United States	Office of Research and	EPA'600 3-90'075
Environmental Protection	Development	September 1990
Agency	Washington, DC 20460
-&EPA Sheepshead Minnow and
Inland Silverside Larval
Survival and Growth
Toxicity Tests
Supplemental Report for
Training Videotape

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Supplemental Report for the
Sbeepshead Minnow and Inland Silverside
Larval Survival and Growth Toxicity Tests
Training Videotape
U.S. Environmental Protection Agency
Center for Environmental Research Information
26 West Martin Luther King Drive
Cincinnati, Ohio 45268
U.S. Environmental Protection Agency
Environmental Research Laboratory
South Ferry Road
Narragansett, RI 02882
This report has been reviewed by the U.S. Environmental Protection Agency and approved for
publication. Mention of trade names, products, or services is not, and should not be interpreted as
conveying official EPA approval, endorsement, or recommendation.
PROPERTY OF
ENVIRONMENTAL PROTECTION AGENCY

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TABLE OF CONTENTS
page
Introduction
1
Care and Feeding of Adults and Larvae
2
Beginning the Test
4
Test Solution Renewal
6
Terminating the Test
8
Final Notes
9
References
10
Appendix A - Preparing Hypersaline Brine
Appendix B - Preparing Brine Shrimp and Rotifers for Feeding
Appendix C - Materials Needed for Testing
Appendix D - Summary of Test Conditions
Appendix E - Sample Data Sheets

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Sheepshead Minnow and Inland Silverside
Larval Survival and Growth Toxicity Tests
INTRODUCTION
The U.S. Environmental Protection Agency's
(EPA's) Environmental Research Laboratory in Duluth,
Minnesota, has developed a series of freshwater toxicity
tests to evaluate effluent toxicity. The tests use freshwater
fish, invertebrates, and plant species as indicators of toxic
effects to lakes, streams, and rivers. The EPA
Environmental Research Laboratory in Narragansett,
Rhode Island (ERL-N), has adapted the freshwater toxicity
testing methods and developed new methods to perform
similar tests for marine or estuarine environments.
This report summarizes methods developed at the
Narragansett laboratory for measuring effects on larval
survival and growth of the sheepshead minnow Cyprinodon
variegatus and the inland silverside Menidia beryllina after
exposure to complex effluents in marine or estuarine
environments. These are short-term tests that span an
exposure time of 7 days, and estimate the chronic toxicity
to larvae in a static renewal exposure system. The
methods described in this report and demonstrated in the
accompanying tape are detailed in the EPA methods
manual, "Short-term Tests for Estimating the Chronic
Toxicity of Effluents and Receiving Waters to Marine and
Estuarine Organisms" (EPA/600/4-87/028).
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CARE AND FEEDING OF ADULTS AND LARVAE
Sheepshead Minnows
ID J	'i
Embryonic development of the
sheepshead minnow (Cyprinidon
varieeatus): A 48-hour embryo;
B. 72-hour embryo; C. newly
hatched fish, actual length 4 mm;
D. 5-day fish (5 mm); E. young
fish (9 mm); F. young fish
(12 mm) (Short-term Methods
manual, p. 103).
Adult sheepshead minnows can be field collected
from Atlantic and Gulf of Mexico coastal estuaries south
of Cape Cod using near-shore nets, purchased from
commercial biological supply houses, or raised from young
fish to maturity in the laboratory. To minimize inbreeding,
use of feral brood stocks or first generation laboratory fish
is recommended. Adult sheepshead minnows are kept in
natural or artificial seawater in a flow-through or
recirculating aerated glass aquarium, and are fed flake food
three or four times a day.
Keep holding tanks at 25 C until the fish reach
sexual maturity and can be used for spawning. When the
fish are sexually mature, maintain the water temperature at
18 to 20 C. About 1 week before the start of the test,
induce the minnows to spawn naturally by raising the
system temperature to 25 C. The females also can be
induced to spawn artificially by intraperitoneal injection
with human chorionic gonadotrophin hormone, though
natural spawning is preferred so that repeated spawnings
can be obtained from the same brood stock. Transfer the
adults to a spawning chamber, or basket, within an
aquarium outfitted with a mesh screen on the bottom.
The eggs will fall through the basket onto the mesh
collecting screen. Collect eggs daily, washing the eggs off
of the screen into a large tray. Strain the collected water
through a 22-micron mesh screen. Roll the eggs gently on
the screen, pressing any food or waste through, leaving the
eggs on top of the screen. Store the collected eggs in
seawater at 25 C.
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Incubate the minnow embryos for 5 to 6 days with
aeration and daily water changes at 30 ppt salinity.
Lowering the salinity or raising the temperature may help
to induce the embryos to hatch. For the sheepshead
minnow growth and survival test, use larvae that hatch less
than 24 hours before the start of the test
Inland Silverside
Inland silversides also can be obtained by beach
seine from Atlantic and Gulf of Mexico coastal estuaries,
from biological supply houses, or by raising young fish in
the laboratory. Only natural seawater is recommended for
the culture and maintenance of the more sensitive
silverside brood stock. Adults should be fed flake food or
frozen brine shrimp twice each day and Artemia nauplii
once each day. Maintain holding and spawning tanks at
25 C.
Embryonic development of the
inland silverside (Menidia
bervlina): A. egg; B. larvae,
actual length 8 mm; C. fry, actual
length 13 mm; D. adult fish
(Fishery Bulletin No. 74 of the
Fish and Wildlife Service, Vol 53,
1953, p. 303)
Inland silversides are encouraged to spawn by
placing polyester aquarium filter fiber in the tanks. When
the fish spawn into the fiber, the hard, light yellow
embryos can be separated from the fibers by hand, or the
eggs and fiber can be placed together into a 10-gallon
aquarium. Larvae will hatch and free themselves from the
fibers. The larvae are then easily identified and should be
removed. Feed silverside larvae the rotifer Brachionus
plicatilis until 6 days post-hatch, and the smallest brine
shrimp nauplii available beginning on day 5. Use 7- to 11-
day-old larvae for the silverside growth and survival test.

