Whole Effluent Toxicity Training
Video Series
Saltwater Series
U.S. Environmental Protection Agency
Office of Wastewater Management
Water Permits Division
1200 Pennsylvania Ave., NW
Washington, DC 20460
EPA 833-C-09-001
March 2009
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If you need additional copies of this document, you can download it at:
www.epa.gov/npdes/wqbasedpermitting
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WHOLE EFFLUENT TOXICITY • TRAINING VIDEO SERIES • saltwater series
Red Algal (Champia parvula)
Sexual Reproduction
Toxicity Tests
Supplement to Training Video
U.S. Environmental Protection Agency
Office of Wastewater Management
Water Permits Division
1200 Pennsylvania Ave., NW
Washington, DC 20460
EPA 833-C-09-001
March 2009
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NOTICE
The revision of this guide has been funded wholly or in part by the
Environmental Protection Agency under Contract EP-C-05-063. Mention of
trade names or commercial products does not constitute endorsement or
recommendation for use.
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U.S. ENVIRONMENTAL PROTECTION AGENCY Red Algal (Champia parvula) Sexual Reproduction Toxicity Tests
Supplement to Training Video
Foreword
This guide serves as a supplement to the video "Red Algal (Champia parvula) Sexual Reproduction Toxicity
Tests" (EPA, 2009). The methods illustrated in the video and described in this guide support the methods
published in the U.S. Environmental Protection Agency's (EPA's) Short-term Methods for Estimating the
Chronic Toxicity of Effluents and Receiving Waters to Marine and Estuarine Organisms, Third Edition (EPA,
2002a), referred to as the Saltwater Chronic Methods Manual. The video and this guide provide details on
preparing for and conducting the test based on the expertise of personnel at the following EPA Office of
Research and Development (ORD) laboratories:
National Health and Environmental Effects Research Laboratory (NHEERL) - Atlantic Ecology Division
in Narragansett, Rhode Island
NHEERL - Gulf Ecology Division in Gulf Breeze, Florida
National Exposure Research Lab (NERL) - Ecological Exposure Research Division (EERD) in
Cincinnati, Ohio
This guide and its accompanying video are part of a series of training videos produced by EPA's Office of
Wastewater Management. This Saltwater Series includes the following videos and guides:
"Mysid (/Americamys/s bahia) Survival, Growth, and Fecundity Toxicity Tests"
"Culturing Amer/camys/s bahia"
"Sperm Cell Toxicity Tests Using the Sea Urchin, Arbacia punctulata"
"Red Algal (Champia parvula) Sexual Reproduction Toxicity Tests"
"Sheepshead Minnow (Cyprinodon variegatus) and Inland Silverside (Menidia beryllina) Larval Survival
and Growth Toxicity Tests"
The Freshwater Series, released in 2006, includes the following videos and guides:
"Ceriodaphnia Survival and Reproduction Toxicity Tests"
"Culturing of Fathead Minnows (Pimephales promelas)"
"Fathead Minnow (Pimephales promelas) Larval Survival and Growth Toxicity Tests"
All of these videos are available through the National Service Center for Environmental Publications
(NSCEP) at 800 490-9198 or nscep@bps-lmit.com.
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U.S. ENVIRONMENTAL PROTECTION AGENCY Red Algal (Champia parvula) Sexual Reproduction Toxicity Tests
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U.S. ENVIRONMENTAL PROTECTION AGENCY Red Algal (Champia parvula) Sexual Reproduction Toxicity Tests
Supplement to Training Video
CONTENTS
Foreword i
Introduction 1
Background 1
Culturing Champia parvula 1
Culture Water 1
Photoperiod 2
Culture Vessels 2
Preparing Algae for Testing 2
Conducting the Test 3
Collecting the Algae 3
Effluent Preparation 3
The Exposure Period 5
The Recovery Period 6
Terminating the Test 6
Test Acceptability and Data Review 9
Citations and Recommended References 9
Glossary Glossary-1
Appendix A: Nutrients and Media A-l
Appendix B: Apparatus and Equipment B-l
Appendix C: Reagents and Consumable Materials C-l
Appendix D: Summary of Test Conditions and Test Acceptability Criteria for the Red Macroalga,
Champ/a parvula, Sexual Reproduction Test With Effluents and Receiving Waters D-l
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U.S. ENVIRONMENTAL PROTECTION AGENCY Red Algal (Champia parvula) Sexual Reproduction Toxicity Tests
, Supplement to Training Video
FIGURES
Figure 1. Life History of the Red Macroalga, Champia parvula. Left: Size and Degree of Branching
in Female Branch Tips Used For Toxicity Tests [[[ ................... 2
Figure 2. Apex of Branch of Female Plant, Showing Sterile Hairs and Reproductive Hairs
(Trichogynes) [[[ . ........... 3
Figure 3. A Portion of the Male Thallus Showing Spermatial Sori. The Sorus Areas Are Generally
Slightly Thicker and Somewhat Lighter in Color [[[ 3
Figure 4. A Magnified Portion of a Spermatial Sorus. Note the Rows of Cells that Protrude from the
Thallus Surface [[[ . [[[ 4
Figure 5. Apex of a Branch on a Mature Female Plant That Was Exposed To Spermatia from a Male
Plant [[[ 4
Figure 6. Receiving Water Data Form for the Red Macroalga, Champia parvula, Sexual
Reproduction Test [[[ 5
Figure 7. A Mature Cystocarp [[[ 6
Figure 8. Comparison of a Very Young Branch and an Immature Cystocarp ..................................... 7
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U.S. ENVIRONMENTAL PROTECTION AGENCY Red Algal (Champia parvula) Sexual Reproduction Toxicity Tests
Supplement to Training Video
^jfcsH
Introduction
This guide accompanies the Environmental Protection Agency's (EPA's) video training for conducting red
algal (Champia parvula) sexual reproduction toxicity tests (EPA, 2009). The test method is found in Section
16 of EPA's Short-term Methods for Estimating the Chronic Toxicity of Effluents and Receiving Waters to
Marine and Estuarine Organisms, Third Edition (EPA, 2002a). The test was developed by EPA's Office of
Research and Development's (ORD's) National Health and Environmental Effects Research Laboratory-
Atlantic Ecology Division (NHEERL-AED) in Narragansett, Rhode Island. The material presented in both the
video and this guide summarizes the methods but does not replace a thorough review and understanding
of the methods by laboratory personnel before conducting the test.
Background
Under the National Pollutant Discharge Elimination System (NPDES) program (Section 402 of the Clean
Water Act), EPA uses toxicity tests to monitor and evaluate effluents for their toxicity to biota and their
impact on receiving waters. By determining acceptable or safe concentrations for toxicants discharged
into receiving waters, EPA can establish NPDES permit limitations for toxicity. These Whole effluent toxic-
ity (WET) permit limitations regulate pollutant discharges on a whole effluent effect basis rather than by a
chemical-specific approach only.
Whole effluent toxicity methods measure the synergistic, antagonistic, and additive effects of all the chemi-
cal, physical, and additive components of an effluent that adversely affect the physiological and biochemi-
cal functions of the test organisms. Therefore, healthy organisms and correct laboratory procedures are
essential for valid test results. Laboratory personnel should be very familiar with the test methods and with
red algae handling techniques before conducting a test.
This supplemental guide covers the procedures for conducting the test according to EPA's promulgated
methods (40 CFR Part 136; EPA, 2002c) and also provides some helpful information that is not presented
in the Saltwater Chronic Methods Manual (EPA, 2002a).
This guide summarizes methods developed at NHEERL-AED for estimating the chronic toxicity of marine
or estuarine effluents and receiving waters on the sexual reproduction of the marine macroalga, Champia
parvula. Males and females are exposed to effluents or receiving waters for 2 days, followed by a 5- to
7-day recovery period for the female plants in a control medium. Cystocarp production by the female, which
indicates sexual reproduction, is used as the endpoint. The test results determine the effluent concentra-
tion causing a statistically significant reduction in the number of cystocarps formed.
This guide and accompanying video describe how the test is set up, initiated, terminated, and reviewed,
including suggestions on maintaining healthy cultures of test organisms.
Culturing Champia parvula
There are three macroscopic stages in the life history of Champia. The adult plant body (thallus) is hollow,
septate, and highly branched. Only the mature male and female plants are used in toxicity testing. Mature
plants are illustrated in Figure 1.
To keep a constant supply of plant material available, maintain several unialgal stock cultures of males
and females simultaneously. Also, new cultures should be started weekly from excised branches so that
cultures are available in different stages of development.
CULTURE WATER
Natural seawater, or a 50-50 mixture of natural and artificial seawater, makes optimal culturing media.
Seawater for cultures is filtered at least to 0.45 urn to remove most particulates and autoclaved for 30
minutes at 15 psi (120°C). Carbon stripping the seawater may be necessary before autoclavingto enhance
I
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Red Algal (Chomp/a parvula) Sexual Reproduction Toxicity Tests
Supplement to Training Video
Figure 1. Life History of the Red Macroalga, C/iamp/a parvula. Left: Size
and Degree of Branching in Female Branch Tips Used For Toxicity Tests
spermatia
fertilization
tetrasporangia
TETRASP'
Source: EPA 1987.
—cystocarp
5 mm
its water quality (EPA, 1990).
Instructions for carbon
stripping are provided in the
Saltwater Chronic Methods
Manual (EPA, 2002a). Nutrients
should be added to the water
to ensure healthy cultures.
Recipes for the culturing
medium and nutrient solutions
are provided in Appendix A. The
water temperature should be
maintained at 23°C + 1°C and
the salinity at 30%0 ± 2%0
Gently aerate the cultures.
Change alternate cultures'
medium every week so that if a
stock solution should become
contaminated, the entire batch
will not be lost. While replen-
ishing the medium, divide the growing algae in half with sharp forceps or discard half of the biomass to prevent
overcrowding. New cultures also can be started at this time using 1 cm branch tips. Add nutrients using a pipet;
NHEERL-AED has found a squeeze bottle is quick and easy to use. At the end of approximately three weeks, there
should be enough plant material to conduct the test.
PHOTOPERIOD
The culture conditions should include a photoperiod 16 hours of light and 8 hours of darkness. The light
level should not exceed 500 ft-candles (75 uE/m2/s) and may have to be adjusted to that level, depending
on the reflecting characteristics of the incubators.
CULTURE VESSELS
Maintain stock cultures of males and females in separate, aerated, 1 L Erlenmeyer flasks containing 800
mL of the culture medium. All glass must be acid-stripped in 15 percent HCI and rinsed in deionized water
before use because some detergent residues can be toxic to the Champia. At least every 6 months, the
glass should be cleaned to remove organic materials that can build up on the surface. Always use sterile
techniques when culturing the algae (i.e., autoclave all stock solutions and flame all tools before cutting or
transferring plants) to guard against microalgal contamination.
PREPARING ALGAE FOR TESTING
Examine the stock cultures to determine their readiness for testing. Place a few female branch tips in
seawater in a petri dish, and examine them under a compound microscope to determine if trichogynes are
present. An inverted scope works best with the petri dishes, although standard slides and microscopes also
can be used. Trichogynes are the short, fine reproductive hairs to which the spermatia attach (see Figure
2). They should be seen easily near the apex of the branch tip. Although both sterile hairs and trichogynes
occur on the apex, sterile hairs occur over the entire plant thallus. Sterile hairs are wider and generally much
longer than trichogynes, and appear hollow, except at their tip, where they seem to be plugged.
Males should be visibly producing spermatia. Sometimes, the presence of spermatia sori can be deter-
mined by placing some male tissue in a petri dish and holding it against a dark background. Mature sori
can be easily identified under a microscope along the edge of the thallus. The sorus areas are generally
thicker and lighter in color than the rest of the plant body. At higher magnification, the spermatia them-
selves can be seen (see Figures 3 and 4).
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Red Algal (Champia parvula) Sexual Reproduction Toxicity Tests
Supplement to Training Video
The readiness of the male stock
culture can also be assessed
by placing a portion of a female
plant into a portion of the solu-
tion from the male culture for
a few seconds. Under a micro-
scope, numerous spermatia
should be seen attached to the
sterile hairs and trichogynes of
the female plant (see Figure 5).
Once readiness is established
for both males and females, the
test can begin.
Conducting the Test
COLLECTING THE ALGAE
Prepare cuttings from the most
healthy-looking plants. Prepare
the female cuttings first to mini-
mize the chances of contaminat-
ing them with water containing
spermatia from the male stock
cultures. Place each plant in
Figure 2. Apex of Branch of Female Plant, Showing Sterile Hairs and
Reproductive Hairs (Trichogynes)
sterile hairs
ichogynes
Sterile hairs are wider and generally much longer than trichogynes, and appear hollow
except at the tip. Roth types of hairs occur on the entire circumference of the thallus, but
are seen easiest at the "edges." Receptive trichogynes occur only near the branch tips.
Source: EPA 1987.
a petri dish containing a small amount of seawater. Using a fine-point forceps or scalpel, prepare five cut-
tings from the female plants for each treatment replicate, severing the plant 7-10 mm from the ends of the
branch. Try to be consistent in the degree of branching in the cuttings, since cystocarps form at the branch
tips.
For male plants, use one cutting for each treatment replicate, severing the plant about 2 - 3 cm from the
end of the branch. If there are few branches, or the spermatial sori appear sparse, larger male cuttings may
be needed. The cuttings can be kept at room temperature for up to an hour.
EFFLUENT PREPARATION
Effluent sampling should be
conducted according to Section
8 of the Saltwater Chronic
Methods Manual (EPA, 2002a)
and any specific requirements of
a NPDES permit. The effluent or
receiving waters should be held
at 0°C - 6°C until used for test-
ing. Under the NPDES program,
lapsed time from sample collec-
tion to first use in the test must
not exceed 36 hours. Under
special conditions or variances,
samples may be held longer but
should never be used for testing
if held for more than 72 hours.
Figure 3. A Portion of the Male Thallus Showing Spermatial Sori. The
Sorus Areas Are Generally Slightly Thicker and Somewhat Lighter in
Color
1 cm
spermatial sorus
Source: EPA 1987.
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Red Algal (Champla parvula) Sexual Reproduction Toxlcity Tests
Supplement to Training Video
Dilution Water
The type of dilution water used
to make the test concentrations
is dependent on the objectives
of the test Any specific require-
ments included in NPDES permits
should be followed. The Saltwater
Chronic Methods Manual (Section
7) provides the following guidelines:
* If the test is conducted to esti-
mate the absolute chronic tox-
ic/ty of the effluent, synthetic
dilution water should be used. If
the cultures were maintained in
different water than used for dilu-
tion water, a second set of control
replicates should be conducted
using the culture water.
• If the test is conducted to
estimate the chronic toxicity
of the effluent in uncon-
taminated receiving waters,
the test can be conducted using
a grab sample of the receiving
waters collected outside the influ-
ence of the outfall, other uncon-
taminated waters, or standard
dilution water with the same
salinity as the receiving waters. If
the cultures were maintained in
different water than used for dilu-
tion water, a second set of control
replicates should be conducted
using the culture water,
• If the test is conducted to
estimate the additive or miti-
gating effects of the effluent
on already contaminated
receiving waters, the test
must be conducted using receiv-
ing waters collected outside the
influence of the outfall. Controls
should be conducted using both
receiving water and culture water.
Figure 4. A Magnified Portion of a Spermatial Sorus. Note the Rows
of Cells that Protrude from the Thallus Surface
cuticle
thallus surface
Source: EPA 1987.
Figure 5. of a on a Plant That
To from a
spermatia
The sterile hairs and trichogynes are covered with spermatia. Note that
few or no spermatia are attached to the older hairs (those more than
I mm from the apex).
Source: EPA 1987.
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Red Algal (Champ/a parvula) Sexual Reproduction Toxicity Tests
Supplement to Training Video
Maintain the salinity of the test samples to 30%o ± 2%o. To do this, effluent samples may need to be
adjusted using hypersaline brine (HSB). A recipe for MSB is provided in Appendix A of this manual.
Approximately 1 hour before the test is to begin, adjust approximately 1 L of effluent to the test tempera-
ture of 23°C ± 1°C and maintain that temperature while preparing the test concentrations. To test a series
of decreasing concentrations of effluent, use a dilution factor of > 0.5. When starting with effluent that has
0%0 salinity, the maximum effluent concentration that can be prepared at 30%o is 70 percent effluent. A
table for preparing the samples is provided in Appendix A.
THE EXPOSURE PERIOD
A 125 ml Erlenmeyer flask is used for each test chamber, but any clean container can be used. The test
chambers should be labeled using colored tape and marking pens to identify each treatment and replicate.
These should be placed in randomized positions for the duration of the test.
Under a hood, prepare five dilutions using a > 0.5 dilution factor in 300 or 400 ml replicates.
Approximately 1800 ml of effluent is required for a test conducted using a 0.5 dilution factor. This allows
for enough of each prepared effluent concentration to provide four replicates at 100 ml and 400 ml for
chemical analyses and water quality data. Record the water quality data on a form such as the one pro-
vided in Figure 6.
Figure 6. Receiving Water Data Form for the Red Macroalga, Champ/a parvula, Sexual Reproduction Test.
Site:
Collection Date:
Test Date:
Locations
Initial
Salinity
Final
Salinity
Source of Salts for Salinity Adjustment1
1Natural seawater, GP2 brine, GP2 salts, etc. (include some indication of amount.)
Source: EPA, 2002a.
The 2-day exposure period starts when the algae are added to the test chambers. Add five female branch-
es and one male branch to each prepared chamber. Pick up the branch at the base or cut end to avoid
injuring the tips. The effluent must be in the test chamber before the algae are added.
Cover the chambers with aluminum foil or a foam stopper, exposing the cultures to 16 hours of cool white
light and 8 hours of darkness each day for the 2-day exposure, as well as the 5- to 7-day recovery peri-
ods. Maintain the temperature at 23°C + 1°C, and the salinity between 28%o and 32%o with the variance
between chambers on any day maintained at < 2%o.
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Red Algal (Champia porvulo) Sexual Reproduction Toxicity Tests
Supplement to Training Video
Check on the chambers twice a day, and gently hand-swirl the chambers, or shake continuously at 100
rpm on a rotary shaker. Spermatia are not motile, so some motion is critical during the exposure period for
reproduction to occur. If desired, the media can be changed after 24 hours. Record the temperature daily
from a thermometer placed in a flask of water among the chambers.
Routine chemical and physical observations should be made during the
test. Dissolved oxygen (DO) is measured at the beginning and end of
each 24-hour exposure period in one test chamber at each concentra-
tion and in the control. Temperature, pH, and salinity are measured at the
end of each 24-hour exposure period, also in one test chamber at each
concentration and in the control. Temperature also should be monitored
continuously, observed and recorded daily for at least two locations in the
environmental control system or the samples. The locations for determin-
ing temperature should be sufficient to indicate any temperature variations in the environmental chamber.
pH should be measured in
the effluent sample before
any new test solutions are
made to determine changes
in the effluent sample.
THE RECOVERY PERIOD
Prepare recovery bottles by labeling clean 100 - 200 ml vessels with the effluent concentrations tested,
and fill them with 150 ml of natural seawater and nutrients. Smaller volumes can be used but may require
changes of the medium to allow for adequate growth.
After the 48-hour exposure period, use forceps to gently remove all of the females from each test chamber,
and place them into recovery bottles. When all the replicates have been transferred, place the vessels
under cool white light and aerate or shake for the 5- to 7-day recovery period. Aeration will enhance the
growth rate of plants in the recovery bottles, although adequate growth will occur using a shaker. Aerate
using plastic tubes held in place by foam stoppers.
TERMINATING THE TEST
At the end of the recovery period, drain the chambers and remove the females with forceps, starting with
the control plants and ending with those in the highest concentration. Place the female plants between the
inverted halves of a petri dish Figure 7. A Mature Cystocarp
containing a small amount
of seawater, and count the
cystocarps under a stereo-
microscope. Cystocarps are «/. ' • . '.*&. ^ . ^ostiole
distinguished from young
branches by the darkly ^i^^Uf fK. ^!:*J^I J ** sP°res
pigmented spores enclosed
in the nodule, and the apical
opening for spore release
(ostiole). Figures 7 through 9
provide illustrations to help
identify cystocarps.
In the controls and lower effluent concentrations, cystocarps often occur in clusters of
10 or 12.
Source: EPA 1987.
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Red Algal (Champia parvula) Sexual Reproduction Toxicity Tests
Supplement to Training Video
Figure 8. Comparison of a Very Young Branch and an Immature
Cystocarp
young branch
cells
immature
cystocarp
Both the young branch and immature cystocarp can have sterile hairs. Trichogynes
might or might not be present on a young branch, but are never present on an imma-
ture cystocarp. Young branches are more pointed at the apex and are made up of
larger cells than immature cystocarps, and never have ostioles.
Source: EPA 1987.
Figure 9. An Aborted Cystocarp.
A new branch will eventually develop at the apex.
Source: EPA 1987.
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Red Algal (Champia parvula) Sexual Reproduction Toxicity Tests
Supplement to Training Video
If there is doubt about the identification of an immature cystocarp, aerate the plants a little longer in the
recovery bottles. Within 24 to 48 hours, the suspected cystocarp will look more like a mature cystocarp
or a young branch, or will have changed very little, if at all, indicating an aborted cystocarp. Occasionally
cystocarps will abort, and these should not be included in the counts. Aborted cystocarps are easily identi-
fied by their dark pigmentation and/or by the formation of a new branch at the apex. Dead plants lose their
pigmentation and appear white.
Record all counts for the test on a form such as the one provided in Figure 10.
Figure 10. Cystocarp Data Sheet for the Red Macroalga, Champia parvula, Sexual Reproduction Test
Collection Date:
Exposure Began (date):
Effluent or Toxicant:
Recovery Began (date):
Counted (date):
Treatment (% Effluent, mg/L, or receiving water sites)
Replicates
A 1
2
3
4
Mean
B 1
2
3
4
Mean
C 1
2
3
4
Mean
D 1
2
3
4
• Mean
Overall
Mean
Control
Temperature:.
Salinity:
Light: :
Source of Dilution Water:
Source: EPA, 1987.
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U'S' ENVIRONMENTAL PROTECTION AGENCY Red Algal (Champia parvula) Sexual Reproduction Toxicity Tests
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Test Acceptability and Data Review
Test data are reviewed to verify that EPA's WET test methods' test acceptability criteria (TAG) requirements
for a valid test have been met. The algal sexual reproduction test requires that several criteria be met
before the test results are considered acceptable.
• Control plants should average 10 or more cystocarps per plant and survival in the control must be
80 percent or greater.
• Control and lowest-concentration exposed algae should be in good physical condition—for exam-
ple, the branches should not be fragmented. Broken or fragmented branches could indicate that
the plants were unhealthy or stressed from the beginning of the test.
• The results from the replicate control chambers should be similar.
• All replicates from the affected concentration chambers should show effect.
The concentration-response relationship generated for each multi-concentration test must be reviewed to
ensure that calculated test results are interpreted appropriately. In conjunction with this requirement, EPA
has provided recommended guidance for concentration-response relationship review (EPA, 2000b).
EPA's promulgated toxicity testing method manuals (EPA, 2002a, b) recommend the use of point estima-
tion technique approaches for calculating endpoints for effluent toxicity tests under the NPDES program.
The promulgated methods also require a data review of toxicity data and concentration-response data, and
require calculating the percent minimum significant difference (PMSD) when point estimation (e.g., LC50,
IC25) analyses are not used. EPA specifies the PMSD must be calculated when NPDES permits require sub-
lethal hypothesis testing. EPA also requires that variability criteria be applied as a test review step when
NPDES permits require sub-lethal hypothesis testing endpoints (i.e., no observed effect concentration
[NOEC] or lowest observed effect concentration [LOEC]) and the effluent has been determined to have no
toxicity at the permitted receiving water concentration (EPA, 2002b). This reduces the within-test variabil-
ity and increases statistical sensitivity when test endpoints are expressed using hypothesis testing rather
than the preferred point estimation techniques.
Citations and Recommended References
EPA. 1979. Methods for chemical analysis of water and wastes. Environmental Monitoring and Support
Laboratory, U.S. EPA, Cincinnati, OH 45268. EPA-600/4- 79/020, revised March 1983.
EPA. 1985. Aquatic Toxicity Testing Seminar Manual. 1985. National Health and Environmental Effects
Research Laboratory-Aquatic Ecology Division, Narragansett, Rl. NHEERL-AED Contribution No.
796.
EPA. 1987. Guidance manual for conducting sexual reproduction test with the marine macroalga Champia
parvula for use in testing complex effluents. Contribution No. X103. Thursby, G.B. and R.L. Steele.
In: Schimmel S.C., ed. Users guide to the conduct and interpretation of complex effluent toxicity
tests at estuarine/marine sites. Environmental Research Laboratory, U.S. EPA, Narragansett, Rl
02882. Contribution No. 796. 265 pp.
EPA. 1989. 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, Rl. EPA/625/8-89/015.
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U.S. ENVIRONMENTAL PROTECTION AGENCY Red Algal (Champia parvula) Sexual Reproduction Toxicity Tests
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',$ , ,-, .ft .-
EPA. 1990. Supplemental methods and status reports for short-term saltwater toxicity tests.
G. Morrison and G. Chapman. ERL Contrib. No. 1199. Environmental Research Laboratory,
U.S. EPA, Narragansett, Rl 02882. 127 pp.
EPA. 1991. Technical Support Document for Water Quality-based Toxics Control. U.S. EPA Office of Water
Enforcement and Permits, Washington, D.C. EPA-505-2-90-001.
EPA. 2000a. Method Guidance and Recommendations for Whole Effluent Toxicity (WET) Testing (40 CFR
Part 136). Office of Water, Washington, D.C. EPA 821-B-00-004.
EPA. 2000b. Understanding and Accounting for Method Variability in Whole Effluent Toxicity Applications
Under the National Pollutant Discharge Elimination System Program. Office of Wastewater
Management, Washington, D.C. EPA 833-R-00-003.
EPA. 2002a. Short-term Methods for Estimating the Chronic Toxicity of Effluents and Receiving Waters to
Marine and Estuarine Organisms, Third Edition. (Saltwater Chronic Methods Manual). Office of
Water, Cincinnati, OH. EPA-821-R-02-014.
EPA. 2002b. Methods for Measuring the Acute Toxicity of Effluents and Receiving Waters to Freshwater
and Marine Organisms, Fifth Edition. (Acute Methods Manual). Office of Water, Cincinnati, OH.
EPA-821-R-02-012.
EPA. 2002c. Final Rule. 40 CFR Part 136. Guidelines Establishing Test Procedures for the Analysis of
Pollutants; Whole Effluent Toxicity Test Methods. 67 FR 69952-69972, November 19, 2002.
EPA, 2009. Red Algal (Champia parvula) Sexual Reproduction Toxicity Tests. Supplement to Training Video.
Whole Effluent Toxicity Training Video Series, Saltwater Series. March 2009.
EPA 833-C-09-001.
Spotte, S., G. Adams, and P.M. Bubucis. 1984. GP2 as an artificial seawater for culture or maintenance of
marine organisms. Zool. Biol. 3:229-240.
EPA references are available online at www.epa.gov/npdes.
If you need additional copies of this document, you can download it at:
www.epa.gov/npdes/wqbasedpermitting.
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yjg| U.S. ENVIRONMENTAL PROTECTION AGENCY Red Algal (Champia parvula) Sexual Reproduction Toxicity Tests
Va»' Supplement to Training Video
Glossary
Acute toxicity. An adverse effect measured on a group of test organisms during a short-term exposure in
a short period of time (96 hours or less in toxicity tests). The effect can be measured in lethality
or any variety of effects.
Champia parvula. The scientific name for red algae. Champia parvula have soft, gelatinous, pinkish red,
much-branched fronds that are densely matted, with blunt apices, to 100 mm high. Their axes are
segmented, with nodal diaphragms. The segments are about as broad as long, filled with a watery
mucilage. Red algae are found epiphytic on smaller algae in lower intertidal pools. They are found
widely distributed in the Atlantic and Pacific marine environments.
Chronic toxicity. An adverse effect that occurs over a long exposure period. The effect can be lethality,
impaired growth, reduced reproduction, etc.
Diluent water. Dilution water used to prepare the effluent concentrations.
Effluent concentrations. Concentrations or dilutions of an effluent sample to which test organisms are
exposed to determine the biological effects of the sample on the test organism.
Effluent sample. A representative collection of the discharge that is to be tested.
Hypothesis testing. Technique (e.g., Dunnett's test) that determines what concentration is statistically
different from the control. Endpoints determined from hypothesis testing are NOEC and LOEC.
IC25 (Inhibition Concentration, 25%). The point estimate of the toxicant concentration that would cause a
25% reduction in a non-quantal biological measurement (e.g., reproduction or growth) calculated
from a continuous model.
LC50 (Lethal Concentration, 50%). The concentration of toxicant or effluent that would cause death to
50% of the test organisms at a specific time of observations (e.g., 96-hour LC50).
Lowest Observed Effect Concentration (LOEC). The LOEC is the lowest concentration of toxicant to
which organisms are exposed in a test, which causes statistically significant adverse effects on
the test organisms (i.e., where the values for the observed endpoints are statistically significantly
different from'the control). The definitions of NOEC and LOEC assume a strict dose-response
relationship between toxicant concentration and organism response.
Minimum Significant Difference (MSD). The MSD is the magnitude of difference from the control where
the null hypothesis is rejected in a statistical test comparing a treatment with a control. MSD
is based on the number of replicates, control performance and power of the test. MSD is often
measured as a percent and referred to as PMSD.
No Observed Effect Concentration (NOEC). The NOEC is the highest tested concentration of toxicant to
which organisms are exposed in a full life-cycle or partial life-cycle (short-term) test, that causes
no observable adverse effect on the test organism (i.e., the highest concentration of toxicant
at which the values for the observed responses are not statistically significantly different from
the controls). NOECs calculated by hypothesis testing are dependent upon the concentrations
selected.
NPDES (National Pollutant Discharge Elimination System) Program. The national program for issuing,
modifying, revoking and reissuing, terminating, monitoring and enforcing permits, and imposing
and enforcing pretreatment requirements, under Sections 307, 318, 402, and 405 of the Clean
Water Act.
