47-minute increment every one or two weeks
until the photoperiod is 15 hr 29 min light, and 8 hr 31 min dark.
Ovarian growth and egg deposition can be accomplished with 100-watt,
1750-lumen incandescent light bulbs, as well as fluorescent and growth-light
lamps. However, the latter types stimulate undesirable algal growth in
tanks, a feature to be avoided in toxicity exposures.
Although spawning can be successfully accomplished in non-recirculating
aquaria, the time period can be shortened in flow-through aquaria. A change
of water in static aquaria containing conditioned shrimp is followed by a
burst of egg deposition. Therefore, I postulate that shrimp produce a sub-
stance that inhibits spawning in overcrowded, stagnant conditions. Thus, I
recommend that spawning in static systems be accomplished by (1) using a re-
circulating system with a biological filter, aW/or, (2) changing the water
at least weekly. Shrimp being conditioned for spawning must be fed a daily
diet of freshly hatched Artemia nauplii and a commercial fish flake food
containing vegetable and animal material, but no detectable pollutants.
69
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Laboratory egg production is similar to natural production: each female
deposits from 100 to 500 eggs each spawning and may spawn every four to six
weeks. Incubation of larvae on the pleopods of the female requires from two
to three weeks. Release of larvae frequently is followed by deposition of a
new egg mass.
LARVAE PRODUCTION
To produce a number of larvae for toxicity testing, rearing, etc.,
transfer a number of ovigerous females from the spawning population to a
hatching apparatus. Larvae may be hatched under static conditions as des-
cribed in the section "Static Bioassay Procedure Using Grass Shrimp (Palae-
monetes sp.) Larvae" or in the flow-through system described here.
Shrimp can be cultured in a hatching apparatus (Figure 1) using filtered,
flowing seawater, and two commercially available 37.8-liter (10 gal) aquaria.
SIPHON FROM
HEAD BOX
OVIGEROUS
FEMALES
SCREEN COVERED
OVERFLOW PIPE
LARVAE
CAPTURING
TANK
OVERFLOW DRAIN
NITEX SCREEN
RING
Figure 1-H A flow-through hatching apparatus for grass shrimp larvae
production.
70
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Filtered seawater (20 ym) is introduced into the first aquarium by calibrated
siphon at a flow-rate of approximately 1 liter/hour. The water is heated to
25eC by a small aquarium heater. Approximately 50 to 75 ovigerous females
are placed in the first aquarium. Chelipeds must be removed with a pair of
fine, surgical scissors to reduce removal of the eggs by the females. They
are fed freshly hatched Artemia nauplii daily. The overflow drain (35 mm
diameter) from the first aquarium is covered with a nylon mesh screen
(2,000 ym) to prevent loss of adult females.
Newly hatched grass shrimp larvae pass through the overflow from the
first aquarium containing ovigerous females into a second aquarium fitted
with a special drain to retain the larvae. The drain pipe consists of a
neoprene stopper bored to hold a length of 10- to 12-mm glass tubing. The
length of the glass tubing determines water level in the larval tank. A disc
of plexiglass on which is cemented a collar of nylon mesh screen (363pm) is
fitted on the glass tubing. The nylon collar is of sufficient mesh size to
prevent larvae from being flushed from the tank. This collar must extend one
or two cm above water level to prevent loss of larvae due to unexpected
change of water depth in the larval aquarium.
Larvae are attracted to a lamp placed at one corner of the tank and then
collected daily. They may be removed by a wide-mouth pipette or a small
piece of nylon screen (363 ym). To obtain larvae of uniform age, drain and
flush the larvae capturing aquarium with freshwater, refill with saltwater,
and collect larvae the next day. Uniform age of larvae is frequently an
important consideration in toxicity testing and culture.
REARING TO SEXUAL MATURITY
Grass shrimp larvae can be reared from larvae to adulthood in the labora-
tory. Newly hatched larvae can be reared in 90-liter (23.7 gal) aquaria (83
cm long x 41 cm wide x 35.5 cm deep). Water depth can be maintained at 30 to
32 cm by using a nylon mesh covered drain as described for the larval aquar-
ium of the hatching apparatus. Temperature-salinity optima for P_. pugjp
larvae, 25°C-25 /oo (Floyd, 1977), should be maintained. ~
We have had success with stocking densities from 20 to 33 larvae/liter
and water flow rates of 1 liter/hour, using calibrated siphons and constant
head boxes. A slow flow rate through the aquaria allows a slower turnover of
food organisms (freshly hatched Artemia nauplii), thus enhancing survival of
the grass shrimp larvae. Maintain dissolved oxygen by use of a small aqua-
rium air pump and one or two airstones.
Metamorphosis to postlarvae occurs within 12 to 35 days after hatching;
thereafter, it is usually advisable to remove postlarvae to less crowded
conditions, i.e., a larger aquarium (51 cm long x 67.5 cm wide x 24 cm
deep). Grass shrimp juveniles can be reared to adulthood on a diet of live
Artemia nauplii, frozen adult Artemia, or fish flake food. Sexual maturity
Should be achieved in one to two months after larvae hatch.
71
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REFERENCES
Floyd, W.R. 1977. The Effects of Temperature and Salinity on the Larval
Development of the Grass Shrimp, Palaemonetes pugio Holthuis, Reared in
the Laboratory. Master's Thesis,' Old Dominion University, Dept. of
Oceanography, Norfolk, VA. 145 p.
Jensen, Jens Peder. 1958. The Relation Between Body Size and Number of
Eggs in Marine Malacostrakes. Meddelelser Fra Danmarks Fiskeri-Og
Havundersogelser. 2(19):1-25.
Little, Georgiandra. 1968. Induced Winter Breeding and Larval Development
in the Shrimp, Palaemonetes pugio Holthuis (Caridea, Palamonidae).
Studies on Decapod Larval Development, Supplement 2. Crustaceana: 19-26.
Meehean, 0. Lloyd. 1936. Notes on the Freshwater Shrimp, Palaemonetes
paludosa (Gibbes). Trans. Amer. Micros. Soc. 55:433-441.
Wood, Carl E. 1967. Physioecology of the Grass Shrimp, Palaemonetes pugio.
in the Calveston Bay Estuarine System. Contr. Mar. Sci. Univ. Tex.
12:54-79.
72
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I. STATIC BIOASdAY PROCEDURE USING GRASS SHRIMP (PALAEMONETES SP.) LARVAE
D. B. Tyler-Schroeder
INTRODUCTION
Procedures for static 96-hour bioassays utilizing grass .shrimp larvae,
Palaemonetes sp., are outlined here. The grass shrimp is an obvious bioassay
choice for several reasons. Three species of the genus, P_. pugio, vulgaris,
and intermedius, are common inhabitants of estuaries along the Gulf and
Atlantic coasts of the United States (Holthuis, 1949, 1952). They are easy
to collect and maintain in the laboratory. Field populations are usually
quite large, allowing greater numbers to be brought into the laboratory for
testing. By manipulating environmental conditions of temperature and photo-
period, it has been possible to induce spawning in the laboratory (Little,
1967), opening the way to laboratory cultures of genetic uniformity. Develop-
ing larvae are also available throughout the year for testing with these
methods.
Larval stages of the three species are hardy and easy to culture in the
laboratory. Developmental stages have been described for all species (Broad,
I957a, b; Broad and Hubschman, 1962; Hubschman and Broad, 1974), and salin-
ity-temperature optima are known for the larval development of P_. vulgaris
(Sandifer, 1973). Developing larvae have demonstrated a susceptibility to
polychlorinated hydrocarbons greater than that demonstrated by adults or
juveniles (Tyler-Schroeder, unpublished manuscript).
CULTURE METHODS
Palaemonetes sp. are easily collected from the field with dip nets or
seines in grassy, shallow estuarine areas. They can also be reared in
enclosed holding ponds.
B
To obtain larvae, 8" glass culture bowls, such as the Carolina culture
dish, containing 1& of filtered sea water are stocked with 3 ovigerous female
shrimp per bowl. In order to produce enough shrimp larvae for a 96-hour test
series (210 per replicate, 630 per test series), at least 17-25 bowls of
ovigerous females (51-75 shrimp) must be maintained continuously in the
laboratory (Figures 1-1 and 2-1). The species of each female is
RCarolina Biological Supply Company, Burlington, North Carolina 27215.
.Mention of commercial products or trade names does not constitute endorsement
by the Environmental Protection Agency.
73
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ConcentrationCs) Control 0.01
(mg/liter - ppm)
Number larvae per
test container 30 30
Total 150 larvae
Replicate
0.1 1.0
30 30
10
30
1st Day larvae
3 replicates
Larval age and
Number of replicates
18 Day larvae
3 replicates
Total number of
larvae
(150 larvae/replicates) X (3 replicates/test) - 450 larvae
(450 larvae/test) X (2 test ages) = 900 larvae
Total = 900 larvae
Test Series
Example mortality:
ppm
0
Control
3
0.01
10
0.1
80
1.0
97
10.0
Estimated LC50 between 0.1 and 1.0 ppm
Figure 1-1. Example of a range-finding bioassay.
-------
Concentrations (ppm)
(chosen from
range-finding
tests, Figure 1-E)
Control
0.1
0.159
0.252
0.399
0.631
1.0
Number of
larvae per
test concentration
30
30
30
30
30
Total number
Larvae
(210 larvae/replicate) X (3 replicates/test) - 630 larvae
(630 larvae) X (3 test ages) » 1890 larvae
Total m 1890 larvae
Test Series
30
30
Larval age
and
number of
replicates
1st Day larvae
3 replicates
18th Day postlarvae
3 replicates
Figure 2-1. Example of a definitive bioassay.
-------
confirmed and the chelipeds removed with fine surgical scissors to prevent
removal of the eggs by the females. Shrimp in culture bowls are fed Artemia
nauplii daily, and water is changed if a slight cloudiness appears. Since
eggs are carried for 2-3 weeks before hatching, it is advisable to select
females with eggs in the more advanced stages of development.
Larvae are removed from bowls containing ovigerous females each morning
and mixed together to insure uniformity of test animals. They are randomly
dispensed into 8" glass culture bowls containing 1& of filtered sea water
(200 larvae/*), fed Artemia nauplii, and reared to the desired test age.
Food is added daily and water changed when a slight cloudiness appears.
There should always be sufficient live food in rearing and test chambers,
since insufficient food accentuates developmental variability (Broad, 1957b)
and produces undesirable variation in test results.
A 10 to 15% mortality must be anticipated in calculating the number of
test larvae that must be reared to a predetermined age. Ideally, the larvae
to be used in a series of 96-hour acute tests should be hatched at one time
and reared in mass culture. Samples of larvae would be removed from this
culture at designated times for testing. This technique would minimize or
circumvent problems due to possible seasonal variation in larval suscepti-
bility to waste material.
Salinity-temperature optima for £. vulgaris larvae indicate a broad
range of tolerance to environmental conditions, which is most likely true
for P_. pugio and £. internedius. Survival of !P. pugio is approximately the
same when reared in the laboratory at a temperature of 25 C and salinities
of from 15-25 loo (A.N. Sastry, personal communication*). Bioassays
should be performed within this range, preferably closer to 15 /oo salinity,
as P^. pugio taken from the field are most commonly found in this salinity,
or lower.
Preparation of Test Media, Selection of Test Containers
The nature of the material to be tested indicates choice of test con-
tainer size and shape, preparation of test concentrations, and frequency of
test media replacements. Problems posed by various wastes include insolu-
bility in sea water, adsorption to exposed surfaces, decomposition by
hydrolysis, photolysis, etc., loss by volatilization, high BOD, and bacterial
growth. Such problems can affect results' by causing variation from the
calculated concentration of waste being tested, changing pH of test medium,
releasing breakdown products which may be more or less toxic than parent
compounds, and causing test animal mortality not related to direct effect of
toxicants. Glass is the recommended material for test containers.
>ft.N. Sastry, Graduate School of Oceanography, University of Rhode Island,
Kingston, Rhode Island 02881
76
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When choosing container size, it is important to choose a small vessel
surface area to volume ratio because of possibility of adsorption. A
larger volume is also important because of stocking density requirements.
The 8" diameter, Carolina culture dish, containing IX, of media, has been
found to be a satisfactory test container for bioassay of Palaemonetes
larvae, allowing maximum volume per vessel surface area and an acceptable
stocking density of 30 larvae/H.