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Note:
The growth and survival tests use 10 to 15 larvae for
each replicate. If there are 5 test concentrations plus 1
control, and 3 replicates per concentration, 180 to 270
larvae will be needed for a complete test.
BEGINNING THE TEST
Store the effluent or receiving waters in an
incubator or refrigerater at 4 C until the tests begin, but
not longer than 48 hours. Prepare dilutions of the
effluent sample using a 0.3 or 0.5 dilution factor. The
tests require about 3 liters of each test solution each day,
enough for renewing 3 replicates of each concentration
and performing chemical analyses. It is essential to
maintain constant salinity among treatments and treatment
replicates throughout the duration of the test. Use
concentrated seawater or hypersaline brine to keep the
salinity of the solutions between 20 and 30 ppt for the
sheepshead minnows, and between 5 and 30 ppt for the
inland silversides. Before adding the solutions to the test
chambers, warm the samples to 25 C in a water bath.
Set out the test chambers. There will generally be
at least 5 dilutions plus 1 control, and 3 or 4 replicates of
each. ERL-N uses glass chambers equipped with a
screened-off sump area; or 1,000-ml glass or disposable
plastic beakers can also be used as test chambers. Add a
small amount of clean seawater to each chamber, enough
to cover the bottom to a depth of about 1 cm. Pipet 2 or
3 larvae at a time into each chamber, adding larvae to all
chambers; then start again to add more, until each
chamber contains the required number of larvae - a
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minimum of 10 and a maximum of IS larvae. Use a
minimum amount of seawater to deposit the animals into
the containers to avoid diluting the effluent samples
further. Using a white background or a light table
facilitates counting the larvae in the chambers. Since
clean seawater is in all of the chambers, larvae can be
exchanged among test chambers until all contain the
correct number. Because the inland silverside larvae are
sensitive to handling, it may be best to distribute the
animals into chambers containing control solution lday
before the start of the exposure period.
Randomly apply colored labels to the chambers to
indicate treatment and replicates, then fill each chamber
with approximately 750 ml of the appropriate test solution,
pouring through the sump area or down the side.
Measure the initial temperature, salinity, and dissolved
oxygen in each chamber. Record all measurements on the
test data sheet. Copies of the data sheets used at ERL-N
are provided in Appendix E.
When all measurements have been taken and
recorded, place the chambers in a 25 C water bath
according to a random numbers table. Keep the chambers
in those same positions for the duration of the test. Feed
the fish Artemia nauplii. The initial amount for feeding is
2 ml of a solution made from 4 ml of concentrated
Artemia nauplii in 80 ml of seawater. Cover the chambers
during incubation to prevent evaporation of the test
solutions.
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TEST SOLUTION RENEWAL
Each day, the test and control solutions must be
replenished. Prepare new dilutions daily from effluent
stored at 4 C. When tests are performed on site, effluent
and receiving water should be collected daily. Off-site
toxicity tests are often performed with effluent collected
on days 1, 3, and 5 of the exposure period. Again, do not
store the effluent samples longer than 48 hours before use.
Warm the solutions to 25 C in a water bath just before
adding to the chambers.
Temperature and salinity should be maintained
under carefully controlled conditions across all test
concentrations and replicates throughout the test. Each
day before changing the solutions, measure and record the
temperature in each chamber. Maintain the chambers at
25 +. 2 C, and supply 14 hours of ambient laboratory light
and 10 hours of darkness each day for both species.
Measure and record the salinity from each chamber every
day as well, before renewing the test solutions. Note that
there should be no more than a 2 ppt salinity difference
between any two chambers on a given day. If receiving
water and effluent tests are conducted concurrently,
conduct the tests at similar salinities. Monitor dissolved
 '
oxygen concentrations each day, recording the reading. If
dissolved oxygen falls below 40% saturation in any one of
the exposure chambers, all samples must be aerated.
Before changing the test solutions, count and record
the number of live larvae in each replicate, discarding any
dead animals. Then clean any uneaten Artemia from the
chamber using a siphon or a large pipet. To avoid
removing test animals along with uneaten food, set the
chambers on a light box or light table to better observe
the larvae. Besides making the larvae more visible, the
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light also serves to concentrate the nauplii on the bottom
of the chamber. Siphon the water and remaining Artemia
into a large beaker or white plastic tray. Individual larvae
that are accidentally removed can be seen easily in the
beaker, and should be returned to their test chambers.
Note the accidental siphoning of any larvae in the test
records. Once the solution in the test chamber is emptied
to a depth of 7 to 10 mm, slowly and carefully add
approximately 500 to 750 ml of new test solution, pouring