Glossary-1
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U.S. ENVIRONMENTAL PROTECTION AGENCY Red Algal (Champia parvula) Sexual Reproduction Toxicity Tests
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Point Estimation Techniques. This technique is used to determine the effluent concentration at which
adverse effects (e.g., fertilization, growth or survival) occurred, such as Probit, Interpolation
Method, Spearman-Karber. For example, a concentration at which a 25% reduction in
reproduction and survival occurred.
Receiving Water Concentration (RWC). The RWC is the concentration of a toxicant or the parameter
toxicity in the receiving water (i.e., riverine, lake, reservoir, estuary.or ocean) after mixing.
Toxicity test. A test to measure the toxicity of a chemical or effluent using living organisms. The test
measures the degree of response of an exposed organism to a specific chemical or effluent.
WET (Whole effluent toxicity). The total toxic effect of an effluent measured directly with a toxicity test.
Glossary-2
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Red Algal (Chompia parvula) Sexual Reproduction Toxicity Tests
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Appendix A:
Nutrients and Media
The following instructions for nutrients are provided in the Saltwater Chronic Methods Manual
(EPA, 2002a). Table A-l lists the additional nutrients to be added to natural or artificial seawater for stock
cultures and test media. The concentrated stock solution is autoclaved at standard temperature and pres-
sure for 15 minutes before the vitamins are added. Adjust the solution to about pH 2 before autoclavingto
minimize the possibility of precipitation.
Table A-l. Nutrient Stock Solution
Nutrient Stock
Solution"
NaNO3
NaH2PO4 • H2O
Na2EDTA • 2 H2O
Na3C6H507 • 2 H2O
lronb
Vitamins0
Amount IL Concentrated Nutrient Stock Solution
Stock Solution for
Culture Medium
6.35 g
0.64 g
133 mg
51 mg
9.75 ml
10 mL
Stock Solution for
Test Medium
l.58g
0.16 g
—
12.8 mg
2.4 mL
2.5mL
3 Add 10 mL of appropriate nutrient stock solution per liter of culture or test medium.
b A stock solution of iron is made that contains 1 mg iron/mL. Ferrous or ferric chloride can be used.
c A vitamin stock solution is made by dissolving 4.88 g thiamine HCI, 2.5 mg biotin, and 2.5 mg Bi2 in 500 mL deionized
water. Adjust vitamin stock to approximately pH 4, divide into 10 mL subsamples, and autoclave for 2 minutes before it is
added to the nutrient stock solution.
Preparing Hypersaline Brine (HSB)
BACKGROUND
Champia parvula cannot be cultured in 100% artificial seawater. However, 100% artificial seawater can be
used during the 2-day exposure period. This allows 100% effluent to be tested.
Salinity adjustments are a vital part of using marine and estuarine species for toxicity testing. The major-
ity 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. The salinity of the effluent can be adjusted by adding HSB pre-
pared from natural seawater (100%o), concentrated (triple strength) salt solution (GP2 described in table
below), or dry GP2 salts (also below). Adjust the salinity of the effluent to 30%o. Control solutions should be
prepared with the same percentage of natural seawater and at the same salinity as the effluent solutions.
Constant salinity should be maintained across all treatments throughout the test for quality control.
Matching the test solutions' salinity to the expected receiving water's salinity may require salinity adjust-
ments. EPA NHEERL-AED uses HSB, prepared from filtered natural seawater, to adjust exposure solution
salinities.
HSB 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
A-l
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Red Algal (Champ/a parvula) Sexual Reproduction Toxicity Tests
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•„•,'.- A. «TvJVSa«
desired salinity with deionized water. The brine can be made from any high quality, filtered seawater supply
through simple heating and aerating.
Table A-2. GP2 Artificial Seawater for Use in Conjunction with Natural Seawater for the Red Macroalga,
Champ/a parvula, Sexual Reproduction Toxicity Test
Compound
NaCI
Na2SO4
KCI
KBr
Na2B4O7- IOH20
MgCI2 • 6 H2O
CaCI2 • 2 H2O
SrCI2 • 6 H2O
NaHCO3a
Concentration (gIL)
21.03
3.52
0.61
0.088
0.034
9.50
1.32
0.02
0.17
Amount (g) Required for 20-L
420.6
70.4
12.2
1.76
0.68
190.0
26.4
0.400
3.40
The original formulation calls for autoclaving anhydrous and hydrated salts separately to avoid precipitation. However, if
the sodium bicarbonate is autoclaved separately (dry), all of the other salts can be autoclaved together. Since no nutrients
are added until needed, autoclaving is not critical for effluent testing. To minimize microalgal contamination, the artificial
seawater should be autoclaved when used for stock cultures. Autoclaving (120°C) should be for at least 10 minutes for
1-L volumes, and 20 minutes for 10- to 20-L volumes.
Artificial seawater should be prepared in 10- to 20-L batches. Effluent salinity adjustment to 30%o can be made by adding
the appropriate amount of dry salts from this formulation, by using a triple-strength brine prepared from this formulation,
or by using a 100%o salinity brine prepared from natural seawater.
Nutrients listed in Table A-l should be added to the artificial seawater in the same concentration described for natural
seawater.
3 A stock solution of 68 mg/mL sodium bicarbonate is prepared by autoclaving it as a dry powder, and then dissolving it in
sterile deionized water. For each liter- of GP2, use 2.5 mL of this stock solution.
Source: EPA, 2002a. Modified from Spotte et a/., 1984. Constituents salts and concentrations were taken from EPA
1990.
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.
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 |jm 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 (or more or less
often, depending on the volume being generated) to ensure that the salinity does not exceed 100%o 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
A-2
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Red Algal (Champia parvula) Sexual Reproduction Toxicity Tests
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of seawater made from brine with salinity greater than 100%o, 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 MSB
To calculate the volume of brine (Vb) to add to 0%o sample to produce a solution at certain salinity (Sf), use
this equation:
Where Vb =
vb * sb = sf * vf
volume of brine, ml
Sb = salinity of brine, %o
Sf = final salinity, %o
Vf = final volume, mL (brine brought to this volume with 0 %o sample).
Table A-3 gives volumes needed to make 30%o test solutions from effluent (0%o), deionized water, and
100%o MSB. At 30%o salinity, the highest achievable concentration is 70% effluent.
Table A-3. Preparation of Test Solutions at a Salinity of 30%o Using HSB for a Final Test Concentration
Volume of 1000 ml.
Exposure
Concentration (%)
70
25
7
2.5
0.7
Control
Effluent
(0%o)
(mL)
700
250
70
25
7
—
Deionized Water
(mL)
—
450
630
675
693
1,000
Hypersaline Brine
(100 %o)
(mL)
300
300
300
300
300
—
A-3
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A-4
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U.S. ENVIRONMENTAL PROTECTION AGENCY Red Algal (Chomp/a parvula) Sexual Reproduction Toxicity Tests
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Appendix B:
Apparatus and Equipment
Air lines, and air stones. For aerating cultures, brood chambers, and holding tanks, and supplying air to
test solutions with low DO.
Air pump. For oil-free air supply.
Balance. Analytical, capable of accurately weighing to 0.00001 g.
Beakers, Class A. Borosilicate glass or non-toxic plasticware, 1000 ml for making test solutions.
Bottles. Borosilicate glass or disposable polystyrene cups (200 - 400 ml) for use as recovery vessels.
Compound microscope. For examining the condition of plants.
Count register. 2-place for recording cystocarp counts.
Dissecting (stereomicroscope) microscope. For counting cystocarps.
Drying oven. To dry glassware.
Erlenmeyer flasks, 250 mL, or 200 ml disposable polystyrene cups, with covers. For use as exposure
chambers.
Environmental chamber or equivalent facility with temperature control (23 ± 1°C).
Facilities for holding and acclimating test organisms.
Filtering apparatus. For use with membrane filters (47 mm).
Forceps, fine-point, stainless steel. For cutting and handling branch tips.
Laboratory red macroalga, Champ/a parvula, culture unit. To test effluent or receiving water toxicity,
sufficient number of sexually mature male and female plants must be available.
Meters: pH and DO, and specific conductivity. For routine physical and chemical measurements.
Micropipettors, digital, 200 and 1000 uL. To make dilutions.
Pipet bulbs and filters. Propipet®, or equivalent.
Pipets, automatic. Adjustable 1 - 100 ml.
Pipets, serological. 1-10 ml, graduated.
Pipets, volumetric. Class A, 1 - 100 mL.
Reference weights, Class S. For checking performance of balance.
Refractometer or other method. For determining salinity.
Rotary shaker. For incubating exposure chambers (hand-swirling twice a day can be substituted).
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U.S. ENVIRONMENTAL PROTECTION AGENCY Red Algal (Champia parvula) Sexual Reproduction Toxicity Tests
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Samplers. Automatic samplers, preferably with sample cooling capability, that can collect a 24-hour
composite sample of 1 L.
Thermometers. National Bureau of Standards Certified (see EPA 2002a). Used to calibrate laboratory
thermometers.
Thermometers. Bulb-thermograph or electronic-chart type for continuously recording temperature.
Thermometers, glass or electronic, laboratory grade. For measuring water temperatures.
Water purification system. Millipore® Milli-Q® deionized water or equivalent.
Wash bottles. For deionized water, for washing organisms from containers and for rinsing small glassware
and instrument electrodes and probes.
Volumetric flasks and graduated cylinders. Class A, borosilicate glass or non-toxic plastic labware,
10 - 1000 ml for making test solutions.
B-2
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U.S. ENVIRONMENTAL PROTECTION AGENCY Red Algal (Champia parvula) Sexual Reproduction Toxicity Tests
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*•>
Appendix C:
Reagents and Consumable Materials
Aluminum foil, foam stoppers, or other closures. To cover cultures and test flasks.
Artificial seawater. A slightly modified version of the GP2 medium (Spotte, et al., 1984) has been used
successfully to perform the red macroalga sexual reproduction test. The preparation of artificial
seawater (GP2) is described in Table A-2.
Buffers pH 4, pH 7, and pH 10. (Or as per instructions of instrument manufacturer) for standards and
calibration check.
Data sheets (one set per test). For data recording (see Figures 6 and 10).
Disposable tips for micropipettors.
Effluent, receiving water, and dilution water. Test waters, including effluent, receiving, and dilution
water should be analyzed to ensure its quality prior to using in tests. Dilution water containing
organisms that might prey upon or otherwise interfere with the test organisms should be filtered
through a fine mesh (with 150 urn or smaller openings).
Laboratory quality assurance samples and standards. For the above methods.
Markers, waterproof. For marking containers, etc.
Mature red macroalga, Champia parvula, plants.
Petri dishes, polystyrene. To hold plants for cystocarp counts and to cut branch tips. Other suitable
containers may be used.
pH buffers pH 4, pH 7, and pH 10. (Or as per instructions of instrument manufacturer) for standards and
calibration check.
Reagent water. Distilled or deionized water that does not contain substances which are toxic to the test
organisms.
Reference toxicant solutions. Reference toxicants such as sodium chloride (NaCI), potassium chloride
(KCI), cadmium chloride (CdCI2), copper sulfate (CuS04), sodium dodecyl sulfate (SDS), and
potassium dichromate (K2Cr207), are suitable for use in the NPDES Program and other Agency
programs requiring aquatic toxicity tests.
Saline test and dilution factor. The use of natural seawater is recommended for this test. A recipe for the
nutrients that must be added to the natural seawater is given in Table A-l. The salinity of the test
water must be 30%o and vary no more than ± 2%o among the replicates. If effluent and receiving
water tests are conducted concurrently, the salinity of these tests should be similar.
The overwhelming majority of industrial and sewage treatment effluents entering marine and
estuarine systems contain little or no measurable salts. Therefore, exposure of the red macroalga,
Champia parvula, to effluents will usually require adjustments in the salinity of the test solutions.
Although the red macroalga, Champia parvula, cannot be cultured in 100% artificial seawater,
100% artificial seawater can be used during the 2-day exposure period. This allows 100%
effluent to be tested. It is important to maintain a constant salinity across all treatments. The
salinity of the effluent can be adjusted by adding MSB prepared from natural seawater (100%o),
concentrated (triple strength) salt solution (GP2 described in Table A-2), or dry GP2 salts (Table
C-l
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U.S. ENVIRONMENTAL PROTECTION AGENCY Red Algal (Champ/a parvula) Sexual Reproduction Toxicity Tests
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-; •* • ,»v - r»t <•,«•* tSsSS-
A-2), to the effluent to provide a salinity of 30%o. Control solutions should be prepared with the
same percentage of natural seawater and at the same salinity (using deionized water adjusted
with dry salts, or brine) as used for the effluent dilutions.
Sample containers. For sample shipment and storage.
Tape, colored. For labeling test chambers.
C-2
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Red Algal (Champia parvula) Sexual Reproduction Toxicity Tests
Supplement to Training Video
Appendix D:
Summary of Test Conditions and Test Acceptability
Criteria for the Red Macroalga, Champia parvula, Sexual
Reproduction Test With Effluents and Receiving Waters
(Note: this test is not listed at 40 CFR Part 136 for nationwide use)
Test type
Salinity
Temperature (C°)
Light source
Light intensity
Photoperiod
Test chamber size
Test solution volume
Number of organisms per test chamber
Number of replicates per concentration
Number of organisms per concentration
Aeration
Dilution water
Test concentrations
Receiving waters
Dilution factor
Test duration
Endpoints
Test acceptability criteria
Sampling requirements
Sample volume required
Static, non-renewal (required)
30%o ± 2 %o of the selected test salinity (recommended)
23°C ± I°C (recommended)
Cool-white fluorescent lights (recommended)
About 75 |jE/m2/s (500 ft-c) (recommended)
16 hr light, 8 hr dark (recommended)
200 mL polystyrene cups (with covers) or 250 mL Erlenmeyer flasks
(recommended)
100 ml (minimum required)
5 female branch tips and one male plant (recommended)
4 (3 required minimum)
24 (18 required minimum)
None; chambers are either shaken at 100 rpm on a rotary shaker or .
hand-swirled twice a day
30%o salinity natural seawater, or a combination of 50% of 30%o salinity
natural seawater and 50% of 30%o salinity GP2 artificial seawater
Effluents: 5 and a control (required minimum)
100% receiving water (or minimum of 5) and a control (recommended)
Effluents: ^ 0.5 (recommended)
Receiving Waters: None or S 0.5 (recommended)
2-day exposure to effluent followed by 7-day recovery period in control
medium for cystocarp development (required)
Reduction in cystocarp production compared to controls (required)
80% or greater survival, and an average of 10 cystocarps per plant in
controls (required)
For on-site tests, one sample collected at test initiation, and used within
24 hr of the time it is removed from the sampling device.
For off-site test, holding time must not exceed 36 hr before test use.
(required)
2 L per test (recommended)
Source: EPA, 2002a. Saltwater Chronic Methods Manual,
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.S. ENVIRONMENTAL PROTECTION AGENCY
Red Algal (Champia parvula) Sexual Reproduction Toxicity Tests
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D-2
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If you need additional copies of this document, you can download it at:
www.epa.gov/npdes/wqbasedpermitting
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WHOLE EFFLUENT TOXICITY • TRAINING VIDEO SERIES • saltwater series
Culturing Americamysis bahia
Supplement to Training Video
U.S. Environmental Protection Agency
Office of Wastewater Management
Water Permits Division
1200 Pennsylvania Ave., NW
Washington, DC 20460
EPA 833-C-09-001
March 2009
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NOTICE
The revision of this guide has been funded wholly or in part by the
Environmental Protection Agency under Contract EP-C-05-063.
Mention of trade names or commercial products does not constitute
endorsement or recommendation for use.
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Culturing Americomys/s bahia
Supplement to Training Video
Foreword
This guide serves as a supplement to the video "Culturing Americamys/s bahia" (EPA, 2009a). The meth-
ods illustrated in the video and described in this supplemental guide support the methods published in
the U.S. Environmental Protection Agency's (EPA's) Methods for Measuring the Acute Toxicity of Effluents
and Receiving Waters to Freshwater and Marine Organisms, Fifth Edition (2002a), referred to as the Acute
Methods Manual. The video and this guide provide details on culturing of mysids for the use in conducting
tests based on the expertise of personnel at the following EPA Office of Research and Development (ORD)
laboratories:
National Health and Environmental Effects Research Laboratory (NHEERL) - Atlantic Ecology Division
in Narragansett, Rhode Island
NHEERL - Gulf Ecology Division in Gulf Breeze, Florida
National Exposure Research Lab (NERL) - Ecological Exposure Research Division (EERD) in
Cincinnati, Ohio
This guide and its accompanying video are part of a series of training videos produced by EPA's Office of
Wastewater Management. The video entitled "Mysid (Americamys/s bahia) Survival, Growth, and Fecundity
Toxicity Tests" (EPA 2009b) complements the material in this video by explaining the 7-day short-term
chronic toxicity test method using mysids. This Saltwater Series includes the following videos and guides:
"Mysid (Americamys/s bahia) Survival, Growth, and Fecundity Toxicity Tests"
"Culturing Americamys/s bahia"
"Sperm Cell Toxicity Tests Using the Sea Urchin, Arbacia punctulata"
"Red Algal (Champia parvula) Sexual Reproduction Toxicity Tests"
"Sheepshead Minnow (Cyprinodon variegatus) and Inland Silverside (Menidia beryllina) Larval Survival
and Growth Toxicity Tests"
The Freshwater Series, released in 2006, includes the following videos and supplemental guides:
"Ceriodaphnia Survival and Reproduction Toxicity Tests"
"Culturing of Fathead Minnows (Pimephales promelas)"
"Fathead Minnow (Pimephales promelas) Larval Survival and Growth Toxicity Tests"
All of these videos are available through the National Service Center for Environmental Publications
(NSCEP) at 800 490-9198 or nscep@bps-lmit.com.
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Culturing Americamysis bahia
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U'S" ENVIRONMENTAL PROTECTION AGENCY
Culturing Americomysis bahia
Supplement to Training Video
Contents
Foreword i
Introduction 1
Water and Light 1
Culture Water 1
Photoperiod 2
Culture Vessels 2
Water Delivery Systems 2
Culture Start Up and Maintenance 3
Starting Cultures 3
Taxonomy 4
Collecting Test Organisms 4
Tank Cleaning 5
Record Keeping : 6
Food Preparation 7
Citations and Recommended References 8
Glossary Glossary-1
Appendix A: Apparatus and Equipment List A-l
FIGURES
Figure 1. The General Morphology of Mysids: (A) Lateral View; (B) Dorsal View 1
Figure 2. Intermittent Flow-Through Water Delivery System 3
Figure 3. Morphological Characteristics Used in Mysid Identification 4
Figure 4. Life Cycle of a Mysid 4
Figure 5. Illustration of Mysid Brood Chamber 5
Figure 6. Illustration of Mysid Generator 6
Figure 7. Data Form for Mysid Cultures 6
TABLE
Table 1. Recommended Culture Conditions for/Amer/camys/s bahia 2
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Culturlng Americamysis bahia
Supplement to Training Video
Introduction
Americamysis bahia, A. almyra, A. bigelowi, Metamysidopsis ilongata, and Neomysis americana, called
mysids or opossum shrimp, have all been used in toxicity tests. This guide focuses on Americamysis bahia,
the EPA-recommended species used in the mysid survival, growth, and fecundity toxicity test (Method
1007 in EPA, 2002b). Americamysis bahia are found in the coastal waters of the Gulf of Mexico and along
the Atlantic coast as far north as Rhode Island.
As shown in Figure 1, mysids usually appear transparent with a yellow, brown, or black tint and range from
4.4 mm to 9.4 mm in length (Molenock, 1969). Americamysis bahia differ from the other Americamysis
species by the armature of the telson and the spine-setaes on the thoracic and uropodal endopods
(Molenock, 1969; Price et al., 1994).
The culturing procedures
presented in this supplemen-
tal guide and illustrated in
the video were developed to
meet the specific needs of
the mysid in each of its life
stages. This guide and the
video "Culturing Americamysis
bahia" (EPA, 2009a) were
produced by EPA to clarify
and expand on methods
explained in the EPA manual
Methods for Measuring the
Acute Toxicity of Effluents
and Receiving Waters to
Freshwater and Marine
Organisms, Fifth Edition (EPA,
2002a). Laboratory person-
nel who are familiar with the
culturing and handling pro-
cedures of the test species
and the use of healthy test
organisms are critical for valid
and successful toxicity test
results.
Figure 1. The General Morphology of Mysids: (A) Lateral View; (B)
Dorsal View.
antennule
antenn'
dorsal process
statocyst
uropod
telson
endopod
exopod
Source: Heard and Price, 2006 as modified from Stuck et al., 1979a.
The first section of this guide covers the selection and preparation of the water for culturing and presents
options for water delivery systems. The second section explains how to set up and maintain mysid cultures
specifically for providing healthy test organisms. The third section provides instructions for collecting young
of the same age for testing. The fourth section provides details on the food preparation methods used at
NHEERL-AED in Narragansett, Rhode Island. This guide also includes a glossary and additional references.
Appendix A provides a list of the apparatus and equipment needed to culture mysids.
Water and Light
CULTURE WATER
Culture water is a primary consideration when starting mysid cultures. EPA recommends using natural
seawater. However, hypersaline brine may be used to make up culture water if natural seawater is not avail-
able. If natural seawater is used, it must be contaminant-free and filtered through a 0.45 urn screen before
use to remove particulates and possible predators. The source of the culture water should be uncontami-
I
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Culturing 4m eric am/sis bahia
Supplement to Training Video
nated, consistent, reliable, and periodically checked to ensure the water supports adequate performance
of the test organisms with respect to survival, growth, and reproduction. More specific instructions for
the preparation of artificial seawater are listed in EPA's Acute Methods Manual (EPA, 2002a) or can be
obtained from commercial suppliers. Optimum culture conditions, including water quality, are provided in
Table 1.
Table 1. Recommended Culture Conditions for Americamysis bahia
Parameter
Salinity
Temperature
PH
Dissolved oxygen
Ammonia
Nitrite
Nitrate
Alkalinity
Photoperiod
Filtration
Tank Size
Substrate
Biological filter / algal mat
Culture Conditions
25 g/l (20%0 - 30%o)
26°C± I°C
7.8-8.2
7.1 mg/L
O.I-0.3mg/L
<0.05 mg/L
<20 mg/L
ISO mg/L
12-hr light: 12-hr dark to 16-hr
light:8-hr dark
20 urn
10-55 gal
Dolomite, oyster shells, coral
Spirulina subsalsa
Source: Lussier et a/., 1988.
Reference toxicant tests should be conducted at least once each month to analyze both the culture water
being used and to check the mysid mass culture's sensitivity. Recommended reference toxicants are cop-
per sulfate, cadmium chloride, or sodium dodecyl sulphate.
PHOTOPERIOD
For optimum growth and fecundity, the photoperiod for mysid cultures should be 16 hours light and 8 hours
dark with a light intensity of about 50 - 100 foot-candles. EPA recommends using a system that turns the
lights on and off gradually so as not to startle the mysids, which can cause them to jump out of the culture
vessels. Alternatively, the light cycle can be provided using overhead room lights (cool-white fluorescent
bulbs, approximately 50 ft-c), supplemented with individual grow lights placed over each tank (approximate-
ly 65 ft-c). This arrangement allows the overhead lights to turn on one hour before the aquaria lights turn
on and to turn off one hour after they are extinguished.
CULTURE VESSELS
Mysids can be cultured in tanks of various sizes. The most commonly used are 20 and 29 gallon aquaria.
Wider tanks are more suitable for culturingthan taller ones because a large surface area to volume ratio
provides both good oxygen exchange and a larger surface area for these epibenthic organisms that prefer
to hover over the bottom of the tank. Tanks, as with all culturing equipment, should be cured in the culture
water for approximately 3-5 days before being used for organisms.
WATER DELIVERY SYSTEMS
Mysids can be cultured in flow-through, recirculating, or static systems. The preferred system is the flow-
through arrangement where water is delivered to the tanks at a measured rate and the runoff is discharged
out of the system (see Figure 2). The flow rate through the culture tanks should be no less than 4-5 liters
per hour or two complete turn-overs per day. Non-toxic materials such as glass, fiberglass, Teflon®, and
polyvinylchloride (PVC) pipe are recommended for the water delivery system. Materials such as rubber, cop-
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Culturing Americomys/s bah/a
Supplement to Training Video
Source: EPA 2002b.
per, brass, or plastic should Figure 2. Intermittent Flow-Through Water Delivery System
not be used because they
could become a source of
toxicity.
Recirculating systems also
can be used to culture mysids
and should be designed to
provide the same flow rate
as the flow-through sys-
tem. However, recirculating
systems must also provide a
biofiltering system that can
be constructed out of any
non-toxic, high-surface-area
material such as crushed
coral, pea gravel, or dolomite.
This biological filtration system
serves to oxidize the ammonia and nitrites that can build up in a closed system. A sand filter also may be
added to the system.
Static systems are made of a series of tanks that are independently filtered and supplied with water. The
advantage of this type of system is that problems such as disease are confined to one tank and complete
culture "crashes" (sudden death of a culture) are less common. Each tank in a static system should be
supplied with an under gravel filter and water changes should be made by replacing one-half of the tank's
volume of water with fresh culture water every other day. Static systems are harder to maintain than flow-
through or recirculating systems due to evaporation. Tanks should be covered and care must be taken to
avoid the concentration of salts as the water evaporates.
Culture Start Up and Maintenance
STARTING CULTURES
Once the culture system and water source are designed, obtained, and seasoned, mysids can be pur-
chased from a number of sources. A reliable supplier will certify that the correct species has been shipped.
Records of the verification should be retained with a few preserved organisms. If test animals are not
needed immediately, cultures should be started with juveniles to allow laboratory personnel to become
familiar with mysid handling and maintenance requirements before learning to collect the young.
Mysids should be shipped in Nalgene® containers packed inside coolers or polyfoam boxes within card-
board shipping cartons. The shipping density should be <100 mysids per liter and the container should
have 2 - 4 cm of airspace to ensure a supply of oxygen throughout the shipping period. No food should
be added to the containers. A reliable overnight delivery service should be used for shipment so that the
mysids are not in transit without food for more than 24 hours.
After the shipment is received, the mysids must be acclimated to the receiving laboratory's culture water
and conditions. The temperature and salinity of the water used for shipping must be measured. Slow
adjustment of the water temperature can be accomplished by placing the container in a water bath. The
salinity can be adjusted by adding new culture water to the water used for shipment. Increases or decreas-
es in temperature and salinity should not exceed 2°C or 2%p - 3%o, respectively, per day.
For optimum growth and reproduction, the stocking density for adult mysids should be approximately 20
mysids per liter. Juveniles can be stocked at higher densities than adults. A healthy, unstressed culture
should have at least 70% of the females carrying eggs in their brood pouch.
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Culturing Americamysis bahia
Supplement to Training Video
TAXONOMY
Mysids usually appear trans-
parent with a yellow, brown,
or black tint and range from
4.4 mm to 9.4 mm in length
(Molenock, 1969). The mor-
phological characteristics used
to distinguish A. bahia from
other mysids are presented in
Figure 3.
Figure 4 shows the life cycle of
a mysid. Mysids produce live
young called early juveniles.
These juveniles are planktonic
for the first 24 hours post-
release and then settle to the
bottom where they orient to
the current in the tank and
begin to feed. Depending on
water temperature and diet,
females reach sexual matu-
rity in about 20 days. Brood
pouches appear at the age
of 12 - 16 days and young
are released at approximately
20 days. A gravid female is
identified by an enlarged and
darkened brood pouch contain-
ing the developing embryos.
The female is ready to release
the young when the eyespots
can be identified in the brood
pouch. Females average 5 -
7 young per brood, but can
produce as many as 20 in one
brood. Broods are produced
for several months at a rate of
one every 4-6 days.
Figure 3. Morphological Characteristics Used in Mysid
Identification
Collecting Test
Organisms
To conduct toxicity tests using
mysids, organisms of the
same age must be collected
and pooled. To accomplish
this, gravid females are col-
lected from the culture tanks
and their young are collected
and held until the proper age
for starting tests. For testing
Morphological features most useful in identifying Americamysis bahia. a.
male; b. female; c. thoracic leg 2; d. telson; e. right uropod, dorsal; f. male,
dorsal (redrawn from Molenock, 1969; Heard et al, 1987). Note testes in
area where marsupium is located on female and length of male pleopods
as compared to female. Also note the three spines on the endopod of the
uropod (e).
Source: Molenock, 1969; Price et al., 1994
Figure 4. Life Cycle of a Mysid
Day 20
First Brood
Release
Day 12
Secondary Sex
Characteristics
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Culturing Americamysis bahia
Supplement to Training Video
needs, assume a reproduc-
tion rate of two juveniles per
female per day because not all
females will release their young
on the same day. Collect the
gravid females from a minimum
of three culture tanks. While
identifying and selecting gravid
females for the brood chamber,
the sex ratio and density of each
tank should be determined and
adjusted, if needed, to maintain
a ratio of 2 females:! male.
Brood chambers such as the one
illustrated in Figure 5 are used
to collect test animals. Gravid
females are collected from a
minimum of three culture tanks
and placed in a 4 L Nalgene®
beaker that is placed inside a
separatory funnel containing
culture water. The solid plastic
bottom of the Nalgene® beaker is
replaced by 1 mm mesh screen.
The screen allows the newly
released young to pass through
while preventing the adults from leaving the beaker.
Figure 5. Illustration of Mysid Brood Chamber
INFLOW
OUTFLOW
NETTED
CHAMBER
SEPARATORY
FUNNEL
NETTED
CULTURE DISH
_-^
Source: Lussier, et a/., 1987.
Once the females are placed in the brood chamber, provide food and gentle aeration by either placing
an airstone in the neck of the separatory funnel or providing water inflow and outflow to the funnel. The
females should be left overnight and the young collected the next day.
