v>EPA
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Environmental Research
Laboratory
Duluth MN 55804
EPA-600/8-81-011
May 1982
Reseaecr. <»«.l Development
User's Guide for
Conducting Life-Cycle
Chronic Toxicity Tests
with Fathead Minnows
(Pimephales promelas]
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EPA-600/8-81-011
July 1981
USER'S GUIDE FOR CONDUCTING LIFE-CYCLE CHRONIC TOXICITY TESTS WITH
FATHEAD MINNOWS (PIMEPHALES PROMELAS)
by
Duane A. Benoit
Environmental Research Laboratory-Duluth
Duluth, Minnesota 55804
ENVIRONMENTAL RESEARCH LABORATORY-DULUTH
OFFICE OF RESEARCH AND DEVELOPMENT
U.S. ENVIRONMENTAL PROTECTION AGENCY
DULUTH, MINNESOTA 55804
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DISCLAIMER
This report has been reviewed by the Environmental Research Laboratory-
Duluth, U.S. Environmental Protection Agency, and approved for publication.
Mention of trade names or commercial products does not constitute endorsement
or recommendation for use.
ii
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FOREWORD
The original chronic bioassay procedures for fathead minnows were compiled in
1971 by John Eaton for the Committee on Aquatic Bioassays at the U.S.
Environmental Research Laboratory-Duluth. These methods were then published
in Standard Methods for the Examination of Water and Wastewater (1).
The current revised procedures for 1981 represent an updated and reorganized
version of the old methods. These new procedures are based on recent
evaluations of toxicity test results and methods used to conduct life-cycle
chronic tests and early life stage tests with fathead minnows.
iii
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ABSTRACT
This paper represents a revised procedural guide for conducting life-cycle
chronic toxicity tests with fathead minnow (Pimephales promelas). These new
procedures are based on recent evaluations of published toxicity tests and
methods used by aquatic toxicologists to conduct life-cycle chronic tests and
early life stage tests with fathead minnows. These published papers are
referenced in the appropriate place throughout the text of this report. If
more detailed information on test apparatus or specific biological and
chemical methods is desired, the reader is encouraged to study the reference
material.
All routine methods not covered in this procedure (e.g., physical and
chemical determinations, handling of fish) should be followed as described in
Standard Methods for the Examination of Water and Wastewater (1).
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CONTENTS
Foreword iii
Abstract iv
Tables vii
1. Physical System
1.1 Diluter 1
1.2 Toxicant mixing and flow splitting 1
1.3 Test tanks 1
1.3.1 Arrangement A 1
1.3.2 Arrangement B 2
1.4 Embryo incubation cups. 2
1.5 Spawning substrates ........... 2
1.5.1 Cement 2
1.5.2 Stainless steel 2
1.6 Diluent water ..... 3
1.7 Flow rate 3
1.8 Aeration 3
1.9 Test water temperature 3
1.10 Light 3
1.11 Photoperiod . 4
1.12 Cleaning 4
1.13 Disturbance 4
1.14 Construction materials 4
2. Chemical System
2.1 Preparing a stock solution 7
2.2 Measurement of toxicant concentration 7
2.3 Measurement of other variables 7
2.4 Residue analysis 8
2.5 Methods 8
3. Biological System
3.1 Source of test fish 9
3.2 Preliminary tests 9
3.3 Embryo exposure 10
3.4 Larval-juvenile exposure 10
3.5 Juvenile-adult exposure 11
3.6 Second generation embryo exposure 12
3.7 Second generation larval-juvenile exposure 12
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3.8 Additional exposures and special examinations .... 13
3.9 Biological data recorded. . , 13
3.9.1 Embryo exposure 13
3.9.2 Larval-juvenile exposure ... 13
3.9.3 Juvenile-adult exposure 14
3.10 Data analysis and evaluation 14
References 15
VI
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TABLES
Number^ Page
1-1 Test Photoperiod (Evansville, Indiana) for Fathead 5
Minnow Life-Cycle Chronic Toxicity Tests
vii
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CHAPTER 1
PHYSICAL SYSTEM
1.1 Diluter
Intermittent-flow proportional diluters (2,3) or continuous-flow serial
diluters (4) should be employed. The operation of the diluter should be
checked daily, either directly or through measurement of toxicant
concentrations. A minimum of five toxicant concentrations with a dilution
factor not greater than 0.50 and one control should be used for each test.