down the side of the chamber or into the sump area to
avoid excessive turbulence. After changing all the
solutions, return the chambers to the water bath and feed
the larvae.
Proven quality Artemia nauplii should be used to
feed the larvae daily throughout the test. Two
concentrations of prepared nauplii are used sequentially
during the exposure period. Detailed instructions for
culturing Artemia are included in Appendix B. The first
solution consists of 4 ml concentrated Artemia nauplii in
80 ml seawater. Feed the larvae 2 ml of this test solution
on the first 2 days of the test For days 3 through 6 of
the test, feed the fish 2 ml of a more concentrated
solution of 6 ml concentrated Artemia in 80 ml of
seawater.
Swirl the Artemia suspension while distributing food
to the test chambers; it is important that all chambers
receive the same amount of food throughout the test. If
the survival rate in any chamber falls below 50 percent,
reduce the amount of food supplied to that chamber by
1/2 for the remainder of' the test Cover the chambers
between feedings.
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TERMINATING THE TEST
At the end of the test, on day 7, the larvae are
counted to determine survival rate. Do not feed the
larvae on the seventh day. Working with groups of
replicates, remove any dead larvae from the chambers,
carefully recording the number of surviving animals.
Record the final temperature, salinity, and dissolved oxygen
for each chamber.
Pour the contents of the chamber into a 500-micron
mesh screen over a large beaker. Quickly submerge the
screen in an ice and deionized water bath. The cold will
immobilize the fish, and swirling the screen in the
deionized water will wash away uneaten Artemia and salts
that may interfere with the weight determination. Then
either dry the animals for immediate weighing or preserve
them for later drying in separate scintillation vials
containing 4% formalin or 70% ethanol. To dry the
surviving animals, place the fish from each replicate into a
pre-weighed aluminum weighing boat, and dry the fish at
60 C for 24 hours, or at 105 C for 6 hours.
After drying, and until they are weighed, place the
dried larvae directly into a dessicator to prevent moisture
from the air from adsorbing to the samples. Weigh each
sample to the nearest 0.01 mg. Determine the weight of
the larvae alone by subtracting the weight of the weigh
boat. Divide the final dry weight by the number of larvae
in the sample to determine the average dry weight of the
surviving larvae. This average weight is then compared
statistically to the control animals' average weight to
identify any effluent effects on the animals' growth.
For the test to be considered acceptable, control
animals' survival must be _> 80 percent for both species.
The average dry weight of unpreserved control larvae must
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be greater than or equal to 0.60 mg for the sheepshead
minnow, and 0.50 mg for the inland silverside. Minimum
dry weights for preserved animals are 0.50 mg for the
sheepshead minnow and 0.43 mg for the inland silverside.
FINAL NOTES
	Keep careful records throughout the test.
	Record any deaths and whether any larvae
were accidentally siphoned out of their
chamber.
	Take special note of any behavioral changes
that the larvae may exhibit, or any physical
abnormalities.
	Note the results of the chemical and physical
measurements taken during the test.
This information should all be carefully compiled, and
considered important clues to how the effluent may affect
marine animals. The methods manual, "Short-term
Methods for Estimating Chronic Toxicity of Effluents and
Receiving Waters to Marine and Estuarine Organisms,"
details the procedure for data analysis.
The larval survival and growth toxicity tests described
here are currently used to assess the potential toxic effects
of complex chemical mixtures on marine and estuarine
organisms. Used in conjunction with chemical-specific
methods, these tests can provide a comprehensive and
effective approach to assessing the impact of complex
effluents discharged to the marine and estuarine
environments.
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REFERENCES
Aquatic Toxicity Testing Seminar Manual. 1985. U.S. EPA Environmental Research
Laboratory, Narragansett, RI. ERL-N Contribution No. 796.
(Provides detailed descriptions of the methods used at Narragansett to evaluate the
toxicity of discharges to marine and estuarine waters. Served as the basis of a series of
seminars conducted by ERL-N personnel.)
Biomonitoring for Control of Toxicity in Effluent Discharges to the Marine Environment. 1989.
U.S. EPA Center for Environmental Research Information, Cincinnati, OH; U.S. EPA
Environmental Research Laboratory, Narragansett, RI. EPA/625/8-89/015.
(Describes the use of biomonitoring as an effective water quality-based approach to
controlling the toxicity of discharges to estuarine and marine waters. Covers regulatory
background, testing methods, and case studies.)
Short-term Methods for Estimating the Chronic Toxicity of Effluents and Receiving Waters to
Marine and Estuarine Organisms. 1987. Environmental Monitoring and Support Laboratory,
Cincinnati, OH. EPA/600/4-87/028.
(Describes methods, quality assurance, laboratory safety, facilities and equipment, data
analysis, report preparation, and organism culture and handling for six short-term tests to
estimate the chronic toxicity of effluents and receiving waters.)