To harvest the young, remove the airstone or stop the flow of water and slowly drain the separatory cham-
ber into a 300 urn mesh cup placed in a culture dish. To prevent injury to the young mysids, partially
submerge the mesh cup in culture water within the culture dish before draining the brood chamber. While
the water is draining, gently lift and dunk the beaker containing the females to wash any remaining young
out through the screen. As the water drains from the funnel, gently rinse the sides 2 - 3 times with clean
seawater to wash out any mysids that may stick to the sides. The females should be placed back into the
culture tanks. The young can be used immediately for testing or grown out in a separate tank to the desired
age. The harvested young should be maintained at conditions similar to the regular cultures.
An alternative system for collecting young is a siphon entrapment system, or a "mysid generator" (see
Figure 6). The siphon inlet is covered by a 750 urn screen that excludes adults and allows juveniles to pass
through to a collection vessel. In the collection vessel juveniles are deposited into a 350 - 370 urn Nitex®
screen cup. The juveniles in the screen cup are collected daily for test use. When using mysid generators,
care must be taken to siphon all of the juveniles out of the tank each day. Otherwise, the collected juve-
niles' ages may not be within 24 hours of each other as test methods require.
TANK CLEANING
Culture tanks should be cleaned at least once each month. The sides of each tank should be scraped to
remove any algal growth and the gravel should be stirred to dislodge the accumulated debris, which will
clear the dolomite filter.
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Culturing Americamysis bahia
Supplement to Training Video
. - ^ ..t. », «
Approximately,twice each year,
the tanks should be completely
emptied and scrubbed. At this
time the gravel should also
be replaced. It is important
to cure any new materials as
described in the previous sec-
tion "Waterand Light, Culture
Vessels," before using them in
culture tanks.
RECORD KEEPING
Culture tanks should be
monitored and all conditions
recorded on data forms that
are kept in a permanent file.
These forms are used to
assess any problems that may
occur with the cultures and
assist in eliminating possible
causes. The forms also serve
as a record for testing labora-
tories to verify that their test
organisms were raised using
proper culture techniques.
Figure 6. Illustration of Mysid Generator
Pump Return
Tube
Filter
Incoming
Siphon
Culture Tank
(75 L)
Overflow
Collection
Tank
Source: Lussier et a/., 1988.
Figure 7 is a data sheet adapted from the one used by AED-Narragansett for mysid cultures. Each of the
conditions is checked daily and initialed by the technician taking the reading, checking the condition, or
performing the task. Daily tasks performed are measurement of temperature, pH, salinity, and dissolved
oxygen; seawater and airflow checks; and feeding (twice daily).
Figure 7. Data Form for Mysid Cultures
Dote
Temp
°C
pH
SU
Salinity
%0
DO
mg/L
SW
Flow
Air
Flow
Mysids.
Fed
Comments
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Culturing Americamysis bahia
Supplement to Training Video
Food Preparation
Mysid cultures are fed Artemia nauplii (newly-hatched brine shrimp) twice each day at a rate that ensures
live Artemia are always available in the tanks (approximately 150 Artemia nauplii per mysid per day). The
Artemia should be cultured in the laboratory in order to provide 24 - 48 hour old nauplii on a daily basis.
Artemia cysts are available from commercial suppliers. Each shipment of Artemia received should be
analyzed for priority pollutants and should be tested on a small batch of mysids to ensure that good mysid
growth and reproduction occur before the Artemia are fed to entire mysid cultures. Food supplements are
commercially available and are used more often when using artificial seawater for culturing.
Culture the Artemia-by adding dry cysts to clean seawater at a rate of approximately 10 ml cysts to 1 L
seawater (ASTM, 1998). A separatory funnel works well for culturing Artemia. Inverted two-liter plastic
bottles also have been used by cutting out their bottoms and inserting a rubber stopper with a flexible tube
and pinch clamp.
After placing the water and cysts into the culture chamber, aerate vigorously to keep the cysts (and eventu-
ally the newly-hatched nauplii) in suspension. Deliver the filtered air through a 1 ml pipet by resting the-tip
of the pipet at the bottom of the neck of the chamber. This keeps the nauplii from settling and depleting
the oxygen supply.
IMPORTANT NOTE:
The nauplii must be aerated
if they remain unused for
more than a few minutes.
Without aeration the nauplii
will begin to die.
The cysts will hatch in approximately 24 hours. Newly-hatched Artemia
nauplii are more nutritious than older ones and are the appropriate
size for feeding early juvenile mysids. To harvest the nauplii for feeding,
remove the air supply and allow the cysts and nauplii to separate for five
minutes. The empty cysts will float and the nauplii will descend to the
neck of the chamber. The nauplii are attracted to light, so a light source
placed at the bottom of the chamber and/or a dark cover or hood
placed on the top will hasten the separation process.
Drain the nauplii through the stop clamp or siphon them from the bottom of the chambers. If the nauplii
are drained through the stop cock, the first plug of unhatched cysts that collect at the neck of the cham-
ber should be discarded and not mixed with the nauplii. Drain only the hatched nauplii (the bright orange
suspension), leaving behind the empty cysts. The nauplii should be drained through a 150 urn screen and
rinsed with clean seawater to remove any chemicals released during hatching.
To determine the correct amount of Artemia for feeding, an aliquot of the hatched Artemia should be
counted under a microscope to determine the density of the culture. This density will serve as a reference
to ensure that future cultures are hatching at the same rate and that mysids are being fed a consistent
amount of food.
Once the Artemia are rinsed, the volume of clean seawater that is added determines the volume of food
provided to each tank. From the calculated and adjusted density of the diluted food supply, determine the
volume of food needed for each tank by estimating a feeding rate of 150 Artemia per mysid per day, or 75
Artemia per mysid per feeding. Feeding the mysids in two feedings, 8-12 hours apart ensures there are
always live Artemia available for the mysids.
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Culturing Americamysis bahia
Supplement to Training Video
Citations and Recommended References
American Society for Testing and Materials. 1998. Standard Practice for Using Brine Shrimp Nauplii as
Food for Testing Animals in Aquatic Toxicology. ASTM Designation E-1203-98. Philadelphia, PA.
Blaxter, J.H.S., F.S. Russell, and M. Yonge, eds. 1980. The biology of mysids and euphausiids. Part 1. The
biology of the mysids. Adv. Mar. Biol. 18:1-319.
Brattegard, T. 1969. Marine Biological Investigations in the Bahamas 10. Mycidacea from shallow water
in the Bahamas and southern Florida, Part 1. Sarsia 39:17-106.
Davey, E.W., J.H. Gentile, S.J. Erickson, and P. Betzer. 1970. Removal of trace metals from marine culture
media. Limnol. Oceanogr. 15:486-488.
EPA. 1991. Technical Support Document for Water Quality-based Toxics Control. U.S. EPA Office of Water
Enforcement and Permits, Washington, D.C. EPA-505-2-90-001.
EPA. 2000a. Method Guidance and Recommendations for Whole Effluent Toxicity (WET) Testing (40 CFR
Part 136). Office of Water, Washington, D.C. EPA 821-B-00-004.
EPA. 2000b. Understanding and Accounting for Method Variability in Whole Effluent Toxicity Applications
Under the National Pollutant Discharge Elimination System Program. Office of Wastewater
Management, Washington, D.C. EPA 833-R-00-003.
EPA. 2002a. Methods for Measuring the Acute Toxicity of Effluents and Receiving Waters to Freshwater
and Marine Organisms, Fifth Edition. (Acute Methods Manual). Office of Water, Cincinnati, OH.
EPA-821-R-02-012.
EPA. 2002b. Short-term Methods for Estimating the Chronic Toxicity of Effluents and Receiving Waters
to Marine and Estuarine Organisms, Third Edition. (Chronic Methods Manual). Office of Water,
Cincinnati, OH. EPA-821-R-02-014.
EPA. 2009a. Culturing Americamysis bahia. Supplement to Training Video. Whole Effluent Toxicity Training
Video Series, Saltwater Series. March 2009. EPA 833-C-09-001.
EPA. 2009b. Mysid (Americamysis bahia) Survival, Growth, and Fecundity Toxicity Tests. Supplement to
Training Video. Whole Effluent Toxicity Training Video Series, Saltwater Series. March 2009. EPA
833-C-09-001.
Farrell, D.H. 1979. Guide to the shallow-water mysids from Florida. Fla. Dept. Environ. Reg., Techn. Ser.
Fotheringham, N., and S.L Brunenmeister. 1975. Common marine invertebrates of the northwestern Gulf
coast. Gulf. Publ. Co., Houston, TX.
Heard, R.W. 1982. Guide to the common tidal marsh invertebrates of the northeastern Gulf of Mexico.
Publ. No. MASGP-79-004, Mississippi-Alabama Sea Grant Consortium, Ocean Springs, MS.
Heard, R.W., W.W. Price, and K.C. Stuck. 1987. Mysid Identification Workshop. The Gulf Coast Research
Laboratory, Ocean Springs, Mississippi (unpublished.)
Heard, R.W. and W.W. Price. 2006. A taxonomic Guide to the Mysids of the South Atlantic Bight. U.S.
Department of Commerce, National Oceanic and Atmospheric Administration.
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Culturing Americamysis bahia
Supplement to Training Video
Johns, D.M., WJ. Berry, and W. Walton. 1981. International study on Artemia. XVI. Survival, growth, and
reproductive potential of the mysid Mysidopsis bahia Molenock fed various geographical strains
of the brine shrimp Artemia. J. Exp. Mar. Biol. Ecol. 53:209-219.
Lawler, A.R. and S.L. Shepard. 1978. Procedures for eradication of hydrozoan pests in closed-system
mysid culture. Gulf Res. Rept.
Lussier, S.M., A. Kuhn, and J. Sewall. 1987. Guidance manual for conducting 7-day mysid survival/
growth/reproduction study using the estuarine mysid, Mysidopsis bahia. Contribution No. X106.
In: Schimmel, S.C., ed. Users guide to the conduct and interpretation of complex effluent toxic-
ity tests at estuarine/marine sites. Environmental Research Laboratory, U.S. Environmental
Protection Agency, Narragansett, Rhode Island. Contribution No. 796, 265 pp.
Lussier, S.M., A. Kuhn and R. Comeleo. 1999. An evaluation of the seven-day toxicity test with
/Amer/camys/s bahia (formerly Mysidopsis bahia). Environ. Toxicol. and Chem. 18:2888-2893.
[Errata: in the section on Experimental Design, the test chamber should read "200-ml plastic cup"
not "30-ml."]
Lussier, S.M., A. Kuhn, MJ. Chammas, and J. Sewall. 1988. Techniques for the laboratory culture of
Mysidopsis species (Crustacea: Mysidacea). Environ. Tox. Chem. 7:969-977.
Molenock, J. 1969. Mysidopsis bahia, a new species of mysid (Crustacea: Mysidacea) from Galveston Bay,
Texas. Tulane Stud. Zool. Bot. 15(3):113-116.
Nimmo, D.R. and T.L. Hamaker. 1982. Mysids in toxicity testing - a review. Hydrobiol. 93:171-178.
Nimmo, D.R., T.L. Hamaker, E. Matthews, and W.T. Young. 1982. The long-term effects of suspended par-
ticulates on survival and reproduction of the mysid shrimp, Mysidopsis bahia, in the laboratory. In:
G.F. Mayer, ed., Ecological Stress and the New York Bight: Science and Management. Estuarine
Res. Found., Columbia, S.C. pp. 41-50.
Nimmo, D.R., T.L. Hamaker, C.A. Sommers. 1978. Culturing the mysid (Mysidopsis bahia) in flowing sea-
water or a static system. In: Bioassay Procedures for the Ocean Disposal Permit Program, U.S.
Environmental Protection Agency, Environmental Research Laboratory, Gulf Breeze, Florida. EPA-
600/9-78-010. pp. 59-60.
Price, W.W., R.W. Heard and L. Stuck. 1994. Observations on the genus Mysidopsis sars. 1864 with the
designation of a new genus, Americamysis, and the descriptions of Americamysis aliens and A.
stuck! (Peracarida: Mysidacea: Mysidae), from the Gulf of Mexico. Proc. Biol. Soc. Wash. 107: 680-
698.
Price, W.W. 1982. Key to the shallow water Mysidacea of the Texas coast with notes on their ecology.
Hydrobiol. 93(l/2):9-21.
Stuck, K.C., H.M. Perry, and R.W. Heard. 1979a. An annotated key to the Mysidacea of the North Central
Gulf of Mexico. Gulf Res. Rept. 6(3):225-238.
Stuck, K.C., H.M. Perry, and R.W. Heard. 1979b. Records and range extensions of Mysiacea from coastal
and shelf water of the Eastern Gulf of Mexico. Gulf Res. Rept. 6(3):239-248.
Venables, B. 1986. Mysidopsis sp.: Life history and culture workshop report. Gulf Breeze, FL. October
15-16,1986, Institute of Applied Sciences, North Texas State University, Denton, TX.
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Culturing Americamysis bahia
Supplement to Training Video
U.S. ENVIRONMENTAL PROTECTION AGENCY
Ward, S.H. 1984. A system for laboratory rearing of the mysid, Mysidopsis bahia Molenock. Progr. Fish-
Cult. 46(3):170-175.
Wigley, R.L. and B.R. Burns. 1971. Distribution and biology of mysids (Crustacea: Mysidacea) from the
Atlantic Coast of the United States in the NMFS Woods Hole Collection. Fish. Bull. 69:717-746.
EPA references are available online atwww.epa.gov/npdes.
If you need additional copies of this document, you can download it at:
www.epa.gov/npdes/wqbasedpermitting.
10
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Culturing Americomysis bahia
Supplement to Training Video
Glossary
Artemia. The marine invertebrate (referred to as brine shrimp) used as the recommended food source for
mysid cultures and test organisms; Brazilian or Colombian strains are preferred because the sup-
plies are found to have low concentrations of chemical residues and nauplii are of suitably small
size.
Crash. Sudden (overnight) death of cultured organisms in a tank.
Cyst. The life stage of unhatched Artemia.
Epibenthic. Pertaining to the area just above the sediment.
Fecundity. Productivity or fertility as measured in the mysid test as the percentage of females with eggs
in the oviduct and/or brood pouch.
Flow-through water delivery system. An open water flow system that delivers fresh water or seawater to
culture tanks, which is disposed of after it leaves those tanks.
Mysid (Americamysis bahia). An estuarine crustacean, formerly known as Mysidopsis bahia, ranging 4.4
mm to 9.4 mm in length found from the Gulf of Mexico and along the Atlantic coast as far north as
Rhode Island; used in test procedures as an indicator species for aquatic toxicity.
Nauplii. Free-swimming microscopic larvae stage characteristic of copepods, ostracods, barnacles, etc.
typically with only three pairs of appendages.
Recirculating water delivery system. A water flow system that treats water after it passes through the
culture tanks (usually with sand and biofilters) and delivers the same treated water back to the
tanks.
Static water system. An enclosed system contained within one culture tank. The water is filtered through
an underground or charcoal filter and is delivered back to the same tank.
WET (Whole effulent toxicty). The total toxic effect of an effluent measured directly with a toxicty text.
Glossary-1
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ISC) U'S- ENVIRONMENTAI- PROTECTION AGENCY Culturing Americamysif bahia
V=«^ Supplement to Training Video
Intentionally Left Blank
Glossary-2
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Culturing Americamysis bahia
Supplement to Training Video
Appendix A
Apparatus and Equipment List
Air line and air stones. For aerating cultures, brood chambers, and holding tanks, and supplying air to test
solutions with low DO.
Air pump. For oil-free air supply.
Balance. Analytical, capable of accurately weighing to 0.00001 g.
Beakers or flasks. Six, borosilicate glass or non-toxic plasticware, 2 - 3 L for making test solutions.
Brine shrimp (Artemia) culture unit. See "Food Preparation" section.
Depression glass slides or depression spot plates. Two for observing organisms.
Dissecting microscope (240 - 400X magnification). For examining organisms to determine their sex and
to check for the presence of eggs in the oviducts of the females.
Droppers, and glass tubing with fire polished edges. 4 mm inner diameter (ID), for transferring
organisms.
Environmental chamber or equivalent facility with temperature control (26 ± 1°C).
Facilities for holding and acclimating test organisms.
Light box. For illuminating organisms during examination.
Meters: pH and DO, and specific conductivity. For routine physical and chemical measurements.
Mysid (Americamysis bahia) culture unit. See "Culture Start Up and Maintenance" section. The test
requires a minimum of 240 7-day old (juvenile) mysids.
NITEX® or stainless steel mesh sieves. 150 urn and 100 Mm for concentrating organisms; 1 mm mesh
and 300 |jrn mesh for collection of juveniles.
Pipet bulbs and fillers. Propipet®, or equivalent.
Pipets, automatic. Adjustable, 1 - 100 ml_.
Pipets, serological. 1-10 mL, graduated.
Pipets, volumetric, Class A. 100 ml.
Reference weights, Class S. For checking performance of balance.
Refractometer or other method. For determining salinity.
Separatory funnels, 2-liters. Two to four for culturing/Artem/a.
Standard or micro-Winkler apparatus. For determining DO and checking DO meters.
Thermometers, bulb-thermograph or electronic-chart type. For continuously recording temperature.
Thermometers, glass or electronic, laboratory grade. For measuring water temperatures.
A-1
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Culturing Amencamysis bahia
Supplement to Training Video
Thermometers. National Bureau of Standards Certified (see EPA, 2002b). Used to calibrate laboratory
thermometers.
Volumetric flasks and graduated cylinders. Class A, borosilicate glass or non-toxic plastic labware,
50 - 2000 ml for making test solutions.
Wash bottles. For deionized water, for washing organisms from containers and for rinsing small glassware
and instrument electrodes and probes.
Water purification system. Millipore® Milli-Q® deionized water or equivalent.
A-2
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If you need additional copies of this document, you can download it at:
www.epa.gov/npdes/wqbasedpermitting
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/HOLE bFFLUENT IOXICITY • I RAINING VIDEO SERIES • Saltwater Series
Mysid (Americamysi's bah/a)
Survival, Growth, and Fecundity
Toxicity Tests
Supplement to Training Video
U.S. Environmental Protection Agency
Office of Wastewater Management
Water Permits Division
1200 Pennsylvania Ave., NW
Washington, DC 20460
EPA 833-C-09-001
March 2009
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NOTICE
The revision of this guide has been funded wholly or in part by the
Environmental Protection Agency under Contract EP-C-05-063.
Mention of trade names or commercial products does not constitute
endorsement or recommendation for use.
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U.S. ENVIRONMENTAL PROTECTION AGENCY Mysid (Xtmericam/sis bah/a) Survival, Growth, and Fecundity Toxicity Tests
Supplement to Training Video
Foreword
This guide serves as a supplement to the video "Mysid (Americamysis bahia) Survival, Growth, and
Fecundity Toxicity Tests" (EPA, 2009a). The methods illustrated in the video and described in this sup-
plemental guide support the methods published in the U.S. Environmental Protection Agency's (EPA's)
Short-term Methods for Estimating the Chronic Toxicity of Effluents and Receiving Waters to Marine and
Estuarine Organisms, Third Edition (EPA, 2002a), referred to as the Saltwater Chronic Methods Manual.
The video and this guide provide details on preparing for and conducting the test based on the expertise of
personnel at the following EPA Office of Research and Development (ORD) laboratories:
National Health and Environmental Effects Research Laboratory (NHEERL) - Atlantic Ecology Division
in Narragansett, Rhode Island
NHEERL - Gulf Ecology Division in Gulf Breeze, Florida
National Exposure Research Lab (NERL) - Ecological Exposure Research Division (EERD) in
Cincinnati, Ohio
This guide and its accompanying video are part of a series of training videos produced by EPA's Office of
Wastewater Management. This Saltwater Series includes the following videos and guides:
"Mysid (/Amer/camys/s bahia) Survival, Growth, and Fecundity Toxicity Tests"
"CulturingX\mer/camys/s bahia"
"Sperm Cell Toxicity Tests Using the Sea Urchin, Arbacia punctulata"
"Red Algal (Champia parvula) Sexual Reproduction Toxicity Tests"
"Sheepshead Minnow (Cyprinodon variegatus) and Inland Silverside (Menidia beryllina) Larval Survival
and Growth Toxicity Tests"
The Freshwater Series, released in 2006, includes the following videos and supplemental guides:
"Ceriodaphnia Survival and Reproduction Toxicity Tests"
"Culturingof Fathead Minnows (Pimephales promelas)"
"Fathead Minnow (Pimephales promelas) Larval Survival and Growth Toxicity Tests"
All of these videos are available through the National Service Center for Environmental Publications
(NSCEP) at 800 490-9198 or nscep@bps-lmit.com.
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U.S. ENVIRONMENTAL PROTECTION AGENCY Mysid (Americamysis bahia) Survival, Growth, and Fecundity Toxicity Tests
Supplement to Training Video
Intentionally Left Blank
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U.S. ENVIRONMENTAL PROTECTION AGENCY Mysid (Americomysis bahia) Survival, Growth, and Fecundity Toxicity Tests
Supplement to Training Video
Contents
Foreword i
Tables iv
Figures iv
Introduction 1
Background 1
Maintaining and Feeding Cultures 1
Culture Maintenance 1
Feeding 2
Collecting Juveniles for Test Use 2
Conducting the Test .3
Effluent Sampling 3
Dilution Preparation 3
Routine Chemistries 4
Test Chambers 5
Test Organisms 6
Feeding 6
Renewals 6
Terminating the Test 8
Test Acceptability and Data Review 11
Citations and Recommended References 12
Glossary Glossary-1
Appendix A: Summary of Test Conditions and Test Acceptability Criteria A-l
Appendix B: Apparatus and Equipment List B-l
Appendix C: Reagents and Consumable Materials C-l
Appendix D: Preparing Hypersaline Brine (MSB) D-l
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U.S. ENVIRONMENTAL PROTECTION AGENCY Mysid (Americomysis bohia) Survival, Growth, and Fecundity Toxicity Tests
Supplement to Training Video
FIGURES
Figure 1. The General Morphology of Mysids. (A) Lateral View; (B) Dorsal View 2
Figure 2. Apparatus for Collection of Juvenile Mysids from Gravid Females 2
Figure 3. Data Form for the Mysid Survival and Fecundity Toxicity Test - Water Quality Data 5
Figure 4. Data Form for the Mysid Survival and Fecundity Toxicity Test - Survival and
Fecundity Data 7
Figure 5. Data Form for the Mysid Survival and Fecundity Toxicity Test - Dry Weight Measures .. 8
Figure 6. Mature Female A. bah/a with Eggs in Oviducts. Lateral view (top) Dorsal view (bottom) 9
Figure 7. Mature Female A. bahia with Eggs in Oviducts and Developing Embryos in Brood Sac.
Lateral view (top) Dorsal view (bottom) 10
Figure 8. Mature Male A. bahia. Lateral view (top) Dorsal View (bottom) ..." 10
Figure 9. Immature A. bahia. Lateral view (top) Dorsal view (bottom) 11
TABLES
Table 1. Monitoring Schedule 4
Table A-l. Summary of Test Conditions and Test Acceptability Criteria for Americamysis bahia
7-day Survival, Growth, and Fecundity Toxicity Test A-l
Table D-l. Preparation of Test Solutions at a Salinity of 20%o Using HSB for a Final Test
Concentration Volume of 2000 mL D-2
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U.S. ENVIRONMENTAL PROTECTION AGENCY Mysid (Americomysis bahia) Survival, Growth, and Fecundity Toxicity Tests
Supplement to Training Video
Introduction
This supplemental guide accompanies the Environmental Protection Agency's (EPA's) video to provide
instructions for conducting the standard 7-day survival, growth, and fecundity toxicity test using the
mysid, Americamysis bahia (EPA, 2009a; EPA, 2009b). The test method is found in Short-term Methods
for Estimating the Chronic Toxicity of Effluents and Receiving Waters to Marine and Estuarine Organisms,
Third Edition (EPA, 2002a). The methods presented in this guide and the video are based on the expe-
rience and standardized practices developed at EPA's Office of Research and Development's (ORD's)
National Health and Environmental Effects Research Laboratory-Atlantic Ecology Division (NHEERL-AED)
in Narragansett, Rhode Island. The material presented in both the video and this guide summarizes the
methods but does not replace a thorough review and understanding of the methods by laboratory person-
nel before conducting the test.
Background
Under the National Pollutant Discharge Elimination System (NPDES) program (Section 402 of the Clean
Water Act), EPA uses toxicity tests to monitor and evaluate effluents for their toxicity to biota and their
impact on receiving waters. By determining acceptable or safe concentrations for toxicants discharged
into receiving waters, EPA can establish NPDES permit limitations for toxicity. These whole effluent toxicity
(WET) permit limitations regulate pollutant discharges on a whole effluent effect basis rather than solely by
a chemical-specific approach.
The mysid survival, growth, and fecundity toxicity test (Test Method 1007.0 in EPA, 2002a) is used by EPA
for determining the toxicity of marine or estuarine discharges by measuring specified endpoints after a
7-day exposure period. Whole effluent toxicity methods measure the synergistic, antagonistic, and addi-
tive effects of all the chemical, physical, and additive components of an effluent that adversely affect the
physiological and biochemical functions of the test organisms. Therefore, healthy organisms and correct
laboratory procedures are essential for valid test results. Laboratory personnel should be very familiar with
the test methods and with mysid handling techniques before conducting a test.
This supplemental guide covers the procedures for conducting the test according to EPA's promulgated
methods (40 CFR Part 136; EPA, 2002c) and also provides some helpful information that is not presented
in the Saltwater Chronic Methods Manual (EPA, 2002-a).
Maintaining and Feeding Cultures
CULTURE MAINTENANCE
/4mericamys/s bahia (mysids, or opossum shrimp) are estuarine invertebrates generally found in the coastal
waters of the Gulf of Mexico and along the Atlantic coast as far north as Rhode Island (see Figure 1). They
usually appear transparent with a yellow, brown, or black tint and range from 4.4 mm to 9.4 mm in length
(Molenock, 1969). Adult mysids can be collected from the field, however, they must be verified taxonomi-
cally as the correct species before being placed in cultures for test use (Price et al., 1994). Alternatively,
commercial suppliers provide adults for cultures and juveniles for cultures or testing. The supplier should
verify that the correct species is sent.
Cultures should be maintained in glass aquaria supplied with flow-through or recirculating seawater
(Lussier et al., 1988). The water temperature should be 26°C and salinity between 20%o to 30%0 and
should not fluctuate more than 2°C or 2%o per day, respectively. The light regime recommended for cultur-
ing is 16 hours light and 8 hours dark. The light should be phased on and off gradually so as not to startle
the mysids.
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Mysid (Americamyfis bahia) Survival, Growth, and Fecundity Toxicity Tests
Supplement to Training Video
FEEDING
Mysids are fed <24-hr old
Artemia nauplii (newly hatched
brine shrimp) twice daily.
Feeding amounts should be
adequate to provide live food
at all times for the mysids
to feed upon. Approximately
150 Artemia per mysid per
day is recommended. Artemia
supplies should be checked
periodically for contamination
and hatch rates.
Detailed instructions on
culturing/4rtem/a are pre-
sented in the video "Culturing
Americamysis bahia," and its
accompanying supplemental
guide, and in the EPA manual
Methods for Measuring the
Acute Toxicity of Effluents
and Receiving Waters to
Freshwater and Marine
Organisms, Fifth Edition
(EPA, 2009b; EPA, 2002b).
Collecting Juveniles
for Test Use
The 7-day survival, growth, and
fecundity toxicity test must be
started with 7-day old mysids that
are all within 24 hours age of each
other. Seven-day old juveniles are
needed in sufficient number to
randomly select five juveniles for
each replicate. For a test with five
effluent concentrations and one
control, with 8 replicates at each
concentration, it is recommended
to have approximately 240 -
300, 7-day old mysids available
to choose from. Avoid using any
mysids that appear injured.
To collect juveniles and to be
assured of their age range (within
24-hours age), a brood chamber
is used (see Figure 2). The brood
chamber is set up eight days
before the start of the test.
Figure 1. The General Morphology of Mysids. (A) Lateral View; (B)
Dorsal View
antennule
antenna
dorsal process
statocyst
thoracic segments dorsal process
Source: Heard and Price, 2006 as modified from Stuck et a/., 1979.
Figure 2. Apparatus for Collection of Juvenile Mysids from Gravid
Females
INFLOW
OUTFLOW
NETTED
CHAMBER
SEPARATORY
FUNNEL
NETTED
'CHAMBER
CULTURE DISH
Source: Lussier et a/., 1987.
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U.S. ENVIRONMENTAL PROTECTION AGENCY
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Supplement to Training Video
Gravid females selected from a minimum of three culture tanks are placed in a netted chamber inside a
funnel. Gravid females are those ready to release their young and are identified by dark spots in their brood
pouches. Because not all of the females will release young on the same day, an estimate of two juveniles
per female per day should be used to determine the number of gravid females needed. Therefore, to have
sufficient mysids for test initiation, approximately 125 - 150 gravid females should be placed in the brood
chamber.
Twenty-four hours after placing the females in the brood chamber, or seven days before the test start date,
remove the netted chamber containing the gravid females from the brood chamber allowing the juveniles to
escape through the screened bottom. Return the females to the culture tanks and drain the juveniles from
the funnel into a mesh cup placed in a dish containing culture water. To prevent injury to the test animals,
gently rinse the sides of the funnel as it drains. These juveniles, all born within the last 24 hours should
be counted and transferred into a separate tank where they will be held for the next seven days. Because
stocking density is very important to the rate of juvenile development, no more than 300 juveniles should
be held in a 10-gallon tank. If the holding tank used is a static system, half of the water must be replaced
every other day with new culture water.
Nutrition and temperature are important factors in mysid development (Lussier et al., 1999). During the
7-day holding period maintain the holding tanks at 26°C - 27°C with a salinity similar to the culture/test
water. If necessary, the salinity should be gradually adjusted (<2%o/day) to the desired test salinity (20%o -
30%o) during this holding period. Feed the juveniles <24-hour old Artemia nauplii twice daily.
Conducting the Test
Under the NPDES program, lapsed
time from sample collection to first
use of that sample in a toxicity
test (i.e., test initiation) must not
exceed 36 hours. If stored correctly,
the sample may be used for test
renewals at 24 hours, 48 hours,
and/or 72 hours after test initiation.