An automatically triggered emergency aeration and alarm system should be
installed to alert staff in case of diluter, temperature control or water
supply failure.
1.2 Toxicant Mixing and Flow Splitting
If a proportional diluter is used, a container to promote mixing of toxicant
and diluent water should be used between the diluter and test tanks for each
concentration (5). Separate flow splitter delivery tubes should run from
this container to each replicate larval and adult tank. If a continuous-flow
serial diluter is used, additional mixing containers are not needed but
separate flow splitter delivery tubes must run from the diluter to all test
tanks. Delivery tubes are allocated to tanks by stratified random
assignment. Flow splitting accuracy must be within 10% and should be checked
periodically to see that the intended amounts of test water are going to each
tank.
1.3 Test Tanks
All test tanks should be made of either glass or stainless steel with glass
ends. Two arrangements of test tanks are recommended.
1.3.1 Arrangement A
Under this arrangement, duplicate adult spawning tanks measuring 30.5 x 30.5
x 91.4 cm long are used. A 30.5 cm square portion at the upper end of each
tank is screened off for larval exposures. Each larval section is divided in
half so that there are two larval growth chambers for each adult spawning
tank (6). Larval chambers should be designed with glass bottoms and drains
that allow water to be drawn down to 3 cm when the chamber is lifted out of
the larval section for photographic growth measurements (see Section 3.4).
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Test water must be delivered separately to each adult tank and larval
section, with one-third of the water volume going to the latter. Test water
delivered to each larval section must also be split evenly to each growth
chamber. Test water depth in adult tanks and larval chambers should be a
minimum of 15 cm.
1.3.2 Arrangement B
Duplicate adult spawning tanks measuring 30.5 x 30.5 x 61 cm long and
separate larval tanks are used in this arrangement. Each larval tank should
be a minimum of 28,373 cubic cm and should be divided in half to form two
larval growth chambers for each adult spawning tank (7). Larval chambers are
designed and test water divided as previously described in Section 1.3.1.
Larval tanks can be conveniently located directly above spawning tanks
containing test solutions of the same concentrations so they can be drained
directly into the spawning tank.
1.4 Embryo Incubation Cups
Embryo incubation cups should be made from 120 ml glass jars (5.1 cm OD) with
the bottoms cut off and replaced with stainless steel or nylon screen (40
meshes per 2.54 cm). Cups should either be oscillated vertically (2.5-4.0
cm) in the test water by means of a rocker arm apparatus driven by a 2 rpm
motor (8) or placed in separate chambers with self-starting siphons. Both
methods will produce a frequent flow of test water around the embryos during
the 4-5 day incubation period. Cups should not be hung in tanks containing
juvenile or adult test fish, unless each cup is designed with an additional
double bottom stainless steel screen (>^ 10 mesh per 2.54 cm) to prevent fish
from sucking embryos through the 40 mesh screen bottom.
1.5 Spawning Substrates
Fathead minnows deposit their adhesive eggs on the underside of submerged
objects. The following types of laboratory spawning substrates for fathead
minnows have been tested and are recommended for use in culture units and
life-cycle chronic tests (9).
1.5.1 Cement
Cement drain tiles (7.6 to 10.2 cm ID) can be used for spawning substrates.
Tiles must be cut into 7 to 10 cm long sections and then halved lengthwise
and inverted to form a semicircular arch.
1.5.2 Stainless Steel
Stainless steel (#316, 16-18 gauge) cut and bent into the same shape and size
as described above with a thin layer of renewable quartz sand coated on the
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underside can also be used as a spawning substrate. Embryos adhering to the
above substrates are rolled off with a gentle circular motion of the finger
while pressing on them lightly (10).