Technical Support Document for Water Quality-based Toxics Control. 1985. U.S. EPA Office
of Water Enforcement and Permits, Washington, D.C.
f
(Provides guidance for each step in the water quality-based toxics control process, from
screening to compliance monitoring.)
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APPENDIX A
PREPARING HYPERS ALINE BRINE
BACKGROUND
Salinity adjustments are a vital part of using marine and estuarine species for toxicity
testing. The majority of industrial and sewage treatment effluents entering marine and
estuarine waters contain little or no measurable salts. Therefore, the salinity of these effluents
must be adjusted before exposing estuarine or marine plants and animals to the solutions. In
addition, it is important to maintain constant salinity across all treatments throughout the test
for quality control. Finally, matching the test solutions' salinity to the expected receiving
water's salinity may require salinity adjustments. ERL-N uses hypersaline brine, prepared from
filtered natural seawater, to adjust exposure solution salinities. Note that commercially available
artificial sea salts have not been sufficiently tested, and therefore are not recommended for all
of the subchronic toxicity tests at this time.
Hypersaline brine has several advantages over artificial sea salts that make it more suitable
for use in toxicity testing. Concentrated brine derived from natural seawater contains the
necessary trace metals, biogenic colloids, and some of the microbial components necessary for
adequate growth, survival, and/or reproduction of test organisms. It may be held for prolonged
periods without any apparent degradation. Brine may be added directly to the effluent to
increase the salinity, or may be used as control water by diluting to the desired salinity with
deionized water. The brine can be made from any high quality, filtered seawater supply
through simple heating and aerating.
GENERATING THE BRINE
The ideal container for making brine from natural seawater has a high surface-to-volume
ratio, is made of a non-corrosive material, and is easily cleaned. Shallow fiberglass tanks are
ideal.
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Collect high quality (and preferably high salinity) seawater on an incoming tide to minimize
the possibility of contamination. Special care should be used to prevent any toxic materials
from coming in contact with the seawater. The water should be filtered to at least 10 um
before placing into the brine tank. Thoroughly clean the tank, aeration supply tube, heater,
and any other materials that will be in direct contact with the brine before adding seawater to
the tank. Use a good quality biodegradable detergent, followed by several thorough deionized-
water rinses. Fill the tank with seawater, and slowly increase the temperature to 40 C. If a
heater is immersed directly into the seawater, make sure that the heater components will not
corrode or leach any substances that would contaminate the brine. A thermostatically
controlled heat exchanger made from fiberglass works well.
Aeration prevents temperature stratification and increases the rate of evaporation. Use an
oil-free air compressor to prevent contamination. Evaporate the water for several days,
checking daily (ormore or less often, depending on the volume being generated) to ensure that
the salinity does not exceed 100 o/oo and the temperature does not exceed 40 C. If these
changes are exceeded, irreversible changes in the brine's properties may occur. One such
change noted in original studies at ERL-N was a reduction in the alkalinity of seawater made
from brine with salinity greater than 100 o/oo, and a resulting reduction in the animals' general
health. Additional seawater may be added to the brine to produce the volume of brine desired.
When the desired volume and salinity of brine is prepared, filter the brine through a 10-
um filter and pump or pour it directly into portable containers (5-gallon Cubitainers or
polycarbonate water cooler jugs are most suitable). Cap the containers, and record the
measured salinity and the date the brine was generated. Store the brine in the dark at room
temperature until used.
SALINITY ADJUSTMENTS USING HYPERSALINE BRINE
To calculate the volume of brine (Vb) to add to 0 o/oo sample to produce a solution at a
certain salinity (Sf), use this equation:
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Vb * Sb = Sf * V,
where Vb	=	volume of brine, ml
Sb	=	salinity of brine, o/oo
Sf	=	final salinity, o/oo
V,	=	final volume, ml (brine brought to this volume with 0 o/oo sample).
To calculate the volume of brine (Vb) required to raise the salinity of an effluent or
receiving water sample (S.) to a certain salinity (Sf), use this equation:
Vb = [(Vf * Sf) - (V, * S.)]/(Sb - Sr)
where Vb	=	volume of brine, ml
Sb	=	salinity of brine, o/oo
V,	=	volume of sample, ml
S,	=	salinity of sample, o/oo
Sf	=	final salinity , o/oo
Vf	=	final volume, ml (final volume is combined brine and deionized water plus the
sample volume; percent original sample in the final sample = VJVt * 100).
The table on the next page gives volumes needed to make 20 o/oo test solutions from effluent
(0 o/oo), deionized water, and 100 o/oo hypersaline brine.
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Quantities of effluent, deionized water and a hypersaline brine of 100 o/oo (only) needed for
conducting daily renewals of test solutions at 20 o/oo salinity.
Exposure