EFFLUENT SAMPLING
Effluent sampling should be conducted according to the EPA Saltwater
Chronic Methods Manual (EPA, 2002a) and any conditions specified
in a regulatory permit. In static renewal tests, each grab or composite
sample may be used to prepare test solutions for renewal at 24, 48,
and/or 72 hours after first use if stored between 0°C - 6°C, with mini-
mum head space. According to the EPA 2002 promulgated methods, for
WET samples with a specified storage temperature of 4°C, storage at a
temperature above the freezing point of water to 6°C shall be acceptable
(0°C - 6°C). EPA has further clarified that hand-delivered samples used
on the day of collection do not need to be cooled to 0°C - 6°C prior to
test initiation (EPA, 2002c).
Dilution Water
The type of dilution water used to make the test concentrations is
dependent on the objectives of the test. Any specific requirements
included in NPDES permits should be followed. The Saltwater Chronic
Methods Manual (Section 7) provides the following guidelines:
• If the test is conducted to estimate the absolute chronic toxicity
of the effluent, synthetic dilution water should be used. If the cultures
were maintained in different water than used for dilution water, a
second set of control replicates should be conducted using the culture
water.
• If the test is conducted to estimate the chronic toxicity of the
effluent in uncontaminated receiving waters, the test (com.)
DILUTION PREPARATION
To start a test, warm the effluent to
26°C ± 1°C slowly to avoid exceed-
ing the desired temperature. This
is accomplished using a water bath
and monitoring the temperature
closely. A temperature of 26°C ± 1°C
should be maintained throughout the
7-day test period and the instanta-
neous temperature must not deviate
by more than 3°C during the test.
Once the effluent and the dilution
water reach the desired tempera-
ture, the dilutions are prepared.
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Mysid (Americomysis bahia) Survival, Growth, and Fecundity Toxicity Tests
Supplement to Training Video
Dilution Water (cont.)
can be conducted using a grab sample of the receiving waters collected
outside the influence of the outfall, other uncontaminated waters, or
standard dilution water with the same salinity as the receiving waters.
If the cultures were maintained in different water than used for dilution
water, a second set of control replicates should be conducted using the
culture water.
• If the test is conducted to estimate the additive or mitigating
effects of the effluent on already contaminated receiving
waters, the test must be conducted using receiving waters collected
outside the influence of the outfall. Controls should be conducted using
both receiving water and culture water.
tions because less dilution is needed to adjust to the proper salinity.
Because the marine/estuarine
species used for testing are salin-
ity sensitive, the effluent must be
adjusted to the proper salinity before
preparing the test concentrations.
Hypersaline brine is recommended
for adjusting the effluent salinity.
Appendix D provides instructions for
preparing the brine solution (EPA,
2002a). To prepare test concentra-
tions at the desired salinity, adjust
the diluent (deionized water) with the
hypersaline brine before adding it to
the effluent. Using hypersaline brine
instead of seawater allows the test to
be run at higher effluent concentra-
Use a minimum of five exposure concentrations and a control with a minimum of eight replicates per
concentration. The Saltwater Chronic Methods Manual recommends the use of a 0.5 dilution factor, which
provides precision of + 100%. Test precision shows little improvement as the dilution factor is increased
beyond 0.5, and declines rapidly if a smaller dilution factor is used. Approximately 3 L of test solution are
needed each day for a test conducted with 8 replicates of 5 concentrations and a control.
ROUTINE CHEMISTRIES
Once the various concentrations are prepared, set aside one aliquot of each for conducting
routine chemistries. By setting these aside, the chemistries can be performed without con-
taminating the actual test solutions with the probe. For test initiation and renewals, measure
and record the dissolved oxygen (DO) at the beginning and end of each 24-hour renewal in
at least one test chamber of each test concentration and in the control. If the DO falls below
4.0 mg/L in any replicate, aerate all concentrations and the control. Take care not to cause
excess turbulence that can cause physical stress to the organisms.
Dissolved oxygen, temperature, pH, and salinity must be measured on each new sample. Dissolved oxygen
is measured at the beginning and end of each 24-hour renewal in at least one test chamber of each test
concentration and in the control. Measuring salinity at the beginning and end of each 24-hour renewal is pre-
ferred but not required. The salinity, temperature, and pH of the effluent sample must be measured at the end
of each 24-hour exposure period in one test chamber at each concentration and in the control. See Table 1.
Table 1. Monitoring Schedule
Parameter
Dissolved Oxygen1 2
Temperature13
pH'3
Salinity1'2
Monitoring Frequency
Each New Sample
X
X
X
X
24-hr Exposure Period
Beginning
X
X
End
X
X
X
X
/ Measured in each new sample (100% effluent or receiving water) and in control.
2 Beginning and end measurement on one replicate in each concentration and the control.
3 End measurement on one replicate in each concentration and the control.
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Mysid (Americamysis bahia) Survival, Growth, and Fecundity Toxicity Tests
Supplement to Training Video
These parameters should fall within the recommended ranges for conducting the test and they should be
recorded on the test data sheet. The recommended test conditions are presented in Appendix A and a
sample water quality data sheet is provided in Figure 3.
Figure 3. Data Form for the Mysid Survival and Fecundity Toxicity Test - Water Quality Data
Test:
Start Date:.
Salinity:.
Day 1
Day 2
Day 3
Day 4
Day 5
Day 6
Day 7
Day 1
Day 2
Day 3
Day 4
Day 5
Day 6
Day?
TRTMT
REP
REP
REP
REP
REP
REP
REP
REP
REP
REP
REP
REP
REP
REP
TRTMT
REP
REP
REP
REP
REP
REP
REP
REP
REP
REP
REP
REP
REP
REP
TEMP
TEMP
Salinity
Salinity
DO
DO
pH
pH
TRTMT
TRTMT
TEMP
TEMP
Salinity
Salinity
DO
DO
pH
pH
Source: EPA, 2002a.
TEST CHAMBERS
The test chambers should be readied before the effluent concentrations are prepared. EPA recommends
using 8 oz disposable plastic drinking cups or 400 ml glass beakers to conduct this test. The test cham-
bers are presoaked in clean seawater and labeled with colored tape. Each concentration is indicated by a
different color tape with the replicate number (1 - 8) written on it. The use of different colored tape makes
renewals easier because all of the replicates of one concentration can be identified quickly.
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U.S. ENVIRONMENTAL PROTECTION AGENCY Mysid (Americamysis boh/a) Survival, Growth, and Fecundity Toxicity Tests
Supplement to Training Video
• •• > •
Once the cups are prepared and the effluent solutions have been adjusted to within the proper parameter
ranges, each test solution is distributed to eight replicate cups. Each replicate should contain approximate-
ly 150 ml. The cups are placed in holding trays that are randomly placed in a temperature-controlled water
bath. The holding trays should be labeled with the same colored tape and replicate numbers as the cups
which allows for easier collection and replacement of the randomized cups during renewals. The cups will
stay in the same randomized positions for the duration of the test. Specific directions for test randomiza-
tion are provided in Appendix A of the Saltwater Chronic Methods Manual (EPA, 2002a).
TEST ORGANISMS
Juvenile mysids should be collected from gravid females obtained from at least three separate culturing
tanks. To begin a test with five effluent concentrations and a control, each with eight replicates, a mini-
mum of 240 juveniles are needed. Having more than 240 juveniles allows for extra juveniles from which to
choose. Select juveniles at random, but avoid using any that appear injured.
Juvenile mysids are assigned to the test chambers at a density of five mysids per chamber. The juveniles
are randomly selected from the 7-day old juvenile pool and pipetted using a large bore (4 mm inner diam-
eter [ID]) pipet into small presoaked ampules, two to three at a time. The open covers of the ampules
serve as handles. This random selection and assignment is continued until all of the ampules contain five
mysids. As the mysids are placed in these ampules, a minimum amount of water should be transferred with
them so that the effluent concentrations are not diluted.
To transfer the mysids to the test chambers, the ampules should be dipped below the water level in each
cup and gently rinsed to deposit the mysids. Pouring the mysids from above the water surface may cause
injury. The test chambers should remain in the water bath while this transfer is made.
FEEDING
Once the test has been set-up, the mysids are fed. The initial feeding rate is 0.5 ml of a food solution made
from 4.0 mL concentrated Artemia nauplii in 80 ml of uncontaminated, filtered seawater. This concentra-
tion of nauplii should yield a level of approximately 150 24-hr old nauplii per mysid per day. This amount
of food solution should provide the test organisms with a sufficient number of live Artemia for the next 24
hours until test renewal. Immediately after renewal each day, feed the mysids 0.25 ml of food solution.
Another 0.25 ml should be fed 8-12 hours later. The food should be dispensed using an automatic pipet
and the food solution should be swirled before pipetting to ensure an even distribution of the Artemia. After
feeding the mysids, cover the test chambers to prevent evaporation or contamination.
RENEWALS
To conduct the daily renewals, collect the test cups from the water bath starting with the control and
working toward the higher concentrations. Measure and record the temperature, salinity, DO, and pH in a
composite aliquot of a minimum of two randomly selected replicates from each concentration (see Figure
3). If the DO concentration falls below 4 mg/L in any one of the exposure chambers, all chambers must be
gently aerated at a rate of approximately 100 bubbles/minute. During renewals the mysids in each cham-
ber should be counted and the survival recorded on the test data sheets. Any dead animals should be
discarded. A sample survival and fecundity data sheet is presented as Figure 4.
To renew the effluent, pour or siphon off the old effluent solution into a white tray or a large beaker placed
on a light table. Either of these receptacles will clearly show any mysids that are accidentally removed.
Slowly pouring the effluent from the cups works well because mysids tend to swim against the current and will
swim towards the back of the cups. If a mysid is poured out with the old effluent it should be pipetted back into
the exposure chamber and recorded as "returned during renewal" on the test data sheet. When removing
the old effluent, a pipet should be used to clean any uneaten Artemia from the bottom of the chamber.
To add the new effluent solution to the chamber, gently pour approximately 150 ml of the appropriate solu-
tion down the side of the chamber avoiding as much turbulence as possible. This renewal procedure must
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Mysid (Americamysis bahia) Survival, Growth, and Fecundity Toxicity Tests
Supplement to Training Video
be repeated on days two through six of the exposure period. All data should be carefully recorded on the
data sheets each day.
Immediately after renewal each day, feed the mysids 0.25 ml of food solution. Another 0.25 mL should
be fed 8-12 hours later. If the survival rate in any replicate drops below 50%, the food provided to that
replicate should be reduced by half. Detailed instructions for culturing Artemia are provided in the video
"Culturing/4mericamys/s bahia" and in its supplemental guide (EPA, 2009b).
Figure 4. Data Form for the Mysid Survival and Fecundity Toxicity Test - Survival and Fecundity Data
Test:
Start Date:
Salinity:.
Treatment/
Replicate
\
2
3
Control 4
5
6
7
8
1
2
3
1 4
5
6
7
8
1
2
3
2 4
5
6
7
8
1
Day 1
Alive
Day 2
Alive
Day 3
Alive
Day 4
Alive
Day5
Alive
Day 6
Alive
Day 7
Alive
Females
vfleggs
Females
No eggs
Males
Imma-
ture!
Source: EPA, 2002a.
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Mysid (Americamysis bahia) Survival, Growth, and Fecundity Toxicity Tests
Supplement to Training Video
Terminating the Test
On the last day of the 7-day exposure, the replicates are checked for survival and fecundity and the animals
are prepared for growth measurements. The mysids are not fed on the last day of the test so that total
weights do not reflect the added weight of any undigested Artemia.
In preparation for the test termination, prepare small pieces (1 cm2) of clean, light-weight aluminum foil by
labeling them with sequential numbers. Gloves should be worn or forceps should be used to handle the
aluminum because oils from skin could affect weight differences. After they are numbered, these pieces
of foil should all be dried, tared, and their weights recorded on the growth-data sheet. The sample growth-
data sheet is presented as Figure 5.
Figure 5. Data Form for the Mysid Survival and Fecundity Toxicity Test - Dry Weight Measures
Test:
Start Date:
Salinity:.
Treatment/
Replicate
1
2
3
Control 4
5
6
7
8
1
2
3
1 4
5
6
7
8
1
2
3
2 4
5
6
7
8
L_
Pan#
Tare Wt.
Total Wt.
Organism Wt.
# of Organisms
Wt./Organism
Source: EPA, 2002a.
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After the aluminum is prepared,
pick up the test chambers
in the same manner as for
conducting a renewal. That
is, collect all of the replicates antennuie
of one concentration at
one time, starting with the
control. Final water quality
measurements, including DO,
temperature, salinity, and pH
should be measured on ali-
quots taken from several test
chambers in each concen-
tration and the control and
recorded (see Figure 3).
First, remove dead mysids
from the test chambers and
record the final survival count
for each replicate on the test
data sheet (see Figure 4).
The minimum requirement
for an acceptable test is 80%
survival in the controls.
Figure 6. Mature Female A. bahia with Eggs in Oviducts. Lateral
view (top) Dorsal view (bottom)
eyestalk
carapace
statocyst
telson
.telson
developing brood sac
oviducts with developing ova
uropod
Source: Lussier, Kuhn, and Sewall, 1987.
Second, determine the sexual
development and fecundity of each mysid in each replicate. The effluent should be poured off in the same
manner as during renewals. For each replicate remove the mysids and place each one in a separate well
of a multi-well slide. Any excess water transferred with the mysid can be removed from the well to make
viewing under a microscope easier.
Using a stereomicroscope at 240X, determine the sexual development of each mysid and record it on
the test data sheet (see Figure 4). This must be conducted while the mysids are alive because they turn
opaque upon dying. Figures 6 through 9 illustrate the sexual characteristics used to determine the matu-
rity and fecundity of the mysids.
Figure 6 is a mature female with eggs in the oviducts. This is most easily determined when viewed from
above and is determined by large, dark, oval-shaped bodies in the mid-section of the thorax.
Figure 7 shows a mature female with eggs in the brood pouch, and is characterized by the presence of
dark pigmented spots on the lateral sides of the body. These can be seen both from above and from
the side. Females that have no eggs or embryos have an empty brood pouch and empty oviducts. These
females can be identified by a single dark spot on each half of the brood pouch. These spots can be seen
from both above and from the side, although from the top is easiest. The video provides examples of
females with, and without, eggs and embryos.
Figure 8 presents a mature male mysid. Males are determined by the presence of testes that appear either as
clear circles, when viewing them from above, or as appendages at the junction of the thorax and abdomen when
viewing them from the side.
Figure 9 presents a diagram of an immature mysid. Immature mysids are those that do not have character-
istics that determine their classification as either mature males or females. Care must be taken, however,
not to mistake a barren female for an immature rrtysid. As the sex of each mysid is determined it should be
recorded on the survival and fecundity data sheet (see Figure 4).
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U.S. ENVIRONMENTAL PROTECTION AGENCY
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After the sex, maturity, and
fecundity of each mysid from
one replicate is determined,
all of the mysids from that
replicate should be placed
on a Nitex® screen that rests antennuie
on top of a beaker. Rinse the
mysids with deionized water
to remove any salts that may
interfere with the dry weights.
After the animals are rinsed
they are placed on the desig-
nated pre-tared piece of alu-
minum foil for that replicate.
Note that all of the mysids
from one replicate are placed
on the same piece of foil.
Once this process has been
repeated for all of the repli-
cates the mysids are dried in
an oven at 60°C for 24 hours
or 105°C for at least six
hours. The mysids must be
completely dried before they
are weighed but they should
not be overdried.
Figure 7. Mature Female A. bahia with Eggs in Oviducts and
Developing Embryos in Brood Sac. Lateral view (top) Dorsal view
(bottom)
eyestalk
carapace
itatocyst
telson
developing brood sac
oviducts with developing ova
.telson
uropod
The mysids should be
transported and stored in a
desiccator when weighing
them. This prevents mois-
ture from reabsorbing into
the mysids. The mysids are
weighed, one replicate at a
time, to the nearest milligram
(0.001 g.). Because small dif-
ferences in weight or appear-
ance can easily change the
test results, it is critical to
record observations and
measurements clearly and
accurately. See Figure 5
for a sample data sheet
for recording weights. The
minimum requirement for an
acceptable test is an average
weight of at least 0.20 mg/
mysid in the controls.
The analysis of this test com-
pares the maturity, fecundity,
growth, and survival of the
Source: Lussier, Kuhn, and Sewall, 1987.
Figure 8. Mature Male A. bahia. Lateral view (top) Dorsal View
(bottom)
eyestalk
carapace
antennuie
statocyst
telson
telson
uropod
Source: Lussier, Kuhn, and Sewall, 1987.
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exposed mysids to the con-
trol mysids. The Saltwater
Chronic Methods Manual
(EPA, 2002a) provides
instructions for statistical antennule
analysis of the survival,
growth, and fecundity data.
TEST ACCEPTABILITY
AND DATA REVIEW
Test data are reviewed to
verify that EPA's WET test
methods' acceptability
criteria (TAG) requirements
for a valid test have been
met. For instance, the TAG
requires 80% or greater
survival in controls with an
average weight of at least
0.20 mg/mysid and 50%
or more of the females in
the controls must have
eggs.
Figure 9. Immature A. bahla. Lateral view (top) Dorsal view
(bottom)
eyestalk
carapace
statocyst
telson
telson
uropod
Source: Lussier, Kuhn, and Sewall, 1987)
The concentration-response relationship generated for each multi-concentration test must be reviewed to
ensure that calculated test results are interpreted appropriately. In conjunction with this requirement, EPA
has provided recommended guidance for concentration-response relationship review (EPA, 2000a).
EPA's promulgated toxicity testing method manuals (EPA, 2002a, b) recommend the use of point estima-
tion technique approaches for calculating endpoints for effluent toxicity tests under the NPDES program.
The promulgated methods also require a data review of toxicity data and concentration-response data, and
require calculating the percent minimum significant difference (PMSD) when point estimation (e.g., LC50,
IC25) analyses are not used. EPA specifies the PMSD must be calculated when NPDES permits require sub-
lethal hypothesis testing. EPA also requires that variability criteria be applied as a test review step when
NPDES permits require sub-lethal hypothesis testing endpoints (i.e., no observed effect concentration
[NOEC] or lowest observed effect concentration [LOEC]) and the effluent has been determined to have no
toxicity at the permitted receiving water concentration (EPA, 2002b). This reduces the within-test variabil-
ity and increases statistical sensitivity when test endpoints are expressed using hypothesis testing rather
than the preferred point estimation techniques.
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Citations and Recommended References
American Society for Testing and Materials. 1987. Practice Guide for Conducting Life Cycle Toxicity Tests
with Saltwater Mysids. ASTM E-1191-87. ASTM, Philadelphia, PA.
American Public Health Association. 1985. Standard Methods for the Examination of Water and
Wastewater. 16th Ed. APHA, Washington, D.C.
Bahner, L.H., C.D. Craft, and D.R. Nimmo. 1975. A saltwater flow-through bioassay method with controlled
temperature and salinity. Progr. Fish-Cult. 37:126-129.
Borthwick, P.W. 1978. Methods for acute static toxicity tests with mysid shrimp (Mysidopsis bahia). In:
Bioassay Procedures for the Ocean Disposal Permit Program. U.S. Environmental Protection
Agency, Environmental Research Laboratory, Gulf Breeze, Florida.
Breteler, R.J., J.W. Williams, and R.L. Buhl. 1982. Measurement of chronic toxicity using the opossum
shrimp Mysidopsis bahia. Hydrobiol. 93:189-194.
Buikema, A.L., B.R. Neiderlehner, and J. Cairns. 1982. Biological monitoring. Part IV. Toxicity Testing.
Water Res. 16:239-262.
EPA. 1991. Technical Support Document for Water Quality-based Toxics Control. U.S. EPA Office of Water
Enforcement and Permits, Washington, D.C. EPA-505-2-90-001.
EPA. 2000a. Understanding and Accounting for Method Variability in Whole Effluent Toxicity Applications
Under the National Pollutant Discharge Elimination System Program. Office of Wastewater
Management, Washington, D.C. EPA 833-R-00-003.
EPA. 2000b. Method Guidance and Recommendations for Whole Effluent Toxicity (WET) Testing (40 CFR
Part 136). Office of Water, Washington, D.C. EPA 821-B-00-004.
EPA. 2002a. Short-term Methods for Estimating the Chronic Toxicity of Effluents and Receiving Waters to
Marine and Estuarine Organisms, Third Edition. (Saltwater Chronic Methods Manual). Office of
Water, Cincinnati, OH. EPA-821-R-02-014.
EPA. 2002b. Methods for Measuring the Acute Toxicity of Effluents and Receiving Waters to Freshwater
and Marine Organisms, Fifth Edition. (Acute Methods Manual). Office of Water, Cincinnati, OH.
EPA-821-R-02-012.
EPA. 2002c. Final Rule. 40 CFR Part 136. Guidelines Establishing Test Procedures for the Analysis of
Pollutants; Whole Effluent Toxicity Test Methods. 67 FR 69952-69972, November 19, 2002.
EPA. 2009a. Mysid (/Americamys/s bahia) Survival, Growth, and Fecundity Toxicity Tests. Supplement to
Training Video. Whole Effluent Toxicity Training Video Series, Saltwater Series. March 2009. EPA
833-C-09-001.
EPA. 2009b. Culturing Americamysis bahia. Supplement to Training Video. Whole Effluent Toxicity Training
Video Series, Saltwater Series. March 2009. EPA 833-C-09-001.
Heard, R.W. and W.W. Price. 2006. A Taxonomic Guide to the Mysids of the South Atlantic Bight. U.S.
Department of Commerce, National Oceanic and Atmospheric Administration.
Lussier, S.M., A. Kuhn and R. Comeleo. 1999. An evaluation of the 7-day toxicity test with Americamysis
bahia (formerly Mysidopsis bahia). Environ. Toxicol. and Chem. 18:2888-2893. [Errata: in the sec-
tion on Experimental Design, the test chamber should read "200-ml plastic cup" not "30-ml."]
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U.S. ENVIRONMENTAL PROTECTION AGENCY Mysid (Americamysis bahia) Survival, Growth, and Fecundity Toxicity Tests
Supplement to Training Video
Lussier, S.M., A. Kuhn, MJ. Chammas, and J. Sewall. 1988. Techniques for the laboratory culture of
Mysidopsis species (Crustacea: Mysidacea). Environ. Tox. Chem. 7:969-977.
Lussier, S.M., A. Kuhn, and J. Sewall. 1987. Guidance manual for conducting 7-day mysid survival/
growth/reproduction study using the estuarine mysid, Mysidopsis bahia. Contribution No. X106.
In: Schimmel, S.C., ed. Users guide to the conduct and interpretation of complex effluent toxic-
ity tests at estuarine/marihe sites. Environmental Research Laboratory, U.S. Environmental
Protection Agency, Narragansett, Rhode Island. Contribution No. 796, 265 pp.
Molenock, J. 1969. Mysidopsis bahia, a new species of mysid (Crustacea: Mysidacea) from Galveston Bay,
Texas. Tulane Stud. Zool. Bot. 15(3):113-116.
Nimmo, D.R. and T.L. Hamaker. 1982. Mysids in toxicity testing — a review. Hydrobiol. 93:171-178.
Nimmo, D.R., T.L. Hamaker, and C.A. Sommers. 1978. Entire life cycle toxicity test using mysids
(Mysidopsis bahia) in flowing water. In: Bioassay Procedures for the Ocean Disposal Permit
Program, U.S. Environmental Protection Agency, Environmental Research Laboratory, Gulf Breeze,
Florida. EPA-600/9-78-010. pp. 64-68.
Personne, G., E. Jaspers, and C. Glaus, eds. 1980. Ecotoxicological testing for the marine environment.
Vol. 1 State University of Ghent and Institute for Marine Scientific Research, Bredene, Belgium.
Price, W.W., R.W. Heard and L. Stuck. 1994. Observations on the genus Mysidopsis sars. 1864 with the
designation of a new genus, Americamys/s, and the descriptions of Americamysis alleni and A.
stucki (Peracarida: Mysidacea: Mysidae), from the Gulf of Mexico. Proc. Biol. Soc. Wash. 107:680-
698.
Schimmel, S.C., ed. 1987. Users guide to the conduct and interpretation of complex effluent toxicity tests
at estuarine/marine sites. Environmental Research laboratory, U.S. Environmental Protection
Agency, Narragansett, Rhode Island. Contribution No. 796, 265 pp.
Stuck, K.C., H.M. Perry and R.W. Heard 1979. An annotated key to the Mysidacea of the North Central Gulf
of Mexico. Gulf Res. Rept. 6(3):255 - 238.
Walters, D.B. and C.W. Jameson. 1984. Health and safety for toxicity testing. Butterworth Publ., Woburn,
Massachusetts.
EPA references are available online at www.epa.gov/npdes.
If you need additional copies of this document, you can download it at:
www.epa.gov/npdes/wqbasedpermitting.
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Glossary
Acute toxicity. An adverse effect measured on a group of test organisms during a short-term exposure in
a short period of time (96 hours or less in toxicity tests). The effect can be measured in lethality
or any variety of effects.
Artem/a. The marine invertebrate (referred to as brine shrimp) used as the recommended food source for
mysid cultures and test organisms; Brazilian or Colombian strains are preferred because the sup-
plies are found to have low concentrations of chemical residues and nauplii are of suitably small
size.
Chronic toxicity. An adverse effect that occurs over a long exposure period. The effect can be lethality,
impaired growth, reduced reproduction, etc.
Cyst. The life stage of unhatched Artemia.
Diluent water. Dilution water used to prepare the effluent concentrations.
Effluent concentrations. Concentrations or dilutions of an effluent sample to which test organisms are
exposed to determine the biological effects of the sample on the test organism.
Effluent sample. A representative collection of the discharge that is to be tested.
Fecundity. Productivity or fertility as measured in this test as the percentage of females.with eggs in the
oviduct and/or brood pouch.
Flow-through water delivery system. An open water flow system that delivers fresh water or seawater to
culture tanks and is disposed of after it leaves those tanks.
Hypothesis testing. Technique (e.g., Dunnett's test) that determines what concentration is statistically
different from the control. Endpoints determined from hypothesis testing are NOEC and LOEC.
IC25 (Inhibition Concentration, 25%). The point estimate of the toxicant concentration that would cause a
25% reduction in a non-quantal biological measurement (e.g., reproduction or growth) calculated
from a continuous model.
LC50 (Lethal Concentration, 50%). The concentration of toxicant or effluent that would cause death to
50% of the test organisms at a specific time of observations (e.g., 96-hour LC50).
Lowest Observed Effect Concentration (LOEC). The LOEC is the lowest concentration of toxicant to
which organisms are exposed in a test, which causes statistically significant adverse effects on
the test organisms (i.e., where the values for the observed endpoints are statistically significantly
different from the control). The definitions of NOEC and LOEC assume a strict dose-response
relationship between toxicant concentration and organism response.
Minimum Significant Difference (MSD). The MSD is the magnitude of difference from the control where
the null hypothesis is rejected in a statistical test comparing a treatment with a control. MSD
is based on the number of replicates, control performance and power of the test. MSD is often
measured as a percent and referred to as PMSD.
Mysid (Amer/camys/s bahia). An estuarine crustacean, formerly known as Mysidopsis bahia, ranging 4.4
mm to 9.4 mm in length found from the Gulf of Mexico and along the Atlantic coast as far north
as Rhode Island, used in test procedures as an indicator species for marine or estuarine aquatic
toxicity.
Glossary-1
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Nauplii. Free-swimming microscopic larvae stage characteristic of copepods, ostracods, barnacles, etc.
typically only with three pairs of appendages.
No Observed Effect Concentration (NOEC). The NOEC is the highest tested concentration of toxicant to
which organisms are exposed in a full life-cycle or partial life-cycle (short-term) test, that causes
no observable adverse effect on the test organism (i.e., the highest concentration of toxicant
at which the values for the observed responses are not statistically significantly different from
the controls). NOECs calculated by hypothesis testing are dependent upon the concentrations
selected.
NPDES (National Pollutant Discharge Elimination System) Program. The national program for issuing,
modifying, revoking and reissuing, terminating, monitoring and enforcing permits, and imposing
and enforcing pretreatment requirements, under Sections 307, 318, 402, and 405 of the Clean
Water Act.
Point Estimation Techniques. This technique is used to determine the effluent concentration at which
adverse effects (e.g., fertilization, growth or survival) occurred, such as Probit, Interpolation
Method, Spearman-Karber. For example, a concentration at which a 25% reduction in
reproduction and survival occurred.
Receiving Water Concentration (RWC). The RWC is the concentration of a toxicant or the parameter
toxicity in the receiving water (i.e., riverine, lake, reservoir, estuary or ocean) after mixing.
Recirculating water delivery system. A water flow system that treats water after it passes through the
culture tanks (usually with sand and biofilters) and delivers the same treated water back to the
tanks.
Static renewal. The exposure medium is replaced each day by moving the test animal to a new test cup
prepared with the proper effluent concentration.
Static water system. An enclosed system contained within one culture tank. The water is filtered through
an underground or charcoal filter and is delivered back to the same tank.
Toxicity test. A test to measure the toxicity of a chemical or effluent using living organisms. The test
measures the degree of response of an exposed organism to a specific chemical or effluent.
WET (Whole effluent toxicity). The total toxic effect of an effluent measured directly with a toxicity test.