1.6 Diluent Water
The water used should be from a well or spring, if at all possible, or
alternatively from a surface water source. Only as a last resort should
dechlorinated water from a municipal water supply be used. If there is any
chance the water supply could be contaminated with fish pathogens, the water
should be passed through an ultraviolet or similar sterilizer immediately
before it enters the test system (6).
1.7 Flow Rate
Flow rates to each adult tank or larval chamber should be equal to at least
6-10 tank or chamber water volumes per 24 hours. Flow rates must be great
enough so that dissolved oxygen does not drop below 75% of saturation (11) or
toxicant concentrations to drop by more than 20% when fish are in the test
tanks. Flow rates can be increased above those specified to maintain proper
dissolved oxygen or toxicant concentrations.
1.8 Aeration
Diluent water should be aerated vigorously (with oil-free air) or passed over
a screen column with a recirculating pump before flowing through the diluter.
Aeration of diluent water will eliminate supersaturation of dissolved gases
and also insure that dissolved oxygen concentrations will be at or near
saturation (90-100%). However, the test tanks and chambers themselves should
not be aerated.
1.9 Test Water Temperature
Test water temperature should not deviate from 25° C by more than 2° C and
should not remain outside the range of 24 to 26° C for more than 48 hours at
a time (12). Temperature should be recorded continuously.
1.10 Light
The lights used should simulate the wavelength spectra of sunlight as nearly
as possible. A combination of Durotest (Optima FS) fluorescent tubes
manufactured by Duro-Test Corporation, Secaucus, NJ 07094, and wide spectrum
Grow-Lux fluorescent tubes manufactured by Sylvania Lighting Center, Danvers,
MA 09123 has proven satisfactory (13). Light intensities at the water
surface during tests have ranged from 10-100 lumens.
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1.11 Photoperiod
In order to maintain standard periods of light, photoperiods (Table 1-1)
should simulate the dawn-to-dusk times of Evansville, Indiana (representing
average light conditions for the middle of the continental United States).
Adjustments in day-length are to be made on the first and fifteenth day of
every Evansville test month (14). The table is arranged so that adjustments
need be made only in the dusk times. Regardless of the actual date that the
experiment is started, the amount of light used should be the same as the
Evansville test photoperiod of December 1. To illustrate this point, an
experiment started with 24-hour-old embryos in Duluth, Minnesota, on August
28 (actual date), would require use of a December 1 Evansville test
photoperiod, and the lights could go on anytime on that day as long as they
remained on for 10 hours and 45 minutes. Fifteen days later (September 12
actual date, December 15 Evansville test date) the day-length would be
changed to 10 hours and 30 minutes. Gradual changes in light intensity at
dawn and dusk (15) may be included within the photoperiods if desired, but
should not last for more than 30 minutes from full-on to full-off and vice
versa.
1.12 Cleaning
All adult tanks and larval chambers (after larvae swim-up) must be siphoned
at least three times a week and brushed when algal or fungal growth becomes
noticeable. Siphoning should be done just before the last feeding of the
day. Incubation cup screen bottoms should also be brushed periodically if
they become clogged.
Siphoning can be done safely with either a large pipette (50 ml) fitted with
a squeeze bulb or a glass tube and siphon hose leading to a white pan. Fish
which are siphoned accidentally can be observed easily in the pipette or
white pan and returned carefully without harm to the chamber.
1.13 Disturbance
Adults and larvae should be shielded from disturbances such as people walking
past the tanks or extraneous lights that might alter the intended
photoperiod.
1.14 Construction Materials
Construction materials which contact the diluent water should not contain
leachable substances and should not absorb significant amounts of substances
from the water. Stainless steel is the preferred construction material.
Glass significantly absorbs some trace organics. Rubber must not be used.