Test media should be prepared fresh at the time of replacement, so that
decomposition of toxicant, adsorption to preparation containers, depletion
of oxygen, and bacterial growth are minimal. Likewise, it is necessary to
change solutions in test containers at least every 24 hours, preferably
every 12 hours.
All sea water to be used should be of natural origin, preferably from
the dumping site. It should be filtered through a filter of ly porosity. To
adjust salinity the addition of either distilled water or a high-salinity
brine is necessary. The high-salinity brine may be of natural or artificial
origin. If natural origin is desired, place a closed container one-half to
three-quarters full of filtered sea water (>30 loo salinity) in a freezer
until solid throughout, usually 2-3 days. Subsequent to removal from the
freezer, the supernatant is drained after the first thaw (2-3 hours).
Supernatant should be 80-110 loo salinity or above and can be stored indefin-
itely.
An artificial brine may be made with any of the commercial artificial
sea salts and distilled water, but should be used with caution. Several of
these preparations contain one or more chelator substances, e.g., EDTA,
which would bias test results with waste material containing heavy metals.
The use of artificial sea water totally in place of natural sea water is not
recommended at this time. In addition to various chelators in commercial
preparation, the presence of high levels of contaminant heavy metals in
artificial or laboratory prepared sea salt mixes should be checked. Several
shelf chemicals are known to have background levels of Cu, for example, as
high as 5-10 ppb (yg/Jl) (Erickson et al., 1970; J.H. Gentile, personal com-
munication*) . Unwanted trace metals can be removed by passing the sea water
through a column containing a deionizing resin (Davey et al., 1970), but
this method may not be practical for large volumes of water.
Many effluents to be tested are complex mixtures having both solid and
liquid components. There may also be gaseous components. The following
guidelines should be followed when preparing test media:
*J.H. Gentile, National Marine Water Quality Laboratory, South Ferry Road,
Narragansett, Rhode Island 02882
77
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If Liquid Only--
Waste material should be stirred or shaken thoroughly before use.
Waste material may be used directly or as in a stock prepared by dilution
with filtered sea water to a desired concentration. All stocks and test
concentrations should be prepared on a weight-to-volume basis (gm/Jl, mg/Jl,
yg/£). If volume/volume basis is used, a correction should be made for
specific gravity of the material being tested, i.e., (weight/volume)
(specific gravity) = volume/volume.
If Solid and Liquid--
It may be desirable to test the effects of solids on Palaemonetes^
larvae. Solids can be added to the test containers by weight and agitated
to keep them in suspension; combined toxic-mechanical effects then are
determined. Alternately, one volume of solid material may be diluted with
four volumes of sea water to prepare a standard elutriate.
All test glassware should be thoroughly washed, using the following
procedure:
1) Empty old test solution and rinse with cold water.
2) Rinse with acetone, followed by a warm water rinse.
3) Wash with laboratory soap and a brush. Rinse thoroughly with
warm water 4-5 times.
4) Rinse with 10% HC1 or HCO_, if the toxicant contains heavy metals,
5) Rinse with distilled water and allow to dry. If an acid rinse is
used, it should be followed by 4-5 thorough rinses with deionized water.
Bioassay Procedures
Because Palaemonetes normally exhibits variability in molting and
developmental rates during larval life, it is not feasible to produce
sufficient larvae of individual stages for testing. Therefore, tests use
larvae of specified ages (e.g., ages 1 and 18 days). Most larvae will
metamorphose to postlarvae (PL) on approximately day 18-21. Hence, one
bioassay is performed on 18-day-old larvae, and one on postlarvae, to deter-
mine if the biochemical and physiological changes accompanying metamorphosis
alter the response to the toxicant. For the same reasons, a bioassay using
day 30 postlarvae may be required.
Palaemonetes larvae are added to test containers, using a method of
random selection (total randomization, stratified randomization, etc.).
Larvae are removed from culture dishes, using a rectangular piece of fine
mesh Nitex nylon net, and stocked in test dishes at a density of 30 larvae/
liter of test media/culture dish. Larvae are fed an excess supply of
Artemia nauplii throughout the test. Artemia are added with each change
78
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of test media. Mortalities of Palaemonetes are recorded at the time of each
test media change (every 12 or 24 hours), and all dead animals removed at
this time. All test and control culture dishes should be maintained at 25°C
in a culture cabinet, BOD incubator, or water table. Tests may be run in
total darkness or on a 12-hr light - 12 hr-dark regime. All tests should
include 4-6 concentrations and a sea water control. Control mortality
exceeding 10% invalidates test results. Because of the inherent variability
of each age group of larvae, 2 to 3 replicates must be run simultaneously
for each test concentration in each experiment. These basic test conditions
are to be followed for both range-finding and definitive bioassays, as
discussed below. (See Figures 1-1 and 2-1.)
Initially, a series of range-finding 96-hour assays are performed, using
1- and 18-day-old larvae to determine the range of toxicity of the material
being examined, and to determine the best test conditions. A broad range of
concentrations covering at least four orders of magnitude should be tested;
e.g., 0.01, 0.1, 1.0, and 10 mg/Jl (ppm), or gm/Z. (%). Temperature, pH, and
dissolved oxygen (DO) levels should be monitored throughout these tests to
help determine need for aeration and frequency of test solution change.
After the range-finding test is completed, a LC50, concentration lethal
to 50% of the shrimp, is approximated and a series of definitive bioassays
performed. The purpose of the definitive bioassay is to more clearly deter-
mine the limits of toxicity of a waste and better estimate the LC50. Concen-
trations chosen for definitive bioassays are determined by results from the
range-finding tests, i.e., the lowest definitive test concentration should
equal or be greater than the greatest concentration in range-finding tests
that killed few or no test organisms. Likewise, the greatest definitive
test concentration should be equal to or less than the least concentration
in range-finding tests that killed all or almost all test organisms. (See
Figures 1-1 and 2-1.) Once upper and lower definitive test concentrations
are chosen, intermediate concentrations are calculated, using progressive
bisection of intervals on a logarithmic scale (Standard Methods, 1965). At
least five, and preferably more, test concentrations are used to yield mor-
tality data on either side of a 50% kill, a condition necessary for statis-
tical treatment of data using Probit Analysis.
Growth is often a more sensitive indication of effect than mortality and
is useful in choosing concentrations to be used for chronic tests. There-
fore, at the end of each test, the rostrum-telson length of surviving
larvae from each test concentration and controls should be measured with an
ocular micrometer. A sample of untreated larvae should be measured at the
beginning of the test for comparative purposes. Additional observations,
such as loss of equilibrium, cessation of feeding, irregular movements, and
other behavioral aberrations, should be noted at the time of each test media
change.
79
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Analysis of Data
Data from 96-hour acute bioassays should be analyzed by Probit Analysis
(Finney, 1964a, b). This method estimates a value for LC30, 70, and 90, as
well as the LC50. Because Probit Analysis is generally performed by com-
puter, it is wise to check the computer output by plotting percentage kill
in probits against logarithm of concentration, and comparing computer and
graphed LC50s. The line thus plotted should closely resemble that deter-
mined by the computer. The Litchfield-Wilcoxon method of LC50 estimation or
graphical interpolation, using Probit graph paper (Standard Methods, 1965),
can be used when data do not meet the more rigorous specifications required
by Probit Analysis (Litchfield, 1949; Litchfield and Wilcoxon, 1949, 1953).
The 95% confidence limits should be indicated for all data.
Reports
At the completion of testing and data analysis, a report is usually
required. Such reports should include the following information:
1. Name of method, investigator, laboratory, and date tests were con-
ducted.
2. Detailed description of material tested, source, date, and time of
collection, composition, known physical and chemical properties.
3. Source of sea water, date, and method or preparation.
4. Detailed information about test animals, including' scientific name,
life stage, age, source, history, and acclimation procedure for larvae, if
appropriate.
5. Experimental design, test containers, volume of test solution,
initial test conditions', number of organisms at each concentration, number
of organisms in each control, and types of controls run.
6. Definitions of response used to determine the effect under investi-
gation and a summary of general observations of related effects or symptoms.
7. Percentage of control organisms that died or were affected during
the test.
8. LC50, with confidence limits. LC30, 70, and 90, if pertinent.
9. Methods used for and results of all DO, pH, and temperature measure-
ments.
10. Any deviations and reasons.
11. Other relevant information.
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REFERENCES
Broad, A. Carter. 1955. Reproduction, Larval Development and Metamorphosis
of Some Natantia from Beaufort, N.C. Ph.D. Thesis. Duke University.
87 pp.
. 1957a. Larval Development of Palaemonetes pugio Holthuis. Biol.
Bull. 112:144-161.
. 1957b. The Relationship Between Diet and Larval Development of
Palaemonetes. Biol. Bull. 112:162-170.
and Jerry H. Hubschman. 1962. A Comparison of Larvae and Larval
Development of Species of Eastern U.S. Palaemonetes With Special
Reference to the Development of Palaemonetes intermedius Holthuis.
Am. Zool. 2(3):172 (Abstr.).
Davey, E.W., J.H. Gentile, S.J. Erickson and P. Betzer. 1970. Removal of
Trace Metals from Marine Culture Medium. Limnol. Oceanog. 15(3):333-
490.
Erickson, S.J., N. Lackie and T.E. Maloney. 1970. A Screening Technique
for Estimating Copper Toxicity to Estuarine Phytoplankton. J. Water
Pollut. Control Fed. 42:R270-R278.
Federal Register, Part II. 1973. U.S. Environmental Protection Agency—
Ocean Dumping Criteria, May 16, 1973. 38(94):12874.
Finney, D.J. 1964a. Probit Analysis: A Statistical Treatment of the
Sigmoid Response Curve. Cambridge at the University Press, Cambridge.
318 pp.
. 1964b. Statistical Method in Biological Assay. 2nd Ed. Hafner,
N.Y. 668 pp.
Holthuis, L.B. 1949. Notes on the Species of Palaemonetes (Crustacea,
Decapoda) Found in the United States of America. K. Ned. Akad. v.
Wet. 52:87-95.
. 1952. A General Revision of the Palaemonidae (Crustacea, Decapoda,
Natantia) of the Americas. II. The Subfamily of Palaemoninae. Occas.
Pap. Allen Hancock Found. 12:1-369.
81
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Hubschman, J.H. and A.C. Broad. 1974. The Larval Development of
Palaemonetes intermedius Holthuis, 1949 (Decapoda, Palaemonidae) Reared
In the Laboratory. Crustaceana 26(1):89-103.
Litchfield, J.T., Jr. 1949. A Method For Rapid Graphic Solution of Time-
percent Effect Curves. J. Pharmacol. Exp. Ther. 97:399-408.
and F. Wilcoxon. 1949. A Simplified Method of Evaluating Dose-effect
Experiments. J. Pharmacol. Exp. Ther. 96:99-113.
. 1953. The Reliability of Graphic Estimates of Relative Potency from
Dose-percent Effect Curves. J. Pharmacol. Exp. Ther. 108:18-25.
Little, Georgiandra. 1968. Induced Winter Breeding and Larval Development
in the Shrimp, Palaemonetes pugio Holthuis (Caridea, Palaemonidae).
Crustaceana, Supplement 2: Studies on Decapod Larval Development. 19-
26 pp.
Sandifer, Paul A. 1973. Effects of Temperature and Salinity on Larval
Development of Grass Shrimp, Palaemonetes vulgaris (Decapoda, Caridea).
U.S. Fish. Wildlf. Serv. Fish Bull. 71(1):115-123.
Standard Methods for the Examination of Water and Wastewater. 12th Ed.
1965. American Public Health Association, Inc. New York, N.Y. 769 pp.
82
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J. ENTIRE LIFE-CYCLE TOXICITY TEST USING GRASS SHRIMP (PALAEMONETES PUGIO
HOLTHUIS)
Dana Beth Tyler-Schroeder
INTRODUCTION
The purpose of this method is to assess toxicity of a material to all
life stages of the grass shrimp in flow-through systems. This experiment
determines effects on survival, growth, and reproduction (including number
of females spawning, number of days before onset of spawning, number of eggs
per female, and hatching success) of parental generation shrimp. Effects on
survival, larval development, and growth are also determined for F^ genera-
tion shrimp. These tests must extend through an entire life-cycle of the
shrimp—from juvenile stage of the parental generation, sexual maturation
and reproduction, through hatching, larval development, and growth of the F^
generation to juvenile stage. Tests may terminate at this point, or expo-
sures can be continued if necessary to determine effect on F^ reproduction
and F£ larval development.