Glossary-2
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Appendix A
Summary of Test Conditions and Test Acceptability
Criteria
Table A-l. Summary of Test Conditions and Test Acceptability Criteria for Americamys/s bahia 7-day
Survival, Growth, and Fecundity Toxicity Test
Test type
Salinity
Temperature (C°)
Photoperiod
Light intensity (quality)
Test chamber size
Test solution volume
Renewal of test solutions
Age of test organisms
Number of concentrations per study
Number of organisms per test chamber
Number of replicate chambers per concen-
tration
Source of food
Feeding regime
Aeration
Dilution water
Effects measured
Cleaning
Sample volume needed
Test concentrations
Dilution factor
Test duration
Endpoints
Test acceptability criteria
Sampling requirements
Static renewal (required)
20%o - 30%o ± 2%0 (recommended)
26 ± 1 °C (recommended) '
16 hours light; 8 hours dark, with phase on/off period (recommended)
10-20 uE/mVs (50 - 100 ft-c) (ambient lab levels) (recommended)
8 oz plastic disposable cups, or 400 mL glass beakers (recommended)
ISO mL per replicate cup (recommended minimum)
Daily (required)
7 days at start of test (required)
Minimum of 5 concentrations and a control (required minimum)
5 (40 per concentration) (required minimum)
8 (required minimum)
Newly hatched Anemia nauplii (<24-hr old; required)
Feed ISO 24-hr old nauplii per mysid daily, half after test solution
renewal and half after 8 - 12 hr (recommended)
None unless DO falls below 4.0 mg/L, then gently aerate all cups
(recommended)
Natural seawater, or hypersaline brine diluted with deionized water,
or artificial seasalts (available options)
Survival and growth (required); egg development (recommended)
Pipet excess food from cups daily immediately before test solution
renewal and feeding (recommended)
3 L per day (recommended)
Effluents: 5 and a control (required)
Receiving waters: 100% receiving water (or minimum of 5) and a con-
trol (recommended)
Effluents: S 0.5 series (required)
Receiving waters: None, or ^ 0.5 (recommended)
7 days (required)
Survival and growth (required); and egg development (recommended)
80% or greater survival, average dry weight 0.20 mg or greater in
controls (required); fecundity may be used if 50% or more of females in
controls produce eggs (required if fecundity endpoint used)
For on-site tests, samples collected daily and used within 24 hr of the
time they are removed from the sampling device. For off-site tests, a
minimum of three samples (e.g., collected on days one, three, and five)
with a maximum holding time of 36 hr before first use (see Saltwater
Chronic Methods Manual, Section 8, Effluent and Receiving Water
Sampling, Sample Handling and Sample Preparation for Toxicity Test,
Subsection 8.5.4) (required)
Source: Adapted from EPA, 2002a.
'Lussier at al, 1999 found that test conducted at 26°C - 27°C exhibited higher probability of meeting test acceptability criteria for fecundity
than tests conducted at 26 ±\°C.
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Appendix B
Apparatus and Equipment List
Air line, and air stones. For aerating cultures, brood chambers, and holding tanks, and supplying air to
test solutions with low DO.
Air pump. For oil-free air supply.
Balance. Analytical, capable of accurately weighing to 0.00001 g.
Beakers or flasks. Six, borosilicate glass or non-toxic plasticware, 2 - 3 L for making test solutions.
Brine shrimp (Artem/a) culture unit. See section on "Maintaining and Feeding Cultures."
Depression glass slides or depression spot plates. Two for observing organisms.
Desiccator. For holding dried organisms.
Dissecting microscope (240 - 400X magnification). For examining organisms in the test vessels to
determine their sex and to check for the presence of eggs in the oviducts of the females.
Droppers, and glass tubing with fire polished edges. 4 mm inner diameter for transferring organisms.
Drying oven. 50 - 105°C, for drying organisms.
Environmental chamber or equivalent facility with temperature control (26 ± 1°C).
Facilities for holding and acclimating test organisms.
Forceps (fine tips such as jewelers forceps). For transferring organisms to weighing boats.
Light box. For illuminating organisms during examination.
Meters: pH and DO, and specific conductivity. For routine physical and chemical measurements.
Mysid (Americamysis bahia) culture unit. See section on "Maintaining and Feeding Cultures". The test
requires a minimum of 240 7-day old (juvenile) mysids.
NITEX® or stainless steel mesh sieves. 150 urn and 100 urn for concentrating organisms; 1 mm mesh
and 300 urn mesh for collection of juveniles.
Pipet bulbs and fillers. Propipet®, or equivalent.
Reference weights, Class S. For checking performance of balance.
Refractometer or other method. For determining salinity.
Samplers. Automatic sampler, preferably with sample cooling capability, that can collect a 24-hour
composite sample of 5 L.
Separatory funnels, 2-liters. Two to four funnels for culturing Artem/a.
Standard or micro-Wlnkler apparatus. For determining DO and checking DO meters.
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Test vessels. 200 ml borosilicate glass beakers or 8 oz disposable plastic cups or other similar
containers. Cups must be rinsed thoroughly in distilled or deionized water and then pre-soaked
(conditioned) overnight in dilution water before use. Forty-eight (48) test vessels are required for
each test (eight replicates at each of five effluent concentrations and a control). To avoid potential
contamination from the air and excessive evaporation of test solutions during the test, the
chambers should be covered with safety glass plates or sheet plastic (6 mm thick).
Thermometers, bulb-thermograph or electronic-chart type. For continuously recording temperature.
Thermometers, glass or electronic, laboratory grade. For measuring water temperatures.
Thermometers. National Bureau of Standards Certified (see EPA 2002a). Used to calibrate laboratory
thermometers.
Trays. For test vessels: one large enough to transport eight vessels at one time; one to hold 56 test
vessels (approximately 90 x 48 cm).
Volumetric flasks and graduate cylinders. Class A. Borosilicate glass or non-toxic plastic labware,
50 - 2000 ml for making test solutions.
Wash bottles. For deionized water, for washing organisms from containers and for rinsing small glassware
and instrument electrodes and probes.
Water purification system. Millipore® Milli-Q® deionized water or equivalent.
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Appendix C: Reagents and Consumable Materials
Data sheets. One set per test for recording data
Effluent, receiving water, and dilution water. Dilution water containing organisms that might prey upon
or otherwise interfere with the test organisms should be filtered through a fine mesh (with 150 pm
or smaller openings).
Saline test and dilution water. The salinity of the test water must be in the range of 20%o - 30%o.
The salinity should vary by no more than ±2%0 among the chambers on a given day. If effluent and
receiving water tests are conducted concurrently, the salinities of these tests should be similar.
It is important to maintain a constant salinity across all treatments during a test. It is desirable
to match the test salinity with that of the receiving water. Two methods are available to adjust
salinities - a hypersaline brine (HSB) derived from natural seawater or artificial sea salts. Both are
described in EPA, 2002a.
Food source. Feed the mysids Artemia nauplii that are less than 24-hour-old.
Laboratory quality assurance samples and standards
Markers, waterproof. For marking containers, etc.
Membranes and filling solutions for DO probe. Or reagents, for modified Winkler analysis
(See EPA, 2002a).
pH buffers 4, 7, and 10 - (Or as per instructions of instrument manufacturer) for standards and
calibration check (see EPA 2002a).
Reagent water Distilled or deionized water that does not contain substances which are toxic to the test
organisms.
Reference toxicant solutions. Reference toxicants such as sodium chloride (NaCI), potassium chloride
(KCI), cadmium chloride (CdCI2), copper sulfate (CuS04), sodium dodecyl sulfate (SDS), and
potassium dichromate (K2Cr207), are suitable for use in the NPDES Program and other Agency
programs requiring aquatic toxicity tests.
Sample containers. For sample shipment and storage.
Tape, colored. For labeling test containers.
Test organisms. The test is begun with 7-day-old juvenile Amer/camys/s bahia (mysids).
Weighing pans, aluminum. To determine the dry weight of the organisms
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Appendix D:
Preparing Hypersaline Brine (HSB)
Salinity adjustments are a vital part of using marine and estuarine species for toxicity testing. Because the
majority of industrial and sewage treatment effluents entering marine and estuarine waters contain little or
no measurable salts, the salinity of these effluents must be adjusted before exposing estuarine or marine
plants and animals to the test solutions. It also is important to maintain constant salinity across all treat-
ments throughout the test for quality control. Finally, matching the test solution's salinity to the expected
receiving water's salinity may require salinity adjustments. NHEERL-AED uses HSB, prepared from filtered
natural seawater, to adjust exposure solution salinities.
, HSB 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. HSB can be held for prolonged periods without any apparent degradation, added directly to
the effluent to increase the salinity, or used as control water by diluting to the desired salinity with deion-
ized 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.
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, fol-
lowed by several thorough deionized-water rinses.
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 urn before placing into the brine tank. 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 could contaminate the brine. A
thermostatically controlled heat exchanger made from fiberglass is suggested.
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 (or more or
less often, depending on the volume being generated) to ensure that the salinity does not exceed 100%o
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 NHEERL-AED was a reduction
in the alkalinity of seawater made from brine with salinity greater than 100%0, 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 1-mm filter and pump
or pour it directly into portable containers (20-L cubitainers or polycarbonate water cooler jugs are most
suitable). Cap the containers, and record the measured salinity and the date generated. Store the brine in
the dark at room temperature.
D-l
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.S. ENVIRONMENTAL PROTECTION AGENCY
Mysid (Americamysis bahia) Survival, Growth, and Fecundity Toxicity Tests
Supplement to Training Video
SALINITY ADJUSTMENTS USING HYPERSALINE BRINE
To calculate the volume of brine (Vb) to add to a 0%o sample to produce a solution at a desired salinity (Sf),
use this equation:
vb * sb = s, * vf
Where:
v«,=
v,=
volume of brine, ml
salinity of brine, %o
final salinity, %0
final volume needed, ml
Table D-l gives volumes needed to make 20%o test solutions from effluent (0%o), deionized water, and
100%o MSB. The highest effluent exposure concentrations achievable are 80% effluent at 20%o salinity
and 70% effluent at 30%o salinity. Test solutions presented in Table D-l are not meant as recommenda-
tions, rather as examples.
Table D-l. Preparation of Test Solutions at a Salinity of 20%o Using HSB for a Final Test Concentration
Volume of 2000 ml.
Exposure Concentration
(% effluent)
80
40
20
10
5
Control
Effluent
(assumes 0%o salinity)
(mL)
1,600
800
400
200
100
—
De/onized
Water (mL)
0
800
1,200
1,400
1,500
2,000
HSB
(f 00%. salinity) (mL)
400
400
400
400
400
400
D-2
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If you need additional copies of this document, you can download it at:
www.epa.gov/npdes/wqbasedpermitting
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WHOLE EFFLUENT ToxiCITY • TRAINING VIDEO SERIES • saltwater series
Sperm Cell Toxicity
Tests Using the Sea Urchin
(Arbacia punctulata)
Supplement to Training Video
U.S. Environmental Protection Agency
Office of Wastewater Management
Water Permits Division
1200 Pennsylvania Ave., NW
Washington, DC 20460
EPA 833-C-09-001
March 2009
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NOTICE
The revision of this guide has been funded wholly or in part by the
Environmental Protection Agency under Contract EP-C-05-063. Mention of
trade names or commercial products does not constitute endorsement or
recommendation for use.
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^,| U.S. ENVIRONMENTAL PROTECTION AGENCY Sperm Cell Toxicity Tests Using the Sea Urchin (Arbacia punctulata)
Supplement to Training Video
Foreword
This guide serves as a supplement to the video "Sperm Cell Toxicity Tests Using the Sea Urchin, Arbacia
punctulata" (EPA, 2009). The methods illustrated in the video and described in this supplemental guide
support the methods published in the U.S. Environmental Protection Agency's (EPA's) Short-term Methods
for Estimating the Chronic Toxicity of Effluents and Receiving Waters to Marine and Estuarine Organisms,
Third Edition (EPA, 2002a), referred to as the Saltwater Chronic Methods Manual. The video and this guide
provide details on preparing for and conducting the test based on the expertise of personnel at the follow-
ing EPA Office of Research and Development (ORD) laboratories:
National Health and Environmental Effects Research Laboratory (NHEERL) - Atlantic Ecology Division
in Narragansett, Rhode Island
NHEERL - Gulf Ecology Division in Gulf Breeze, Florida
National Exposure Research Lab (NERL) - Ecological Exposure Research Division (EERD) in
Cincinnati, Ohio
This guide and its accompanying video are part of a series of training videos produced by EPA's Office of
Wastewater Management. This Saltwater Series includes the following videos and guides:
"Mysid (Americamysis bahia) Survival, Growth, and Fecundity Toxicity Tests"
"Culturing Americamysis bahia"
"Sperm Cell Toxicity Tests Using the Sea Urchin, Arbacia punctulata"
"Red Algal (Champia parvula) Sexual Reproduction Toxicity Tests"
"Sheepshead Minnow (Cyprinodon variegatus) and Inland Silverside (Menidia beryllina) Larval Survival
and Growth Toxicity Tests"
The Freshwater Series, released in 2006, includes the following videos and guides:
"Ceriodaphnia Survival and Reproduction Toxicity Tests"
"Culturing of Fathead Minnows (Pimephales promelas)"
"Fathead Minnow (Pimephales promelas) Larval Survival and Growth Toxicity Tests"
All of these videos are available through the National Service Center for Environmental Publications
(NSCEP) at 800 490-9198 or nscep@bps-lmit.com.
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Sperm Cell Toxicity Tests Using the Sea Urchin (Arbacia punctulota)
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U.S. ENVIRONMENTAL PROTECTION AGENCY Sperm Cell Toxicity Tests Using the Sea Urchin (Arbacia punctulata)
Supplement to Training Video
Contents
Foreword i
Introduction 1
Background : 1
Water and Light 1
Obtaining and Maintaining Sea Urchins 1
Culture Water 1
Photoperiod 1
Culture Vessels 2
Water Delivery Systems 2
Food Preparation 2
Test Method 2
Obtaining Gametes 2
Making Stock Solutions of Sperm and Eggs 3
Effluent Preparation 5
Beginning the Test 5
Routine Chemistries 6
Terminating the Test 6
Test Acceptability and Data Review 6
Citations and Recommended References 7
Glossary Glossary-1
Appendix A: Preparing Hypersaline Brine (MSB) A-l
Appendix B: Apparatus and Equipment B-l
Appendix C: Reagents and Consumable Materials C-l
Appendix D: Summary of Test Conditions and Test Acceptability Criteria D-l
Appendix E: Data Sheets E-l
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U.S. ENVIRONMENTAL PROTECTION AGENCY Sperm Cell Toxicity Tests Using the Sea Urchin (Arbacia punctulata)'
Supplement to Training Video
FIGURES
Figure 1. Schematic of the aboral surface of Arbacia punctulata, with spines partly removed to
show structure, especially the genital pores 2
Figure E-l. Sperm Cell Toxicity Test, Sample Data Sheet #1 E-l
Figure E-2. Sperm Cell Toxicity Test, Sample Data Sheet #2 - Raw Data E-2
TABLES
Table 1. Fifty Percent Serial Dilution Method for Counting Sperm Cell Density 3
Table A-l. Preparation of Test Solutions at a Salinity of 30%o Using MSB for a Final Test
Concentration Volume of 1000 ml A-2
Table A-2. Preparation of Test Solutions at a Salinity of 30%o Using Natural Seawater, Hypersaline
Brine, or Artificial Sea Salts A-2
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U S ENVIRONMENTAL PROTECTION AGENCY Sperm Cell Toxicity Tests Using the Sea Urchin (Arbacia punctulata)
Supplement to Training Video
Introduction
This supplemental guide accompanies the Environmental Protection Agency's (EPA's) video training for
conducting sea urchin (Arbacia punctulata) fertilization toxicity tests (EPA,. 2009). The test method is found
in Section 15 of EPA's Short-term Methods for Estimating the Chronic Toxicity of Effluents and Receiving
Waters to Marine and Estuarine Organisms, Third Edition (EPA, 2002a). The test was developed at EPA's
Office of Research and Development's (ORD's) National Health and Environmental Effects Research
Laboratory-Atlantic Ecology Division (NHEERL-AED) in Narragansett, Rhode Island, and is based on the
freshwater tests developed at the EPA Mid-Continent Ecology Division (MED) in Duluth, Minnesota. The
material presented in both the video and this guide summarizes the methods but does not replace a thor-
ough review and understanding of the methods by laboratory personnel before conducting the test.
Background
Under the National Pollutant Discharge Elimination System (NPDES) program (Section 402 of the Clean
Water Act), EPA uses toxicity tests to monitor and evaluate effluents for their toxicity to biota and their
impact on receiving waters. By determining acceptable or safe concentrations for toxicants discharged
into receiving waters, EPA can establish NPDES permit limitations for toxicity. These WET (Whole effluent
toxicity) permit limitations regulate pollutant discharges on a whole effluent effect basis rather than by a
chemical-specific approach only.
Whole effluent toxicity methods measure the synergistic, antagonistic, and additive effects of all the chemi-
cal, physical, and additive components of an effluent that adversely affect the physiological and biochemi-
cal functions of the test organisms. Therefore, healthy organisms and correct laboratory procedures are
essential for valid test results. Laboratory personnel should be very familiar with the test methods and with
sea urchin handling techniques before conducting a test.
This supplemental guide covers the procedures for conducting the test according to EPA's promulgated
methods (40 CFR Part 136; EPA, 2002c) and also provides some helpful information that is not presented
in the Saltwater Chronic Methods Manual (EPA, 2002a).
This test method examines the effect of effluent or receiving waters on the reproduction of sea urchin
gametes after exposure in a static system for 1 hour and 20 minutes. Sperm cells are exposed to a series
of effluent concentrations for 1 hour. The eggs are then introduced to the test chambers which contain
the sperm cells. After 20 minutes, the test is ended and the effects on exposed gametes are compared to
controls to determine if the effluent concentrations had any effect on fertilization.
This guide and the accompanying video describe how the test is set up, initiated, terminated, and reviewed,
including suggestions on maintaining healthy cultures of test animals.
Water and Light
OBTAINING AND MAINTAINING SEA URCHINS
Before conducting tests, healthy sea urchin cultures should be established. Adult sea urchins can be
ordered from commercial biological supply houses, or collected along the Atlantic coast. Keep male and
female animals in separate tanks. To determine the sex of each animal, briefly stimulate each with a
12-volt transformer. This causes the immediate release of masses of gametes from genital pores on the
top of the animal. The eggs are red and the sperm are white. Separate the animals into 20 L aerated fiber-
glass tanks; each can hold about 20 adults.
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Sperm Cell Toxicity Tests Using the Sea Urchin (Arbacia punctulata)
Supplement to Training Video
CULTURE WATER
The quality of water used for
maintaining sea urchins is very
important. Culture water and
all water used for washing and
dilution steps and for control
water in the tests should be
maintained at a salinity of
30%o ± 2%o using natural
seawater, hypersaline brine
(MSB), or artificial sea salts.
Instructions for making dilution
water and MSB are provided in
Appendix A of this document
and Section 7 of the Saltwater
Chronic Methods Manual (EPA,
2002a).
PHOTOPERIOD
Figure 1. Schematic of the aboral surface of Arbacia punctulata,
with spines partly removed to show structure, especially the genital
pores
anus
suranal
plate
ocular
plate
genital
plate
genital
pore
madreporite
madreporic
plate
The sea urchin conditions
should include a photoperiod
of 16 hours light and 8 hours
darkness. The light quality and
intensity should be at ambient laboratory levels, which is approximately 10 - 20 E/um2/s or 50 to 100 foot
candles (ft-c) (EPA, 2002a).
CULTURE VESSELS
Adult sea urchins are kept in natural or artificial seawater in a flow-through or recirculating aerated 40-L
glass aquarium.
Allow filtered seawater to flow into the tanks at a rate of 5 L per minute and maintain the temperature at
15°C±3°C.
WATER DELIVERY SYSTEMS
Equip the adult sea urchin aquarium with an under-gravel or outside biological filter, or cartridge filter. A
stock of at least 12 males and 12 females are needed for routine testing. If the animals will be used for an
on-site test, transport them separated by sex in separate or partitioned coolers packed with wet kelp and
paper towels. Once on site, the sea urchins should be transferred into separate 10-gallon aquarium tanks
with gravel-bed filtration. Even with filtration, the water should be changed periodically to maintain good
water quality.
Collect eggs first to avoid any
possible pre-fertilization.
FOOD PREPARATION
Sea urchins are fed kelp of the species Laminaria obtained from uncori-
taminated coastal waters or ordered from commercial supply houses, or
romaine lettuce. Supply the urchins with ample food, renewing the kelp
each week and removing decaying kelp as necessary. Healthy sea urchins will attach to kelp or aquarium
walls within hours — any unhealthy animals should be removed and should not be used for testing. Every 1
to 2 weeks, empty and clean the tanks.
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Sperm Cell Toxicity Tests Using the Sea Urchin (Arfaocia punctulata)
Supplement to Training Video
At AED, staff use the data
sheet included in Appendix E for
calculating and recording dilutions
Test Method
OBTAINING GAMETES
To prepare for the test, all vials, pipets, and pipet tips should be soaked in clean, 30%o seawater overnight.
Collect eggs and sperm from healthy animals by transferring the animals into a shallow bowl filled with
enough control seawater to just cover their shells. Eggs are obtained from female sea urchins using electri-
cal stimulation by touching the shells close to the genital pores with
electrodes from a 10 - 12-volt transformer for about 30 seconds.
The red eggs pool on the sea urchin shell above the genital pores.
These are collected from the shell using a 10 ml disposable syringe
with an 18-gauge, blunt-tipped needle with the tip cut off so that it
will rest on the shell without puncturing it. After collection, the needle
is removed and the eggs emptied into conical centrifuge tubes. Pool the eggs and keep them at room tem-
perature until use, but not longer than a few hours. Four females should yield enough eggs to test five test
dilutions plus one control, with four replicates.
Obtain sperm from four male sea urchins. Again, place the animals in a shallow bowl with their shells barely
covered with control seawater. Like the females, the males are induced to spawn by placing electrodes
from a 10 - 12-volt transformer against their shells for 30 seconds. The sperm appear white. Collect the
concentrated sperm that pools on top of the shell using a syringe fitted with an 18-gauge, blunt-tipped
needle. Pool the sperm, keep the sample on ice, and record the collection time. The sperm must be used in
a toxicity test within 1 hour of collection.
MAKING STOCK SOLUTIONS OF SPERM AND EGGS
To ensure reproducibility in the test results, the sperm and eggs must be concentrated to known dilutions
using the 30%o seawater. During the exposure period, 2,500 sperm should be present for every one egg.
Figure E-l, presented in Appendix E, provides a sample data sheet used to calculate the sperm and egg
deliveries.
After collection, the sperm should be in a volume of about 0.5 to 1 mL of control water in the collecting
syringe. This is called the "sperm stock" solution. Perform a 50 percent serial dilution for counting the
sperm cell density using the following dilution method (see Table 1).
Add sperm from Vial E to both sides of a Neubauer hemacytometer. Let the sperm settle 15 minutes. Count
the number of sperm in the central 400 squares on both sides of the hemacytometer under a compound
microscope (100X).
The average of the two sperm cell counts (sperm/mL or SPM) from Vial E = # x 104.
Calculate the SPM in all the other suspensions based on this count:
Vial A = 40 x SPM of Vial E
Vial B = 20 x SPM of Vial E
Vial D = 5xSPMofVial E
SPM of original sample = 2000 x SPM of Vial E
The egg solution can be prepared
during the first hour of the test after
the sperm exposure has started.
To prepare the sperm suspension for the test, select the vial containing an SPM greater than 5 x 107 SPM.
To determine the dilution needed for the test:
The calculated SPM
(5 x 107)
[(DF) x 10] -10
= Dilution Factor (DF)
= mL of seawater to add to selected vial
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Sperm Cell Toxicity Tests Using the Sea Urchin (Arbacia punctulata)
Supplement to Training Video
Table 1. Fifty Percent Serial Dilution Method for Counting Sperm Cell Density.
'•
2.
3.
4.
5.
6.
7.
Add 400 uL of sperm stock to 20 mL of seawater to create Vial A.
using a 5-mL pipettor, or by inversion.
Add 10 ml from Vial A to 10 ml of seawater to create Vial B. Mix
5-mL pipettor, or by inversion.
Add 10 mL from Vial B to 10 mL of seawater to create Vial C. Mix
5-mL pipettor, or by inversion.
Add 10 mL from Vial C to 10 mL of seawater to create Vial D. Mix
a 5-mL pipettor, or by inversion.
Mix by gently pipetting
by gently pipetting using a
by gently pipetting using a
by gently pipetting using
Discard 10 mL from Vial D so that all vials now contain 10 mL.
Vial C is used to create a final dilution that is killed and counted. Add 10 mL 10% acetic acid
in seawater to Vial C; cap the vial and mix by inversion.
Add 1 mL of the killed sperm in Vial C to 4 mL of seawater in Vial
using a 4-mL pipettor.
E. Mix by gently pipetting
1. 400 Ml
sperm stock
2. 10ml
3. 10 ml
4. 10 ml
5. Discard 10 ml
Vial A
20 mL stock
seawater
Vial B 7 1 m/ Vial C
10 ml stock / lOmLstock
seawater / seawater
VialD
10 mL stock
seawater
6. 10 ml 10% acetic
acid in saltwater
Mix well
before each
transfer.
Vial C
The sperm cell count in the test stock should be confirmed. Add 0.1 mL of test stock to 9.9 mL of 10
percent acetic acid in seawater and count the sperm cells using a hemacytometer. This count
should average 50 ±5 cells. Only about 2.5 mL of sperm test stock solution is
needed for testing 5 test solutions and a control, with 4 or more replicates. Hold the
test stock on ice until the test begins, but no longer than 1 hour.
The eggs must be washed before preparing the standard egg dilution needed for
the test (2,000 eggs/mL). To wash the eggs, first remove the supernatant water
from the settled eggs. Add seawater and mix carefully by inversion. Spin the vial in a tabletop centrifuge at
the lowest possible setting (e.g., 500xg) for 3 minutes to form a lightly packed pellet. Wash and spin the
eggs twice more. If at any time the wash water appears red the eggs are lysing (the membranes have been
disturbed) and the eggs are unsuitable for testing; discard these eggs and start again.
After washing, transfer the washed eggs to a beaker containing 200 mL of control seawater. This is called
the "egg test stock." Mix the stock solution using gentle aeration until the egg solution is homogenous. The
aeration device used in Narragansett is a 3-pronged diffuser attached by flexible tubing to an air pump.
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Sperm Cell Toxicity Tests Using the Sea Urchin (Arbacia punctulata)
Supplement to Training Video
Make a 1:10 dilution of the test stock for the purpose of counting the eggs. Cut the point from a wide-
mouth pipet tip to make sure the eggs will not be damaged and transfer 1 ml of egg solution to a vial
containing 9 ml of control water. Mix by inversion.
Transfer 1 ml of the egg solution to a Sedgewick-Rafter counting chamber. Count the number of eggs
under a dissecting microscope at 25X magnification. Ten times the number of eggs in that milliliter equals
the number of eggs/mL in the egg stock. The target concentration for test initiation is 2,000 eggs/ml.
If the egg count is greater than or equal to 200 eggs, add the proper volume of water:
(# of eggs counted) - 200 = volume (ml) of control water to add
If less than 200 eggs were
counted, allow the eggs to
settle in the beaker, remove
the supernatant water to con-
centrate the eggs to greater
than 200, repeat the count,
and dilute the egg test stock
as described above.
Verify the concentration by
counting 1 ml of a 1:10
dilution of the adjusted stock
solution. The count for the
final dilution should equal
100 ± 20 eggs/mL. The test
requires 24 ml of egg test
stock for a control and five
exposure concentrations.
EFFLUENT PREPARATION
Dilution Water
The type of dilution water used to make the test concentrations ,>s
dependent on the objectives of the test Any specific requirements
included in NPDES permits should be followed. The Saltwater Chronic
Methods Manual (Section 7) provides the following guidelines:
* If the test is conducted to estimate the absolute
of the effluent, synthetic dilution water should be used. If the1 cultures
were maintained in different water than used for dilution wafer, a
second set of control replicates should be conducted using the culture
water.
• If the test is conducted to estimate the chronic toxicity of the
effluent in uncontaminated receiving waters, the test con be
conducted using a grab sample of the receiving waters collected outside
the influence of the outfall, other uncontaminated waters, or standard
dilution water with the same salinity as the receiving waters, if the
cultures were maintained in different water than used for dilution water,
a second set of control replicates should be conducted using the culture
• If the test is conducted to estimate the additive or mitigating
effects of the effluent on already contaminated receiving
waters, the test must be conducted using receiving waters collected
outside the influence of the outfall. Controls should be conducted us/'r
both receiving water and culture water.
Effluent sampling should
be conducted according to
Section 8 of the Saltwater
Chronic Methods Manual
(EPA, 2002a) and any
specific requirements of a
NPDES permit. The effluent
or receiving waters should be
held at 0°C - 6°C until used
for testing. Under the NPDES
program, lapsed time from
sample collection to first use in the test must not exceed 36 hours. Under special conditions or variances,
samples may be held longer but should never be used for testing if held for more than 72 hours.
Maintain the salinity of the test samples to 30%o ± 2%o. To do this, effluent samples may need to be
adjusted using hypersaline brine (MSB). A recipe for MSB is provided in Appendix A of this manual.
Approximately 1 hour before the test is to begin, adjust approximately 1 L of effluent to the test tempera-
ture of 20°C ± 1°C and maintain that temperature while preparing the test concentrations. To test a series
of decreasing concentrations of effluent, use a dilution factor of > 0.5. When starting with effluent that
has 0%o salinity and using MSB, the maximum effluent concentration that can be prepared at 30%o is 70
percent effluent. Table A-l presents the volumes needed for the test concentrations using MSB.
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U'S- ENVIRONMENTAL PROTECTION AGENCY Sperm Cell Toxicity Tests Using the Sea Urchin (Arbacia punctulata)
Supplement to Training Video
BEGINNING THE TEST
In Narragansett, disposable glass vials are used as test chambers. They are labeled with concentration
and replicate numbers and arranged in the partitioned cardboard box in which they are shipped. Prepare
the effluent dilutions for four replicates of each concentration and the control solution to reduce variability
among replicates. Each concentration should be prepared in one beaker and 5 ml distributed to each of the
test chambers. Be sure the effluent temperature has been brought up to 20°C before beginning the test.
Within 1 hour of collecting and preparing the sperm test stock, add 100 pL of the well-mixed sperm test
stock to each test and control vial. Cover the chambers, record the time, and maintain the chambers at
20°C + 1°C for 1 hour.