Plastic containing fillers, additives, stabilizers, plasticizers, etc.,
must not be used. Teflon, nylon, and their equivalents are not known to
contain leachable materials, nor do they absorb significant amounts of test
substances. All batches of neoprene stoppers should be checked for toxicity
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TABLE 1-1
TEST PHOTOPERIOD (EVANSVILLE, INDIANA) FOR
FATHEAD MINNOW LIFE-CYCLE CHRONIC TOXICITY TESTS
Dawn to Dusk
Time
6:00
6:00
6:00
6:00
6:00
6:00
6:00
6:00
6:00
6:00
6:00
6:00
6:00
6:00
6:00
6:00
6:00
6:00
6:00
6:00
6:00
6:00
6:00
6:00
- 4:45
- 4:30
- 4:30
- 4:45
- 5:15
- 5:45
- 6:15
-7:00
- 7:30
- 8:15
- 8:45
- 9:15
- 9:30
- 9:45
- 9:45
- 9:30
-9:00
- 8:30
-8:00
- 7:30
- 6:45
- 6:15
- 5:30
- 5:00
Date
Dec.
Jan.
Feb.
Mar.
Apr.
May
June
July
Aug.
Sept.
Oct.
Nov.
1
15
1
15
1
15
1
15
1
15
1
15
1
15
1
15
1
15
1
15
1
15
1
15
Day-length (hour and minute)
10:45
10:30
10:30
10:45
11:15
11:45
12:15
13:00
13:30
14:15
14:45
15:15
15:30
15:45
15:45
15:30
15:00
14:30
14:00
13:30
12:45
12:15
1 1 : 30
11:00
Day one of life-cycle
chronic test
5-month pre-spawning
growth period
4-month spawning
period
Post-spawning period
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prior to their use in the diluter and exposure chambers. Recent static tests
at the U.S. EPA, Environmental Research Laboratory, Duluth have shown that
certain lots of neoprene stoppers are acutely toxic to fathead minnow larvae
(S. J. Broderius, personal communication, U.S. Environmental Protection
Agency, Duluth, MN 55804).
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CHAPTER 2
CHEMICAL SYSTEM
2.1 Preparing a Stock Solution
Distilled or diluent water should be used in making-up the test stock
solutions. The recent development of several chemical saturators for use
with hydrophobic chemicals has eliminated the need to use carrier solvents
with most test chemicals (16,17,18,19).
If carrier solvents other than water are absolutely necessary, reagent grade
or better should be used, but amounts must be kept to a minimum. Triethylene
glycol (TEG) and dimethyl formamide (DMF) are preferred, but methanol,
ethanol or acetone can also be used. The calculated solvent concentration to
which any test organisms are exposed must never exceed 0.1 ml/liter.
When a carrier is used, use two sets of duplicate controls. One set should
contain no solvent and one set should contain the highest concentration of
solvent to which any organisms in the test are exposed.
2.2 Measurement of Toxicant Concentration
As a minimum, the concentration of toxicant must be measured in one tank at
each toxicant concentration every week, alternating between duplicate tanks
at each concentration from week to week. Water samples should be taken about
midway between the top and bottom and the sides of the tank and should not
include any surface scum or material stirred up from the bottom or sides of
the tank.
2.3 Measurement of Other Variables
Dissolved oxygen must be measured at each concentration at least once a week.
In alternating weeks, the opposite tank at each concentration should be
measured for dissolved oxygen.
A control and one test concentration must be analyzed at least weekly for pH,
alkalinity, hardness, and conductance to show the variability in the test
water. If any of these characteristics is affected by the toxicant, that
characteristic must be measured at each concentration at least once a week
and alternated between duplicate tanks.
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At a minimum, the test water must be analyzed twice during the test period
for calcium, magnesium, sodium, potassium, chloride, sulfate, total solids,
and total dissolved solids.
2.4 Residue Analysis
Mature fish, and possibly eggs, larvae, and juveniles, obtained from the
test, must be analyzed for toxicant residues. Since fathead minnows usually
are consumed as whole organisms by predators, residues should be determined
for whole bodies rather than individual tissues.
2.5 Methods
Methods described in Methods for Chemical Analysis of Water and Wastes (20)
and Manual of Analytical Methods for the Analysis of Pesticides in Human and
Environmental Samples (21) should be used for chemical analysis when
possible. Accuracy should be measured using the method of known additions
for all analytical methods for toxicants. Reference samples should be
analyzed periodically for each analytical method.