Basic methodology for flow-through toxicity tests has been described in
the section, "Entire Life-Cycle Toxicity Test Using Sheepshead Minnows
(Cyprinodon variegatus)" (Hansen et al., this publication) and by the Com-
mittee on Methods for Toxicity Tests with Aquatic Organisms (1975). The
following procedures describe only aspects unique to toxicity testing with
the grass shrimp.
PHYSICAL SYSTEMS
Salinity of test water should be 20 /oo, to be near the salinity optima
for larval development (25 /oo) (Floyd, 1977). Test water temperatures are
controlled to 25°C ± 1°C. Photoperiod regimes for life-cycle toxicity tests
vary and are discussed here and in the section describing laboratory culti-
vation of grass shrimp.
A Mount and Brungs diluter (1967) with appropriate modifications has
been successfully employed in life-cycle toxicity tests. The diluter should
deliver one-half liter to each exposure aquaria for each cycle. If acetone,
triethylene glycol, ethanol, or other solvents are used as carriers, the
diluter must be modified to provide equal carrier concentrations in all
exposure water. A carrier control and a control without carrier must be
provided (Schimmel et al., 1974).
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Exposure aquaria, 1/4-inch plate or 1/8-inch double strength glass, are
constructed with a clear, silicone rubber sealant. All materials in expo-
sure aquaria or equipment must be glass, or another inert material, such as
nylon, teflon, etc.
Test aquaria of 40 cm x 61 cm x 22 cm deep with a water depth of 17 cm
were successfully utilized in a life-cycle toxicity test with grass shrimp
exposed to the chlorinated hydrocarbon pesticide, endrin (Tyler-Schroeder,
in press). The drain is constructed to resemble that in the tank used to
capture larvae. (See "Culture of Grass Shrimp (Palaemonetes pugio) in the
Laboratory.") The plexiglas disc is of approximately 5-cm diameter, and the
drain pipe must bend at a 90° angle to drain through the front rather, than
the bottom of the tank. The nylon screen collar is constructed of 363 ym
mesh and must extend one to two centimeters above water level.
BIOLOGICAL ASPECTS
Grass shrimp for toxicity testing may be obtained from natural or labor-
atory populations. Test animals must be uncontaminated, i.e., whole body
residue analyses must be free of unacceptable concentrations of pesticides,
PCB's, heavy metals, or other pollutants of concern.
Juvenile grass shrimp are acclimated in the laboratory under conditions
described for the life-cycle toxicity test. Mortality can be no greater
than two percent during the 4-day acclimation period. Near darkness (15-
watt bulb turned on only for feeding, daily cleaning, and observations) is
maintained during acclimation to inhibit premature reproduction.
Adult and juvenile shrimp are fed fish flakes (such as MetaframeR
HiProMin Tropical Flakes or TetraMinR) that contain both plant and animal
material during acclimation and testing. A supplement of frozen adult brine
shrimp or newly hatched brine shrimp nauplii is added during growth until
sexual maturity and induction of spawning. Grass shrimp larvae must be fed
newly hatched brine shrimp nauplii several times a day. At the time of
metamorphosis to postlarvae, the diet may be changed to the fish flake.
Foods must not be contaminated with pesticides, PCB's, heavy metals, or
other pollutants of concern.
TEST REGIME
Initial Stage
Start the life-cycle toxicity test with 100 juvenile shrimp (less than
15 mm rostrum-telson length) randomly distributed in each test concentration.
Gonad development must not be obvious at the start of the test. Exposure
aquaria are examined daily to count and remove dead shrimp. Daily counts of
surviving shrimp are impractical; however, individual shrimp are to be
counted every four to six weeks. Lengths of 30 shrimp per concentration
are measured at the start of the test and every four weeks until termination
to determine effects on growth. All shrimp lengths discussed here refer to
rostrum-telson length. (The shrimp is extended to its full length and mea-
sured from the tip of the rostrum to the end of the uropods of the telson).
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During the time required for pollutant uptake and 'growth of shrimp to
sexual maturity Cusually two to three weeks), the photoperiod is held con-
stant at 8L:16D, using 15-watt, 125-lumen incandescent bulbs to prevent pre-
mature induction of gonad development and spawning. The length of this
initial exposure is based on the time necessary for the pollutant to reach
equilibrium between shrimp tissue and the concentration in water. Prelimi-
nary bioconcentration exposures are conducted according to methods pre-
scribed by Hamelink (1977).
Induction of Spawning
After approximately a two- to ,three-week exposure, shrimp should average
20 to 25 mm in length. Spawning is induced, using 100-watt, 1750-lumen
incandescent lamps and 10L:14D photoperiod. Thereafter, photoperiod is
increased by 47-minute increments every two weeks to a maximum 15-hr, 29-min
light:8-hr, 31-min darkness.
When production of ovigerous females is first noticed, a partition of
2.0 mm nylon mesh is installed in the tank at a distance of 11 cm from the
front. Ovigerous females are separated from the rest of the population to
allow an accurate count of number of females spawning per day.
i
Effects on Reproductive Success
Number of ovigerous females produced in each exposure concentration must
be recorded daily. Eggs from a£ least 10 females per concentration must be
counted. Rostrum-telson length 'of each female is recorded with the respec-
tive egg count because egg production is proportional to length. This
recorded length must be used as a covariate in statistical analyses.
Effects on hatching success of embryos is determined in a hatching
apparatus and larval rearing tray that are installed in each aquarium once
females are ovigerous and eggs have darkened. Each hatching apparatus con-
sists of a 6.35-mm, I.D. tubing manifold that delivers water to five glass
spawning chambers. A spawning chamber is constructed of a 30-mm diameter
glass tube, 8.5-mm long fitted with neoprene stoppers and input and output
tubing on either end (Tyler-Schroeder, in press). One ovigerous female is
placed in each spawning chamber to facilitate individual egg counts. Water
from the diluter flushes newly hatched larvae from each chamber into an egg
cup constructed of a 100-mm diameter Petri dish top fitted with a nylon mesh
collar (363ym) 13 mm high. Larvae hatched from at least 10 females in each
exposure concentration are counted and compared to egg counts to estimate
hatching success.
The egg cups are held in larval hatching and rearing trays placed in
each aquarium during reproductive and larval development phases of the test.
Each tray consists of an elevated tank supported by rectangular glass
legs on two ends; the glass legs form each end of the tank. Hatching trays,
34 cm x 43 cm x 13 cm high, are constructed of 1/8-inch double-strength
glass. Water depth is controlled by a self-starting siphon, causing the
water level to fluctuate from 8 to 11.5 cm. Thus, water from the diluter
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flows in and out of egg cups.
Larval Development and Metamorphosis
After recording hatching success, the hatching apparatus is removed from
the larval-rearing tray. Larvae hatched from several females are placed in
larger diameter egg cups (150-mm Petri dish tops, with 363 ym nylon mesh
collar 15 mm high) to observe effects on larval development (20 larvae/egg
cup; four to five egg cups per concentration). Larvae are counted daily to
record effects on survival, length of larval development, and metamorphosis.
Larvae metamorphose to postlarvae within 12 to 20 days of hatching.
Effects on Growth
Thirty-five days after hatching, the rostrum-telson length of all post-
larvae are measured to determine pollutant effects on growth. Postlarvae
are subsequently released from the egg cups into the larval-rearing tray.
The self-starting siphon is replaced with a screened drain to prevent escape
of small shrimp from the hatching and rearing tray. Lengths of at least 30
shrimp per concentration are recorded at weekly intervals to evaluate effect on
growth.
Additional Tests
In order to develop application factors used in setting water quality
criteria, it is necessary to determine "effect" and "no-effect" concentra-
tions in a life-cycle toxicity test and compare these to the 96-hr LC50 for
juvenile shrimp determined in an acute toxicity test (Eaton, 1973). Methods
for acute toxicity tests in flow-through systems have been described by the
Committee on Methods for Toxicity Tests with Aquatic Organisms (1975) and by
other authors in this manual. These methods should be consulted and a 96-
hour LC50 obtained.
Test Termination
Termination of a life-cycle toxicity test depends upon nature of the
pollutant being tested, its intended use and disposal pattern, and the kind
of information desired. Technically, if the test began with 15-mm juve-
niles, a full life-cycle has been completed by the time FI larvae metamor-
phose to postlarvae and grow to 15 mm long juveniles. Some chemicals are
particularly persistent in the environment or are released into the environ-
ment in consistent amounts over a long period of time. When testing such
pollutants, it may be desirable to continue the life-cycle toxicity test
exposures until effects on FI reproduction and F£ hatching success, larval
development, and growth can be assessed. In either case, the test is termin-
ated after shrimp of the youngest desired generation (F^ and F£) have com-
pleted larval development, metamorphosed to postlarvae, and grown to juve-
niles approximately 15 mm long.
86
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At test termination, all surviving shrimp (parental generation shrimp
that spawned, parental generation shrimp that did not spawn, F! generation
shrimp) are individually measured (rostrum-telson length). Shrimp from each
test concentration are grouped as above (parental shrimp that spawned,
parental shrimp that did not spawn, FI generation shrimp) and a composite
weight recorded for each grouping. The number of shrimp in each group is
recorded to calculate individual weight per shrimp in each group or to
calculate average weight of individual shrimp in each test concentration.
Residue analyses of whole body shrimp are made with the same groupings per
test concentration as above. Shrimp are quickly killed by brief exposure to
a stream of steaming hot water.
A final count of surviving shrimp is made for each test concentration
(using the same groupings as above) to determine effects on survival.
STATISTICAL ANALYSES
Data from the life-cycle toxicity test are analyzed by analysis of vari-
ance and multiple comparison methods to determine differences between means.
All data compared as percentages (e.g., percent survival, percent metamorpho-
sis, etc.) should be transformed, using the arc sine transformation (Winer,
1971).
The data related to effects on egg production and larval hatching are
compared by analysis of covariance, the covariate being the rostrum-telson
length of the female from which eggs or hatched larvae were produced.
Appropriate multiple comparison methods are used to determine differences
among means.
The data from the juvenile 96-hour acute toxicity test are analyzed by
linear regression after probit transformation to determine LCSO's and 95
percent confidence limits. Uptake and depuration rates from bioconcen-
tration studies are calculated by the nonlinear statistical model of Bahner
and Oglesby (in press).
87
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REFERENCES
Bahner, L.H., and J.L. Oglesby. (In Press). Test of Model for Predicting
Kepone Accumulation in Selected Estuarine Species. American Society for
Testing and Materials.
Committee on Methods for Toxicity Tests with Aquatic Organisms. 1975.
Methods for Acute Toxicity Tests with Fish, Macroinvertebrates, and
Amphibians. EPA Report No. EPA-660/3-75-009. U.S. Environmental Pro-
tection Agency, Washington, DC. pp. 61.
Eaton, J.G. 1973. Recent Developments in the Use of Laboratory Bioassays
to Determine "Safe" Levels of Toxicant. G.E. Glass (ed.) Ann Arbor
Science Publishers, Inc., Ann Arbor, MI. 1Q7-115.
Floyd, W.R. 1977. The Effects of Temperature and Salinity on the Larval
Development of the Grass Shrimp, Palaemonetes pugio Holthuis, Reared
in the Laboratory. Master's Thesis, Old Dominion University, Dept. of
Oceanography, Norfolk, VA. 145 pp.
Hamelink, J.L. 1977. Current Bioconcentration Test Methods and Theory in
Aquatic Toxicology and Hazard Evaluation, ASTM STP 634, F.L. Mayer and
J.L. Hamelink, eds., American Society for Testing and Materials,
pp. 149-161.
Mount, Donald I., and William Brungs. 1967. A Simplified Dosing Apparatus
for Fish Toxicology Studies. Water Res. 2: 21-29.
Schimmel, Steven C., David J. Hansen, and Jerrold Forester. 1974. Effects
of Aroclor^ 1254 on Laboratory-Reared Embryos and Fry of Sheepshead
Minnows (Cyprinodon variegatus). Trans. Am. Fish. Soc. 103(3): 582-586.
Tyler-Schroeder, D.B. (In Press). Use of the Grass Shrimp, Palaemonetes
pugio, in a Life-Cycle Toxicity Test. American Society for Testing and
Materials.