At the end of the hour, mix the egg test stock using gentle aeration and add 1 mL of the egg solution to
each exposure vial using a wide-mouth pipet. When all of the vials contain eggs, lift the storage box and
gently move it in circles to "swirl" the egg-sperm suspension. Cover the chambers, record the time, and
incubate the eggs and sperm at 20°C + 1°C for 20 minutes.
ROUTINE CHEMISTRIES
At the beginning of the exposure period, DO, pH, temperature and salinity are measured in one chamber at
each test concentration and the control.
TERMINATING THE TEST
After 20 minutes, end the test and preserve the samples by adding 2 mL of 1% formalin in seawater to
each vial. .Cap the vials and record the time. The test should be evaluated immediately but can be evalu-
ated up to 48 hours later.
Test Acceptability and Data Review
This test demonstrates the effluent or receiving water's effect on sea urchin fertilization. To evaluate this,
exposed and control eggs are examined under a microscope and the number of unfertilized eggs in each
test chamber is recorded.
For each replicate, transfer about 80 - 120 uL of the preserved eggs to a multiple-chamber counting slide.
If a Sedgewick-Rafter counting chamber is used, transfer about 1 mL. Using a compound microscope at
100X magnification, observe 100 - 200 eggs per sample. This should be done with adequate ventilation,
preferably under a hood, to reduce exposure to the formalin fumes.
For each test chamber, record the total number of eggs counted, and the number that were not fertilized.
Fertilized eggs are surrounded by a fertilization membrane, while unfertilized eggs lack this membrane.
Abnormal eggs are not counted. Figure E-2 in Appendix E provides a sample data collection sheet.
Test data are reviewed to verify that test acceptability criteria (TAG) requirements for a valid test have been
met. For the test to be acceptable, the control chambers are required to have between 70% and 90% fer-
tilization of the eggs. The concentration-response relationship generated for each multi-concentration test
must be reviewed to ensure that calculated test results are interpreted appropriately. In conjunction with
this requirement, EPA has provided recommended guidance for concentration-response relationship review
(EPA, 2000b).
EPA's promulgated toxicity testing method manuals (EPA, 2002a, b) recommend the use of point estima-
tion technique approaches for calculating endpoints for effluent toxicity tests under the NPDES program.
The promulgated methods also require a data review of toxicity data and concentration-response data, and
require calculating the percent minimum significant difference (PMSD) when point estimation (e.g., LC50,
IC25) analyses are not used. EPA specifies the PMSD must be calculated when NPDES permits require sub-
lethal hypothesis testing. EPA also requires that variability criteria be applied as a test review step when
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U.S. ENVIRONMENTAL PROTECTION AGENCY Sperm Cell Toxicity Tests Using the Sea Urchin (Arbacia punctulata)
Supplement to Training Video
NPDES permits require sub-lethal hypothesis testing endpoints (i.e., no observed effect concentration
[NOEC] or lowest observed effect concentration [LOEC]) and the effluent has been determined to have no
toxicity at the permitted receiving water concentration (EPA, 2002b). This reduces the within-test variabil-
ity and increases statistical sensitivity when test endpoints are expressed using hypothesis testing rather
than the preferred point estimation techniques.
The sea urchin sperm cell test is currently used to assess the potential toxic effects of complex chemical
mixtures on marine and estuarine organisms. Used in conjunction with chemical-specific methods, this test
can provide a comprehensive and effective approach to assessing the impact of complex effluents dis-
charged to the marine and estuarine environments.
Citations and Recommended References
EPA. 1991. Technical Support Document for Water Quality-based Toxics Control. U.S. EPA Office of Water
Enforcement and Permits, Washington, D.C. EPA-505-2-90-001.
EPA. 2000a. Method Guidance and Recommendations for Whole Effluent Toxicity (WET) Testing (40 CFR
Part 136). Office of Water, Washington, D.C. EPA 821-B-00-004.
EPA. 2000b. Understanding and Accounting for Method Variability in Whole Effluent Toxicity Applications
Under the National Pollutant Discharge Elimination System Program. Office of Wastewater
Management, Washington, D.C. EPA 833-R-00-003.
EPA. 2002a. Short-term Methods for Estimating the Chronic Toxicity of Effluents and Receiving Waters to
Marine and Estuarine Organisms, Third Edition. (Saltwater Chronic Methods Manual). Office of
Water, Cincinnati, OH. EPA-821-R-02-014.
EPA. 2002b. Methods for Measuring the Acute Toxicity of Effluents and Receiving Waters to Freshwater
and Marine Organisms, Fifth Edition. (Acute Methods Manual). Office of Water, Cincinnati, OH.
EPA-821-R-02-012.
EPA. 2002c. Final Rule. 40 CFR Part 136. Guidelines Establishing Test Procedures for the Analysis of
Pollutants; Whole Effluent Toxicity Test Methods. 67 FR 69952-69972, November 19, 2002.
EPA. 2009. Sperm Cell Toxicity Tests Using the Sea Urchin (Arbacia punctulata). Supplement to Training
Video. Whole Effluent Toxicity Training Video Series, Saltwater Series. March 2009. EPA 833-C-
09-001.
EPA references are available online atwww.epa.gov/npdes.
If you need additional copies of this document, you can download it at:
www.epa.gov/npdes/wqbasedperrnitting.
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Glossary
Acute toxicity. An adverse effect measured on a group of test organisms during a short-term exposure in
a short period of time (96 hours or less in toxicity tests). The effect can be measured in lethality
or any variety of effects.
Arbacia punctulata. A species of Arbacia genus of purple-spined sea urchins. Its natural habitat is in the
Western Atlantic Ocean. Arbacia punctulata can be found in shallow water from Massachusetts
to Cuba and the Yucatan Peninsula, from Texas to Florida in the Gulf of Mexico, the coast from
Panama to French Guiana and in the Lesser Antilles, usually on rocky, sandy, or shelly bottoms.
Chronic toxicity. An adverse effect that occurs over a long exposure period. The effect can be lethality,
impaired growth, reduced reproduction, etc.
Diluent water. Dilution water used to prepare the effluent concentrations.
Effluent concentrations. Concentrations or dilutions of an effluent sample to which test organisms are
exposed to determine the biological effects of the sample on the test organism.
Effluent sample. A representative collection of the discharge that is to be tested.
Flow-through water delivery system. An open water flow system that delivers fresh water or seawater to
culture tanks and is disposed of after it leaves those tanks.
Hypothesis testing. Technique (e.g., Dunnett's test) that determines what concentration is statistically
different from the control. Endpoints determined from hypothesis testing are NOEC and LOEC.
IC2s (Inhibition Concentration, 25%). The point estimate of the toxicant concentration that would cause a
25% reduction in a non-quantal biological measurement (e.g., reproduction or growth) calculated
from a continuous model.
Laminaria. The scientific name for a species of kelp given as food to laboratory sea urchins.
LCSO (Lethal Concentration, 50%). The concentration of toxicant or effluent that would cause death to
50% of the test organisms at a specific time of observations (e.g., 96-hour LC50).
Lowest Observed Effect Concentration (LOEC). The LOEC is the lowest concentration of toxicant to
which organisms are exposed in a test, which causes statistically significant adverse effects on
the test organisms (i.e., where the values for the observed endpoints are statistically significantly
different from the control). The definitions of NOEC and LOEC assume a strict dose-response
relationship between toxicant concentration and organism response.
Minimum Significant Difference (MSD). The MSD is the magnitude of difference from the control where
the null hypothesis is rejected in a statistical test comparing a treatment with a control. MSD
is based on the number of replicates, control performance and power of the test. MSD is often
measured as a percent and referred to as PMSD.
No Observed Effect Concentration (NOEC). The NOEC is the highest tested concentration of toxicant to
which organisms are exposed in a full life-cycle or partial life-cycle (short-term) test, that causes
no observable adverse effect on the test organism (i.e., the highest concentration of toxicant
at which the values for the observed responses are not statistically significantly different from
the controls). NOECs calculated by hypothesis testing are dependent upon the concentrations
selected.
Glossary-1
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U.S. ENVIRONMENTAL PROTECTION AGENCY Sperm Cell Toxicity Tests Using the Sea Urchin (Arbacia punctulata)
Supplement to Training Video
NPDES (National Pollutant Discharge Elimination System) Program. The national program for issuing,
modifying, revoking, and reissuing, terminating, monitoring and enforcing permits, and imposing
and enforcing pretreatment requirements, under Sections 307, 318, 402, and 405 of the Clean
Water Act.
Point Estimation Techniques. This technique is used to determine the effluent concentration at which
adverse effects (e.g., fertilization, growth or survival) occurred, such as Probit, Interpolation
Method, Spearman-Karber. For example, a concentration at which a 25% reduction in
reproduction and survival occurred.
Receiving Water Concentration (RWC). The RWC is the concentration of a toxicant or the parameter
toxicity in the receiving water (i.e., riverine, lake, reservoir, estuary or ocean) after mixing.
Recirculating water delivery system. A water flow system that treats water after it passes through the
culture tanks (usually with sand and biofilters) and delivers the same treated water back to the
tanks.
Toxicity test. A procedure to measure the toxicity of a chemical or effluent using living organisms. The
test measures the degree of response of an exposed organism to a specific chemical or effluent.
WET (Whole effluent toxicity). The total toxic effect of an effluent measured directly with a toxicity test.
Glossary-2
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U.S. ENVIRONMENTAL PROTECTION AGENCY Sperm Cell Toxicity Tests Using the Sea Urchin (Arbacia punctulata)
Supplement to Training Video
Appendix A:
Preparing Hypersaline Brine (HSB)
Salinity adjustments are a vital part of using marine and estuarine species for toxicity testing. Because the
majority of industrial and sewage treatment effluents entering marine and estuarine waters contain little or
no measurable salts, the salinity of these effluents must be adjusted before exposing estuarine or marine
plants and animals to the test solutions. It also is important to maintain constant salinity across all treat-
ments throughout the test for quality control. Finally, matching the test solution's salinity to the expected
receiving water's salinity may require salinity adjustments. NHEERL-AED uses HSB, prepared from filtered
natural seawater, to adjust exposure solution salinities.
HSB 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. HSB can be held for prolonged periods without any apparent degradation, added directly to
the effluent to increase the salinity, or used as control water by diluting to the desired salinity with deion-
ized 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.
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, fol-
lowed by several thorough deionized-water rinses.
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 urn before placing into the brine tank. 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 could contaminate the brine. A
thermostatically controlled heat exchanger made from fiberglass is suggested.
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 (or more or
less often, depending on the volume being generated) to ensure that the salinity does not exceed 100%o
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 NHEERL-AED was a reduction
in the alkalinity of seawater made from brine with salinity greater than 100%o, 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 1-mm filter and pump
or pour it directly into portable containers (20-L cubitainers or polycarbonate water cooler jugs are most
suitable). Cap the containers, and record the measured salinity and the date generated. Store the brine in
the dark at room temperature.
A-1
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.S. ENVIRONMENTAL PROTECTION AGENCY
Sperm Cell Toxicity Tests Using the Sea Urchin (Arbacia punctulata)
Supplement to Training Video
SALINITY ADJUSTMENTS USING HYPERSALINE BRINE
To calculate the volume of brine (Vb) to add to a 0%o sample to produce a solution at a desired salinity (Sf),
use this equation: .
Where:
sf =
vf =
w * c
vb 3t
volume of brine, ml
salinity of brine, %o
final salinity, %o
final volume needed, ml
Table A-l presents volumes needed to make 30%o test solutions from effluent (0%o), deionized water, and
100%o MSB. At 30%o salinity, the highest achievable concentration is 70% effluent.
Table A-l. Preparation of Test Solutions at a Salinity of 30%o Using MSB for a Final Test Concentration
Volume of 1000 ml.
Exposure
Concentration (%)
70
25
7
2.5
0.7
Control
Effluent
(0 %o)
(mL)
700
250
70
25
7
—
Deionized Water
(mL)
—
450
630
675
693
1,000
Hypersaline Brine
(I00%o)
(mL)
300
300
300
300
300
—
Table A-2 gives examples of attainable exposure concentrations and dilution volumes needed when an
effluent salinity is raised to 30%o using artificial sea salts and using 0.5 serial dilution.
Table A-2. Preparation of Test Solutions at a Salinity of 30%o Using Natural Seawater or Artificial Sea Salts.1
Effluent Solution
1
2
3
4
5
Control
Total
Effluent Concentration
(%)
100
50
25
12.5
6.25
0.0
Solutions To Be Combined
Volume of Effluent
Solution (mL)
840
420
420
420
420
Volume of Diluent
Seawater (30%o) (mL)
— •
Solution 1 + 420
Solution 2 + 420
Solution 3 + 420
Solution 4 + 420
420
2,080
^-Tiii's illustration assumes: /) the use of 5 ml of test solution in each of four replicates (total of 20 mL) for the control and five concentra-
tions of effluent, 2) an effluent dilution factor of 0.5, 3) the effluent lacks appreciable salinity, and 4) 400 mL of each test concentration is
used for chemical analysis. A sufficient initial volume (840 mL) of effluent is prepared by adjusting the salinity to 30%o. In this example, the
salinity is adjusted by adding artificial sea salts to the 100% effluent, and preparing a serial dilution using 30%o seawater (natural seawater,
HSB, or artificial seawater). Stir solutions I hour to ensure that the salts aVssolve. The salinity of the initial 840 mL of 100% effluent is
adjusted to 30%<> by adding 25.2 g of dry artificial sea salts (FORTY FATHOMS®). Test concentrations are then made by mixing appropri-
ate volumes of salinity adjusted effluent and 30%0 salinity dilution water to provide 840 mL of solution for each concentration. IfHSB alone
(I00%o) is used to adjust the salinity of the effluent, the highest concentration of effluent that could be tested would be 70% at 30%o salinity.
Source: EPA, 2002a.
A-2
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U.S. ENVIRONMENTAL PROTECTION AGENCY Sperm Cell Toxicity Tests Using the Sea Urchin (Arbacia punctulata)
• Supplement to Training Video
Appendix B:
Apparatus and Equipment
Air lines, and air stones. For aerating water containing adults, or for supplying air to test solutions with
low DO.
Air pump. For oil-free air supply.
Balance. Analytical, capable of accurately weighing to 0.00001 g.
Beakers or flasks. Six, borosilicate glass or non-toxic plasticware, 1000 ml for making test solutions.
Centrifuge. Bench-top, slant-head, variable speed for washing eggs.
Centrifuge tubes. Conical for washing eggs.
Compound microscope. For examining and counting sperm cells and fertilized eggs (25X and 100X).
Count register. 2-place for recording sperm and egg counts.
Cylindrical glass vessel. 8-cm diameter for maintaining dispersed egg suspension.
Dissecting microscope. For counting diluted egg stock (100X).
Environmental chamber or equivalent facility with temperature control (20°C + 1°C).
Fume hood. To protect from formaldehyde fumes.
Glass dishes. Flat bottomed, 20-cm diameter for holding sea urchins during gamete collection.
Hemacytometer, Neubauer. For counting sperm.
Ice bucket. Covered for maintaining live sperm after collection until test initiation.
Laboratory sea urchins, Arbacia punctulata, culture unit. To test effluent or receiving water toxicity,
sufficient eggs and sperm must be available from healthy adult animals.
Meters: pH and DO, and specific conductivity. For routine physical and chemical measurements.
Pipets, automatic. Adjustable 1 - 100 ml.
Pipets, serological. 1-10 mL, graduated.
Pipets, volumetric. Class A, 1 - 100 ml.
Pipet bulbs and filters. Propipet®, or equivalent.
Reference weights, Class S. For checking performance of balance. Weights should bracket the expected
weights of materials to be weighed.
Refractometer or other method. For determining salinity.
Samplers. Automatic sampler, preferably with sample cooling capability, that can collect a 24-hour
composite sample of 5 L.
B-l
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U.S. ENVIRONMENTAL PROTECTION AGENCY Sperm Cell Toxicity Tests Using the Sea Urchin (Arbacia punctulata)
Supplement to Training Video
Sedgwick-Rafter counting chamber. For counting egg stock and examining fertilized eggs.
Syringes. 1 ml, and 10 ml, with 18 gauge, blunt-tipped needles (tips cut off) for collecting sperm and
eggs.
Thermometers. National Bureau of Standards Certified (see EPA 2002a). Used to calibrate laboratory
thermometers.
Thermometers, glass or electronic, laboratory grade. For measuring water temperatures.
Transformer, 10-12 Volt. With steel electrodes for stimulating release of eggs and sperm.
Vacuum suction device. For washing eggs.
Volumetric flasks and graduated cylinders. Class A, Borosilicate glass or non-toxic plastic labware,
10 - 1000 ml for making test solutions.
Wash bottles. For deionized water, for washing organisms from containers and for rinsing small glassware
and instrument electrodes and probes.
Water purification system. Millipore® Milli-Q® deionized water or equivalent.
B-2
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U.S. ENVIRONMENTAL PROTECTION AGENCY Sperm Cell Toxicity Tests Using the Sea Urchin (Arbacia punctulata)
Supplement to Training Video
if'. '«>.«,* *•*.,.
Appendix C:
Reagents and Consumable Materials
Acetic acid. 10%, reagent grade, in seawater for preparing killed sperm dilutions.
Buffers pH 4, pH 7, and pH 10. (Or as per instructions of instrument manufacturer) for standards and
calibration check.
Data sheets (one set per test). For data recording (see Appendix E).
Effluent, receiving water, and dilution water. Test waters, including effluent, receiving, and dilution
water should be analyzed to ensure its quality prior to using in tests. Dilution water containing
organisms that might prey upon or otherwise interfere with the test organisms should be filtered
through a fine mesh (with 150 urn or smaller openings).
Food. Kelp, Laminaria sp., or romaine lettuce for the sea urchin, Arbacia punctulata.
Formalin. 1%, in 2 ml of seawater for preserving eggs at end of test.
Gloves, disposable; lab coat and protective eyewear. For personal protection from contamination.
Laboratory quality assurance samples and standards. For calibration of the above methods.
Markers, waterproof. For marking containers, etc.
Parafilm. To cover tubes and vessels containing test materials.
Reagent water Distilled or deionized water that does not contain substances which are toxic to the test
organisms.
Reference toxicant solutions. Reference toxicants such as sodium chloride (NaCI), potassium chloride
(KCI), cadmium chloride (CdCI2), copper sulfate (CuS04), sodium dodecyl sulfate (SDS), and
potassium dichromate (K2Cr207), are suitable for use in the NPDES Program and other Agency
programs requiring aquatic toxicity tests.
Saline test and dilution water. The salinity of the test water must be in the range of 20%o - 30%o. The
salinity should vary by no more than + 2%o among the chambers on a given day. If effluent and
receiving water tests are conducted concurrently, the salinities of these tests should be similar.
It is important to maintain a constant salinity across all treatments during a test. It is desirable
to match the test salinity with that of the receiving water. Two methods are available to adjust
salinities — a hypersaline brine (MSB) derived from natural seawater or artificial sea salts. Both
are described in EPA, 2002.
Sample containers. For sample shipment and storage.
Sea Urchins. Arbacia punctulata, minimum of 12 of each sex.
Scintillation vials. 20 ml, disposable; to prepare test concentrations.
Standard salt water aquarium or Instant Ocean Aquarium. Capable of maintaining seawater at 15°C,
with appropriate filtration and aeration system.
Tape, colored. For labeling tubes.
C-l
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U.S. ENVIRONMENTAL PROTECTION AGENCY Sperm Cell Toxicity Tests Using the Sea Urchin (Arbacia punctulata)
Supplement to Training Video
Intentionally Left Blank
C-2
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Sperm Cell Toxicity Tests Using the Sea Urchin (Arboc/a punctufato)
Supplement to Training Video
Appendix D:
Summary of Test Conditions and Test Acceptability
Criteria
Summary of Test Conditions and Test Acceptability Criteria for Sea Urchin, Arbacia punctulata, Fertilization Test with Effluent and
Receiving Waters (Test Method I008.0)1
Test type
Salinity
Temperature (C°)
Light quality
Light intensity
Test chamber size
Test solution volume
Number of sea urchins
Number of eggs and sperm cells per chamber
Number of replicate chambers per concentration
Dilution water
Test concentrations
Dilution factor
Test duration
Endpoint
Test acceptability criteria
Sampling requirements
Sample volume required
Static, non-renewal (required)
30%0 ± 2%o of the selected test salinity (recommended)
20°C ± I°C (recommended) Test temperatures must not deviate by more
than 3°C during the test (i.e., max. temp - min. temp S 3°C) (required)
Ambient laboratory light during test preparation (recommended)
10-20 uE/m2/s, or 50 - 100 ft-c (Ambient laboratory levels) (recommend-
ed)
Disposable (glass) liquid scintillation vials (20 mL capacity), pre-soaked in
control water (recommended)
5 mL (recommended)
Pooled eggs from 4 females and pooled sperm from 4 males per test
(recommended)
About 2,000 eggs and 5,000,000 sperm cells per vial (recommended)
4 (required minimum)
Uncontaminated source of natural seawater; deionized water mixed with
MSB or artificial sea salts (available options)
Effluents: 5 and a control (required minimum) Receiving waters: 100%
receiving water (or minimum of 5) and a control (recommended)
Effluents: ^ 0.5 (recommended)
Receiving Waters: None or > 0.5 (recommended)
1 hour and 20 minutes (required)
Fertilization of sea urchin eggs (required)
70% - 90% egg fertilization in controls (required)
For on-site tests, one sample collected at test initiation, and used within
24 hr of the time it is removed from the sampling device. For off-site tests,
holding time must not exceed 36 hr before first use for NPDES compliance
testing, (required)
1 L per test (recommended)
^-Source: EPA, 2002a. For the purposes of reviewing WET test data submitted under NPDES permits, each test condition
listed above is identified as required or recommended. Additional requirements may be provided in individual permits,
such as specifying a given test condition where several options are given in the method.
D-l
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U.S. ENVIRONMENTAL PROTECTION AGENCY Sperm Cell Toxicity Tests Using the Sea Urchin (Arbacia punctulata)
Supplement to Training Video
,",„:,, t
Intentionally Left Blank
D-2
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Sperm Cell Toxicity Tests Using the Sea Urchin (Arbacia punctulata)
Supplement to Training Video
Appendix E:
Data Sheets
Figure E-l. Sperm Cell Toxicity Test, Sample Data Sheet #1
Test ID:
Performed By:.
Sperm Dilutions:
Hemacytometer Count, E:
Sperm Concentrations
Solution Selected for Test (>5X 107 SPM):
Dilution: SPM/(5 X 107) =
X 104 = SPM "E"
"E"X40 = A = SPM
"E"X20 = B = SPM
"E"X5 = D = SPM
DF
((DF) X 10) -10 = _
Final Sperm Counts = _
Egg Dilutions:
Initial Egg Count: = _
Egg Stock Concentration = Egg Count (1 ml of 1:10 dilution) X 10: = _
(Allow eggs to resettle and recount until count < 200)
Volume of SW to Add to Dilute Egg Stock to 2000/mL: Egg Count - 200: =
Verify Final Egg Count (in 1 ml of 1:10 dilution): = _
(Count should = 100 + 20 eggs/mL)
Test Stocks:
Sperm Stock:
+ SW, ml
Egg Stock:
Volume Added/Test Vial:
Volume Added/Test Vial
Test Times:
Sperm Collection:
Egg Collection:
Sperm Added:
Eggs Added:
Fixative Added:
Samples Read:
(5 X 107 SPM)
(100 uL)
(2000/mL)
(ImL)
Salinities:
E-l
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Sperm Cell Toxicity Tests Using the Sea Urchin (Arbacio punctulata)
Supplement to Training Video
Figure E-2. Sperm Cell Toxicity Test, Sample Data Sheet #2 - Raw Data
Test ID:
Performed by:
Time:.
Date:
Egg Counts at End of Test
Cone. (%)
Replicate 1
Total
Unfert
Replicate 1
Total
Unfert
Replicate 3
Total
Unfert
Replicate 4
Total
Unfert
Statistical Analysis:
Analysis of variance:
Control:
Different from Control (P):
Comments:
E-2
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If you need additional copies of this document, you can download it at:
www.epa.gov/npdes/wqbasedpermitting
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WHOLE EFFLUENT TOXICITY • TRAINING VIDEO SERIES • saltwater series
Sheepshead Minnow (Cyprinodon
variegatus) and Inland Silverside
(Men/d/o beryllina) Larval Survival
and Growth Toxicity Tests
Supplement to Training Video
U.S. Environmental Protection Agency
Office of Wastewater Management
Water Permits Division
1200 Pennsylvania Ave,, NW
Washington, DC 20460
EPA 833-C-09-001
March 2009
-------
NOTICE
The revision of this guide has been funded wholly or in part by the
Environmental Protection Agency under Contract EP-C-05-063. Mention of
trade names or commercial products does not constitute endorsement or
recommendation for use.
-------
U.S. ENVIRONMENTAL PROTECTION AGENCY Sheepshead Minnow (Cyprinodon variegatus) and Inland Silverside (Menidia beryllina)
Larval Survival and Growth Toxicity Test • Supplement to Training Video
Foreword
This supplemental guide serves as a supplement to the video "Sheepshead Minnow (Cyprinodon varie-
gatus) and Inland Silverside (Menidia beryllina) Larval Survival and Growth Toxicity Tests" (EPA, 2009).
The methods illustrated in the video and described in this guide support the methods published in the
U.S. Environmental Protection Agency's (EPA's) Short-term Methods for Estimating the Chronic Toxicity of
Effluents and Receiving Waters to Marine and Estuarine Organisms, Third Edition (EPA, 2002a), referred
to as the Saltwater Chronic Methods Manual. The video and this guide provide details on preparing for
and conducting the test based on the expertise of personnel at the following EPA Office of Research and
Development (ORD) laboratories:
National Health and Environmental Effects Research Laboratory (NHEERL) - Atlantic Ecology Division
in Narragansett, Rhode Island
NHEERL - Gulf Ecology Division in Gulf Breeze, Florida
National Exposure Research Lab (NERL) - Ecological Exposure Research Division (EERD) in
Cincinnati, Ohio
This guide and its accompanying video are part of a series of training videos produced by EPA's Office of
Wastewater Management. This Saltwater Series includes the following videos and guides:
"Mysid (/Amen'camys/s bahia) Survival, Growth, and Fecundity Toxicity Tests"
"Culturing/Americamys/s bahia"
"Sperm Cell Toxicity Tests Using the Sea Urchin, Arbacia punctulata"
"Red Algal (Champia parvula) Sexual Reproduction Toxicity Tests"
"Sheepshead Minnow (Cyprinodon variegatus) and Inland Silverside (Menidia beryllina) Larval Survival
and Growth Toxicity Tests"
The Freshwater Series, released in 2006, includes the following videos and guides:
"Ceriodaphnia Survival and Reproduction Toxicity Tests"
"Culturing of Fathead Minnows (Pimephales promelas)"
"Fathead Minnow (Pimephales promelas) Larval Survival and Growth Toxicity Tests"
All of these videos are available through the National Service Center for Environmental Publications
(NSCEP) at 800 490-9198 or nscep@bps-lmit.com.
-------
U.S. ENVIRONMENTAL PROTECTION AGENCY Sheepshead Minnow (Cyprinodon vor/egotus) and Inland Silverside (Men/did beryllina)
Larval Survival and Growth Toxicity Test • Supplement to Training Video
• : .>-* v-**'S'-;,-<*r
Intentionally Left Blank
-------
U.S. ENVIRONMENTAL PROTECTION AGENCY Sheepshead Minnow (Cyprinodon variegotus) and Inland Silverside (Menidia beryffina)
Larval Survival and Growth Toxicity Test • Supplement to Training Video
Contents
Foreword i
Introduction 1
Background 1
Care and Feeding of Adults and Larvae 1
Culture Water 2
Photoperiod '. 2
Culture Vessels 2
Water Delivery Systems 2
Food Preparation 2
Obtaining Larvae for Toxicity Tests 3
Culture Water 4
Photoperiod 5
Culture Vessels 5
Water Delivery Systems 5
Food Preparation 5
Obtaining Larvae for Toxicity Tests 5
Test Method 6
Effluent Sampling 6
Dilution Preparation 6
Routine Chemistries , 7
Renewals 7
Feeding 8
Test Termination 8
Test Acceptability and Data Review 9
Other Procedural Considerations 9
Citations and Recommended References 9
Glossary Glossary-1
Appendix A: Preparing Hypersaline Brine (HSB) A-l
Appendix B: Preparing Brine Shrimp and Rotifers for Feeding B-l
Appendix C: Apparatus and Equipment - Sheepshead Minnow and Inland Silverside Tests.... C-l
Appendix D: Reagents and Consumable Materials D-l
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U.S. ENVIRONMENTAL PROTECTION AGENCY Sheepshead Minnow (Cyprinodon variegatus) and Inland Silverside (Menidia beryllina)
Larval Survival and Growth Toxicity Test • Supplement to Training Video
Appendix E: Summary of Test Conditions and Test Acceptability Criteria E-l
Appendix F: Data Sheets F-l
FIGURES
Figure 1. Embryonic development of sheepshead minnow, Cyprinodon variegatus: A. Mature
unfertilized egg, showing attachment filaments and micropyle, X33; B. Blastodisc fully devel-
oped; C/D. Blastodisc, 8 cells; E. Blastoderm, 16 cells; F. Blastoderm, late cleavage stage; G.
Blastoderm with germ ring formed, embryonic shield developing; H. Blastoderm covers over % of
yolk, yolk noticeably constricted; I. Early embryo. (Continued, J - 0 on page 4) 3
Figure 1 (continued). Embryonic development of sheepshead minnow, Cyprinodon variegatus: J.