Deionized
Hypersaline
Concentration
Effluent
Water
Brine

(0 o/oo)

(100 o/oo)
(%)
(ml)
(ml)
(ml)
32
640
960
400
10
200
1,400
400
3.2
65
1,535
400
1.0
20
1,580
400
0.32
7
1,593
400
Control
~
1,600
400
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APPENDIX B
PREPARING BRINE SHRIMP AND ROTIFERS
FOR FEEDING
INTRODUCTION
The brine shrimp (Artemia sp.) is used to feed larval Menidia beryllina and Cyprinodon
variegatus in the 7-day effluent toxicity tests. At hatching, however, M beryllina are too small
to ingest Artemia, and must be fed rotifers (B. plicatilis). Preparation and culture of rotifers is
described at the end of this Appendix.
Brine shrimp are highly suited to this testing protocol because: 1) the naupliar stages are
nutritionally acceptable to these species; 2) they may be obtained from cysts within 24 hours
after immersion in seawater; and 3) the cysts are readily available and can be stored for
prolonged periods of time. There are some disadvantages to keep in mind, as well. For
example, it may be difficult to obtain large quantities of cysts. In addition, the shrimp's
nutritional quality may vary considerably from batch to batch because they are obtained from
diverse geographical areas.
Rates of fish growth and survival differed when fed strains of brine shrimp from various
geographic locations (Klein-MacPhee, 1982; Johns et al., 1981; Leger and Sorgeloos, 1984).
Reference brine shrimp have, therefore, been recommended for use in toxicity testing or as a
standard for comparison against other geographic strains of brine shrimp (Sorgeloos, 1981).
Brine shrimp normally hatch after incubation for 24 to 48 hours at room temperature.
Different geographical strains may differ somewhat in time-to-hatch (Vanhaecke and Sorgeloos,
1983) and may diminish in nutritional quality after 48 hours (Vanhaecke et al., 1983).
Therefore, it is important to harvest the nauplii as soon as possible after approximately 90%
have hatched.
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CULTURING ARTEMIA
A batch of cysts should be started every 24 hours (for feeding the following day) with the
same proportion of cysts to seawater so that consistent densities of nauplii are obtained daily
(Persoone et al., 1980).
1.	Fill a 2- to 4-liter separatory funnel (or other appropriate container) with enough 25-
30C sea water to ensure adequate hatching. Add 10 cc brine shrimp cysts per liter, and aerate
for at least 24 hours at 25C. (Two separatory funnels are recommended, started on alternate
days, since it may require more than 24 hours to hatch certain strains of brine shrimp.)
2.	Nauplii will hatch from brine shrimp cysts within 24 to 48 hours, but before nauplii are
fed to the fish they should be separated from the cysts by taking advantage of their phototactic
response or by straining the culture. After removing the source of air, the nauplii's phototactic
response is stimulated by covering the top of the funnel with a dark cloth or paper towel for 5
minutes. The nauplii will concentrate at the bottom, however, leaving nauplii longer than 5
minutes without aeration may cause mortality. Another way to stimulate phototactic response is
to rinse the nauplii into a beaker (500 ml) or a black separator box (15 x 8 x 8 cm high), place
a light source at one end, and leave for no more than 10 to 15 minutes. After live nauplii
migrate toward the light, they can be pipetted or siphoned out of the container, leaving the
unhatched cysts behind. The nauplii can also be separated from the cysts using a sieve.
3.	Pour the nauplii onto a nylon screen (mesh <150 um), rinse with filtered control
seawater, and drain off most of the water.
4.	On days 0, 1, and 2, weigh 4 g (wet weight) or pipette 4 ml of concentrated, rinsed
Artemia nauplii from the quantity of Artemia on the screen. On days 3 through 6, weigh 6 g
(wet weight) or pipette 6 ml nauplii from the quantity of Artemia on the screen. Resuspend
the Artemia in 80 ml of sea water in a 100 ml beaker. Aerate or swirl the Artemia to equally
distribute the nauplii; then withdraw and dispense individual 2 ml portions of Artemia to each
test chamber using a pipette or adjustable syringe. Uniform distribution of food to all replicates
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is critical to minimize the variability of larval weight, important for successful tests. If the
replicate chambers are subdivided, divide the 2 ml equally among the compartments; if the
survival rate of any replicate on any day falls below 50%, reduce the volume of Artemia
dispensed to that replicate by one-half.
Some live Artemia should remain overnight in test chambers, however, excessive Artemia