Winer, B.J. 1971. Statistical Principles in Experimental Design. McGraw-
Hill Book Company. New York, NY. 907 pp.
88
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K. STATIC METHOD FOR ACUTE TOXICITY TESTS USING FISH AND MACROINVERTEBRATES
(See list of contributors.)
EQUIPMENT
Facilities
For maximum convenience and versatility, the facilities should
include tanks or cages to hold and acclimate test animals, a tank for salt
water, a temperature-controlled recirculating water bath, or an environ-
mentally controlled room for the test containers. The holding and acclima-
tion tanks should be equipped for temperature control, and the holding tank
should be equipped for aeration. Because air used for aeration must not
contain oil or fumes, it must be taken from a well-ventilated, fume-free
area, and powered by a surface aerator or an oil-free rotary or piston-type
air compressor. During holding, acclimation, and testing, test animals
should be shielded from disturbances.
Construction Materials
Construction materials and commercial equipment that might contact
water in which test animals will be placed should not contain any substances
that can be leached or dissolved by the water. In addition, materials and
equipment should be chosen to minimize sorption of toxicants from water. It
is suggested that glass, #316 stainless steel, or perfluorocarbon plastics
be used when possible.
Test Containers
1. Type: The test solution for fish and invertebrates should be
placed in containers measuring between 15 and 20 centimeters (cm) deep.
These animals can be tested in 19.6£ (5-gallon) wide-mouthed soft glass
bottles containing 15I of solution. Alternatively, test containers can be
made by welding (not soldering) stainless steel or by gluing double-strength
window glass with clear silicone adhesive. As little adhesive as possible
should be in contact with the water; extra beads of adhesive should be on
the outside of the containers rather than on the inside. Some invertebrates
can be tested in 3.9£ (1-gallon) wide-mouthed soft glass bottles or battery
jars containing 3£ of solution.
2. Cleaning: Test containers must be cleaned before use. New
containers must be washed with detergent and rinsed with 10% hydrochloric
acid, acetone, and tap or other clean water. The containers, if reused
after a test, should be (1) emptied; (2) rinsed with water; (3) cleaned by
an appropriate procedure to remove the test toxicant, e.g., acid to remove
metals and bases; detergent, organic solvent, or activated charcoal to
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remove organic compounds; and (4) rinsed with water. Acid is useful for
removing scale, and hypochlorite (bleach) is useful in removing organic
matter and as a disinfectant. All test containers must be rinsed with salt
water immediately before use.
Salt Water
Acute toxicity tests require salt water in which healthy animals can
survive throughout acclimation and testing without sign of stress, such as
unusual behavior or discoloration. Salt water is prepared from commercially
available formulations or from ingredients listed in Table 1-K, using deion-
ized or glass-distilled water. Deionized or distilled water is used to
dilute the salt water to a salinity of 30 parts per thousand ( /oo). Natural
salt water that satisfies the acclimation requirement also can be used.
TABLE 1-K. STANDARD SALT WATER*
Ingredient
Amount (g)
Ingredient
Amount (g)
SrCl0.6H00
2 2
H.,BO_
3 3
KBr
CaCl0.2H00
2 2
Na SO
0.02 MgCl9.6H_0
L *•
0.03 NaCl
0.10 Na2Si03.9H20
1.10 EDTAt
4.00
10.0
23.50
0.02
0.003
*To formulate this water, mix technical grade salts with 900 mi of distilled
or demineralized water in the order and quantities listed. Then add enough
distilled or demineralized water to make the final volume 1£. Dilute the
water with distilled or demineralized water to achieve a salinity of 30
°/oo. If necessary, add NaHCO to adjust final pH of water to between 8.0
and 8.2. Before use, filter water through a 0.22-micron membrane filter.
tEthylenediaminetetracetate.
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Test Organisms
Species—
The Regional Administrator shall designate the appropriate test
animals to be used in a particular region. Test animals are as
follows (specific names must be reported):
Invertebrates:
White sea urchin, Tripneustes esculentus
White shrimp, Penaeus setiferus
Pink shrimp, P_. duorarum
Brown shrimp, P_. aztecus
Grass shrimp, Palaemonetes sp.
Shrimp, Crangon sp.
Oceanic shrimp, Pandalus jordani
Blue crab, Callinectes sapidus
Dungeness crab, Cancer magister
Vertebrates:
Sheepshead minnow, Cyprinodon variegatus
Mummichog, Fundulus heteroclitus
Silverside, Menidia sp.
Threespine stickleback, Gasterosteus aculeatus
Pinfish, Lagodon rhomboides
Spot, Leiostomus xanthurus
Shiner perch, Cymatogaster aggregata
Buffalo sculpin, Enophrys bison
Pacific staghorn sculpin, Leptocottus armatus
English sole, Parophrys vetulus
Other species indigenous to the dumping area can be used and are
preferred, if approved by EPA. The specific name of the animals must be
verified and reported. Samples of the test animals can be requested by EPA.
Tests on other animals under other experimental conditions may be required
by EPA.
Source—
Test animals are usually collected from wild populations in rela-
tively unpolluted areas. (Collecting permits may be required by local or
state agencies.) Some animals can be purchased from commercial suppliers or
reared in the laboratory. (See culture techniques.) All animals should be
healthy and as uniform in size and age as possible.
Size—
1. Fish: Fish that weigh between 0.5 and 5.0 grams each are
usually desirable. In any single test, the standard length (tip of snout to
end of caudal peduncle) of the longest fish should be no more than two times
the standard length of the shortest fish.
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2. Maximum size of invertebrates:
shrimp—less than 10-cm rostrum-telson length,
crabs—less than 10-cm carapace width.
Since cannibalism occurs in many species, the claws of crabs should be
banded, or the individuals should be physically isolated.
Care and handling—
If the animals are to be tested at a temperature or salinity other
than that at which they are collected, they should not be subjected to more
than a 3°C change in water temperature in any 24-hour period or to more than
a 5 °/oo change in salinity in any 24-hour period. Crowding should be
avoided to maintain animals in good condition during holding and acclima-
tion. Animals should be fed at least once a day if held for an extended
period, and tanks should be cleaned after feeding.
Animals should be handled as little as possible. Any necessary
handling should be done as gently, carefully, and quickly as possible.
Test animals cannot be taken from any group of organisms apparently
diseased or otherwise stressed, or from any group in which more than 3
percent of the individuals died during the 48 hours immediately prior to
establishing test containers.
Recommended Procedure for Testing Material
Experimental Design—
At least 10 control and 10 test animals must be exposed to each
concentration or dilution of the test material. (They can be in two or more
containers.) However, use of additional animals and replication of treat-
ments are desirable. Replicates, if used, should have no water connection
between replicate test containers. Exposures can be conducted by stratified
randomization (random assignment of one test container for each treatment in
a row, followed by random assignment of a second test container for each
treatment in another or extended row) or by total randomization. A repre-
sentative sample of test animals should be distributed impartially to test
containers by adding one animal (when less than 11 are used) or two animals
(when more than 11 are used), and then repeating this process until desired
number of test animals is reached in each container. Animals can be assigned
alternatively either by total randomization or by stratified randomization
(random assignment of one animal to each test container, random assignment
of a second animal to each test container, etc.).
Controls for every test must duplicate the salt water conditions,
and animals used in containers with test material. A test is not acceptable
if more than 10 percent of the control animals die.
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Temperature—
Test water temperature must be maintained within 1°C of the water
temperature listed in Table 2-K.
TABLE 2-K. SUGGESTED SEA WATER TEST TEMPERATURES FOR VERTEBRATES AND
INVERTEBRATES*
Region Temperature
I 20°C
lit and III 25°C
IV, VI and IX 30 °C
X 15°C
*Temperatures in this table should be revised to the highest average monthly
temperature of oceanic surface waters at dump sites in each region.
tPuerto Rico and Virgin Islands are in Region II administratively but
should use temperatures suggested for Region IV.
Salinity test— Q
Test water salinity should be 30 /oo before the material to be
tested is added.
Loading—
The mass of animals in each test container must be limited so
that the animal's oxygen requirements do not influence the test results. In
general, test containers should not contain more than one gram of animals
per liter. Tests at high temperature may require reduced loading. Proper
loading can be confirmed by measuring dissolved oxygen concentration in the
water of the unaerated control containers. It must not be less than 402
saturation.
preparation of Material to be Tested (See other section of manual on this
subject)—
Samples, whether liquid waste or sludge, must be stirred to a
uniform consistency before dilutions are made.
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Concentrations—
Dilutions of samples, by volume, of 10% (100,000 ppm, 100 ml/Z)
1% (10,000 ppm, 10 ml/O, 0.1% (1,000 ppm, 1 ml/A), 0.01% (100 ppm, 0.1
ml/fc), 0.001% (10 ppm, .01 ml/a), and 0.0001% (1 ppm, 0.001 ml/A) are
recommended as initial test concentrations. In some instances, concentra-
tions of >10% must be tested and resultant salinity adjusted to that of
control.
The highest concentration (dilution) is prepared as follows: 9
volumes of salt water are added to 1 volume of the stirred sample. (Ade-
quate space should be reserved in the test container for stirring and
addition of animals.)
Each succeeding concentration is prepared by a similar l-in-10
serial dilution from the previous test container. Adequate stirring of
the contents of the test' container is essential before each dilution.
Transfer of Animals—
Animals must be added to the test containers within 1 hour
after the proper dilutions of the material to be tested have been made.
Feeding—
The organisms must not be fed while in the test containers
except for Mysidopsis, and these organisms must be fed daily. (See pages
59 and 62.)
Measurements—
The dissolved oxygen concentration, pH, and temperature must be
measured (1) before adding animals and (2) at 24-hour intervals thereafter
in the highest and lowest concentration and in the control. Additional
measurements are required in containers in which animals die. Water
samples should bej taken midway between the top, bottom, and sides of the
test containers and should not include any surface scum of material
stirred up from the bottom or sides.
Observations—
At a minimum, the number of dead or affected animals must be
recorded at 24-hour intervals throughout the test. More observations are
often desirable, especially near the beginning of the test. Dead animals
must be removed as soon as they are observed.
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The adverse effect most often used to study acute toxicity with
aquatic animals is death. Criteria for death are no movements, especially
no opercular movement in fish, and no reaction to gentle prodding. However
death is not easily determined for some invertebrates; thus an EC50 (effec-'
tive concentration to 50% of test animals) is often measured rather than an
LC50 (lethal concentration to 50% of test animals). The effect generally
used for determining an EC50 with invertebrates is immobilization, which is
defined as the inability to move, except for minor activity of appendages,
or loss of equilibrium. Other effects can be used for determining an EC50,
but the effect and its definition must always be reported. General observa-
tions on such things as erratic swimming, loss of reflex, discoloration,
changes in behavior, excessive mucous production, hyperventilation, opaque
eyes, curved spine, hemorrhaging, molting, and cannibalism should be reported.
Calculation and Reporting
At the end of the test period, the bioassays are terminated and the LC50
or EC50 values are determined.
Calculations—
An LC50 is a concentration at which 50% of the experimental animals
died, and an EC50 is a concentration at which 50% of the experimental
animals were affected. Either may be an interpolated value based on per-
centages of animals dying or affected at two or more concentrations.
Estimating the LC50 or EC50 by interpolation involves plotting the data on
semilogarithmic coordinate paper with concentrations on the logarithmic axis
and percentages of dead or affected animals on the arithmetic axis. A
straight line is drawn between two points representing death or effect in
concentrations that were lethal to or effective against more than half and
less than half of the organisms. The concentration at which the line
crosses the 50% mortality or effect line is the LC50 or EC50 value. If 50%
of the test animals are not affected by the highest concentration, the
percentage affected should be reported.
Reports—
Any deviation from this method must be noted in all reports of
results. A report of the results of both aerated and unaerated tests must
include:
1. name of method, author, laboratory, and date tests were
conducted;
2. a detailed description of the material tested, including its
source, date and time of collection, composition, known physical and chemi-
cal properties, and variability;
3. the source of the salt water, date prepared, and method of
preparation;
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4. detailed information about the test animals, including name,
standard length, weight, source, history, and acclimation procedure used;
5. a description of the experimental design, the test containers,
the volume of test solution, initial test conditions, the number of organ-
isms per concentration, and the loading;
6. definitions of the criteria used to determine the effect and
a summary of general observations on other effects or symptoms;
7. percentage of control organisms that died or were affected
in each test container;
8. the 24-, 48-, and 96-hour LC50, or EC50;
9. methods used for and the results of all dissolved oxygen, pH
and temperature measurements; and
10. any other relevant information.