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CHAPTER 3
BIOLOGICAL SYSTEM
3.1 Source of Test Fish
Sufficient numbers of embryos are transported easily from a brood stock
culture unit to other laboratories or field sites to initiate a life-cycle
chronic test. These embryos usually are shipped in well-oxygenated water in
insulated containers.
Laboratory brood stock culture units can be started by obtaining embryos from
a well-established culture unit such as is maintained at the Environmental
Research Laboratory in Duluth, MM. This laboratory periodically mixes their
brood stock with healthy wild minnows to eliminate the risk of developing a
homogeneous strain.
At 25° C and a constant 16-hour day-light photoperiod, fish fed unrestricted
quantities of live brine shrimp nauplii and frozen adult brine shrimp will
mature in 5-6 months. With proper care and maintenance these adult fish will
produce large numbers of embryos for 6-8 months.
Paired spawning, as opposed to group spawning, eliminates fighting and
competition. This method is recommended for use in brood stock culture units
and life-cycle chronic tests. Experiments conducted by Benoit and Holcombe
(7) have demonstrated the successful use of paired spawning in individual,
screened, spawning chambers. These investigators correctly determined the
sex of 40 pairs of mature minnows. Results of their test showed that 38 out
of 40 pairs of adults spawned repeatedly. When the U.S. Environmental
Research Laboratory-Duluth, MM, began to use this approach, the number of
embryos produced in the stock culture units almost doubled. Each culture
unit consists of one tank measuring 30.5 cm x 30.5 cm x 61 cm long with a
water depth of 18 cm and four individual spawning chambers (15.2 x 30.5 cm)
formed by stainless steel screen dividers (5 mesh, 0.89 mm wire).
3.2 Preliminary Tests
Selection of the test concentration for the life-cycle exposure should be
determined on the basis of the results of a 6-10 day range-finding test and a
96-hour toxicity test (22) with either larval or juvenile fathead minnows.
Unless the test chemical is extremely toxic, the highest test concentration
selected for the chronic exposure should usually be no less than the 96-hour
LC20 and no greater than the 96-hour LC50. All fish used in preliminary and
life-cycle tests should be from the same adult stock.
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3.3 Embryo Exposure (4-5 days)
The life-cycle chronic toxicity test must begin with embryos from at least
three separate spawnings that are jŁ 24-hours-old and have soaked in
dilution water for at least 2 hours. Prior to the start of the test all
embryos also must be viewed carefully with a dissecting scope or magnifying
viewer to remove empty shells and opaque, or abnormal appearing embryos. If
less than 50 percent of the embryos from a substrate appear to be healthy and
fertile, all embryos from that substrate should be discarded. Embryos stuck
together in clumps of four or more are either manually separated or
discarded.
The test is started by impartially distributing 50 embryos (<_ 24-hour-old) to
each of the four replicate larval growth chambers using the following
suggested method: 10 embryos are impartially selected and transferred with a
large bore eye dropper to successive incubation cups which are standing in
dilution water. This process is repeated until 50 embryos are in each cup.
The incubation cups are then distributed by stratified random assignment to
each replicate larval chamber.
Dead embryos usually will turn opaque and must be counted and removed each
day until hatching is complete. Live fungused embryos also must be removed
daily, and are subtracted from the original total when calculating percentage
hatch. Embryos found floating in the incubation cup can be submerged by
gently proding with a glass rod or by gently swirling the cup in the test
water.
Upon completion of hatching, the total number of larvae, in each replicate,
including those dead or deformed, are counted. Dead or deformed larvae are
subtracted from the total in determining the number of normal larvae at
hatch. Time to complete hatch in each cup is recorded to the nearest day.
3.4 Larval-Juvenile Exposure (8 weeks)
After hatching, each group of larvae is randomly reduced to 25, and released
in replicate larval growth chambers. All live fish that are lethargic or
deformed must also be included in the random selection. Another option to
the above method would be to impartially reduce the embryos to 25 before
hatching and release all of the larvae into the growth chamber after
hatching.