Embryo 48 h after fertilization, no segmented throughout, pigment on yolk sac and body, otoliths
formed; K. Posterior portion of embryo free from yolk and moves freely within egg membrane, 72
h after fertilization; L. Newly hatched fish, actual length 4 mm; M. Larval fish 5 days after hatch-
ing, actual length 5 mm; N. Young fish 9 mm in length; 0. Young fish 12 mm in length 4
Figure 2. Inland silverside, Menidia beryllina: A. Adult, ca. 64 mm SL; B. Egg (diagrammatic), only
bases of filaments shown; C. Egg, 2-cell stage; D. Egg, morula stage; E. Advanced embryo, 2Vz
days after fertilization 6
Figure 3. Glass test chamber with sump area. Modified from Norberg and Mount (1985) 7
Figure C-l. Glass test chamber with sump area. Modified from Norberg and Mount (1985). ... C-2
Figure F-l. Data Form for the Sheepshead Minnow and Inland Silverside, Larval Survival and
Growth Toxicity Test. Daily Record of Larval Survival and Test Conditions F-l
Figure F-2. Data Form for the Sheepshead Minnow and Inland Silverside, Larval Survival and
Growth Toxicity Test. Summary of Test Results F-3
Figure F-3. Data Form for the Sheepshead Minnow and Inland Silverside, Larval Survival and
Growth Toxicity Test. Dry Weights of Larvae .. F-4
TABLES
Table A-l. Preparation of Test Solutions at a Salinity of 20%0 Using MSB for a Final Test
Concentration Volume of 4000 mL A-2
Table E-l. Summary of Test Conditions and Test Acceptability Criteria for the Sheepshead
Minnow, Cyprinodon variegatus, Larval Survival and Growth Test with Effluents and Receiving
Waters (Test Method 1004.0) E-l
Table E-2. Summary of Test Conditions and Test Acceptability Criteria for the Inland Silverside,
Menidia beryllina, Larval Survival and Growth Test with Effluents and Receiving Waters (Test
Method 1006.0) E-2
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U.S. ENVIRONMENTAL PROTECTION AGENCY Sheepshead Minnow (Cyprinodon variegotus) and Inland Silverside (Menidia beryllina)
Larval Survival and Growth Toxicity Test • Supplement to Training Video
Introduction
This guide accompanies the Environmental Protection Agency's (EPA's) video training for conducting
sheepshead minnow (Cyprinodon variegatus) and inland silverside (Menidia beryllina) larval survival and
growth toxicity tests (EPA, 2009). The test methods are found in Short-term Methods for Estimating the
Chronic Toxicity of Effluents and Receiving Waters to Marine and Estuarine Organisms, Third Edition (EPA,
2002a). The tests were developed by EPA's Office of Research and Development's (ORD's) National Health
and Environmental Effects Research Laboratory - Aquatic Ecology Division (NHEERL-AED) in Narragansett,
Rhode Island. The material presented in both the video and this guide summarizes the methods but does
not replace a thorough review and understanding of the methods by laboratory personnel before conduct-
ing the test.
Background
Under the National Pollutant Discharge Elimination System (NPDES) program (Section 402 of the Clean
Water Act), EPA uses toxicity tests to monitor and evaluate effluents for their toxicity to biota and their
impact on receiving waters. By determining acceptable or safe concentrations for toxicants discharged
into receiving waters, EPA can establish NPDES permit limitations for toxicity. These whole effluent toxicity
(WET) permit limitations regulate pollutant discharges on a whole effluent effect basis rather than solely by
a chemical specific approach.
Whole effluent toxicity methods measure the synergistic, antagonistic, and additive effects of all the chemi-
cal, physical, and additive components of an effluent that adversely affect the physiological and biochemi-
cal functions of the test organisms. Therefore, healthy organisms and correct laboratory procedures are
essential for valid test results. Laboratory personnel should be very familiar with the test methods and with
sheepshead minnows and inland silverside handling techniques before conducting a test.
This supplemental guide covers the procedures for conducting the test according to EPA's promulgated
methods (40 CFR Part 136; EPA, 2002c) and also provides some helpful information that is not presented
in the Saltwater Chronic Methods Manual (EPA, 2002a).
This guide summarizes methods developed at ORD 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 short-term tests span an exposure time
of 7 days to estimate the chronic toxicity of effluent or receiving water on newly-hatched larvae in a static
renewal exposure system. The methods described in this guide and demonstrated in the accompanying
video 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, Third Edition (EPA, 20023)1.
Care and Feeding of Adults and Larvae
SHEEPSHEAD MINNOWS
Adult sheepshead minnows (Cyprinodon variegatus) can be field collected from Atlantic and Gulf of Mexico
coastalestuaries 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. Fish that are field-caught should be held for a
minimum of 2 weeks before use in testing to determine that they are healthy and not injured.
1 The methods for these two species are presented together in the video and this guide because they are conducted in a very similar man-
ner. The complete methods in the Saltwater Chronic Methods Manual are presented in Section II (Sheepshead Minnows) and Section 13
(Inland Silverside).
I
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U.S. ENVIRONMENTAL PROTECTION AGENCY Sheepshead Minnow (Cyprinodon voriegotus) and Inland Silverside (Men/did beryllina)
Larval Survival and Growth Toxicity Test • Supplement to Training Video
- >u - ;,ft, s* r f- • . j-
CULTURE WATER
The quality of water used for test organism culturing and for dilution water in toxicity tests is extremely
important. Water for these two uses should come from the same source. Holding and rearing tanks and any
area used for manipulating live sheepshead minnows should be located in a room or space separated from
that in which toxicity test are to be conducted.
The salinity of the culture systems should between 20%o and 30%o. Water temperature for the brood stock
should be maintained at 24°C - 26°C. The holding and rearing tanks should be aerated so that the dis-
solved oxygen is not less than 4.0 ppm.
Replace approximately 10% of the culture water every 2 weeks, or 25% monthly. The culture water should
be clear. If the water appears cloudy of discolored, replace at least 50% of it. Replacement water should be
well oxygenated and at the same temperature and salinity as the existing culture water. Salinity is main-
tained at the proper level by adding deionized water to compensate for evaporation. Artificial seawater is
prepared by dissolving artificial sea salts in deionized water to a salinity of 20%o - 30%o (see Appendix A
for preparation of hypersaline brine solution [MSB]).
PHOTOPERIOD
The culture conditions should include a photoperiod of 16 hours light and 8 hours dark (EPA, 2002a). The
light quality and intensity should be at ambient laboratory levels, which is approximately 10 - 20 uE/m2/s
or 50 to 100 foot candles (ft-c) (EPA, 2002a).
CULTURE VESSELS
Holding tanks are kept at ambient laboratory temperature (25°C) until the fish reach sexual maturity (3-5
months post hatch) at which time they can be used for spawning. Mature sheepshead minnows have an
average length of approximately 27 mm for females and 34 mm for males. Once mature, males will begin
to exhibit sexual dimorphism and initiate territorial behavior. Once sexually mature, hold the adults in water
reduced to 18°C - 20°C.
To avoid excessive build up of algal growth, periodically scrape the walls of the culture system. Some of the
algae will serve as a supplement to the diet of the fish'. A partial activated carbon "charcoal" change in the
filtration systems should be done monthly or as needed. The detritus (dead brine shrimp nauplii and cysts,
adult brine shrimp, other organic material accumulation) should be siphoned from the bottom of rearing
and holding aquaria or tanks each week or as needed.
WATER DELIVERY SYSTEMS
Adult sheepshead minnows (>1 month) are kept in natural or artificial seawater in a flow-through or recircu-
lating aerated glass aquarium that is equipped with an undergravel or outside biological filter, or cartridge
filter. Static systems are equipped with an undergravel filter. Recirculating systems are equipped with an
outside biological filter constructed in the laboratory using a reservoir system of crushed coral, crushed
oyster shells or dolomite and gravel, charcoal, floss, or a commercially available cartridge filter or an
equivalent system.
FOOD PREPARATION
The adult sheepshead minnows are fed flake food three to four times daily, supplemented with frozen adult
brine shrimp.
The larvae are fed newly hatched Artemia nauplii and crushed flake food, ad libitum, daily. The Artemia
should be cultured in the laboratory in order to provide 24 - 48 hour old nauplii. Appendix B describes in
detail how to culture Artemia.
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Sheepshead Minnow (Cyprinodon variegatus) and Inland Silverside (Men/did beryffina)
Larval Survival and Growth Toxicity Test • Supplement to Training Video
Figure 1. Embryonic development of Sheepshead minnow, Cyprinodon
variegatus: A. Mature unfertilized egg, showing attachment filaments
and micropyle, X33; B. Blastodisc fully developed; C/D. Blastodisc, 8
cells; E. Blastoderm, 16 cells; F. Blastoderm, late cleavage stage; G.
Blastoderm with germ ring formed, embryonic shield developing; H.
Blastoderm covers over 3A of yolk, yolk noticeably constricted; I. Early
embryo. (Continued, J - 0 on page 4).
OBTAINING LARVAE FOR
TOXICITY TESTS
For the Sheepshead larval
survival and growth toxicity
test, larvae that are less than
24 hours old are needed at the
start of the test. To have the
appropriate age larvae at the
start of a test, induce the min-
nows to spawn by raising the
To keep the egg collecting
screens clean, feed the
spawning fish while the
collecting screen is removed
for egg collection.
system temperature to 25°C
approximately 1 week before
the start of the test. This
gradual temperature increase
is started in the morning. By
afternoon, transfer the adults
(at least five females and three
males) to a spawning chamber,
or basket made from 3-5
mm NITEX® screen, within an
aquarium outfitted with a mesh
screen (150 - 250 urn mesh)
under the basket or on the
bottom. The fish will begin to
spawn within 24 hours and the eggs will fall through the basket onto the mesh collecting
G H
Source: Kuntz, 1916 in EPA, 2002a.
screen.
Collect eggs daily by washing the eggs off of the screen into a large tray. Roll the eggs gently on the screen
during collection, pressing any food or waste through, leaving the eggs on top of the screen. Embryos will
tend to stick together due to the presence of adhesive threads. After embryos have been manipulated,
wash them by placing them in a 250-um sieve and rinsing them with seawaterfrom a squeeze bottle. This
should reduce any fungal contamination of the embryos.
Females also can be induced to spawn artificially by intraperitioneal injection with human chorionic gonad-
otrophin (HCG) hormone. Natural spawning is preferable because repeated spawnings can be obtained
from the same brood stock. Additional details on forced spawning are provided in section 11.6.15 of the
Saltwater Chronic Methods Manual (EPA, 2002a).
The collected embryos should be checked under a dissecting microscope to identify any abnormal or unfer-
tilized eggs. The embryos should be in stages C - G as illustrated in Figure 1.
After collection, incubate the collected minnow embryos in seawater at 25°C, 20%o - 30%o salinity, and
16-hour light and 8-hour dark photoperiod for 5 - 6 days with aeration and daily water changes.
At 48 hours after collection, check the embryos under a dissecting microscope and discard any abnormal or
unfertilized eggs. At this time, the embryos should be at stages I or J as illustrated in Figure 1. To conduct one
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Sheepshead Minnow (Cyprinodon variegatus) and Inland Silverside (Menidia beryllina)
Larval Survival and Growth Toxicity Test • Supplement to Training Video
test with four replicates of
15 larvae and five efflu-
ent concentrations plus
a control, collect approxi-
mately 400 viable embryos
for incubation at this stage.
Reducing the salinity, raising
the temperature, or chang-
ing the water can help
induce hatching. If culture
dishes are used, they should
be covered to reduce evapo-
ration which could increase
salinity.
For the sheepshead minnow
growth and survival toxicity
test, use larvae that hatch
less than 24 hours before
the start of the test. If some
embryos hatch earlier than
24 hours prior to the test
start, remove them but keep
them to supplement the
younger larvae in case there
are not be enough larvae at
the start of the test. If this is
done, larvae should not be
more than 48 hours old and
should all be within 24 hours
of the same age. Selection
of the older larvae should be
randomized by placing them
back into the pool before
selection.
INLAND SILVERSIDE
Figure 1 (continued). Embryonic development of sheepshead
minnow, Cyprinodon variegatus: J. Embryo 48 h after fertilization, no
segmented throughout, pigment on yolk sac and body, otoliths formed;
K. Posterior portion of embryo free from yolk and moves freely within
egg membrane, 72 h after fertilization; L. Newly hatched fish, actual
length 4 mm; M. Larval fish 5 days after hatching, actual length 5 mm;
N. Young fish 9 mm in length; 0. Young fish 12 mm in length.
From Kuntz, 1916 in EPA, 2002a.
Inland silversides (Menidia beryllina) 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. Gravid females
can be found in low salinity waters along the Atlantic coast during April to July. If beach seines (3 mm - 6
mm mesh) are used, silversides should not be landed onto the beach as they are very sensitive to handling
and should not be removed from water by net — only by bucket or beaker. Several species of silversides
may be included in field caught specimens (e.g., M. beryllina, M. menidia, and M. peninsulas); care should
be taken to identify and separate the species.
If fish are collected from the field, record the temperature and salinity at each collection site so that the
conditions can be maintained in the culture tanks. After transfer to laboratory culture tanks, slowly intro-
duce laboratory water (maximum change of 2°C/day and 5%o salinity/day) to bring the water up to 25°C and
20%o - 32%o.
CULTURE WATER
Only natural seawater is recommended for the culture and maintenance of the more sensitive silverside
brood stock. Maintain holding and spawning tanks at a temperature of 25°C and a salinity of 20%o - 32%o.
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U.S. ENVIRONMENTAL PROTECTION AGENCY Sheepshead Minnow (Cyprinodon variegatus) and Inland Silverside (Menidia beryl/ina)
Larval Survival and Growth Toxicity Test • Supplement to Training Video
PHOTOPERIOD
The culture conditions should include a photoperiod of 16 hours light and 8 hours dark (EPA, 2002a). The
light quality and intensity should be at ambient laboratory levels, which is approximately 10 - 20 uE/m2/s
or 50 - 100 foot candles (ft-c).
CULTURE VESSELS
Adult inland silverside should be stocked in tanks of a minimum volume of 150L at a density of 50 fish/
tank. Detritus should be siphoned off from the bottom weekly, or as needed.
WATER DELIVERY SYSTEMS
Adult inland silversides are kept in a flow-through or recirculating aerated glass aquarium that is equipped
with an undergravel or outside biological filter, or cartridge filter. Static systems are equipped with an
undergravel filter. Recirculating systems are equipped with an outside biological filter constructed in the
laboratory using a reservoir system of crushed coral, crushed oyster shells or dolomite and gravel, char-
coal, floss, or a commercially available cartridge filter or an equivalent system.
FOOD PREPARATION
Feed silverside larvae the rotifer Brachionus plicatilis until 4-6 days post-hatch, and the smallestX\rtem/a
nauplii available (<12 hour old) beginning on day 5. After day 7, feed the larvae with Artemia only and
increase the size to 12 - 24 hours old. Food preparation instructions are provided in Appendix B.
The adult inland silversides should be fed flake food or frozen brine shrimp twice daily and Artemia nauplii
once daily.
The larvae are fed newly hatched Artemia nauplii and crushed flake food, ad libitum, daily. The Artemia
should be cultured in the laboratory in order to provide 24 - 48 hour old nauplii. Appendix B describes in
detail how to culture Artemia.
OBTAINING LARVAE FOR TOXICITY TESTS
Inland silversides are sexually mature after 1-2 months. In the wild, eggs are adhered to submerged
vegetation. In the laboratory, silversides are encouraged to spawn by placing polyester aquarium filter fiber
in the tanks. The fiber (~ 15 cm x 10 cm x 10 cm) is suspended on a string 8 cm - 10 cm below the surface
of the water and in contact with the side of the tank. These should be placed into the tank 14 days prior to
the beginning of a test. Place the floss directly above an airstone to keep it aerated, and weigh it down to
keep it from floating on the surface.
When the fish spawn into the fiber, the hard, light yellow embryos (-0.75 mm in diameter) can be separated
from the fibers by hand, or the eggs and fiber can be placed together into a 10-gallon aquarium. The floss
should be suspended 8 cm - 10 cm below the surface of the water and should be stretched to keep the
embryos from being-crowded. Lightly aerate the tank and hold the temperature at 25°C.
Larvae will hatch in 6 - 7 days when incubated at 25°C and maintained in seawater ranging from 5%o -
30%0. The larvae will free themselves from the fibers at which time they are easily identified and should be
removed. The newly hatched larvae will range from 3.5 mm - 4.0 mm in total length. Figure 2 illustrates
the life stages of the inland silverside.
For the inland silverside larval survival and growth toxicity test, use 7- to 11-day-old larvae. For one test
using 15 larvae for each of four replicates and five test concentrations plus a control, approximately 400
larvae are needed.
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Sheepshead Minnow (Cyprinodon variegatus) and Inland Silverside (Menidia faery/lino)
Larval Survival and Growth Toxicity Test • Supplement to Training Video
Test Method
EFFLUENT SAMPLING
For both species, handle efflu-
ent and receiving water sam-
ples in the same manner. Store
effluent or receiving waters in
an incubator or refrigerator at
0°C - 6°C until the tests begin,
but not longer than 36 hours
if being used for compliance
fora NPDES permit. Prepare
dilutions of the effluent sample
using a 0.5 dilution factor (e.g.,
6.25%, 12.5%. 25%, 50%, and
100%). If a high rate of mortal-
ity is observed during the first
1-2 hours, additional repli-
cates in the lower ranges of
effluent concentration should
be added.
The tests require about 5-6
L of each effluent or receiv-
ing water sample each day,
enough for renewing four
replicates of each concentra-
tion plus the control and for
performing chemical analyses.
It is essential to maintain
constant salinity among
treatments and treatment
replicates throughout the
test. Use concentrated sea-
water or hypersaline brine
(MSB) to keep the salinity
of the solutions between
20%o and 30%o for the
sheepshead minnows, and
between 5%o and 30%o for
the inland silversides. Before
adding the solutions to the
test chambers, wartn the
samples to 25°C in a water
bath. Keep the temperature
constant (25°C + 1°C) for the
duration of the test.
DILUTION PREPARATION
Figure 2. Inland silverside, Menidia berylllna: A. Adult, ca. 64 mm SL;
B. Egg (diagrammatic), only bases of filaments shown; C. Egg, 2-cell
stage; D. Egg, morula stage; E. Advanced embryo, 2V2 days after
fertilization..
G Larva 8.9 mm TL
D E
From Martin and Drewry, (978 in EPA, 2002a.
Dilution Water
The type of dilution water used to make the test concentrations is
dependent on the objectives of the test. Any specific requirements
included in NPDES permits should be followed. The Saltwater Chronic
Methods Manual (Section 7) provides the following guidelines:
• If the test is conducted to estimate the absolute chronic toxicity
of the effluent, synthetic dilution water should be used. If the cultures
were maintained in different water than used for dilution water, a
second set of control replicates should be conducted using the culture
water.
• If the test is conducted to estimate the chronic toxicity of the
effluent in uncontaminated receiving waters, the test can be
conducted using a grab sample of the receiving waters collected outside
the influence of the outfall, other uncontaminated waters, or standard
dilution water with the same salinity as the receiving waters. If the
cultures were maintained in different water than used for dilution water,
a second set of control replicates should be conducted using the culture
water.
• If the test is conducted to estimate the additive or mitigating
effects of the effluent on already contaminated receiving
waters, the test must be conducted using receiving waters collected
outside the influence of the outfall. Controls should be conducted using
both receiving water and culture water.
Set out the test chambers.
Typically, there will be at least five dilutions plus one control, and a minimum of four replicates. For both
species NHEERL-AED uses glass chambers equipped with a screened-off sump area (see Figure 3). One
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Sheepshead Minnow (Cyprinodon voriegotus) and Inland Silverside (Menidia bery/Jina)
Larval Survival and Growth Toxicity Test • Supplement to Training Video
Figure 3. Glass test chamber with sump area. Modified from Norberg
and Mount (1985).
Glass
Reinforcements
Sump
Source: EPA, 2002a.
thousand ml glass or
disposable plastic beakers
also can 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 1cm.
Pipet two or three larvae at
a time into each chamber,
adding larvae to all cham-
bers; then start again, add-
ing more until each cham-
ber contains the required
number of larvae — a
minimum of 10. Use a mini-
mum 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 facili-
tates 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 them into chambers containing control solution 1 day
before the start of the exposure period.
Randomly apply colored labels to the chambers to indicate treatment and replicates. Fill each chamber
with approximately 750 ml of the appropriate test solution, pouring through the sump area or down the side.
Each test chamber should contain a minimum of 50 ml of test solution/larvae and a depth of at least 5 cm.
ROUTINE CHEMISTRIES
Measure the initial temperature, salinity, and dissolved oxygen (DO) in each chamber. Record
all measurements on the test data sheet. Copies of the data sheets used at NHEERL-AED are
provided in Appendix F.
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 posi-
tions for the duration of the test.
RENEWALS
Each day, the test and control solutions must be replenished. Prepare new dilutions daily from effluent
stored at 0° - 6°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 36 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°C + 1°C, and supply 16 hours of
ambient laboratory light and 8 hours of darkness each day for both species. Measure and record the salin-
ity from each chamber every day as well, before renewing the test solutions. Note that there should be no
more than a 2%o salinity difference between any two chambers on a given day. If receiving water and efflu-
ent tests are conducted concurrently, the effluent salinity should be adjusted to match the receiving water
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U.S. ENVIRONMENTAL PROTECTION AGENCY Sheepshead Minnow (Cyprinodon variegatus) and Inland Silverside (Men/did beryl/ino)
Larval Survival and Growth Toxicity Test • Supplement to Training Video
- -- - h .". - - -..V*! V3*4
sample if possible. Monitor DO concentrations each day and record the data on the data sheet. If DO falls
below 40% saturation in any one of the exposure chambers, all chambers must be aerated.
Before changing the test solutions, count and record the number of live larvae in each replicate, discard-
ing any dead animals. Then remove 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 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 respective 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 - 10 mm, slowly and carefully add approxi-
mately 500 - 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 their same randomized
positions in the water bath and feed the larvae.
FEEDING
Proven quality/Artem/a nauplii should be used to feed the larvae daily throughout the test. Two concentra-
tions of prepared nauplii are used sequentially during the exposure period. Detailed instructions for cultur-
ing Artemia are included in Appendix B. The first food solution used for day 0 - 2 consists of 4 mL concen-
trated Artemia nauplii in 80 mL seawater. Feed each replicate 2 mL of this solution on the first 2 days of
the test. The 2 mL volume should yield approximately 0.10 g wet weight of Artemia nauplii. Care should be
taken to swirl the solution to maintain a constant distribution of Artemia and each 2 mL portion should be
drawn individually to avoid differences in feeding rates due to the settling of Artemia in the dropper.
For days 3 - 6 of the test, feed the larvae 2 mL per replicate of a more concentrated solution of 6 mL of
concentrated Artemia in 80 mL of seawater. This 2 mL volume should yield approximately 0.15 g wet weight
Artemia nauplii. Uneaten Artemia should be siphoned out of the chambers each day so that the larvae eat
newly hatched Artemia and to avoid depletion of DO within the chamber. On day 7, the larvae are not fed.
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%, reduce the amount of food supplied to that chamber by Vb for the remainder
of the test. Cover the chambers between feedings to reduce evaporation.
TEST TERMINATION
At the end of the test, on day 7, the larvae are counted to determine survival rate. Working with groups of
replicates, remove any dead larvae from the chambers, carefully recording the number of surviving ani-
mals. Record the final temperature, salinity, and DO for each chamber.
Pour the contents of each chamber through a 500-um 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 determi-
nation. 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 all of the fish from each
replicate into a labeled, pre-weighed aluminum weighing boat, and dry the fish at 60°C for 24 hours, or
at 105°C for 6 hours. Gloves should be worn or forceps should be used to handle the aluminum weighing
boats because oil from skin could affect weight differences.
After drying, and until they are weighed, place the dried larvae directly into a desicatorto prevent moisture
from the air adsorbing to the samples. Weigh each sample to the nearest 0.01 mg. Because small differ-
ences in weight or appearance can easily change the test results, it is critical to record observations and
measurements clearly and accurately. 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 aver-
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U'S' ENVIRONMENTAL PROTECTION AGENCY Sheepshead Minnow (Cyprinodon variegatus) and Inland Silverside (Menidia beryllina)
Larval Survival and Growth Toxicity Test • Supplement to Training Video
age 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 fishes' growth.
TEST ACCEPTABILITY AND DATA REVIEW
Test data are reviewed to verify that EPA's WET test methods' test acceptability criteria (TAG) requirements
fora valid test have been met. For the test to be considered acceptable, control survival must be > 80% for
both species. The average dry weight of unpreserved control larvae must be > 0.60 mgforthesheepshead
minnow, and > 0.50 mgforthe inland silverside. Minimum dry weights for preserved animals are > 0.50 mg
for the sheepshead minnow and > 0.43 mg for the inland silverside.
The concentration-response relationship generated for each multi-concentration test must be reviewed to
ensure that calculated test results are interpreted appropriately. In conjunction with this requirement, EPA
has provided recommended guidance for concentration-response relationship review (EPA, 2000a).
EPA's promulgated toxicity testing method manuals (EPA, 2002a, b) recommend the use of point estima-
tion technique approaches for calculating endpoints for effluent toxicity tests under the NPDES program.
The promulgated methods also require a data review of toxicity data and concentration-response data, and
require calculating the percent minimum significant difference (PMSD) when point estimation (e.g., LC50,
IC25) analyses are not used. EPA specifies the PMSD must be calculated when NPDES permits require sub-
lethal hypothesis testing. EPA also requires that variability criteria be applied as a test review step when
NPDES permits require sub-lethal hypothesis testing endpoints (i.e., no observed effect concentration
[NOEC] or lowest observed effect concentration [LOEC]) and the effluent has been determined to have no
toxicity at the permitted receiving water concentration (EPA, 2002b). This reduces the within-test variabil-
ity and increases statistical sensitivity when test endpoints are expressed using hypothesis testing rather
than the preferred point estimation techniques.
OTHER PROCEDURAL CONSIDERATIONS
• 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.
These data should be carefully compiled and are 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, Third Edition (EPA, 2002a) 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 marine and estuarine environments.
Citations and Recommended References
Gripe, G.M., B.L. Hemmer, L.R. Goodman, and J.C.Vennari. 2009. Development of a Methodology for
Successful Multigeneration Life-Cycle Testing of the Estuarine Sheepshead Minnow, Cyprinodon
i variegatus. Archives of Environmental Contamination and Toxicology. April 56(3):500-8.
EPA. 1985. Aquatic Toxicity Testing Seminar Manual. National Health and Environmental Effects Research
Laboratory - Aquatic Ecology Division, Narragansett, Rl. NHEERL-AED Contribution No. 796.
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UiS' ENVIRONMENTAL PROTECTION AGENCY Sheepshead Minnow (Cyprinodon variegatus) and Inland Silverside (Menidio beryllina)
Larval Survival and Growth Toxicity Test • Supplement to Training Video
' <••'.. -'*•
EPA. 1987a. Guidance manual for rapid chronic toxicity tests on effluents and receiving waters with
larval inland silversides (Menidia beryllina). Contribution No. 792. Heber, M.A., M.M. Hughes,
S.C. Schimmel, and D.A. Bengston. In: Schimmel, S.C. ed., Users guide to the conduct and
interpretation of complex effluent toxicity tests at estuarine/marine sites. Environmental Research
Laboratory, U.S. EPA, Narragansett, Rl 02882. Contribution No. 796, 265 pp.
EPA. 1987b. Guidance manual for conducting complex effluent and receiving water larval fish growth-survival
studies with the sheepshead minnow (Cyprinodon variegatus). Contribution No. x!04. Hughes, M.M.,
M.A. Heber, S.C. Schimmel, and WJ. Berry. In: Schimmel, S.C. ed., Users guide to the conduct and
interpretation of complex effluent toxicity tests at estuarine/marine sites. Environmental Research
Laboratory, U.S. EPA, Narragansett, Rl 02882. Contribution No. 796, 265 pp.
EPA. 1989. Biomonitoring for Control of Toxicity in Effluent Discharges to the Marine Environment.
U.S. EPA Center for Environmental Research Information, Cincinnati, OH. U.S. EPA National Health
and Environmental Effects Research Laboratory - Aquatic Ecology Division, Narragansett, Rl.
EPA/625/8-89/015.
EPA. 1991. Technical Support Document for Water Quality-based Toxics Control. U.S. EPA Office of Water
Enforcement and Permits, Washington, D.C. EPA-505-2-90-001.
EPA. 2000a. Method Guidance and Recommendations for Whole Effluent Toxicity (WET) Testing (40 CFR
136). Office of Water, Washington, D.C. EPA 821-B-00-004.
EPA. 2000b. Understanding and Accounting for Method Variability in Whole Effluent Toxicity Applications
Under the National Pollutant Discharge Elimination System Program. Office of Wastewater
Management, Washington, D.C. EPA 833-R-00-003.
EPA. 2002a. Short-term Methods for Estimating the Chronic Toxicity of Effluents and Receiving
Waters to Marine and Estuarine Organisms, Third Edition (Saitwater Chronic Methods Manual).
Environmental Protection Agency, Washington, DC. EPA-821-R-02-014.
EPA. 2002b. Methods for Measuring the Acute Toxicity of Effluents and Receiving Waters to Freshwater
and Marine Organisms, Fifth Edition. (Acute Methods Manual). Office of Water, Cincinnati, OH.
EPA-821-R-02-012.
EPA. 2002c. Final Rule. 40 CFR Part 136. Guidelines Establishing Test Procedures for the Analysis of
Pollutants; Whole Effluent Toxicity Test Methods. 67 FR 69952-69972, November 19, 2002.
EPA. 2009. Sheepshead Minnow (Cyprinodon variegatus) and Inland Silverside (Menidia beryllina) Larval
Survival and Growth Toxicity Test. Supplement to Training Video. Whole Effluent Toxicity Training
Video-Saltwater Series. March 2009. EPA 833-C-09-001
Johns, D.M., WJ. Berry, and W. Walton. 1981. International Study on Artemia, XVI. Survival, growth, and
reproduction potential of the mysid, Mys/dops/s bah/a Molenock fed various geographical collec-
tions 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 nau-
plii from various geographical origins for the mysid, Mysidopsis bahia (Molenock). Mar. Ecol. Prog.
Ser. Vol. 15, pp. 307-309.