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L. FLOW-THROUGH METHODS FOR ACUTE TOXICITY TESTS USING FISHES AND
MACROINVERTEBRATES (See list of contributors.)
INTRODUCTION
Continuous- flow (often referred to as "flow- through") bioassays have
definite advantages over static tests in evaluating certain types of wastes
to be disposed of at sea, particularly in testing waste chemicals that have
high biochemical oxygen demands, and are unstable or volatile. Many test
species of fish and macroinvertebrates have high rates of metabolism and are
difficult to maintain in jars or tanks of standing sea water. Continuous-
flow bioassays, conducted under proper conditions, provide for well-oxygen-
ated test solutions, nonf luctuating concentrations of the toxicant, and
,
°1fQ™*abolic wastes of ^e test organisms (Standard Meth-
ods, 13th Edition, 1971).
This method provides general procedures for conducting a 96-hour, flow-
through bioassay on marine fish and macroinvertebrates such as shrimp and
crabs. Evaluation of different types of waste will require some modifica-
tion of these procedures.
Equipment
Facilities —
For maximum convenience and versatility, the facilities should
include tanks or aquaria for holding and acclimating test animals, a tank
for sea water, and a temperature-controlled recirculating water bath or
controlled- environment room for the test containers. The holding and accli-
mation tanks should be equipped for temperature control and the holding tank
should be equipped for aeration for emergency use. During holding, acclima-
tion, and testing, test animals should be shielded from unnecessary distur-
bances.
Construction Materials —
Construction materials and commercially purchased equipment that may
contact any water into which test animals are to be placed should not
contain any toxic substances that can be leached, corroded, or dissolved by
the water In addition, materials and equipment should be chosen to minimize
sorption of toxicants from water. It is suggested that glass, #316 stain-
less steel, or perfluorocarbon plastics be used whenever possible
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Test Containers—
Type: The test solution for fish and invertebrates usually should
be placed in containers measuring between 15 and 20 cm deep. Test containers
can be made by welding (not soldering) stainless steel or by gluing double-
strength window glass with clear silicon adhesive. As little adhesive as
possible should be in contact with the water; extra beads of adhesive should
be on the outside rather than inside the containers. Plywood tanks coated
with fiberglass resin are also acceptable.
Cleaning: Test containers must be cleaned before use. New contain-
ers must be washed with detergent and rinsed with 10% hydrochloric acid,
acetone, and tap or other clean water. Test containers, if reused, should
be (1) emptied; (2) rinsed with water; (3) cleaned by an appropriate pro-
cedure to remove the toxicant tested, e.g., acid to remove metals and
bases; detergent, organic solvent, or activated charcoal to remove organic
compounds; and (A) rinsed with water. Acid is also useful for removing
scale and hypochlorite (bleach) is useful for removing organic matter and
for disinfecting. All test containers must be rinsed with uncontaminated
sea water immediately before use.
Sea Water
Acute toxicity tests, require acceptable sea water in which healthy test
animals can survive throughout acclimation and testing without sign of
stress, such as unusual behavior or discoloration. Natural sea water
(particularly from the dump site) is preferable to artificial sea water;
however, artificial sea water is sometimes more practical due to logistics
or costs. Salinity of test water ideally should duplicate the dump site;
however, requirements of the individual species to be tested must be con-
sidered. See page 21 for composition of artificial sea water.
Test Organisms
Species—
Recommended species are as follows (specific name must be verified
and reported):
Invertebrates:
White sea urchin, Tripneustes esculentus
White shrimp, Penaeus setiferus
Pink shrimp, P_. duorarum
Brown shrimp, £. aztecus
Grass shrimp, Palaemonetes sp.
Shrimp, Crangon sp.
Oceanic shrimp, Pandalus jordani
Blue crab, Callinectes sapidus
Dungeness crab, Cancer magister
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Vertebrates:
Sheepshead minnow, Cyprinodon variegatus
Mummichog, Fundulus heteroclitus
Longnose killifish, F_. similis
Silverside, Menidia sp.
Threespine stickleback, Gasterosteus aculeatus
Pinfish, Lagodon rhomboides
Spot, Leiostomus xanthurus
Shiner perch, Cymatogaster aggregata
Buffalo sculpin, Enophrys bison
Pacific staghorn sculpin, Leptocottus armatus
English sole, Parophrys vetulus
Other species indigenous to the dumping area can be used if approved
by EPA and if the specific name of the organism is verified and reported.
Samples of the test animals can be requested by EPA. Tests on other organ-
isms under other experimental conditions can be required by EPA.
Source—
Test animals are usually collected from wild populations in
relatively unpolluted areas. (Collecting permits may be required by local
or state agencies.) Some animals can be purchased from commercial suppliers.
All animals should be healthy and as uniform in size and age as possible.
Juvenile stages are preferable.
Size—
1. Fish: Fish that weigh between 0.5 and 5.0 g each are
usually desirable. In any single test, the standard length (tip of snout to
end of caudal peduncle) of the longest fish should be no more than two times
the standard length of the shortest fish.
2. Size requirements for invertebrates:
Palaemonetes—10-20 mm rostrum-telson length
shrimp—5-8 cm rostrum-telson length (5-8 g live
weight)
crabs—less than 7 cm carapace width
Acclimation—
Conditions of acclimation should be related to test requirements.
Organisms should be subjected to as little stress as possible. Initially,
temperature and salinity in the laboratory should resemble those of the
medium used to transport test animals. During acclimation, mortality should
not exceed 10 percent. Fishes should be held in the laboratory at least 14
days and invertebrates 4 days prior to testing. If the acclimation tempera-
ture and salinity differ from those of the test, they should be adjusted
gradually (at least 48 hours prior to testing) to the test conditions.
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Care and Handling
Animals should be handled as little as possible. Any necessary
handling should be done with a dip net as gently, carefully, and quickly as
possible. Animals should be fed daily during acclimation, but fish should
not be fed for a period of 48 hours before or during the actual test. It^
may be necessary, however, to feed certain invertebrates during the actual
test.
Crowding should be avoided. Cannibalism occurs in many species of
arthropods; therefore, in some cases it may be necessary to isolate indi-
viduals in compartmented aquaria by such techniques as banding the crab
claws, and placing a 2-3 cm (about 1 in) layer of sand in the bottom of the
aquaria used for testing shrimp.
Recommended Procedure for Testing Materials
Experimental Design
The recommended test procedure consists of a 96-hour bioassay, using
a control and at least five concentrations of the material to be tested.
Acute static tests are useful in determining range of toxicity of the mate-
rial and selecting concentrations for the flow-through tests. (See Section
5-f, Range-finding and Definitive Tests).
In the definitive test, a minimum of 20 organisms is required for
the control and each concentration or dilution of the material to be tested.
(They can be divided in two or more test containers.) However, use of
additional organisms and replication of treatments are desirable, but "load-
ing" must be considered. Replicates, if used, should have no water connec-
tion between the replicate test containers. Stratified randomization (ran-
dom assignment of one test container for each treatment in a row, followed
by random assignment of a second test container for each treatment in another
or extended row) or total randomization of the treatments is recommended.
The test animals should be distributed impartially to test containers
by adding no more than 10 percent to each container, repeating the process
until the desired number of test animals is reached in each test container.
Animals can be assigned alternatively either by total randomization or by
stratified randomization (random assignment of one animal to each test
container, random assignment of a second animal to each test container,
etc.).
Controls for every test must duplicate the salt water, conditions,
and animals (species and size) used in containers with test material. Test
results are unacceptable if mortality of control animals exceeds 10 percent.
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Toxicant Delivery System—
Flowing sea water tests are preferable to static tests because
test solutions are renewed continually, assuring a steady concentration of
the toxicant. However, these tests require metering pumps or other devices
to deliver the toxicant or test material into the sea water flowing through
the test aquaria.
Most toxicant delivery systems have been designed to test toxicants
and solvents in fresh water, and may not be applicable in studies of all
wastes. Many materials proposed for disposal at sea are not homogenous mix-
tures; therefore, innovative toxicant delivery systems are required to
introduce representative samples of the materials into test containers.
Stirring may be required to maintain suspended solids in nonhomogenous dump
material.
Many toxicant delivery systems have been described and used in
various types of bioassays (Sprague, 1969; Freeman, 1971; Bengtsson, 1972;
Cline and Post, 1972; Granmo and Kollberg, 1972; Lowe et al., 1971 and 1972;
Lichatowich et al., 1973; Abram, 1973). The proportional diluter (Mount and
Brungs, 1967) has probably been used routinely (in fresh water) more than
any other system. A small chamber to mix toxicant-bearing water and dilution
water should be placed between the diluter and the test containers for each
concentration. If duplicate test containers are used, separate delivery
tubes can be run from this mixing chamber to each duplicate. Alterations in
the design of the proportional diluter have been found useful (Esvelt and
Conners, 1971; McAllister, Mauch, and Mayer, 1972; Benoit and Pulglisi,
1973; Schimmel, Hansen and Forester, 1974).
The rate for which water flows through the test containers must be
at least five tank-water volumes per 24 hours. It often is desirable to
construct a toxicant delivery system that provides 10 or more volumes of
tank water per 24 hours. Some systems may provide a continuous flow of sea
water. The rate of flow should not vary by more than 10 percent from any
test container or for any time period within a given test.
The calibration of the toxicant delivery system should be checked
carefully before, during, and after each test. The volume of stock solution
and dilution water used in each portion of the toxicant delivery system and
the flow rate through each test container must be determined and operation
of the toxicant delivery systems checked daily during the test.
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Temperature—
Test water temperatures should be the highest average monthly
temperature at the discharge site. Suggested water temperatures in Table 1-
L represent maximum for surface waters in the coastal regions.
Test water temperature should be maintained within 1 C of average
maximum monthly temperature at the dump site or temperature listed in Table
1-L (unless seasonal bioassays are performed). This may be accomplished by
preheating the sea water before it enters the test containers, by immersing
the test containers in a constant temperature water bath, or by a combina-
tion of these methods.
TABLE 1-L. MAXIMUM SEA WATER TEST TEMPERATURES FOR VERTEBRATES AND
INVERTEBRATES*
Region Temperature
I 20 °C
lit and III 25°C
IV, VI and IX 30°C
X 15°C
*Temperature in this table should be revised to the highest average monthly
temperature of oceanic surface waters at dump sites in each region.
tPuerto Rico and Virgin Islands are in Region II administratively but should
use temperatures suggested for Region IV.
Salinity—
The salinity of test water should be that of the dump site if:
(a) dump site water is used or (b) artificial sea water is prepared. The
salinity of any other natural sea water should be ^15 loo.
Loading—
Excessive weight (grams/liter) of organisms in a test container
may adversely affect results of test. Therefore, the loading must be limited
so that:
1. the concentration of dissolved oxygen in the control container
does not fall below 60% saturation;
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2. the concentration of metabolic products does not become too
high; specifically, the concentration of non-ionized ammonia does not
exceed 20
3. the concentration of toxicant is not lowered by more than 20%
because of uptake by the test organisms; and
4. the organisms are not stressed by overcrowding.
Loading in the test containers must not exceed 2 g/^/day for
species listed under Section 4, "Test Organisms." Lower loadings must be
used when necessary to meet the four criteria listed above.
Range-Finding and Definitive Tests —
Time and effort may be saved by "range-finding," static tests
using a few animals and a wide range of concentrations, as a preliminary to
"definitive" flow- through tests which will be used to calculate the final
LC50 or EC50. (See Standard Methods, 14th Edition, 1975 for details.) For
example, waste concentrations of 10, 1, 0.1, and 0.01% might be tested first
by volume and with two or three animals in each concentration for a period
of 24 hours. Definitive test concentrations should then fall between the
highest concentration at which all animals survive and the lowest concentra-
tion at which all or most animals die.
Observations —
At a minimum, the number of dead or affected animals must be
recorded at 24-hour intervals throughout the test. More observations are
often desirable, especially in the beginning stage of the test. Dead animals
must be removed immediately after observed and their deaths recorded.