Larvae should be fed within two days after hatching. Each group must be fed
live, newly hatched, brine shrimp nauplii three times a day at least four
hours apart (or two times a day about six hours apart) during the first 3
weeks after hatching (23). Beginning four weeks after hatching, all fish
should be fed frozen adult brine shrimp at least twice a day (7). The amount
of food provided to each chamber must be proportional to the number of fish
in the chamber. Each batch of brine shrimp eggs and adults should be
analyzed for pesticides.
10
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Survival should be determined in each replicate growth chamber at least once
a week. Larvae which die during the first two weeks after hatching
generally deteriorate so rapidly that they cannot be observed easily in the
test chambers. Therefore, survival during this period is ^et^rmined by
counting the number of live fish.
Record the number of abnormal fish at four and eight weeks after hatching and
measure total lengths of all fish in each replicate growth chamber using the
photographic methods of McKim and Benoit (24). Each glass bottom growth
chamber containing fish is removed from the larval section or tank and test
water drained to a depth of 3 cm. The chamber then is transferred to a light
box having fluorescent lights under a squared millimeter grid of adequate.
size to accommodate the growth chamber. Photos then are taken of the fish
over the grid and enlarged into 20 x 25 cm prints. The length of each fish
is determined subsequently determined by comparing it to the grid. This
method allows growth measurements to be made without handling or removing
test fish from the water, and accuracy is within 1% of the actual total
length.
3.5 Juvenile-Adult Exposure (32-40 weeks)
Eight weeks after hatching, all fish in each growth chamber must be
transferred to the adult spawning tank of the same concentration, and
randomly reduced to 25 fish per tank. Deformed individuals also must be
included in the random selection. If necessary, in order to obtain 25 fish
in each spawning tank, several fish may be selected randomly for transfer
from one tank to another of the same concentration. Record the length and
weight of all fish discarded from each spawning tank.
Four spawning substrates also should be placed in each spawning tank at this
time. Substrates should be separated widely to reduce interaction and
situated so that the underside of the substrates can be viewed from the enri
of the spawning tanks.
Continue routine feeding and cleaning and checking mortality throughout the
juvenile-adult exposure. Handle disease outbreaks according to their nature,
with all experimental and control tanks receiving the same treatment whether
or not there seems to be sick fish in all of them. The frequency of
treatment should be held to a minimum (22).
When secondary sexual characteristics are well-developed and males begin to
establish territories (approximately 20-24 weeks after hatch), separate
males, females and undeveloped fish in each spawning tank (7). Sex can be
determined on maturing fish by viewing each group in a glass aquarium with a
lighted background. Mature males will exhibit tubercles, pads and body color
(25,26); mature females will exhibit extended, transparent anal canals
(urogenital papilla) as described by Flickinger (27).
At this time, four individual spawning chambers (see Section 3.1) are formed
in each spawning tank and one spawning substrate placed in each chamber.
11
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Four males and four females from each spawning tank then are chosen randomly
and assigned to spawning chambers. If necessary, in order to establish four
mature pairs in each spawning tank, several fish should be selected randomly
for transfer from one tank to another of the same concentration. However, if
there are not enough mature fish in either spawning tank, these fish must be
placed back into their respective tanks, without screen dividers, and
observed daily for sexual development and/or spawning activity.
All surplus adults and undeveloped fish are weighed, measured and cut open
for positive sexual identification. Males and females that are readily
distinguishable from one another because of their external characteristics
should be selected initially for determining how to differentiate between
testes and ovaries. The gonads of both sexes will be located just ventral to
the kidneys.
Substrates are checked for spawnings daily, including weekends, (preferably
during the Evansville afternoon hours) and embryos removed as described in
Section 1.5. Substrates must be replaced immediately after embryos are
removed. All embryos produced in each spawning are counted and recorded
separately for each pair.
The adult exposure should be terminated when, during the decreasing
day-length photoperiod, a one-week period passes in which no spawning occurs
in any of the tanks. Record total lengths, weights and sex of parental fish.