may decrease dissolved oxygen concentrations to below the acceptable limit. Siphon the
uneaten Artemia from each chamber prior to test solution renewal to ensure that the fish larvae
eat principally newly-hatched nauplii.
BRINE SHRIMP QUALITY CONTROL
At a minimum, each batch of purchased brine shrimp should be tested to ensure that they
provide the nutrients necessary for adequate fish growth. Before use, individual lot numbers of
cysts are fed to the test organisms in 7-day studies to confirm that the diet is adequate for the
purposes of the test. The shelf-life of an opened container of cysts may be affected by
humidity and temperature, so they should be tested each time a test is started. As long as
more than 90% of the cysts hatch in 24 to 48 hours and the control responses are acceptable,
the cysts may be used (refer to the EPA manual, "Short-term Methods for Estimating the
Chronic Toxicity of Effluents and Receiving Waters in Marine and Estuarine Organisms" for
acceptability parameters).
PREPARING BRACHIONUS PLICATILIS
M. beryllina larvae are too small at hatching to ingest Artemia and must be fed rotifers (B.
plicatilis). Brachionus plicatilis can be cultured continuously when fed algae or yeast in 10- to
15-liter Pyrex carboys at 25 to 28C, 25 to 35 o/oo salinity. Four 12-liter culture carboys, with
an outflow spout near the bottom, should be maintained simultaneously to optimize production.
Fill clean carboys with autoclaved seawater. (Alternatively, heat filtered seawater by
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placing an immersion heater in the carboy, and maintain the temperature at 70 to 80 C for 1
hr.). When the seawater has cooled to 25 to 28 C, aerate and add a start-up sample of
rotifers (50 rotifers/mL) and food (about 1 L of a dense algal culture or 0.1 g yeast per liter of
seawater). Yeast should be dissolved in a minimum of tap water or deionized water before
adding it to the culture.
Check the carboys daily to ensure that adequate food is available and that the rotifer
density is adequate. If the water appears clear, add yeast (0.1 g/L) or remove 1 L of water and
replace it with algae. Remove the water via the bottom spigot, filtering it through a < 60 um
mesh screen. Return any rotifers collected on the screen to the culture. Keeping the carboys
away from light will reduce the amount of algae that attaches to the carboy walls. If detritus
accumulates, populations of ciliates, nematodes, or harpacticoid copepods that may have been
inadvertently introduced can rapidly take over the culture. If this occurs, discard the cultures.
If a precise measure of the rotifer population is needed, resuspend rotifers collected from
a known volume of water in a smaller volume, preserve them with formalin, and count them in
a Sedgwick-Rafter chamber. As the density exceeds 50 rotifers/ml, the amount of food per day
should be increased to 2 L of algae or 0.2 g/L of yeast. The optimum density, 300 to 400
rotifers/ml, will be reached in about 7 to 10 days. This density is sustainable for 2 to 3 weeks.
Once that is attained, the rotifers should be cropped daily.
About 5 days after hatching, M. beryllina larvae can begin feeding on newly hatched
Artemia nauplii; they are fed Artemia daily throughout the 7-day test.
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REFERENCES
Johns, D.M., W.J. Berry, and W. Walton. 1981. International Study on Artemia, XVI.
Survival, growth, and reproduction potential of the mysid, Mysidopsis bahia Molenock fed
various geographical collections of the brine shrimp, Artemia. J. Exp. Mar. Biol. Ecol. Vol.
53, pp. 209-219.
Klein-MacPhee, G., W.H. Howell, and A.D. Beck. 1982. International Study on Artemia, XX.
Comparison of a reference and four geographical strains of Artemia as food for winter
flounder (Pseudopleuronectes americanus) larvae. Aquaculture. Vol. 29, pp. 279-288.
Leger, P., and P. Sorgeloos. 1984. International Study on Artemia, XXIX. Nutritional value of
Artemia nauplii from various geographical origins for the mysid Mysidopsis bahia
(Molenock). Mar. Ecol. Prog. Ser. Vol. 15, pp. 307-309.
Sorgeloos, P. 1981. Availability of reference Artemia cysts. Aquaculture. Vol. 23, pp. 381-
382.
Vanhaecke, P., P. Lavens, and P. Sorgeloos. 1983. International Study on Artemia, XVII.
Energy consumption in cysts and early larval stages of various geographical strains of
Artemia. Ann. Soc. R. Zool. Belg. Vol. 113, pp. 155-165.
Vanhaecke, P., and P. Sorgeloos. 1983. International Study on Artemia, XIX. Hatching data
for ten commercial sources of brine shrimp cysts and revaluation of the "hatching
efficiency" concept. Aquaculture. Vol. 30, pp. 43-52.
B-5