Death is the adverse effect most often used to study acute toxicity
with aquatic animals. Criteria for death are no movements, especially no
opercular movement in fish, and no reaction to gentle prodding. Because
death is not easily determined for some invertebrates, an EC50 (effective
concentration to 50% of test animals) is often measured rather than an LC50
(lethal concentration to 50% of test animals) . The effect generally used
for determining an EC50 with invertebrates is immobilization, which is
defined as the inability to move except for minor activity of appendages, or
loss of equilibrium. Other effects can be used for determining an EC50, but
the effect and its definition must always be reported. General observations
on such criteria as erratic swimming, loss of reflex, discoloration, changes
in behavior, excessive mucous production, hyperventilation, opaque eyes,
curved spine, hemorrhaging, molting, and cannibalism should be reported.
Calculations and Reporting
At the end of the test period, the bioassays are terminated and the LC50
or EC50 values are determined.
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Calculations—
An LC50 is the concentration expected to result in 50 percent
mortality of the experimental animals and an EC50 the concentration expected
to affect 50% of the experimental animals. Either value may be interpolated
from percentages of animals dying or affected at two or more concentrations.
In interpolating LC50 or EC50, plot data on logarithmic-probability graph
paper, placing concentrations on the logarithmic axis and percentage of dead
or affected animals on the probability axis. A line is drawn between all
data points. The concentration at which the line crosses the 50% mortality
or effect line is the LC50 or EC50 value. In fitting the line,points
nearest the 50% effect level should be given more weight. Ideally, data
should consist of enough intermediate (between 0 and 100%) effects to deter-
mine confidence limits by statistical tests (such as probit analysis).
If 50% of the test animals are not affected by the highest con-
centration, the percentage affected at each concentration must be reported.
Reports—
The final report should include:
1. name of method, author, laboratory, and date tests were
conducted;
2. a detailed description of the material tested, including its
source, date, and time of collection, composition, known physical and chemi-
cal properties, and variability of the material tested;
3. the source of the sea water, date prepared, and method of
preparation;
4. detailed information about the test animals, including name,
standard length of fishes, carapace width of crabs, total length of shrimp,
weight, source, history, and acclimation procedure;
5. a description of the experimental design, the test containers,
the volume of test solution, the number of organisms per concentration, and
the loading (water flow to each tank);
6. definitions of the criteria used to determine the effect and
a summary of general observations on other effects or symptoms;
7. percentage of control organisms that died or were affected in
each test container;
8. the 24-, 48-, and 96-hour LC50 or EC50 values;
9. methods used for the results of all dissolved oxygen, pH, and
temperature measurements; and
10. any other relevant information.
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REFERENCES
Abram, F.S.H. 1973. Apparatus for Control of Poison Concentration in
Toxicity Studies With Fish. Water Res. (Oxford) 7(12):1875-1879.
American Public Health Association. 1975. Standard Methods for the
Examination of Water and Wastewater, 14th Edition. Am. Pub. Health
Assoc., Wash., DC. 874 p.
Bengtsson, B.E. 1972. A Simple Principle for Dosing Apparatus in Aquatic
Systems. Arch. Hydrobiol. (Stuttgart) 70(3):413-415.
Benoit, D.A, and F.A. Puglisi. 1973. A Simplified Flow-splitting Chamber
and Siphon for Proportional Diluters. Water Res. (Oxford) 7(12):1915-
1916.
Cline, T.F.5and G. Post. 1972. Therapy for Trout Eggs Infected With
Saprole'gnia. Prog. Fish-Cult. 34 (3) : 148-151.
Esvelt, L.A., and J.D. Conners. 1971. Continuous-flow Fish Bioassay
Apparatus for Municipal and Industrial Effluents. In: L.A. Esvelt, W.J.
Kaufman, and R.E. Selleck. Toxicity Removal from Municipal Wastewaters.
Volume IV of "A Study of Toxicity and Biostimulation in San Francisco
Bay-Delta Waters." Sanitary Engineering Research Laboratory, Univ.
California, Berkeley, pp. 155-182.
Freeman, R.A. 1971. A Constant Flow Delivery Device for Chropic Bioassay.
Trans. Am. Fish Soc. 100(1):135-136.
Granmo, A., and S.C. Kollberg. 1972. A New Simple Water Flow System for
Accurate Continuous Flow Tests. Water Res. 6(9):1597-1599.
Lichatowich, J.A., P.W. O'Keefe, J.A. Strand, and W.L. Templeton. 1973.
Development of Methodology and Apparatus for the Bioassay of Oil. In:
Proceedings of Joint Conference on Prevention and Control of Oil Spills.
American Petroleum Institute, U.S. Environmental Protection Agency, and
U.S. Coast Guard, Washington, DC. pp. 659-666.
Lowe, J.I., P.O. Wilson, A.J. Rick, and A.J. Wilson, Jr. 1971. Chronic
Exposure of Oysters to DDT, Toxaphene and Parathion. Proc. Natl.
Shellfish Assoc. 61:71-79
105
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we J.I«» P'R' Parrisht J-M. Patrick, Jr., and J. Forester. 1972. Effects
of the Polychlorinated Biphenyl Aroclor^ 1254 on the American Oyster,
crassostrea vlrginica. Mar. Blol. (Berl.) 17:209-214.
McAllister, W.A., Jr. W.L. Mauch, and F.L. Mayer, Jr. 1972. A Simplified
Device for Metering Chemicals in Intermittent-flow Bioassays. Trans.
Am. Fish. Soc. 101 (3).-555-557.
Mount, D.I., and W.A. Brungs. 1967. A Simplified Dosing Apparatus for Fish
Toxicological Studies. Water Res. (Oxford) 1(1):21-29.
Schimmel, S.C., D.J. Hansen, and J. Forester. 1974. Effects of Aroclor 1254
on Laboratory-reared Embryos and Fry of Sheepshead Minnows (Cyprinodon
variegatus). Trans. Am. Fish. Soc. 103(3):582-586.
Sprague, J.B. 1969. Review Paper: Measurement of Pollution Toxicity to
Fish. 1. Bioassay Methods for Acute Toxicity. Water Res. 3(11):793-821.
106
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M. LABORATORY CULTURE OF SHEEPSHEAD MINNOWS (CYPRINODON VARIEGATUS)
D. J. Hansen
Sheepshead minnows can be readily cultured in the laboratory in aquaria
with under-substrate filters or in aquaria receiving flowing salt water.
The following discussion presents culture techniques used successfully at
the U.S. Environmental Protection Agency, Environmental Research Laboratory,
Gulf Breeze, Florida 32561.
Spawning Method Using Human Chorionic Gonadotrophic Hormone (HCG)
Although sheepshead minnows spawn naturally in the laboratory, in some
instances it is desirable to obtain large numbers of eggs on one particular
day. To do this, adult fish ^27 mm standard length should be acclimated for
at least two weeks in >_15 /oo salinity water at 30°C. Conditions during
acclimation should not vary from those recommended by the Committee on
Methods for Toxicity Tests with Aquatic Organisms (1975). Photoperiod
should consist of 12 hours of light and 12 hours dark. Fish should be fed
ad_ libitum on frozen adult brine shrimp supplemented with dry food. After
acclimation each adult female should be injected intraperitoneally with 50
International Units (IU) of HCG to enhance egg production. The next day all
females should again be injected with 50 IU of HCG. Three days after the
first injection, manually strip, or dissect, eggs from females and deposit
them in 25-50 ml salt water of acclimation conditions. Remove testes from
five or more males and macerate in a few ml of sea water to free sperm. Mix
sperm with eggs in a beaker and place in 30°C water bath for one hour.
Embryos then are placed in egg chambers for a life-cycle toxicity test using
sheepshead minnows, or placed in suitable hatching chambers.
The advantage of this procedure is that tests can be planned to assure
availability of sufficient embryos for life-cycle tests or sufficient
juveniles for acute static or flow-through tests after 2 weeks acclimation.
However, because females are usually killed or their normal egg production
patterns disrupted, this method should be used only occasionally when sur-
plus females are available. If this procedure is followed, the number of
eggs produced per female usually averages between 100 and 200, depending on
size of females. Fertilization success should be >90 percent.
Spawning Method Using Natural Spawning
It is sometimes desirable to obtain a continuous supply of sheepshead
minnows for toxicity tests by using natural reproduction of laboratory-held
fish. To use this method, adult fish ^27 mm standard length should be
acclimated for at least two weeks in ^15 °/oo salinity water at 30 C.
Conditions during acclimation should be identical to those described for
spawning using HCG. Fish are placed in spawning chambers spacious enough to
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prevent deaths due to aggressive territorlality by males and cannibalism of
eggs. Spawning chambers as described in the section, "Entire Life-Cycle
Toxicity Test Using Sheepshead Minnows," have proven successful at ERL, Gulf
Breeze.
The number of spawning chambers and fish to be spawned should be based
on the requirements for providing sufficient embryos. A pair of fish will
generally produce an average 10 to 30 eggs each day while held in spawning
chambers in water of ^.15 loo salinity and temperature of 30 C. The number
of fish successfully held in our spawning chambers ranged from one pair to
two males and five females.
Rearing Methods for Embryo, Fry, and Juvenile Sheepshead Minnows
Two hatching techniques that have proven most successful require place-
ment of embryos in (1) flowing salt water aquaria chambers as described in
the life-cycle toxicity test method or (2) in static salt water in separa-
tory funnels which are supplied air through a 2.5-mm ID glass tube that
extends to the bottom of the funnel. Air flow to the tube should be only
fast enough to keep embryos and hatched fish suspended in the water column.
Hatching time required for sheepshead minnows depends on temperature of
water, and survival depends on temperature and salinity (Schimmel and
Hansen, 1974). Embryos hatch most rapidly and their survival is optimum in
water _>15 /oo salinity and 30 C temperature. Unfortunately, we have yet to
find an artificial sea salt that can be used in rearing embryos.
After embryos hatch, the fish are removed from embryo and fry chambers,
or separatory funnels, and placed in aquaria for acclimation for toxicity
tests. Salinity and temperature should be adjusted for acclimation as
suggested for acute toxicity test methods. Fish are fed live brine shrimp
nauplii.
REFERENCES
Committee on Methods for Toxicity Tests with Aquatic Organisms. 1975.
Methods for Acute Toxicity Tests with Fish, Macroinvertebrates, and
Amphibians. EPA-660/3-75-009. U.S. Environmental Protection Agency,
Cincinnati, OH.
Schimmel, Steven C., and David J. Hansen. 1974. Sheepshead Minnow
(Cyprinodon variegatus); An Estuarine Fish Suitable for Chronic
(Entire Life-cycle) Bioassays. Proc. 28 Annu. Conf. Southeast. Assoc. Game
Fish Comm. pp. 392-398.
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N. LIFE-CYCLE TOXICITY TEST USING SHEEPSHEAD MINNOWS (CYPRINODON VARIEGATUS)
D.J. Hansen, P.R. Parrish, S.C. Schimmel, and L.R. uoodman.
PURPOSE AND LIMITATIONS
This procedure provides a method to determine the effect of continuous
exposure of a toxic material on sheepshead minnow embryos and fry: their
survival and growth to adulthood, and spawning success. Spawning success is
measured by the ability of fish to spawn naturally, number of eggs spawned,
fertilization success, and survival of embryos and fry. The experiment
requires from 4 to 6 months.
The primary advantage of this test is that results, when compared with
those of acute tests with this species, can be used to calculate an appli-
cation factor (Mount & Stephan, 1967). This factor, used to assess relative
chronic hazards of materials, is important in establishing water quality
criteria (Eaton, 1973; Hansen and Parrish, 1977).
This test has several limitations and should not be considered valid in
assessing toxicity of all materials. Sheepshead minnows can tolerate low
dissolved oxygen and wide ranges of temperature and salinity. Therefore,
toxicity tests using this fish may underestimate the toxicity of materials
that alter these environmental conditions. Materials tested should mix well
with water. Insoluble or highly turbid materials mix poorly, and their
toxicity may be under- or overestimated.
Physical Systems
a. Test Water
1. The source of test water should be (1) from the dump site or (2)
natural seawater with salinity ^15 /oo.
2. Sea water must be filtered to remove particles 15v and larger,
but filtration should not affect the chemical composition of the natural sea
water. Filtration must remove planktonic larvae which prey upon eggs, fry,
and juvenile fish.
3. Any sea water source proposed must be analyzed for possible
pollutants (e.g., pesticides, PCB's, heavy metals, and the material to be
tested).