At this time the gonads of most parental fish will have begun to regress from
the spawning condition, and both external and internal differences between
the sexes will be less distinct. The ovaries are generally larger than
testes and will appear transparent, but may still contain some coarsely
granular yellow pigment. The testes will be quite slender with very fine
granular strands and may be slightly milky when squeezed.
3.6 Second Generation Embryo Exposure (4-5 days)
Fifty embryos from the first five spawnings (>50 embryos) and every third
spawning (>_5o embryos) thereafter from each pair are selected impartially
and transferred to incubation cups for hatch. Those embryos not selected for
incubation are discarded.
Test procedures used during the embryo incubation and hatching of offspring
are described in the last two paragraphs of Section 3.3.
3.7 Second Generation Larval-Juvenile Exposure (4-8 weeks)
Two groups of 25 larvae produced from different pairs in each spawning tank
are released in replicate growth chambers for eight week exposures.
Selection of each group should be from early spawnings so that the chambers
will be available for additional exposures of fish, if desired.
12
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Test procedures used during the selection and exposure of offspring are
described in Section 3.4.
Each group of second generation fish is terminated eight weeks after
hatching. Individual fish are blotted, weighed, and measured before
discarding or freezing.
If the test is continued beyond the four week period or fish are transferred
to clean water for residue half-life studies, individual live fish can
be blotted through a small net and weighed by water displacement. Length can
be determined photographically on live fish as described in Section 3.4.
3.8 Additional Exposures and Special Examinations
Important information on hatchability and larval survival can be gained by
transferring, immediately after spawning, embryos from the control tanks to
tanks having toxicant concentrations in which spawning is known to be reduced
or absent or to tanks in which an effect is seen on survival of embryos or
larvae. Information also can be gained by transferring embryos from these
high toxicant concentrations to the control tanks.
Fish and embryos obtained from the test can be used for physiological,
biochemical, histological, and other tests that may indicate certain toxicant
related effects. Egg adhesiveness can also be evaluated with the use of egg
traps placed under selected spawning substrates (7).
Extended life-cycle chronic tests may be conducted through several
generations by using the test procedures described in sections 3,3 through
3.7.
3.9 Biological Data Recorded
3.9.1 Embryo Exposure
Days to complete hatching; total number of embryos hatched; and number of
normal larvae at hatching in each incubation cup.
3.9.2 Larval-Juvenile Exposure
Survival, deformities, and growth of fish at the end of the 4 and 8 week
exposure periods in each replicate growth chamber.
3.9.3 Juvenile-Adult Exposure
Survival and deformities of fish at the time of selection for paired
spawning; growth and sex of discarded fish not selected for spawning; and
survival, growth, and deformities of male and female fish at the end of the
13
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spawning test period in each duplicate spawning tank. In addition to the
foregoing data, the number of embryos per spawn and the total number of
spawnings by each pair of adults in individual spawning chambers also are
recorded.
3.10. Data Analysis and Evaluation
Test results can be analyzed statistically by standard techniques. A simple
one-way analysis oŁ variance may be used initially to assess whether
significant (P ^0.05) differences have been found (28).
If differences exist, the investigator can then determine whether responses
for a given concentration are significantly (P >^ 0.05) different frora
responses for the control. Two particularly appropriate methods for this
type of evaluation are Dunnett's test (29) and William's test (30,31).
Duncan's new multiple-range test also can be used to analyze statistically
differences between all treatments (32).
14
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REFERENCES
1. American Public Health Association, American Water Works Association,
and Water Pollution Control Federation. 1980. Standard methods for the
examination of water and wastewater. 15th ed., APHA, Washington, DC
20005.
2. Mount, D. I., and W. A. Brungs. 1967. A simplified dosing apparatus
for fish toxicology studies. Water Res. 1: 21-29.
3. Lemke, A. E., W. A. Brungs, and B. J. Halligan. 1978. Manual for
construction and operation of toxicity-testing proportional diluters.
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*USGPO: 1982 559-092/3396
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