10
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U'S- ENVIRONMENTAL PROTECTION AGENCY Sheepshead Minnow (Cyprinodon vor/egatus) and Inland Silverside (Men/did beryllina)
Larval Survival and Growth Toxicity Test • Supplement to Training Video
Norberg, TJ. and DJ. Mount. 1985. A new fathead minnow (Pimephales promelas) Subchronic toxicity
test. Environ. Toxicol. Chem. 4(5):711-718.
Persoone, G., P. Sorgeloos, 0. Roels, and E. Jaspers eds. 1980. The brine shrimp Artemia. Vols. 1-3.
Proceedings of the International Symposium on the brine shrimp Artemia satina, Corpus Christi,
Texas. 1979. Universal Press, Wetteren, Belgium.
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 consump-
tion 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 10 com-
mercial sources of brine shrimp cysts and reevaluation of the "hatching efficiency" concept.
Aquaculture. Vol. 30, pp. 43-52.
EPA references are available online at www.epa.gov/npdes.
If you need additional copies of this document, you can download it at:
www.epa.gov/npdes/wqbasedpermitting.
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Glossary
Acute toxicity. An adverse effect measured in a short period of time (96 hours or less in toxicity tests).
The effect can be measured in lethality or any variety of effects.
Algae. Rotifers are fed the algae Tetraselmus suec/ca or Chlorella sp.
Artemia. The marine invertebrate (referred to as brine shrimp) used as the recommended food source for
culture and test species; Brazilian or Colombian strains are preferred because the supplies are
found to have low concentrations of chemical residues and nauplii are of suitably small size.
Average mean dry weight. All the fish exposed in a given test chamber (replicate) are weighed together.
Trie total dry weight is divided by the number of surviving fish in the replicate to obtain the
average mean dry weight.
Chronic toxicity. An adverse effect that occurs over a long exposure period. The effect can be lethality,
impaired growth, reduced reproduction, etc.
Crash. Sudden (overnight) death of cultured organisms in a tank.
Cyprinodon variegatus. The scientific name for the fish species, sheepshead minnow. The sheepshead
minnow is a short, deep-bodied, compressed fish. It has large scales and a dark marginal band
on its tail. It occurs in hypersaline lagoons and connecting channels, and is found on muddy
bottoms in turbid waters from North and South America: Massachusetts, USA to northeastern
Mexico; also West Indies; northern coast of South America, Bahamas, Antilles, Gulf of Mexico,
Yucatan and Venezuela. It is omnivorous, consuming organic detritus and algae, as well as
microcrustaceans, and dipteran larvae. Sheepshead minnows are very abundant and easily
reproduced in captivity.
Cyst. The life stage of unhatched Artemia.
Diluent water. Dilution water used to prepare the effluent concentrations.
Effluent sample. A representative collection of a NPDES permitted facility's discharge that is to be tested.
Effluent concentration. Different dilutions, or concentrations, of an effluent used to determine the
biological effects on test organisms (i.e., inland silversides or sheepshead minnows).
Flow-through water delivery system. An open water flow system that delivers fresh water or seawater to
culture tanks, which is disposed of after it leaves those tanks.
Hypothesis testing. Technique (e.g., Dunnett's test) that determines what concentration is statistically
different from the control. Endpoints determined from hypothesis testing are NOEC and LOEC.
\C25 (Inhibition Concentration, 25%). The point estimate of the toxicant concentration that would cause a
25% reduction in a non-quantal biological measurement (e.g., reproduction or growth) calculated
from a continuous model.
Larvae. Post-hatch fish that are not free-swimming and are morphologically immature (i.e., <24 hr-old).
LC50 (Lethal Concentration, 50%). The concentration of toxicant or effluent that would cause death to
50% of the test organisms at a specific time of observations (e.g., 96-hour LC50).
Lowest Observed Effect Concentration (LOEC). The LOEC is the lowest concentration of toxicant to
which organisms are exposed in a test, which causes statistically significant adverse effects on
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U.S. ENVIRONMENTAL PROTECTION AGENCY Sheepshead Minnow (Cvprinodon variegatus) and Inland Silverside (Menidia beryllina)
Larval Survival and Growth Toxicity Test • Supplement to Training Video
the test organisms (i.e., where the values for the observed endpoints are statistically significantly
different from the control). The definitions of NOEC and LOEC assume a strict dose-response
relationship between toxicant concentration and organism response.
Minimum Significant Difference (MSD). The MSD is the magnitude of difference from the control where
the null hypothesis is rejected in a statistical test comparing a treatment with a control. MSD
is based on the number of replicates, control performance and power of the test. MSD is often
measured as a percent and referred to as PMSD.
Menidia beryllina. The scientific name for the fish species, inland silverside. It is a marine/estuarine
species that ascends rivers. In fresh water, inland silverside usually occurs at the surface of clear,
quiet water over sand or gravel. It feeds on zooplankton and is found in coastal waters from the
Western Atlantic: Massachusetts to southern Florida in the USA and around the Gulf of Mexico to
northeastern Mexico.
Nauplii. Free-swimming microscopic larvae stage characteristic of copepods, ostracods, barnacles, etc.
typically only with three pairs of appendages.
No Observed Effect Concentration (NOEC). The NOEC is the highest tested concentration of toxicant to
which organisms are exposed in a full life-cycle or partial life-cycle (short-term) test, that causes
no observable adverse effect on the test organism (i.e., the highest concentration of toxicant
at which the values for the observed responses are not statistically significantly different from
the controls). NOECs calculated by hypothesis testing are dependent upon the concentrations
selected.
NPDES (National Pollutant Discharge Elimination System) Program. The national program for issuing,
modifying, revoking and reissuing, terminating, monitoring, and enforcing permits, and imposing
and enforcing pretreatment requirements under Sections 307, 3±8, 402, and 405 of the Clean
Water Act.
Point Estimation Techniques. This technique is used to determine the effluent concentration at which
adverse effects (e.g., fertilization, growth or survival) occurred, such as Probit, Interpolation
Method, Spearman-Karber. For example, a concentration at which a 25% reduction in
reproduction and survival occurred.
Receiving Water Concentration (RWC). The RWC is the concentration of a toxicant or the parameter
toxicity in the receiving water (i.e., riverine, lake, reservoir, estuary or ocean) after mixing.
Recirculating water delivery system. A water flow system that treats water after it passes through the culture
tanks (usually with sand and biofilters) and delivers the same treated water back to the tanks.
Rotifer. The rotifer, Brachionus plicatilis is fed to newly-hatched inland silverside larvae until they are large
enough to be fed Artemia.
Static renewal. The daily replacement of effluent medium in the test chamber.
Static water system. An enclosed system contained within one culture tank. The water is filtered through
an underground or charcoal filter and is delivered back to the same tank.
Toxicity test. A procedure to measure the toxicity of a chemical or effluent using living organisms. The test
measures the degree of response of an exposed organism to a specific chemical or effluent..
WET (Whole effluent toxicity). The total toxic effect of an effluent measured directly with a toxicity test.
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=/ Larval Survival and Growth Toxicity Test • Supplement to Training Video
Appendix A:
Preparing Hypersaline Brine (MSB)
Salinity adjustments are a vital part of using marine and estuarine species for toxicity testing. Because the
majority of industrial and sewage treatment effluents entering marine and estuarine waters contain little or
no measurable salts, the salinity of these effluents must be adjusted before exposing estuarine or marine
plants and animals to the test solutions. It also is important to maintain constant salinity across all treat-
ments throughout the test for quality control. Finally, matching the test solution's salinity to the expected
receiving water's salinity may require salinity adjustments. NHEERL-AED uses MSB, prepared from filtered
natural seawater, to adjust exposure solution salinities.
MSB has se'veral 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. MSB can be held for prolonged periods without any apparent degradation, added directly to
the effluent to increase the salinity, or used as control water by diluting to the desired salinity with deion-
ized 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.
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, fol-
lowed by several thorough deionized-water rinses.
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 urn before placing into the brine tank. 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 could contaminate the brine. A
thermostatically controlled heat exchanger made from fiberglass is suggested.
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 (or more or
less often, depending on the volume being generated) to ensure that the salinity does not exceed 100%o
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 NHEERL-AED was a reduction
in the alkalinity of seawater made from brine with salinity greater than 100%o, 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 1-mm filter and pump
or pour it directly into portable containers (20-L cubitainers or polycarbonate water cooler jugs are most
suitable). Cap the containers, and record the measured salinity and the date generated. Store the brine in
the dark at room temperature.
SALINITY ADJUSTMENTS USING HYPERSALINE BRINE
To calculate the volume of brine (Vb) to add to a 0%o sample to produce a solution at a desired salinity (Sf),
use this equation:
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U.S. ENVIRONMENTAL PROTECTION AGENCY
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Larval Survival and Growth Toxicity Test • Supplement to Training Video
vb * sb = sf * vf
Where: Vb = volume of brine, mL
Sb = salinity of brine, %o
Sf = final salinity, %o
Vf = final volume needed, mL
Table A-l gives volumes needed to make 20%o test solutions from effluent (0%o), deionized water, and
100%o HSB. Quantities of effluent, deionized water and a MSB of 100%o (only) needed for conducting daily
renewals of test solutions at 20%o salinity. The highest concentration achievable is 80% effluent at 20%o
salinity and 70% effluent at 30%o. ,
Table A-l. Preparation of Test Solutions at a Salinity of 20%> Using HSB for a Final Test Concentration
Volume of 4000 mL.
Exposure Concentration
80
40
20
10
5
Control
Effluent
(0 %o mL)
3200
1600
800
400
200
—
Deionized
Water (mL)
—
1,600
2,400
2,800
3,000
4,000
HSB
(100%.) (mL)
800
800
800
800
800
0
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ussX Larval Survival and Growth Toxicity Test • Supplement to Training Video
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. However, just after hatching, M. beryllina are too small to ingest Artemia, and
must be fed rotifers (6. plicatilis). Preparation and culture of Artemia and rotifers are described below.
CULTURING ARTEMIA
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 seawa-
ter; 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 offish growth and survival differed when fed strains of brine shrimp from various geographic loca-
tions (Klein-MacPhee, et. al., 1982; Johns et al., 1981; Leger and Sorgeloos, 1984). Therefore, reference
brine shrimp have 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 - 48 hours at room temperature. Different geographi-
cal 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.
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 - 30°C seawa-
ter to ensure adequate hatching. Add 10 cc brine shrimp cysts per liter, and aerate for at least 24
hours at 25°C. (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 - 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 aer-
ation 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 - 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 urn), 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 nau-
plii from the quantity of Artemia on the screen. On days 3-6, 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. For days 0-2, the final suspension yields 0.10 g wet weight of Artemia
nauplii whereas for days 3-6, the final suspension yields 0.15 g wet weight of Artemia nauplii.
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Larval Survival and Growth Toxicity Test • Supplement to Training Video
Aerate or swirl the Anemia 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 is critical to minimize the variability of larval weight, which is
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 ¥2.
Some live Artemia should remain overnight in test chambers. However, excessive Artemia can decrease DO
concentrations to below the acceptable limit. Siphon the uneaten Artemia from each chamber prior to test
solution renewal to ensure that the fish larvae mainly eat 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 - 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
[EPA, 2002a] for acceptability parameters).
PREPARING ROTIFER CULTURES (BRACHIONUS PLICATILIS)
Newly hatched Menidia beryllina larvae are too small to ingest Artemia and must be fed rotifers
(Brachionus plicatilis). B. plicatilis can be cultured continuously in the laboratory when fed algae or yeast in
10- to 15-L Pyrex carboys at 25°C - 28°C, 25%o - 35%o salinity. Four 12-L 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 placing an immersion
heater in the carboy, and maintain the temperature to 70°C - 80°C for 1 hour.) When the seawater has
cooled to 25°C - 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 urn mesh screen. Return any rotifers collected on the
screen to the culture.
Keep the carboys away from light to 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 - 400 rotifers/mL, will be reached in about 7-10 days and
should then be cropped daily. This density is sustainable for 2 - 3 weeks. Once that is attained, the rotifers
should be cropped daily.
These rotifers are fed to M. beryllina larvae after hatching until about 5 days old. About 5 days after hatch-
ing, the larvae can begin feeding on newly hatched Artemia nauplii. They are fed Artemia daily throughout
the 7-day test.
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Sheepshead Minnow (Cyprinodon variegotus) and Inland Silverside (Menidia beryllina)
Larval Survival and Growth Toxicity Test • Supplement to Training Video
ALGAL CULTURES
Algae for feeding the rotifers, Tetraselmus suecica or Chlorella sp., can be cultured in 20-L plastic water
bottles. Autoclave the bottles (at 110° for 30 minutes) after adding filtered seawater. Cool the bottle to
room temperature and place them in a temperature controlled chamber at 18°C - 20°C. Each bottle or
carboy should contain 1 L of T. suecica or Chlorella sp. starter culture and 100 mL of nutrients.
The nutrient formula for the algal culture is:
Mix into 12-L deionized water:
Mix on a magnetic stirrer at least 1 hour or until all salts are
dissolved.
Add and stir again:
!80gNaNO3
!2gNaH2PO4
6.l6gEDTA
3.78gFeCI3-6H2O
(Solution should be bright yellow)
Aerate the algal culture vigorously by inserting a pipette through a foam stopper at the top of the bottle or carboy. A
dense algal culture will develop in 7 - 10 days and should be used by day 14. For continuous supply of algal cultures for
rotifer feeding, new cultures should be started every 1 or 2 days. For four 12-L rotifer cultures, 6-8 continuous algal
cultures are needed.
Clean bottles or carboys thoroughly with soap and water, rinsing with deionized water between uses.
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Intentionally Left Blank
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U.S. ENVIRONMENTAL PROTECTION AGENCY Sheepshead Minnow (C/prinodon variegatus) and Inland Silverside (Menidia beryllina)
Larval Survival and Growth Toxicity Test • Supplement to Training Video
Appendix C:
Apparatus and Equipment - Sheepshead Minnow and
Inland Silverside Tests
Air lines and air stones. For aerating water containing embryos or larvae, or for supplying air to test
solutions with low DO.
Air pump. For oil-free air supply.
Balance. Analytical, capable of accurately weighing to 0.00001 g.
Beakers, six Class A. Borosilicate glass or non-toxic plasticware, 1000 ml for making test solutions,
Brine shrimp, Artem/a, culture unit.
Crystallization dishes, beakers, culture dishes (1L), or equivalent. For incubating embryos.
Desiccator. For holding dried larvae.
Dissecting microscope. For checking embryo viability (Sheepshead test only).
Droppers, and glass tubing with fire polished edges, 4 mm ID. For transferring larvae.
Drying oven. 50-105°C range, for drying larvae.
Environmental chamber or equivalent facility with temperature control (25 ± 1° C).
Facilities for holding and acclimating test organisms.
Forceps. For transferring dead larvae to weighing boats.
Inland Silverside culture unit. The test requires approximately 400, 7-11 day old larvae. It is preferable
to obtain the test organisms from an in-house culture unit. If it is not feasible to culture fish
in-house, embryos or larvae can be obtained from other sources by shipping them in well
oxygenated saline water in insulated containers.
Light box. For counting and observing larvae.
Meters: pH and DO. For routine physical and chemical measurements.
NITEX® or stainless steel mesh sieves (< 150 um, 500 um, 3-5 mm). For collecting Artem/a naupili
and fish embryos, and for spawning baskets, respectively.
Pipet bulbs and filters. PROPIPET®, or equivalent.
Pipets, automatic. Adjustable, 1 - 100 mL
Pipets, volumetric. Class A, 1 - 100 ml.
Pipets, serological. 1-10 ml, graduated.
Reference weights, Class S. For checking performance of balance. Weights should bracket the expected
weights of the weighing paris and the expected weights of the pans plus fish.
Refractometer. For determining salinity.
C-l
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U.S. ENVIRONMENTAL PROTECTION AGENCY
Sheepshead Minnow (Cyprinodon variegatus) and Inland Silverside (Men/did bery/l/no)
Larval Survival and Growth Toxicity Test • Supplement to Training Video
Samplers. Automatic sampler, preferably with sample cooling capability, that can collect a 24-hour
composite sample of 5 L.
Separatory funnels, 2 L. Two to four for culturing/Artem/a naupili.
Sheepshead minnow culture unit. The maximum number of larvae required per test will range from a
maximum of 360, if 15 larvae are used in each of four replicates, to a minimum of 240 per test,
if 10 larvae are used in each of four replicates. It is preferable to obtain the test organisms from
an in-house culture unit. If it is not feasible to culture fish in-house, embryos or newly hatched
larvae can be obtained from other sources if shipped in well oxygenated saline water in insulated
containers.
Siphon with bulb and clamp. For cleaning test chambers.
Standard or micro-Winkler apparatus. For determining DO (optional).
Test chambers.
Sheepshead. Four chambers are required for each concentration and the control. Borosilicate
glass 1000 mL beakers or modified Norberg and Mount (1985) glass chambers used in the
short-term inland silverside test may be used. It is recommended that each chamber contain a
minimum of 50 mL/larvae and allow adequate depth of test solution (5.0 cm). To avoid potential
contamination from the air and excessive evaporation of test solutions during the test, the
chambers should be covered with safety glass plates or sheet plastic (6 mm thick).
Inland Silverside. Four chambers are required for each concentration and the control. The
chambers should be borosilicate glass or nontoxic disposable plastic labware. To avoid potential
contamination from the air and excessive evaporation of test solutions during the test, the
chambers should be covered with safety glass plates or sheet plastic (6 mm thick).
Each test chamber for the inland silverside should contain a minimum of 750 mL of test solution.
A chamber such as the one in Figure C-l constructed of glass and silicone cement has been
used successfully for this test. This chamber holds an adequate column of test solution and
incorporates a sump area where test solutions can be siphoned and renewed without disturbing
the fragile inland silverside larvae.
When constructing
the chamber it
is recommended
that the screen be
a 200-um Nitex®
screen (rather than
stainless steel) and
thin pieces of glass
rods be silicone
cemented to the
screen to reinforce
the bottom and
sides of the screen
to create the sump
area. A minimum
of silicone should
be used while sill
ensuring that the
Figure C-l. Glass test chamber with sump area. Modified from
Norberg and Mount (1985).
Glass
Reinforcements
Sump
Source: EPA, 2002a.
C-2
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U S ENVIRONMENTAL PROTECTION AGENCY Sheepshead Minnow (Cyprinodon variegatus) and Inland Silverside (Menidia beryl/ina)
Larval Survival and Growth Toxicity Test • Supplement to Training Video
larvae cannot get trapped or drawn into the sump area. All new chambers should be soaked
overnight in seawater (preferably in flowing seawater) to cure the silicone cement before use.
Other types of glass chambers can be used such as 1000 ml beakers. However, each chamber
should contain a minimum of 50 ml of test or control solution per larvae and allow adequate
depth of test solution (5.0 cm).
Thermometers. National Bureau of Standards Certified (see EPA 2002a). Used to calibrate laboratory
thermometers.
Thermometers, bulb-thermograph or electronic-chart-type. For continuously recording temperature.
Thermometers, glass or electronic, laboratory grade. For measuring water temperatures.
Volumetric flasks and graduated cylinders. Class A, borosilicate glass or non-toxic plastic labware,
10 - 1000 ml for making test solutions.
Wash bottles. For deionized water, for washing embryos from substrates and containers, and for rinsing
small glassware and instrument electrodes and probes.
Water purification system. Millipore® Milli-Q®, deionized water (Dl) or equivalent.
C-3
-------
U.S. ENVIRONMENTAL PROTECTION AGENCY Sheepshead Minnow (Cypr/nodon vor/egatus) and Inland Silverside (Menidia beryl/mo)
Larval Survival and Growth Toxicity Test • Supplement to Training Video
Intentionally Left Blank
C-4
-------
U.S. ENVIRONMENTAL PROTECTION AGENCY Sheepshead Minnow (Cypr/nodon rariegatus) and Inland Silverside (Menldia beryllina)
? Larval Survival and Growth Toxicity Test • Supplement to Training Video
Appendix D:
Reagents and Consumable Materials
Buffers, pH 4, pH 7, and pH 10. (Or as per instructions of instrument manufacturer). For standards and
calibration check (see EPA 2002a).
Data sheets (one set per test). For data recording.
Ethanol (70%) or formalin (4%). For use as a preservative for the fish larvae.
Laboratory quality control samples and standards. For calibration of the above methods.
Markers, waterproof. For marking containers, etc.
Membranes and filling solutions for DO probe, or reagents. For modified Winkler analysis (see
EPA 2002a).
Sample containers. For sample shipment and storage.
Tape, colored. For labeling test chambers.
Vials, marked. Twenty-four per test, containing 4% formalin or 70% ethanol, to preserve larvae (optional).
Reference toxicant solutions. Reference toxicants such as sodium chloride (NaCI), potassium chloride
(KCI), cadmium chloride (CdCI2), copper sulfate (CuS04), sodium dodecyl sulfate (SDS), and
potassium dichromate (K2Cr207) are suitable for use in the NPDES Program and other Agency
programs requiring aquatic toxicity tests.
Reagent water. Defined as distilled or deionized water that does not contain substances which are toxic
to the test organisms.
Weighing pans, aluminum. Twenty-four per test (one for each replicate.)
D-l
-------
U.S. ENVIRONMENTAL PROTECTION AGENCY Sheepshead Minnow (Cyprinodon voriegotus) and Inland Silverside (Menidia beryl/ma)
Larval Survival and Growth Toxicity Test • Supplement to Training Video
. ' ' • ' •' 'Vrf.i.'JPS'io
Intentionally Left Blank
D-2
-------
U.S. ENVIRONMENTAL PROTECTION AGENCY Sheepshead Minnow (Cyprinodon voriegotus) and Inland Silverside (Menidio beryllina)
Larval Survival and Growth Toxicity Test • Supplement to Training Video
1?A-!5 •,"-,-, -.- • . "
Appendix E:
Summary of Test Conditions and Test Acceptability
Criteria
Table E-l. Summary of Test Conditions and Test Acceptability Criteria for the Sheepshead Minnow,
Cyprinodon variegatus, Larval Survival and Growth Test with Effluents and Receiving Waters
(Test Method 1004.0)
Test type
Salinity
Temperature (C°)
Light quality
Light intensity
Photoperiod
Test chamber size
Test solution volume
Renewal
Age of test organisms
Number of larvae per test chamber
Number of replicate chambers per
concentration
Number of larvae per concentration
Source of food
Feeding regime
Cleaning
Aeration
Dilution water
Test concentrations
Dilution factor
Test duration
Endpoints
Test acceptability criteria
Sampling requirement
Sample volume required
Static, with 24-hr renewal (required)
20%o - 32%o (maintained at ± 2%o of the selected test salinity) (recommended)
25°C ± I°C (recommended). Test temperatures must not vary by more than
3°C during the test (required)
Ambient laboratory (covered, soft white) light (recommended)
10-20 uE/m2/s (ambient laboratory: 50 - 100 ft-c) (recommended)
16 hr light/8 hr dark (recommended)
600 mL - 1 L containers (recommended)
500 - 750 mL/replicate (loading and DO restrictions must be met)
(recommended)
Daily (required)
Newly hatched larvae (less than 24-hr old; within 24-hr age of each other)
(required)
1 0 (required minimum)
4 (required minimum)
40 (required minimum)
Newly hatched Artemio nauplii (less than 24-hr old) (required)
Feed once per day 0. 10 g wet weight Anemia nauplii per replicate on days
0-2; feed 0.15 g wet weight Artemio nauplii per replicate on days 3-6 (recom-
mended)
Siphon daily, immediately before test solution renewal and feeding (required)
None, unless DO concentration falls below 4.0 mg/L, then aerate all cham-
bers. Rate should be less than 100 bubbles/min. (recommended)
Uncontaminated source of natural seawater, artificial seawater, deionized
water mixed with MSB or artificial sea salts (available options)
Effluent: Five and a control (required). Receiving waters: 100% receiving water
(or minimum of five) and a control (recommended)
Effluents: 50.5 (recommended)
Receiving waters: None, or 50.5 (recommended)
7 days (required)
Survival and growth (weight) (required)
80% or greater survival in controls, average dry weight per surviving organism
in control chambers must be 0.60 mg, if unpreserved or 0.50 mg or greater
average dry weight per surviving control larvae after not more than 7 days in
4% formalin or 70% ethanol (required)
For on-site tests, samples collected daily and used within 24 hr of the time
they are removed from the sampling device.
For off-site tests, a minimum of three samples (e.g., collected on days 1, 3, and
5) with a maximum holding time of 36 hr before first use. (required)
6 L per day (recommended)
Source: EPA, 2002a, Saltwater Chronic Methods Manual.
E-l
-------
U.S. ENVIRONMENTAL PROTECTION AGENCY
Sheepshead Minnow (Cyprinodon variegatus) and Inland Silverside (Menidia bery//ina)
Larval Survival and Growth Toxicity Test • Supplement to Training Video
Table E-2. Summary of Test Conditions and Test Acceptability Criteria for the Inland
Silverside, Menidia beryllina, Larval Survival and Growth Test with Effluents and Receiving Waters
(Test Method 1006.0)
Test type
Salinity •
Temperature (C°)
Light quality
Light intensity
Photoperiod
Test chamber size
Test solution volume
Renewal
Age of test organisms
Number of larvae per test chamber
Number of replicate chambers per
concentration
Number of larvae per concentration
Source of food
Feeding regime
Cleaning
Aeration
Dilution water
Test concentrations
Dilution factor
Test duration
Endpoints
Test acceptability criteria
Sampling requirement
^
Sample volume required
Static, with 24-hr renewal (required)
5%o - 32%o (maintained at ± 2%o of the selected test salinity) (recommended)
25°C ± I°C (recommended). Test temperatures must not vary by more than
3°C during the test (required)
Ambient laboratory (covered, soft white) light (recommended)
10-20 pE/m2/s (ambient laboratory: 50 - 100 ft-c) (recommended)
16 hr light/8 hr dark (recommended)
600 mL - 1 L containers (recommended)
500 - 750 mL/replicate (loading and DO restrictions must be met)
(recommended)
Daily (required)
7-11 days post-hatch; within 24-hr age of each other (required)
1 0 (required minimum)
4 (required minimum)
40 (required minimum)
Newly hatched Artemio nauplii; survival of 7-9 day old M. beryllina larvae
improved by feeding 24-hr old Anemia (required)
Feed 0.10 g wet weight Artemia nauplii per replicate on days 0—2; feed 0.15 g
wet weight Artemia nauplii per replicate on days 3-6 (recommended)
Siphon daily, immediately before test solution renewal and feeding (required)
None, unless DO concentration falls below 4.0 mg/L, then aerate all cham-
bers. Rate should be less than 100 bubbles/min. (recommended)
Uncontaminated source of natural seawater, artificial seawater, deionized
water mixed with MSB or artificial sea salts (available options)
Effluent: Five and a control (required). Receiving waters: 100% receiving water
(or minimum of five) and a control (recommended)
Effluents: £ 0.5 (recommended).
Receiving waters: None, or ^ 0.5 (recommended)
7 days (required)
Survival and growth (weight) (required)
80% or greater survival in controls, 0.50 mg average dry weight of control
larvae where test starts with 7-day old larvae and dried immediately after
test termination, o_r 0.43 mg or greater average dry weight per surviving
control larvae, preserved not more than 7 days in 4% formalin or 70% ethanol
(required)
For on-site tests, samples collected daily and used within 24 hr of the time
they are removed from the sampling device. For off-site tests, a minimum of
three samples (e.g., collected on days 1, 3, and 5) with a maximum holding
time of 36 hr before first use. (required)
6 L per day (recommended)
Source: EPA, 2002a. Saltwater Chronic Methods Manual.
E-2
-------
U.S. ENVIRONMENTAL PROTECTION AGENCY
Sheepshead Minnow (Cyprinodon variegotus) and Inland Silverside (Menidia beryllina)
Larval Survival and Growth Toxicity Test • Supplement to Training Video
Appendix F: Data Sheets
Figure F-l. Data Form for the Sheepshead Minnow and Inland Silverside, Larval Survival and Growth
Toxicity Test. Daily Record of Larval Survival and Test Conditions.
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-------
fjirt U.S. ENVIRONMENTAL PROTECTION AGENCY Sheepshead Minnow (Cyprinodon variegatus) and Inland Silverside (Menidia beryllina)
^u^ Larval Survival and Growth Toxicity Test • Supplement to Training Video
Figure F-l (continued). Data Form for the Sheepshead Minnow and Inland Silverside, Larval Survival and
Growth Toxicity Test. Daily Record of Larval Survival and Test Conditions..
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F-2
-------
U.S. ENVIRONMENTAL PROTECTION AGENCY Sheepshead Minnow (Cyprinodon vor/egotus) and Inland Silverside (Men/dm beryl/ino)
Larval Survival and Growth Toxicity Test • Supplement to Training Video
Figure F-2. Data Form for the Sheepshead Minnow and Inland Silverside, Larval Survival and Growth
Toxicity Test. Summary of Test Results
Test Dates:
. Species:
Effluent Tested:
Treatment
No. Live
Larvae
Survival
(%)
Mean Dry
Wt/Larvae
(mg) ± SD
Signif. Diff.
from Control
(o)
Mean Temp.
(°C) ± SD
Mean Salinity
%0±SD
Ave. DO
(mg/L) ± SD
Comments:
Source: EPA, 1987a.
F-3
-------
U.S. ENVIRONMENTAL PROTECTION AGENCY Sheepshead Minnow (Cyprinodon voriegotus) and Inland Silverside (Men/din beryl/ino)
Larval Survival and Growth Toxicity Test • Supplement to Training Video
Figure F-3. Data Form for the Sheepshead Minnow and Inland Silverside, Larval Survival and Growth
Toxicity Test. Dry Weights of Larvae.
Test Dates:
Species:.
Pan.
No.
Cone. &
Rep.
Initial Wt.
Final Wt.
(mg)
Diff.
No.
Larvae
Avg. Wt./
Larvae (mg)
Source: EPA, 1987b.
F-4
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If you need additional copies of this document, you can download it at:
www.epa.gov/npdes/wqbasedpermitting
-------
-------
If you need additional copies of this document,
you can download it at: http://cfpub.epa.gov/npdes/wqbasedpermitting/wet.cfm
------- |