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b. Dosing Apparatus
A number of apparatus are acceptable for this bioassay. For example,
use the device described by Mount and Brungs (1967) or Hansen et al. (1971)
for substances not requiring a solvent. However, if a solvent is required,
use the device described by Hansen et al. (1974) or Schimmel et al. (1974).
c. Toxicant Mixing
A mixing chamber is necessary to assure adequate mixing of the test
material. Aeration should not be used for mixing. Mixing is extremely
important because if materials are not adequately mixed with water, toxicity
cannot be properly assessed. Improper mixing can either expose the animal
to too much or too little of the material, causing toxicity to be over- or
underestimated. Therefore, scientific judgment should be used for designing
and selecting appropriate dosing apparatus and mixing systems.
d. Duplicates
True duplicates are used for each concentration in all tests (no
water connection between aquaria). Aquaria are located by random selection.
e. Aquaria
Glass aquaria, 45 x 90 x 26 cm high and with a water depth of 19 cm,
have been used successfully.
f. Embryo and Fry Chambers
1. Embryo and fry chambers must allow for adequate exchange of
water and insure that the proper quantity of material enters the chambers.
Chambers can be constructed from Petri dishes to which 40-mesh nylon or a
stainless steel screen is glued (Schimmel et al, 1974). The Petri dish
chambers are placed in aquaria that have a self-starting siphon. Water from
the dosing apparatus fills the aquaria to the level required to start the
siphon. Water then drains from the aquaria, flowing in and out of the
embryo and fry chambers. Chambers can also be constructed from 5-cm OD
round glass or beakers without bottoms. The bottoms are replaced with 40-
mesh stainless steel or nylon screening. Chambers are suspended in the test
water on an oscillating rocker arm apparatus that is driven by a 1-5 rpm
electric motor (Mount, 1968). These chambers must be brushed daily to
prevent clogging.
2. Embryo and fry chambers must be designed so water can be drained
to 1 cm, or the fry removed for observations and measurement.
3. Embryo and fry chambers may be supplied test water by: (1)
separate delivery tubes from the mixing chamber, (2) self-starting siphons
in the aquaria, or (3) an oscillating rocker arm apparatus.
110
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g. Flow Rate
1. Flow rates to each duplicate aquarium must: (1) provide 90%
replacement in 8 to 12 hours (Sprague, 1969); (2) maintain dissolved oxygen
60% saturation; and (3) maintain the toxicant concentration.
2. The test system is equipped with an alarm system to insure con-
tinuation of water flow, toxicant flow, and temperature regulation.
h. Photoperiod
A 12-hour light/12-hour dark cycle is maintained throughout the
test. It may be desirable to control lights by a timing switch (Drummond
and Dawson, 1970). Lighting above each replicate must be balanced.
i. Temperature
Test temperature is maintained at 30°C (+1°C) by either (1) pre-
heating the diluted water to the prescribed temperature, and/or (2) placing
test aquaria in a temperature-controlled water bath. A continuous record of
temperature of test water must be kept.
j. Cleaning
All aquaria are cleaned whenever organic material builds up.
Aquaria are brushed down and siphoned to remove accumulated material. Fish
can be left in the aquaria, but the end of the siphon is covered with
screen. Care should be exercised in cleaning to prevent loss of or damage
to the fry, juveniles, or adults. Embryo and fry chambers may have to be
replaced or cleaned frequently if screens clog or organic material collects.
When a chamber is cleaned, it can be reused only in the aquarium from which
it was removed. Special care is required to prevent injury to fry.
k. Spawning Chambers
Chambers are constructed of either glass or #316 stainless steel
(Hansen et al., 1977). Chambers 20 x 35 x 22 cm high have been used suc-
cessfully. A 2.0-mm mesh screen is attached 1 cm above the bottom of a
removable "drawer" to facilitate passage of eggs, thereby reducing canni-
balism of eggs by parents. A "drawer" of 0.5-nm mesh nylon or 316 stainless
steel screen will catch eggs falling through the screen to the bottom of
spawning chambers. Fish in the test aquarium outside the spawning chamber
must be prevented from eating the eggs. This is accomplished by a. partition
or by a drawer constructed so that fish have no access to eggs.
1. Disturbance
Fish are shielded from excessive outside disturbances. Tanks will
eliminate outside light sources that interfere with the photoperiod.
Preferably, an opaque curtain surrounds the entire test apparatus.
Ill
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m. Concentrations
1. A minimum of five concentrations of toxicant and a control, all
duplicated, are utilized in all chronic tests. When a solvent is used, the
control contains the solvent.
2. One concentration selected must adversely affect a life stage
of the sheepshead minnow and one concentration must not affect any life
stage. Concentrations selected for chronic toxicity tests are based on
results of acute flow-through toxicity tests. Selection of test concentra-
tions is difficult because chronic effects on survival, growth, or repro-
duction of sheepshead minnows can occur at concentrations that range from
0.5 to 0.0001 of the 96-hour LC50. The accuracy of the selection process
can be improved by conducting preliminary tests such as: (a) acute 96-hour
flow-through tests using different life stages (e.g., fry, juvenile, and
adults), (b) acute test to determine incipient LC50 (Sprague, 1969), or (c)
embryo-fry tests (Schimmel & Hansen, 1974). The highest concentration in
the life-cycle test, in most instances, should be the lowest concentration
affecting survival or growth in preliminary tests.
3. Chemical analyses are required to interpret results of this
complex bioassay: Water and a minimum of 10 or more fish should be analyzed
for each aquarium, but preferably water and muscle and gametes of fish in
each life cycle should be chemically analyzed weekly.
n. Acute Tests
Acute flow-through toxicity tests using juvenile fish must be con-
ducted. Consult section on suggested acute flow-through bioassay methods.
Biological Systems
a. Source of Adult Fish
Adult fish are obtained from the one source, either from wild popu-
lations or suitable culture laboratories; wild stocks may be preferable.
They are held in flowing 30°C sea water of >15 loo salinity for at least
two weeks before the eggs are removed. Neither fish nor eggs should contain
excessive contaminants nor exhibit excessive mortality; fish should demon-
strate normal behavior. (Committee on Methods for Toxicity Tests with
Aquatic Organisms, 1975.)
b. Eggs from Adult Fish
To obtain a sufficient number of eggs for a chronic exposure, two
methods may be employed: (1) natural spawning from laboratory stocks; and
(2) artificial inducement, in which egg production is stimulated by injec-
tion of human gonadotrophic hormone. Eggs are removed from females and are
fertilized in salt water with sperm excised from males (Schimmel et al.,
1974). Consult section on culturing sheepshead minnows.
112
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c. Test Implementation
The test begins after 50 microscopically confirmed embryos are
placed in two or more embryo and fry chambers in each duplicate aquarium.
Survival of embryos and fry (which constitute the parental stock) are
observed and recorded daily. Occurrence of abnormalities in embryos and try,
and their frequency, are important indicators of teratogenicity. Also,
signs of poisoning should be observed and recorded as indicators of mode of
action of the toxicant. Effects of toxicants on behavior of this fish can
be as significant as, or more significant than, effects on survival, growth,
or reproduction.
d. Food
1. Fry are fed equal portions of live brine shrimp nauplii two or
more times daily for about two weeks. (Do not use frozen nauplii.)
2. Juveniles and adults can be fed twice daily on equal portions of
dry food (e.g. BiOrell^ or Tetramin^) supplemented with frozen adult brine
shrimp. Each batch of food should be checked for pesticides (DDT, dieldrin,
endrin, etc.) and metals. In addition, chemical analysis should also include
chemicals in the material to be tested.
e. Disease
If disease occurs, a test preferably is terminated and started
again. If diseased animals are treated, they should be handled according to
their nature. Each aquarium is treated identically even though disease is
not evident in all aquaria. Treatments should be kept to the minimum and
recorded as to type, amount of medication, and frequency.
f. Measurements
Fish of the parental generation are measured in mm standard lengths
at four weeks before removal of extra fish. Therefore, measurements are
taken at four-week intervals and at adult termination. Juvenile (F.) fish
are to be measured at week four (termination of test). Techniques suggested
for measuring fish include a photographic method outlined by McKim and
Benoit (1971) and direct measurement at termination.
g. Thinning
At day 28, juvenile fish are randomly reduced to 25 fish per dupli-
cate aquaria, providing enough fish for at least two spawning groups of
three adult females and two males in each duplicate aquarium for obser-
vations on effects on spawning.
h. Spawning
When mature adults begin courtship (indicated by sexual dimorphism,
territoriality, and aggressive behavior by the male), and attain a minimum
standard length of 27 mm, three females and two males are placed in individual
113
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spawning chambers in the test aquaria. Fish from each spawning group are
left in chambers for a minimum 14 days. All (possible) fish in the 2:3
ratios in each aquarium are spawned and extra, unspawned fish from each
duplicate aquarium are combined whenever feasible to form additional 2:3
spawning groups. Adult deaths during spawning should be noted; dead animals
are removed, measured, but not replaced. At termination of each spawning
group, lengths and weights of individual fish are measured.
i. Egg Removal
Records of egg numbers and egg fertility are maintained daily. All
eggs must be removed daily, examined for fertility, reserved for survival
studies or residue analyses, or discarded. Eggs are removed at a fixed time
each day so spawning activity is not disturbed unnecessarily.
j. Egg Incubation
1. Fifty embryos are collected and incubated from adults in each
aquarium. It may be desirable to obtain 25 from one day's spawning by each
of two spawning groups. If spawns are small, the 50 embryos can be collected
over an extended period.
2. If no spawning occurs in the highest concentration, embryos are
transferred from control spawns and incubated in the highest concentration
to gain additional information.
3. Groups of 50 embryos are divided into two-egg cups. Survival of
embryos, time required to hatch, hatching success, and survival of fry for
four weeks are determined and recorded.
4. Additional groups of 50 embryos from fish from contaminated
aquaria should be rinsed with control water and then placed in control
aquaria to determine if the eggs contain chemicals toxic to embryos or fry.
k. Embryo, Fry, and Juveniles (the FI generation)
Survival of embryos and fry is recorded daily for four weeks. Daily
observations are made of embryos and fry; mortalities, development of
abnormalities, and signs of poisoning are recorded. Length and weight of
juvenile fish is measured at test termination (day 28); weight may represent
the total for all fish in each fry chamber. Fish may be saved for chemical
analyses.
1. Termination of Adults
1. In many chronic procedures utilizing other fishes, tests are
terminated when no spawning activity occurs for a two-week interval. Tests
using the sheepshead minnow, however, should terminate after a spawning is
observed for two weeks because this fish spawns readily and almost daily
unless immature or affected by a pollutant. The effect of the toxicant on
each group spawning in the 2:3 ratio is tested and each group then termi-
nated. Final termination follows tests of all spawning groups.
114
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2. Adult fish are weighed, measured for standard length, sexed, and
retained for residue analysis.
m. Additional Tests
Certain materials may contain substances that require additional
tests to determine physiological or pathological effects on one or more life
stage of the sheepshead minnow.
Statistical Analyses
The LC50's and 95% confidence limits are calculated on data from acute
tests by probit analysis. Data from life-cycle bioassays are analyzed by
analyses of variance, or chi-square tests. Post hoc tests are conducted on
treatment means using the Newman-Keuls range test.
115
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REFERENCES
Committee on Methods for Toxicity Tests with Aquatic Organisms, 1975.
Methods for Toxicity Tests with Fish, Macroinvertebrates, and Amphi-
bians. EPA-660/3-75-009. U.S. Environmental Protection Agency,
Cincinnati, OH.
Drummond, Robert A., and Walter F. Dawson. 1970. An Inexpensive Method for
Simulating Diel Patterns of Lighting in the Laboratory. Trans. Amer.
Fish Soc. 99(2):434-435.
Eaton, J.G. 1973. Recent Developments in the Use of Laboratory Bioassays to
Determine "Safe" Levels of Toxicants for Fish. G.E. Glass, Ed.
Ann Arbor Science Publishers, Inc., Ann Arbor, Mich. pp. 107-115.
Hansen, D.J., and P.R. Parrish. 1977. Suitability of Sheepshead Minnows
(Cyprinodon variegatus) for Life-cycle Toxicity Tests. Aquatic
Toxicology and Hazard Evaluation. ASTM STP 634. F.L. Mayer and
J.L. Hamelink, Eds. American Society for Testing and Materials,
pp. 117-126.