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APPENDIX C
MATERIALS NEEDED FOR TESTING
Lab Supplies
2-liter beakers for test solutions
1-liter	graduated cylinders for control and effluent solutions
250-mI graduated cylinders for control and effluent solutions
18 aluminum foils or weigh boats/test in larger marked aluminum boats
Marked vials containing 4% formalin (optional)
Forceps
Eyedroppers
Dessicator with dessicant
18 exposure chambers/test
Colored tape
Water-proof markers
Air lines
Air stones
Crystallization dishes
Thermometer
2-	or 4-liter separatory funnels for Artemia
Mesh seives (150 and 505 um)
1 to 5-ml pipet with siphon bulb or adjustable syringe for feeding
Squirt bottle for deionized water
Siphon with bulb and clamp
3-liter	beakers for effluent drainage
Standard or micro-Winkler apparatus
Refractometer (salinometer)
Data sheets (Appendix E)
C-l

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Lab Equipment
Air pump
Dissecting microscope
Light table
Dissolved oxygen meter
MATERIALS FOR MAINTAINING TEST ORGANISMS
Standard salt water aquarium w/air pump
Artemia (brine shrimp) cysts (see Appendix B)
B. plica tills (rotifers) for larvae (M.beryllina)
C-2

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APPENDIX D
SUMMARY OF TEST CONDITIONS
Sheepshead Minnow
Test duration:
Effects measured:
Number of test organisms
per test chamber:
Number of replicate
chambers per treatment:
Test type:
Salinity:
Temperature:
Photoperiod:
Light intensity:
Light quality:
Test solution volume:
Test chamber size:
Renewal of test solutions:
Age of test organisms:
Number of treatments per study:
Source of food:
7 days
Survival and growth
10 to 15 larvae/replicate
Minimum of 3
Static, with 24-hr renewal
20 o/oo to 32 o/oo +. 2 o/oo
25 + 2C
14 h light:10 h dark
50 to 100 ft-candles
Covered, soft white light
Minimum of 50 ml/larvae (approximately
750 ml per replicate)
Glass dish at least 8.0 cm high, to contain test
solution volume to a depth of >. 5.0 cm, preferably
with a screened-off sump area
Daily
<. 24-hours old
Minimum of 5 effluent concentrations and a
control
<. 24-hour-old Artemia nauplii
D-l

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Feed 0.10 g wet weight Artemia nauplii per
replicate on days 0 to 2, 0.15 g per replicate on
days 3-6
Siphon daily, before renewal and feeding
None, unless D.O. concentration falls below 40
percent of saturation, then aerate all chambers
Uncontaminated seawater; deionized water mixed
with hypersaline brine
Inland Silverside
Sheepshead Minnow Test Conditions, cont
Feeding regime:
Cleaning:
Aeration:
Dilution water:
Test duration:
Effects measured:
Number of test organisms
per test chamber:
Number of replicate
chambers per treatment:
Test type:
Salinity:
Temperature:
Photoperiod:
Light intensity:
Light quality:
Test solution volume:
7 days
Survival and growth
10 to 15 larvae/replicate
Minimum of 3
Static, with 24-hr renewal
5 o/oo to 32 o/oo +. 2 o/oo
25 2C
14 h light: 10 h dark
50 to 100 ft-candles
Covered, soft white light
Minimum of 50 ml/larvae (approximately
750 ml per replicate)
D-2

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Inland Silverside Test Conditions, cont
Test chamber size:
Renewal of test solutions:
Age of test organisms:
Number of treatments per study:
Source of food:
Feeding regime:
Cleaning:
Aeration:
Dilution water:
Glass dish at least 8.0 cm high, to contain
test solution volume to depth of >. 5.0 cm,
preferably with a screened-off sump area
Daily
7 to 11 days post-hatch
Minimum of 5 effluent dilutions and one or
more controls
B. plicatilis (rotifers), newly hatched
Artemia nauplii
Feed rotifers from hatching until five days
old. During test, feed 0.10 g wet weight
Artemia nauplii per replicate on days 0 to 2,
0.15 g per replicate on days 3 to 6.
Siphon daily, before renewal and feeding.
None, unless D.O. concentration falls below
40 percent of saturation, then aerate all
chambers.
Uncontaminated seawater; deionized water
mixed with hypersaline brine.
D-3

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APPENDIX E
DATA SHEETS

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Fish Growth/Survival
Study
Test Dates:	
Date of Hatch:	
Type of Effluent:.
Effluent Tested:	
Species:	
Field:	Lab:
.Test:
Comments:
Treatment: Control
Rep: a
Pan#:
Pan wt.:
Day
0
1
2
3
4
5
6
7
Live