Hansen, D.J., S.C. Schimmel, and J. Forester. 1974. AroclorR 1254 in Eggs
of Sheepshead Minnows (Cyprinodon variegatus). Effect of Fertilization
Success and Survival of Embryos and Fry. Proc. 27th Ann. Conf. South-
east. Assoc. Game Fish Comm. Oct. 1973. Hot Springs, Arkansas: 420-426.
Hansen, D.J., S.C. Schimmel, and J. Forrester. 1977. Endrin: Effects on
the Entire Life-cycle of a Salt Water Fish. J. Toxicol. Environ. Health
3:721-733.
Hansen, D.J., P.R. Parrish, J.I. Lowe, A.J. Wilson, Jr., and g.D. Wilson.
1971. Chronic Toxicity, Uptake, and Retention of Aroclor 1254 in Two
Estuarine Fishes. Bull. Environ. Contarn. Toxicol. 6:113-119.
McKim, J.M., and D.A. Benoit. 1971. Effect of Long-term Exposures to
Copper on Survival, Growth, and Reproduction of Brook Trout (Salvelinus
fontinalis). J. Fish. Res. Board Can. 28(5):655-662.
Mount, Donald I. 1968. Chronic Toxicity of Copper to Fathead Minnows
(Pimephales promelus, Rafinesque). Water Research 2:21-29.
Mount, D.I., and C.E. Stephan. 1967. A Method for Establishing Acceptable
Toxicant Limits for Fish-malathion and the Butoxyethanol Ester of
2,4-D. Trans. Amer. Fish. Soc. 96(2):185-193.
116
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Mount, Donald I. and William Brungs. 1967. A Simplified Dosing Apparatus
for Fish Toxicology Studies. Water Research 2:21-29.
o
Schimmel, S.C. and D.J. Hansen. 1974. Effects of Aroclor 1254 on the
Embryo and Fry of Sheepshead Minnows. Trans. Amer. Fish. Soc. 103(3):
522-586.
Sprague, J.B. 1969. Review Paper: Measurement of Pollution Toxicity to
Fish. 1. Bioassay Methods for Acute Toxicity. Water Research 3(11):
793-821.
117
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O. FISH BRAIN ACETYLCHOLINESTERASE INHIBITION ASSAY
D. L. Coppage
INTRODUCTION
This procedure provides a method for determining the effect of materials
to be dumped in the ocean on acetylcholinesterase (AChE) in fish brains.
This test is appropriate for nerve poisons which disrupt nerve impulse
transmission by inhibiting AChE, the enzyme that modulates levels of the
neurotransmitter acetylcholine (Koelle, 1963; Karczmar, 1970). This pro-
cedure is not necessary for materials that contain no AChE inhibiting poisons,
It has been shown that brain-AChE of fishes is inhibited by in vivo
exposure to organophosphate and carbamate pesticides under laboratory con-
ditions (Weiss, 1958, 1961; Carter, 1971; Coppage, 1972). Furthermore,
environmental water pollution by these pesticides has been monitored by
measuring AChE activity in fish brains (Williams and Sova, 1966; Holland et
al., 1967; Coppage and Duke, 1971). Coppage (1971) defined the conditions
necessary for obtaining reliable and reproducible data in the laboratory
AChE assays and reported j.n vitro effects of four pesticides on AChE activ-
ity in brains of sheepshead minnows (Cyprinodon variegatus). Coppage and
Matthews (1974) further refined assay techniques and reported acute effects
°^ 111 vivo exposure to organophosphate pesticides on cholinesterases of four
estuarine fishes and a shrimp.
Recommended Procedure for Exposing Animals
Fish should be exposed to the material as recommended in the definitive
test of the continuous-flow method for acute toxicity tests, using fish and
macroinvertebrates as described in this manual. Fish to be assayed for AChE
should be from control aquaria and, if possible, from three contaminated
aquaria in which some fish have died. Live control fish should be divided
into three groups of three to six fish for assay. Three to six fish from
the contaminated aquaria should also be assayed by the method described
below.
Recommended Procedure for AChE Assay
Preparation of Fish Brains (3 to 6 brains are pooled for each
sample)—
1. Weigh 5 cm square of aluminum foil in following manner: pick up
and place foil on balance pan with forceps. (Fingers can leave enough
moisture to cause weight error at this low weight.) Weigh, then leave on
pan at full rest.
118
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2. Place another larger piece of foil in dissecting area.
3. Kill fish by placing them in a clean beaker containing acetone
for about 3 minutes.
4. Pour fish into clean sink or pan and scale heads under running
water with scalpel. (Check to ascertain that all dissecting equipment has
been cleaned and rinsed with acetone. As heads are scaled, place fish in
another beaker containing acetone.)
5. Pour fish into clean sink or pan, and then blot fish dry on
paper.
6. Clip top of skull from the brain with scissors.
7. After all skulls have been clipped, remove brains by pulling
off bone flap with forceps and digging bone and flesh away from spinal cord
with probe if necessary. Cut spinal cord about 2 mm behind brain.
8. Strip brain from optic nerves, and place on larger piece of
foil.
9. After removal, transfer brains with forceps to the preweighed
foil on the balance pan and determine weight in milligrams. Divide weight by
five.
10. Transfer weighed brains to nylon cup (see next section) and add
about 4 ml of distilled water.
11. Homogenize for 1 minute, then pour into graduated cylinder.
Rinse cup several times with distilled water and pour into cylinder.
12. Add distilled water to cylinder until total volume (in ml)
equals the amount found by dividing the brain weight by five. Pour this
into beaker to gently mix. Assay within 30 minutes after preparation.
Assay for AchE—
AChE activity should be determined by using an automated recording
pH stat to measure normal and irt vivo-inhibited brain AChE. The following
procedure applies: mix 2 ml of diluted brain homogenate with 2 ml of 0.03 M
acetylcholine iodide in distilled water; titrate the liberated acetic acid
with carbonate-free 0.01 N NaOH; carry out the reaction at pH 7 and 22"" C
while passing nitrogen over the liquid to prevent adsorption of atmospheric
carbon dioxide. Calculate the micromoles of substrate hydrolyzed per unit
of time from the number of raicromoles of NaOH required to neutralize the
liberated acetic acid per unit of time; express AChE activity as micromoles
of ACh hydrolyzed per hour per mg brain tissue in reaction vessel.
119
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Calculations and Reporting
Assay results of exposed and control fishes are compared, and percent-
ages of normal brain AChE activity of exposed fish are reported. Results
should be subjected to statistical analysis (Student's t-test, for example)
to determine statistical validity. Original control fish may be divided
into groups of five and brains pooled for each group of five to obtain
samples for normal AChE and statistical comparisons with exposed fish repli-
cates.
Report8
Any deviation from this method must be noted in all reports of results.
A report of the results of a test must include:
1. name of method, author, laboratory, and date tests were conducted;
2. a detailed description of the material tested. Including its source,
date and time of collection, composition, known physical and chemical
properties, and variability of the material tested;
3. the source of the salt water, its date and method of preparation;
4. detailed information about the test animals, including name, standard
length, weight, age, source, history, and acclimation procedure used;
5. a description of the experimental design, the test containers, the
volume of test solution, the number of organisms per concentration, and the
loading;
6. period of exposure and number of animals dead at end of exposure;
7. percentage of control organisms that died or were affected during
the test;
8. methods used for and the results of all test material, dissolved
oxygen, pH, and temperature measurements; and
9. any other relevant information.
120
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REFERENCES
Carter, F.L. 1971. In vivo Studies of Brain Acecylcholinesterase
Inhibition by Organophosphate and Carbamate Insecticides in Fish.
Unpublished Ph.D. dissertation, Louisiana State Univ., Baton Rouge,
Louisiana.
Coppage, D.L. 1971. Characterization of Fish Brain Acetylcholinesterase
With an Automated pH Stat for Inhibition Studies. Bull. Environ.
Contam. Toxicol. 6(4):304-310.
Coppage, D.L. 1972. Organophosphate Pesticides: Specific Level of Brain
AChE Inhibition Related to Death in Sheepshead Minnows. Trans. Am.
Fish. Soc. 101(3):534-536.
Coppage, D.L., and T.W. Duke. 1971. Effects of Pesticides in Estuaries
along the Gulf and Southeast Atlantic Coasts. In: Proceedings of the
2nd Gulf Coast Conference on Mosquito Suppression and Wildlife Manage-
ment, pp. 24-31. (C.H. Schmidt, Ed.) National Mosquito Control-
'Fish and Wildlife Management Coordinating Committee, Washington, D.C.
Coppage, D.L., and E. Matthews. 1974. Short-term Effects of Organophosphate
Pesticides on Cholinesterase of Estuarine Fishes and Pink Shrimp. Bull.
Environ. Contam. Toxicol. 11(5):483-488.
Holland, H.T., D.L. Coppage, and P.A. Butler. 1967. Use of Fish Brain
Acetylcholinesterase to Monitor Pollution by Organophosphorus
Pesticides. Bull. Environ. Contam. Toxicol. 2(3):156-162.
Karczmar, A.G. (Ed.). 1970. Anticholinesterase Agents. Perganon Press,
New York.
Koelle, G.B. (Ed.). 1963. Cholinesterases and Anticholinesterase Agents.
Springer-Verlag, Berlin.
Weiss, C.M. 1958. The Determination of Cholinesterase in the Brain Tissue
of Three Species of Fresh Water Fish and Its Inactivation ir± vivo.
Ecology 39:194-199.
Weiss, C.M. 1961. Physiological Effect of Organic Phosphorus Insecticides
On Several Species of Fish. Trans. Am. Fish. Soc. 90:143-152.
Williams, A.K. and R.C. Sova. 1966. Acetylcholinesterase Levels in Brains
of Fishes From Polluted Waters. Bull. Environ. Contam. Toxicol
1:198-204.
121
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TECHNICAL REPORT DATA
(Please read Instructions on the men* be fort compiettntt
EPA-600/9-76-010
|4. TITLE AND SUBTITLE
Bioassay Procedures for the Ocean Disposal
Permit Program
7. AUTHOR(S)
3 RECIPIENT'S ACCESSION NO
5 REPORT DATE
* PERFORMING ORGANIZATION COCt
s PERFORMING ORGANIZATION REPORT
|0. PERFORMING ORGANIZATION NAME AND ADDRESS
Environmental Protection Agency Ocean Disposal Bio-
assay Working Group
Environmental Research Laboratory, Gulf Breeze, FL.
10 PROGRAM ELEMENT NO
1EA714
\ C6NTRACY CRANt Ng
12. SPONSORING AGENCY NAME AND ADDRESS
Environmental Research Laboratory
Office of Research and Development
U.S. Environmental Protection Agency
Gulf Breeze, Florida 32561
13 TYPE OF REPORT AND PERIOD COVtRtO
In-house/Final
74 SPONSORING AGENCY COOC
EPA/600/08
The bioassay procedures given in this manual were developed to provide tests
for conducting toxicity evaluations of waste materials considered for ocean disposal
under EPA's Ocean Disposal Permit Program.
i 8Pecify the use of various organisms representing several troph'
levels. Both flow-through and static tests are included. Methods given varv Tn
utility and complexity of performance. The procedures are not intended to be con
sidered "standard methods," but, depending on the judg«nt of the EPA Regional Ad
istrator responsible for the managing of the permit program, are to be used a,
reference methods or official methods.
This manual is a revision of EPA-600/9-76-010 published May 1976.
7.
KEY WORDS AND DOCUMENT ANALYSIS
DESCRIPTORS
[b IDENTIFIERS.QPtN ENDED TERMS
Bioassay, Oysters, Marine fishes,
Algae, Crustacea
«. DISTRIBUTION STATEMENT
Unlimited
For* 2220-1 (R«». 4-77) PREVIOUS COITION n OBSOLETE
Bioassay procedures,
Ocean Disposal Permit
Program, Marine organisms
Marine phytoplankton,
Brine shrimp, Calanoid
copepods, Macroinverte-
brates, Fish brain
jCOSATt
6F
6T
-------
INSTRUCTIONS
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zation.
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17. KEY WORDS AND DOCUMENT ANALYSIS
(a) DESCRIPTORS - Select from the Thesaurus of Engineering and Scientific Terms the proper authorized terms that identify the rriaior
concept of the research and are sufficiently specific and precise to be used as index entries for cataloging.
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• PA Form 2220-1 (Rev. 4-77) (Reverie)
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