Temp. (C)








Salinity (%>)








D.O. (ma/1)








#Fish:
Wt., Fish + Pan:

Rep: b
Pan#:
Pan wt.:
Day
0
1
2
3
4
5
6
7
Live








Temp. (C )








Salinity (<&>)








D.O. (maA)








# Fish:
WtM Fish + Pan:

Rep: c
Pan#:
Pan wt.:
Day
0
1
2
3
4
5
6
7
Live








Temp. (C)








Salinity (<&>)








D.O. (mg/l)








#Fish:
Wt., Fish + Pan:
Treatment:
Rep: a
Pan#:
Pan wt.:
Day
0
1
2
3
4
5
6
7
Live








Temp. (C)








Salinity (<&>)








D.O. (ma/1)








# Fish:
Wt, Fish + Pan:

Rep: b
Pa
a#:
Pan wt.:
Day
0
1
2
3
4
5
6
7
Live








Temp. (C)








Salinity (<&>)








D.O. (ma/1)








# Fish:
Wt., Fish + Pan:

Rep: c
Pan#:
Pan wt.:
Day
0
1
2
3
4
5
6
7
Live








Temp. (C)








Salinity (*&>)








D.O. (maA)








#Fish:
Wt., Fish + Pan:
Day
0
1
2
3
4
5
6

Time Fed







Amt Fed








-------
Page 2
Test Dates:		 Species:
Effluent Tested:		
Treatn
lent:
Rep: a
Pa

Pan wt.:
Day
0
1
2
3
4
5
6
7
Live








Temp. (C)








Salinity (<&>)








D.O. (ma/tf








#Fish:
Wt., Fish + Pan:

Rep: b
Pa
ntf:
Pan wt.:
Day
0
1
2
3
4
5
6
PM
i
Live








Temp. (C)








Salinity (<&>)








D.O. (mg/l)








#Fish:
Wt., Fish + Pan:

Rep: c
Pa
ntf:
Pan wt.:
Day
0
1
2
3
4
5
6
7
Live








Temp. (C)








Salinity (<&>)








D.O. (mg/l)








#Fish:
Wt., Fish + Pan:
Treatment:
Rep: a |Pa
a#:
Pa
n vt.:
Day
0
1
2
3
4
5
6
7
Live








Temp. (C)








Salinity (*&>)








D.O. (mg/l)








# Fish:
Wt., Fish + Pan:

Rep:b
Pa
n#:
Pan wt.:
Day
0
1
2
3
4
5
6
7
Live


i





Temp.(C)








Salinity








D.O. (ma/1)








 Fish:
WtM Fish + Pan:

Rep: c
Pa
a#:
Pan wt.:
Day
0
1
2
3
4
5
6
7
Live








Temp.(C)








Salinity








D.O. (ma/I)








# Fish:
Wt., Fish + Pan:
Comments:

-------
Page 3
Test Dates:	 Species:
Effluent Tested:		
Treatn
lent:
Rep: a
Pa
i#:
Pa
a wt.:
Day
0
1
2
3
4
5
6
7
Live








Temp. (c)








Salinity (<&>)








D.D. (ma/1)








#Fish:
Wt., Fish + Pan:

Rep: b
Pa
Q#:
Pan wt.:
Day
0
1
2
3
4
5
6
7
Live








Temp. (C)








Salinity (*&>)








D.O. (ma A)








# Fish:
Wt, Fish + Pan:

Rep: c
Pa
n#:
Pa
n wt.:
Day
0
1
2
3
4
5
6
7
Live








Temp. (c)








Salinity (<&>)








D.O. (mg/l)








#Fish:
Wt., Fish + Pan:
Treatment:
Rep: a
Pa
a#:
Pan ivt.:
Day
0
1
2
3
4
5
6
7
Live








Temp.(C)








Salinity (%>)








D.O. (ma A)








# Fish:
Wt., Fish + Pan:

Rep: b
Pa
n#:
Pan wt.:
Day
0
1
2
3
4
5
6
7
Live



A




Temo. (C)








Salinity C&)








D.O. (ma/1)








# Fish:
Wt., Fish + Pan:

Rep: c
Pa
n#:
Pa
l wt.:
Day
0
1
2
3
4
5
6
ri
Live








Temn. (C)








Salinity








D.O. (ma/1)








#Fish:
Wt., Fish + Pan:
Comments:

-------
Page 4
Test Dates:	 S pecies:
Effluent Tested:	
Treatment






Number
Live






Percent
Live






Ave. Vt./
Larvae (mg]






Sig. Diff.
from Ctl.






Average
Temp. (C)






Average
Sal. (%>)






Average
D.O. (mg/i)






Comments:
* U.S. GOVERNMENT PRINTING OFFICE: 1990 - 748-159/20498

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